This research has been supported by grants from the National Institute on Deafness and Other Communication Disorders of the National Institutes of Health, the Kellogg Foundation, and private donations from Bridget Sperl and John McCormick. TSLIM has been developed with the excellent assistance of Matthias Hillenbrand, Kerstin John, Meike Lawin, Michel Layher, Tobias Schroeter, Peter Schacht, Oliver Dannberg, and Julian Wuester from the Technical University of Illmenau, Germany, supervision by their mentors (Stefan Sinzinger and Rene Theska) and James Leger.

All the procedures and the use of live animals have been reviewed and approved (Protocol ID #2010-38573A) by the University of Minnesota Institutional Care and Use Committee (IACUC) and investigators who use these animals have been thoroughly trained and tested by the Research Animal Resources (RAR) Veterinarians before they have access to the animal facilities. Both male and female mice were used in this study.
1. Cochlea removal for fixation and tissue processing for imaging
<ol>
	<li>Euthanize a mouse using CO2 inhalation. Decapitate the mouse with scissors and make a dorsal-ventral incision through the brain to hemisect the skull. Remove the brain, identify the round bulla in the baso-ventral part of the skull, open the bulla with rongeurs, and visualize and remove the cochlea.</li>
	<li>Fixation: Perform this procedure under a fume hood and using a dissection microscope at 5x magnification. Wear gloves and protective clothing. Puncture the oval window and remove the stapes with a sharp pick. Insert a pick into the round window to puncture the membrane.</li>
	<li>Cover the opened round window with the cut tip of an infusion set attached to a 1 mL syringe filled with 2 mL of formalin. Slowly infuse formalin through the perilymphatic spaces of the cochlea over a 2 min period, noting that formalin is exiting the cochlea via the opened oval window. Trim excess tissue off the cochlea and immerse in a bottle containing 10% formalin, and place on a rotator overnight.</li>
	<li>Decalcification: Rinse the cochlea in PBS 3x for 5 min each and immerse in a bottle containing 10% solution of disodium ethylenediaminetetraacetic acid (EDTA) with rotation for 4 days, changing the solution daily.</li>
	<li>Dehydration: Perfuse the cochlea with PBS 3x and immerse for 15 min between changes. Dehydrate the cochlea with ascending concentrations of ethanol 10%, 50%, 70%, 95%, 95%, 100%, 100%; for 30 min in each concentration. 
	&#8203;NOTE: It is important to remove all the EDTA before dehydration as EDTA precipitates in ethanol. Also, cochleae can be left in any concentration of ethanol greater than 70% overnight.</li>
	<li>Staining: Stain the whole cochlea by immersion in a solution of Rhodamine B isothiocynate (5 &#181;g/mL in 100% ethanol) overnight with rotation. Remove excess dye from the cochlea with two changes of 100% ethanol, 5 min each change.</li>
	<li>Clearing: Transfer the stained cochlea into two changes of Spalteholz12 solution (5:3 methyl salicylate:benzyl benzoate), 30 min each change and leave overnight in the clearing solution with rotation. Cochleae can be left in Spalteholz solution indefinitely.</li>
</ol>
2. Imaging of cochleae
<ol>
	<li>Attach the cochleae to a specimen rod at the oval and round window membrane end so that the clearing solution remains within the cochlea and bubbles are not formed (Figure 2). Care must be taken not to let bubbles form within the cochlea as they are difficult to remove and if left in the tissue, they will cause imaging artifacts.</li>
	<li>Use a UV activating glue to attach the wet cochlea to the dry specimen rod (Figure 2). Attach the cochlea at the oval and round window ends. Cure the UV glue for 10 s by moving around the cochlea with the UV light. 
	NOTE: A loose attachment of the cochlea to the specimen rod will result in imaging defects. This rod is specifically manufactured for this protocol (see Santi et al.8 for details) and is specific to our light-sheet microscope.</li>
	<li>Suspend the cochlea into the imaging chamber filled with Spalteholz solution for imaging. The specimen chamber is an optically clear quartz fluorometer cell (Video 1).</li>
	<li>Attach the specimen rod to a rotating holder that is also attached to XZ translation stage. Most stacks are obtained by translating the specimen in the XZ planes, but rotational stacks can also be obtained.</li>
	<li>TSLIM optical sectioning: Use a blue or green laser for excitation depending upon the type of fluorochrome staining. Position the light sheet in the middle of the tissue for focusing and determine the magnification that will be used to illuminate the full width of the cochlea. Then, use a custom-designed program to move the specimen through the light-sheet across the specimen in the X-axis (stitching the image) and in Z-steps to make a stack of 2D images throughout the cochlea.</li>
	<li>For the first image, the beam waist of the light-sheet is positioned at the edge of the specimen and the program scans the full width of the specimen collecting columns of data (see image stitching; Santi et al.8) that are the width of the confocal parameter (Figure 3) to produce a composite 2D image of maximum resolution across the width of the specimen. The program automates image stitching for each Z-step until the specimen is completely imaged.</li>
	<li>Image processing: Transfer the image stack to another computer and load it into a 3D rendering program for 3D reconstruction and quantification.</li>
</ol>

