The research leading to these results has received funding from the European Union Seventh Framework Programme (FP7/2007-2013) under grant agreements n. 228334 and 241526. This research project has been also supported by Human Frontier Science Program research grant (RGP0027/2009), and by the Italian Ministry for Education, University and Research in the framework of the Flagship Project Nanomax and by Italian Ministry of Health in the framework of the "Stem Cells Call for proposals". This research has been carried out in the framework of the research activities of ICON foundation supported by "Ente Cassa di Risparmio di Firenze".

1. Tissue Preparation and Storage
<ol>
	<li>Fix mouse brain through standard transcardial perfusion13 with 20 ml 0.01 M phosphate buffered saline (PBS) solution, followed by 100 ml of paraformaldehyde 4% in 0.01 M PBS. Carefully adjust the pH of both solutions to 7.6: slightly lower values of the pH may quench the fluorescence of EGFP.</li>
	<li>Post-fix the excised brains overnight in paraformaldehyde 4% at 4 &#176;C.</li>
	<li>Rinse two times (30 min each) in 0.01 M PBS and store in 0.01 M PBS at 4 &#176;C.</li>
</ol>
2. Dehydration and Clearing
Note: the protocol for dehydration and clearing closely follows the one described by Becker et al.10
<ol>
	<li>To remove peroxides from the dehydration solvent (tetrahydrofuran, THF), which strongly quench XFP fluorescence, filter THF with basic activated aluminum oxide (activity grade Brockman I, about 250 g/L of THF) using a chromatography column. Avoid cracking of the column, which would reduce peroxide removal.</li>
	<li>Add stabilizer (butylated hydroxytoluene, 250 mg/L) to the final solution, even if the THF was already stabilized before filtration. Stabilizer is in fact retained by aluminum oxide: THF without stabilizer may develop large amounts of peroxides and can be explosive and dangerous.</li>
	<li>Dehydrate the whole mouse brain in a graded series of THF in pure water: 50%, 70%, 80%, 90%, 96%, 100%, 1 hr each, then 100% overnight. Place the vial with the sample on a rotating wheel. To avoid light exposure, wrap the vials with an aluminum foil.</li>
	<li>Filter the clearing solvent (dibenzyl ether, DBE) with aluminum oxide (activated basic, activity grade Brockman I, about 250 g/L of DBE) using a filtering funnel.</li>
	<li>Clear the sample in 100% DBE. Change the solution after 3 and 6 hr.</li>
</ol>
3. Specimen Mounting
<ol>
	<li>When the brain looks transparent by eye (typically 1 hr after the second DBE change), mount it on the tipped imaging plate by one of the following methods:</li>
	<li>
	<ol>
		<li>Direct mounting - Gently plunge the sample on one of the tips (Figure 3a).</li>
		<li>Agarose disc - Plunge a disc made of 6% agarose gel on the tips. Gently fix the sample on the agarose disc with acrylic glue (Figure 3b). Wait about 1 min for curing.</li>
		<li>Agarose beaker - Plunge a beaker made of 3% agarose gel on the tips. Gently place the sample on the bottom of the beaker (Figure 3c).</li>
	</ol>
	</li>
</ol>
Note 1: Mounting in agarose beaker should be preferred when it is important imaging the sample in its entirety; however, the agarose gel surrounding the specimen gives rise to additional light scattering which could be detrimental to image quality. If a small part of the sample could be excluded from imaging, it is better to mount the specimen on agarose disk or directly on the tip. The former method is best suited to keep the brain horizontal (excluding from imaging a ventral or dorsal portion of the brain), the latter to keep the brain vertical (excluding from imaging a part of olfactory bulbs or of spinal cord).
Note 2: the agarose gel should be prepared in water, then dehydrated with THF (50% and 100% 4 hr each) and cleared in DBE 100% for 4 hr or until it looks clearly transparent.
<ol start="3">
	<li>After mounting the specimen following one of the previous methods, position the imaging plate inside the specimen chamber using tweezers. Fill the chamber with clearing solution.</li>
</ol>
4. Tomography Preparation
<ol>
	<li>Remove the slit to collect as much fluorescence as possible.</li>
	<li>Illuminate the sample with low laser power to prevent photobleaching.</li>
	<li>Move the sample in the three directions using the software-driven micropositioning system. Identify for each direction the minimum and maximum coordinates for which the sample is illuminated and is within the field of view of the camera (Figure 4a).</li>
	<li>Insert the above coordinates as parameters for the &#39;Pre-tomography&#39; subroutine. Specify also the distance between adjacent stacks to be used in the tomography, and the pre-tomography sampling step in z direction.</li>
	<li>Start the pre-tomography, which collects the images in every stack with the z sampling specified at (4.4).</li>
</ol>
Note: The collected images are used by a software to prepare a rough map of specimen shape and position (Figure 4b), which allows restricting tomography acquisition only to the volume actually occupied by the sample, reducing imaging time and thus photobleaching.
5. Tomography Acquisition
<ol>
	<li>Move the sample outside of the light sheet using the micropositioning system.</li>
	<li>Increase laser power and stop the motion of all the scanning systems, in order to illuminate only a fixed line at the center of the field of view.</li>
	<li>Mount the slit and align it using autofluorescence signal from the clearing solution. You should clearly see the slit line at the center of the field of view.</li>
	<li>Re-activate the scanning system, reduce laser power and move the sample inside the light sheet using the micropositioning system. Note that the laser power used with the slit will be higher than that used without it, since the spatial filter blocks a non-negligible fraction of light.</li>
	<li>Adjust the amplitude and offset of the scanning and de-scanning systems with the control software, until the images look clear and bright.</li>
	<li>Run the tomography acquisition software, after setting the appropriate z step for the experiment. The volume will be imaged in many parallel, partially overlapping stacks (Figure 5a), and the images will be saved in a hierarchical folder structure.</li>
</ol>
6. Stitching and Visualization
<ol>
	<li>Complete the acquired volume with synthetic black images (i.e. images of the same type and dimensions of the collected ones-, but with zero intensity) using an automated software (Figure 4c). This step is mandatory in order for the stitching software to deal with a complete cubic volume.</li>
	<li>Launch Vaa3D software (freely downloadable from <a href="http://www.vaa3d.org/" target="_blank">http://www.vaa3d.org/</a>) with the plugins TeraStitcher and TeraManager installed.</li>
	<li>Load the TeraStitcher plugin. Select the directory containing the imaged volume, indicate the relative orientations of the axes (with respect to a reference right-handed coordinate system) and the voxel size.</li>
	<li>Launch the first part of the stitching. The software will calculate the relative displacement between couples of stacks, and find an overall optimal placement for all