<ol>
	<li>Deafness and hearing loss. World Health Organization Available from: <a target="_blank" href="https://www.who.int/news-room/fact-sheets/deafness-and-hearing-loss">https://www.who.int/news-room/fact-sheets/deafness-and-hearing-loss</a> (2021)</li><li>Vater, M., K&#246;ssl, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+aspects+of+cochlear+functional+organization+in+mammals.">Comparative aspects of cochlear functional organization in mammals.</a> Hearing Research. 273 (1-2), 89-99 (2011).</li><li>Santi, P. A., Blair, A., Bohne, B. A., Lukkes, J., Nietfeld, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+digital+cytocochleogram.">The digital cytocochleogram.</a> Hearing Research. 192 (1-2), 75-82 (2004).</li><li>Voie, A. H., Burns, D. H., Spelman, F. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orthogonal-plane+fluorescence+optical+sectioning:+three-dimensional+imaging+of+macroscopic+biological+specimens.">Orthogonal-plane fluorescence optical sectioning: three-dimensional imaging of macroscopic biological specimens.</a> Journal of Microscopy. 170, 229-236 (1993).</li><li>Voie, A. H., Spelman, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Three-dimensional+reconstruction+of+the+cochlea+from+two-dimensional+images+of+optical+sections.">Three-dimensional reconstruction of the cochlea from two-dimensional images of optical sections.</a> Computerized Medical Imaging and Graphics. 19 (5), 377-384 (1995).</li><li>Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+sectioning+deep+inside+live+embryos+by+selective+plane+illumination+microscopy.">Optical sectioning deep inside live embryos by selective plane illumination microscopy.</a> Science. 305 (5686), 1007-1009 (2004).</li><li>Santi, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+sheet+fluorescence+microscopy:+a+review.">Light sheet fluorescence microscopy: a review.</a> The Journal of Histochemistry and Cytochemistry. 59 (2), 129-138 (2011).</li><li>Santi, P. 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=Thin-sheet+laser+imaging+microscopy+for+optical+sectioning+of+thick+tissues.">Thin-sheet laser imaging microscopy for optical sectioning of thick tissues.</a> BioTechniques. 46 (4), 287-294 (2009).</li><li>Schr&#246;ter, T. J., Johnson, S. B., John, K., Santi, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Scanning+thin-sheet+laser+imaging+microscopy+(sTSLIM)+with+structured+illumination+and+HiLo+background+rejection.">Scanning thin-sheet laser imaging microscopy (sTSLIM) with structured illumination and HiLo background rejection.</a> Biomedical Optics Express. 3 (1), 170-177 (2012).</li><li>Keller, P. J., Schmidt, A. D., Wittbrodt, J., Stelzer, E. H. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstruction+of+zebrafish+early+embryonic+development+by+scanned+light+sheet+microscopy.">Reconstruction of zebrafish early embryonic development by scanned light sheet microscopy.</a> Science. 322 (5904), 1065-1069 (2008).</li><li>Buytaert, J. A. N., Dirckx, J. J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Design+and+quantitative+resolution+measurements+of+an+optical+virtual+sectioning+three-dimensional+imaging+technique+for+biomedical+specimens,+featuring+two-micrometer+slicing+resolution.">Design and quantitative resolution measurements of an optical virtual sectioning three-dimensional imaging technique for biomedical specimens, featuring two-micrometer slicing resolution.</a> Journal of Biomedical Optics. 12 (1), 014039 (2007).</li><li>Spalteholz, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> On making human and animal preparations transparent. , (1914).</li><li>Johnson, S., Schmitz, H., Santi, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TSLIM+imaging+and+a+morphometric+analysis+of+the+mouse+spiral+ganglion.">TSLIM imaging and a morphometric analysis of the mouse spiral ganglion.</a> Hearing Research. 278 (1-2), 34-42 (2011).</li><li>Santi, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organ+of+Corti+surface+preparations+for+computer-assisted+morphometry.">Organ of Corti surface preparations for computer-assisted morphometry.</a> Hearing Research. 24 (3), 179-187 (1986).</li><li>Brown, D., Pastras, C., Curthoys, I., Southwell, C., Van Roon, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endolymph+movement+visualized+with+light+sheet+fluorescence+microscopy+in+an+acute+hydrops+model.">Endolymph movement visualized with light sheet fluorescence microscopy in an acute hydrops model.</a> Hearing Research. 339, 112-124 (2016).</li><li>White, J. A., Burgess, B. J., Hall, R. D., Nadol, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pattern+of+degeneration+of+the+spiral+ganglion+cell+and+its+processes+in+the+C57BL/6J+mouse.">Pattern of degeneration of the spiral ganglion cell and its processes in the C57BL/6J mouse.</a> Hearing Research. 141 (1-2), 12-18 (2000).</li><li>Grierson, K. E., Hickman, T. T., Liberman, M. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dopaminergic+and+cholinergic+innervation+in+the+mouse+cochlea+after+noise-induced+or+age-related+synaptopathy.">Dopaminergic and cholinergic innervation in the mouse cochlea after noise-induced or age-related synaptopathy.</a> Hearing Research. 422, 108533 (2022).</li></ol>

The authors have nothing to disclose.

Optical sectioning by light sheet microscopy for examination of cochlear structures is not mechanically destructive like other more traditional histological methods, and it provides a complete digital view of cochlear structures relative to one another. Previous methods such as surface preparations of the organ of Corti14 provided a map of hair cell loss along the length of the basilar membrane, but SGN loss could not be assessed since the tissue had been dissected away to reveal the organ of Cort...

The cochlea is the peripheral sensory organ for hearing in mammals. It has a complex spiral anatomy of repeating sensory and supporting cells that are anatomically specialized to detect sound vibrations and transmit them to the brain for the perception of hearing. The main sensory elements are the inner and outer hair cells and their innervating nerve fibers whose cell bodies compose the spiral ganglion, which resides within Rosenthal&#39;s canal (Figure 1). These sensory and neural structures are tonotopically arranged such that high-frequency sounds are transduced in the cochlear base and low-frequency sounds are transduced in the cochlea apex2. An anatomical map of this sensory cell distribution along the spiral length of the supporting basilar membrane is called a cytocochleogram3 and can be compared with hearing loss as a function of frequency as depicted in an audiogram.
The membranous labyrinth of the cochlea, which is surrounded by dense bone, makes it technically difficult to examine more than one cochlear structure at a time. Therefore, the rationale for developing a light-sheet microscope is to produce well-aligned serial sections of the complete cochlea so that all cochlear structures can be examined relative to one another in 3D reconstructions. Voie et al.4 and Voie and Spelman5 designed the first light-sheet microscope, called orthogonal plane fluorescence optical sectioning (OPFOS) microscope, to optically section the whole cochlea. However, this microscope was never commercially developed; so, our aim was to construct a light sheet microscope called a thin sheet laser imaging microscope (TSLIM; Figure 2). The design and construction details for TSLIM have previously been published8. TSLIM made several improvements over the OPFOS, including using a low-light digital camera versus a CCD camera for image collection, optically encoded micropositioners for accurate and reproducible movement of the specimen through the light-sheet, use of a commercially available, optically clear specimen chamber, and Rhodamine staining in ethanol rather than in the clearing solution to prevent stain precipitation within the tissue. Commercial development of light-sheet microscopes such as SPIM6 have focused on high-resolution imaging of live, small transparent specimens but are unsuitable for whole cochlear imaging as they lack adequate working distance. A review of the development of other light-sheet microscopes was published by Santi7. TSLIM&#39;s primary advantage over other histological methods to examine the cochlea is to optically section tissues for 3D reconstruction while preserving the integrity of the specimen so that it can be used by other histological methods. Another advantage of TSLIM imaging is that only a thin light-sheet produced by a laser is exposed to the tissue, compared with whole tissue thickness exposure to the laser as in confocal microscopy. Tissue clearing to minimize light scatter and the fact that only a small portion of the tissue is exposed to the laser results in minimal fluorochrome fading (photobleaching) with light-sheet laser imaging. However, the process of fixation, dehydration, and clearing does alter the morphology of cochlear structures and results in tissue shrinkage compared with living tissue. The actual amount of tissue shrinkage that occurs was not determined.
TSLIM was developed by Shane Johnson and eight German optical engineering students (see Acknowledgements). TSLIM construction details were provided by Santi et al.8 and a scanning version (sTSLIM) by Schr&#246;ter et al.9. TSLIM functions as a nondestructive microtome to optically section specimens and as a microscope to collect 2D serial sections through the full width and thickness of the cochlea. TSLIM can image both small (mm) and large (cm), thick specimens. Lenses are air mounted to allow for long working distances with collection objectives of 1x and 2x on a dissection microscope. The dissection microscope also has zoom optics that allow TSLIM to resolve subcellular and synaptic structures on cells. TSLIM is equipped with both a blue (473 nm) and green (532 nm) laser for illumination that allows for a variety of fluorescent probes to be used for imaging. The goal of TSLIM is to produce well-aligned 2D optical sections through a whole cochlea for a complete digital reconstruction of cochlear tissues. Since it is a fluorescent method, ligands and immunohistochemistry can also be used to identify specific cochlear structures.
Initially, a cylindrical lens was used to produce two opposed Gaussian light sheets, but it produced absorption imaging artifacts. Due to the work of Keller et al.10, the fixed cylindrical lens was replaced by a scanning galvanometer mirror to produce the light sheet9. In addition, since the center of the light sheet is the thinnest at the beam waist, sTSLIM 2D images are produced by collecting a composite of X-axis columns of data across the specimen&#39;s width (Figure 3). This method was first described by Buytaert and Dircks11. TSLIM custom software to drive and collect images was developed using a graphical program for instrument control. The light sheet travels through the specimen and illuminates a fluorescent plane within the tissue. This fluorescent plane is projected orthogonally through the transparent specimen and is collected by a dissection microscope. Optically encoded micropositioners allow scanning through the beam waist in the X-axis to collect a single composite 2D image and, subsequently, the Z-axis micropositioner moves the specimen to a deeper plane within the tissue to obtain a stack of serial, sectioned 2D images (Video 1, Figure 4). A stack of translational images is collected through the entire width, thickness, and length of the cochlea, and stitching of images is not required (Video 2). The image stack is transferred to another computer and loaded into a 3D rendering program for 3D reconstruction and quantification. The image stacks contain all the digital information about the morphology of a cochlea at the resolution of the microscope. However, if a higher resolution is required, the intact cochlea can be further processed by destructive histological methods such as microtome sectioning, scanning, and transmission electron microscopy.
The 3D rendering program is used to segment different cochlear structures for 3D rendering and quantitative analysis. For segmentation, each structure in every 2D image of the stack is traced using a different color by a graphics tablet and pen (Figure 5). To date, 20 different cochlear structures have been segmented (Figure 6). After segmentation, a variety of 3D analyses can be performed. For example, 3D rendering software can virtually resection the cochlea in any plane along the structure&#39;s centroid. Video 3 shows sectioning tangential to the organ of Corti, which reveals the hair cells along the length of the basilar membrane. This process first requires manual segmentation of the structure of interest. Next, the structure&#39;s centroid is calculated based on the least squares fit of spline points placed along the center of the structure from its base to its apex, thus allowing an approximation of the structure&#39;s length (Video 4). A similar process called skeletonization can be used to visualize the radial width of the structure along its length using a color map (Video 4). The total volume of each structure is calculated by the program after segmentation, but relative distances can also be quantified and visualized with color maps in a 3D rendering software (Figure 7). Segmented structures can also be exported to produce enlarged, solid-plastic model renderings (Figure 8). In addition, semi-automated cell counting can also be performed using 3D rendering software (Figure 9). Immunohistochemistry and ligand binding can be used to stain specific cochlear structures and these structures can be isolated from other cochlear structures for morphometrical assessment such as producing a cytocochleogram (Figure 10). Length, width, surface, volume, and number of all cochlear structures can be determined from the 3D models, making this approach ideal for mapping cochlear damage to functional impairments. Specifically, cochlear damage due to aging, noise-induced trauma, or other insults can be shown and quantified in 3D cochlear reconstructions from 2D optical sections. Once a cochlea has been digitized there are numerous imaging algorithms that can be used to assess cochlear damage of any tissue within the cochlea in the anatomical registry to other cochlear tissues.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Amira 3D Rendering Software</td>
			<td>ThermoFisher Scientific</td>
			<td></td>
			<td>Address: 501 90th Ave NW, Coon Rapids, MN 55433</td>
		</tr>
		<tr>
			<td>benzyl benzoate (W213810)</td>
			<td>Sigma-Aldrich, Inc.&#160;</td>
			<td></td>
			<td>Address: PO Box, 14508, St. Louis, MO 68178</td>
		</tr>
		<tr>
			<td>Bondic&#160;</td>
			<td>Bondic&#160;</td>
			<td></td>
			<td>Address: 235 Industrial Parkway S., Unit 18 Aurora, ON L4G 3V5 Canada</td>
		</tr>
		<tr>
			<td>Ethanol 95% and 100%&#160;</td>
			<td>University of Minnesota</td>
			<td></td>
			<td>Address: General Storehouse, Minneapolis, MN 55455</td>
		</tr>
		<tr>
			<td>Ethylenediaminetetraacetic acid disodium salt dihydrate (EDTA)&#160; (E5134)</td>
			<td>Sigma-Aldrich, Inc.&#160;</td>
			<td></td>
			<td>Address: PO Box, 14508, St. Louis, MO 68178</td>
		</tr>
		<tr>
			<td>LabVIEW graphical program and Vision</td>
			<td>National Instruments</td>
			<td></td>
			<td>Address: 11500 N Mopac Expwy Austin, TX 78759-3504</td>
		</tr>
		<tr>
			<td>methyl salicylate (M6742)</td>
			<td>Sigma-Aldrich, Inc.&#160;</td>
			<td></td>
			<td>Address: PO Box, 14508, St. Louis, MO 68178</td>
		</tr>
		<tr>
			<td>Olympus MVX10 dissection microscope</td>
			<td>Olympus Corp</td>
			<td></td>
			<td>Address: 3500 Corporate Parkway, Center Valley, PA 18034</td>
		</tr>
		<tr>
			<td>Rhodamine B isothiocynate, (283924)&#160;</td>
			<td>Sigma-Aldrich, Inc.&#160;</td>
			<td></td>
			<td>Address: PO Box, 14508, St. Louis, MO 68178</td>
		</tr>
		<tr>
			<td>Starna Flurometer Cell (3-G-20)</td>
			<td>Starna Cells</td>
			<td></td>
			<td>Address: PO Box 1919, Atascadero, CA 82423</td>
		</tr>
	</tbody>
</table>

imaging the aging cochlea with light-sheet fluorescence microscopy

Deafness is the most common sensory impairment, affecting approximately 5% or 430 million people worldwide as per the World Health Organization1. Aging or presbycusis is a primary cause of sensorineural hearing loss and is characterized by damage to hair cells, spiral ganglion neurons (SGNs), and the stria vascularis. These structures reside within the cochlea, which has a complex, spiral-shaped anatomy of membranous tissues suspended in fluid and surrounded by bone. These properties make it technically difficult to investigate and quantify histopathological changes. To address this need, we developed a light-sheet microscope (TSLIM) that can image and digitize the whole cochlea to facilitate the study of structure-function relationships in the inner ear. Well-aligned serial sections of the whole cochlea result in a stack of images for three-dimensional (3D) volume rendering and segmentation of individual structures for 3D visualization and quantitative analysis (i.e., length, width, surface, volume, and number). Cochleae require minimal processing steps (fixation, decalcification, dehydration, staining, and optical clearing), all of which are compatible with subsequent high-resolution imaging by scanning and transmission electron microscopy. Since all the tissues are present in the stacks, each structure can be assessed individually or relative to other structures. In addition, since imaging uses fluorescent probes, immunohistochemistry and ligand binding can be used to identify specific structures and their 3D volume or distribution within the cochlea. Here we used TSLIM to examine cochleae from aged mice to quantify the loss of hair cells and spiral ganglion neurons. In addition, advanced analyses (e.g., cluster analysis) were used to visualize local reductions of spiral ganglion neurons in Rosenthal&#39;s canal along its 3D volume. These approaches demonstrate TSLIM microscopy&#39;s ability to quantify structure-function relationships within and between cochleae.

Since the theme of this special issue is imaging the effects of aging in the cochlea, a young (3-month-old, HS2479, CBA strain mouse) and aged (23-month-old, HS2521, C57 strain mouse) cochleae will be used as examples. It should be noted that TSLIM is capable of imaging a variety of specimens, including cochleae from humans, mammals, other rodents, and fish, as well as other organs such as the brain.
Johnson et al. 13 published an article on SGNs in young (3-week-old) C...

Watch this Scientific Journal Video about Imaging the Aging Cochlea with Light-Sheet Fluorescence Microscopy at JoVE.com

Imaging the Aging Cochlea with Light-Sheet Fluorescence Microscopy

A light-sheet microscope was developed to image and digitize whole cochlea.

Deafness is the most common sensory impairment, affecting approximately 5% or 430 million people worldwide as per the World Health Organization1. Aging or ...

imaging-the-aging-cochlea-with-light-sheet-fluorescence-microscopy

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2.0K Views.  University of Minnesota. A light-sheet microscope was developed to image and digitize whole cochlea.

Video: Imaging the Aging Cochlea with Light-Sheet Fluorescence Microscopy

Micron-scale Resolution Optical Tomography of Entire Mouse Brains with Confocal Light Sheet Microscopy

Understanding the architecture of mammalian brain at single-cell resolution is one of the key issues of neuroscience. However, mapping neuronal soma and projections throughout the whole brain is still challenging for imaging and data management technologies. Indeed, macroscopic volumes need to be reconstructed with high resolution and contrast in a reasonable time, producing datasets in the TeraByte range. We recently demonstrated an optical method (confocal light sheet microscopy, CLSM) capable of obtaining micron-scale reconstruction of entire mouse brains labeled with enhanced green fluorescent protein (EGFP). Combining light sheet illumination and confocal detection, CLSM allows deep imaging inside macroscopic cleared specimens with high contrast and speed. Here we describe the complete experimental pipeline to obtain comprehensive and human-readable images of entire mouse brains labeled with fluorescent proteins. The clearing and the mounting procedures are described, together with the steps to perform an optical tomography on its whole volume by acquiring many parallel adjacent stacks. We showed the usage of open-source custom-made software tools enabling stitching of the multiple stacks and multi-resolution data navigation. Finally, we illustrated some example of brain maps: the cerebellum from an L7-GFP transgenic mouse, in which all Purkinje cells are selectively labeled, and the whole brain from a thy1-GFP-M mouse, characterized by a random sparse neuronal labeling.

In this article we describe the full experimental procedure to reconstruct, with high resolution, the fine brain anatomy of fluorescently labeled mouse brains. The described protocol includes sample preparation and clearing, specimen mounting for imaging, data post-processing and multi-scale visualization.

Understanding the architecture of mammalian  brain at single-cell resolution is one of the key issues of neuroscience.  However, mapping neuronal soma and ...

A Dual Tracer PET-MRI Protocol for the Quantitative Measure of Regional Brain Energy Substrates Uptake in the Rat

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the rat. The developed method is a small-animal positron emission tomography (PET) protocol, in which&#160;11C-AcAc and 18F-fluorodeoxyglucose (18F-FDG) are injected sequentially in each animal. This dual tracer PET acquisition is possible because of the short half-life of 11C (20.4 min). The rats also undergo a magnetic resonance imaging (MRI) acquisition seven days before the PET protocol. Prior to image analysis, PET and MRI images are coregistered to allow the measurement of regional cerebral uptake (cortex, hippocampus, striatum, and cerebellum). A quantitative measure of 11C-AcAc and 18F-FDG brain uptake (cerebral metabolic rate; &#956;mol/100 g/min) is determined by kinetic modeling using the image-derived input function (IDIF) method. Our new dual tracer PET protocol is robust and flexible; the two tracers used can be replaced by different radiotracers to evaluate other processes in the brain. Moreover, our protocol is applicable to the study of brain fuel supply in multiple conditions such as normal aging and neurodegenerative pathologies such as Alzheimer&#39;s and Parkinson&#39;s diseases.

Small-animal positron emission tomography enables the assessment of the brain&#39;s two main energy substrates:&#160;glucose and ketones. In the present method, 11C-acetoacetate and 18F-fluorodeoxyglucose are injected sequentially in each animal, and their uptake is measured quantitatively in specific brain regions determined from the magnetic resonance images.

We present a method for comparing the uptake of the brain&#39;s two key energy substrates: glucose and ketones (acetoacetate [AcAc] in this case) in the ...

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

The Neuromuscular Junction: Measuring Synapse Size, Fragmentation and Changes in Synaptic Protein Density Using Confocal Fluorescence Microscopy

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. Structural changes at the NMJ can result in neurotransmission failure, resulting in weakness, atrophy and even death of the muscle fiber. Many studies have investigated how genetic modifications or disease can alter the structure of the mouse NMJ. Unfortunately, it can be difficult to directly compare findings from these studies because they often employed different parameters and analytical methods. Three protocols are described here. The first uses maximum intensity projection confocal images to measure the area of acetylcholine receptor (AChR)-rich postsynaptic membrane domains at the endplate and the area of synaptic vesicle staining in the overlying presynaptic nerve terminal. The second protocol compares the relative intensities of immunostaining for synaptic proteins in the postsynaptic membrane. The third protocol uses Fluorescence Resonance Energy Transfer (FRET) to detect changes in the packing of postsynaptic AChRs at the endplate. The protocols have been developed and refined over a series of studies. Factors that influence the quality and consistency of results are discussed and normative data are provided for NMJs in healthy young adult mice.

The neuromuscular junction (NMJ) is altered in a variety of conditions that can sometimes culminate in synaptic failure. This report describes fluorescence microscope-based methods to quantify such structural changes.

The neuromuscular junction (NMJ) is the large, cholinergic relay synapse through which mammalian motor neurons control voluntary muscle contraction. ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in vertebrates. Thus, it is critical to uncover the molecular mechanisms of how motor axons navigate to, arborize, and innervate their peripheral muscle targets during development and degeneration. Although transgenic Hb9::GFP mouse lines have long served to visualize motor axon trajectories during embryonic development, detailed descriptions of the full spectrum of axon terminal arborization remain incomplete due to the pattern complexity and limitations of current optical microscopy. Here, we describe an improved protocol that combines light sheet fluorescence microscopy (LSFM) and robust image analysis to qualitatively and quantitatively visualize developing motor axons. This system can be easily adopted to cross genetic mutants or MN disease models with Hb9::GFP lines, revealing novel molecular mechanisms that lead to defects in motor axon navigation and arborization.

Here, we describe a protocol for visualizing motor neuron projection and axon arborization in transgenic Hb9::GFP mouse embryos. After immunostaining for motor neurons, we used light sheet fluorescence microscopy to image embryos for subsequent quantitative analysis. This protocol is applicable to other neuron navigation processes in the central nervous system.

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in ...

Visualization of Amyloid &#946; Deposits in the Human Brain with Matrix-assisted Laser Desorption/Ionization Imaging Mass Spectrometry

The neuropathology of Alzheimer&#39;s disease (AD) is characterized by the accumulation and aggregation of amyloid &#946; (A&#946;) peptides into extracellular plaques of the brain. The A&#946; peptides, composed of 40 amino acids, are generated from amyloid precursor proteins (APP) by &#946;- and &#947;-secretases. A&#946; is deposited not only in cerebral parenchyma but also in leptomeningeal and cerebral vessel walls, known as cerebral amyloid angiopathy (CAA). While a variety of A&#946; peptides were identified, the detailed production and distribution of individual A&#946; peptides in pathological tissues of AD and CAA have not been fully addressed. Here, we develop a protocol of matrix-assisted laser desorption/ionization-based imaging mass spectrometry (MALDI-IMS) on human autopsy brain tissues to obtain comprehensive protein mapping. For this purpose, human cortical specimens were obtained from the Brain Bank at the Tokyo Metropolitan Institute of Gerontology. Frozen cryosections are cut and transferred to indium-tin-oxide (ITO)-coated glass slides. Spectra are acquired using the MALDI system with a spatial resolution up to 20 &#181;m. Sinapinic acid (SA) is uniformly deposited on the slide using either an automatic or a manual sprayer. With the current technical advantages of MALDI-IMS, a typical data set of various A&#946; species within the same sections of human autopsied brains can be obtained without specific probes. Furthermore, high-resolution (20 &#181;m) imaging of an AD brain and severe CAA sample clearly shows that A&#946;1-36 to A&#946;1-41 were deposited into leptomeningeal vessels, while A&#946;1-42 and A&#946;1-43 were deposited in cerebral parenchyma as senile plaque (SP). It is feasible to adopt MALDI-IMS as a standard approach in combination with clinical, genetic, and pathological observations in understanding the pathology of AD, CAA, and other neurological diseases based on the current strategy.

Visualization of Amyloid &amp;#946; Deposits in the Human Brain with Matrix-assisted Laser Desorption/Ionization Imaging Mass Spectrometry

Molecular imaging with matrix-assisted laser desorption/ionization-based imaging mass spectrometry (MALDI-IMS) allows the simultaneous mapping of multiple analytes in biological samples. Here, we present a protocol for the detection and visualization of amyloid &#946; protein on brain tissues of Alzheimer&#39;s disease and cerebral amyloid angiopathy samples using MALDI-IMS.

The neuropathology of Alzheimer&#39;s disease (AD) is characterized by the accumulation and aggregation of amyloid &#946; (A&#946;) peptides into ...

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

Monitoring Neuronal Survival via Longitudinal Fluorescence Microscopy

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to assess factors that influence neuronal fate. These assays require the observation of separate populations of cells at discrete time points. As a result, individual cells cannot be followed prospectively over time, severely limiting the ability to discriminate whether subcellular events, such as puncta formation or protein mislocalization, are pathogenic drivers of disease, homeostatic responses, or merely coincidental phenomena. Single-cell longitudinal microscopy overcomes these limitations, allowing the researcher to determine differences in survival between populations and draw causal relationships with enhanced sensitivity. This video guide will outline a representative workflow for experiments measuring single-cell survival of rat primary cortical neurons expressing a fluorescent protein marker. The viewer will learn how to achieve high-efficiency transfections, collect and process images enabling the prospective tracking of individual cells, and compare the relative survival of neuronal populations using Cox proportional hazards analysis.

Here, we present a protocol to monitor survival on a single-cell basis and identify variables that significantly predict cell death.

Standard cytotoxicity assays, which require the collection of lysates or fixed cells at multiple time points, have limited sensitivity and capacity to ...

Whole-Brain Single-Cell Imaging and Analysis of Intact Neonatal Mouse Brains Using MRI, Tissue Clearing, and Light-Sheet Microscopy

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of structural changes caused by genetic or environmental perturbations. Whole-brain imaging results in more accurate quantification of cells and the study of region-specific differences that may be missed with commonly used microscopy of physically sectioned tissue. Using light-sheet microscopy to image cleared brains greatly increases acquisition speed as compared to confocal microscopy. Although these images produce very large amounts of brain structural data, most computational tools that perform feature quantification in images of cleared tissue are limited to counting sparse cell populations, rather than all nuclei.
Here, we demonstrate NuMorph (Nuclear-Based Morphometry), a group of analysis tools, to quantify all nuclei and nuclear markers within annotated regions of a postnatal day 4 (P4) mouse brain after clearing and imaging on a light-sheet microscope. We describe magnetic resonance imaging (MRI) to measure brain volume prior to shrinkage caused by tissue clearing dehydration steps, tissue clearing using the iDISCO+ method, including immunolabeling, followed by light-sheet microscopy using a commercially available platform to image mouse brains at cellular resolution. We then demonstrate this image analysis pipeline using NuMorph, which is used to correct intensity differences, stitch image tiles, align multiple channels, count nuclei, and annotate brain regions through registration to publicly available atlases.
We designed this approach using publicly available protocols and software, allowing any researcher with the necessary microscope and computational resources to perform these techniques. These tissue clearing, imaging, and computational tools allow measurement and quantification of the three-dimensional (3D) organization of cell-types in the cortex and should be widely applicable to any wild-type/knockout mouse study design.

This protocol describes methods for conducting magnetic resonance imaging, clearing, and immunolabeling of intact mouse brains using iDISCO+, followed by a detailed description of imaging using light-sheet microscopy, and downstream analyses using NuMorph.

Tissue clearing followed by light-sheet microscopy (LSFM) enables cellular-resolution imaging of intact brain structure, allowing quantitative analysis of ...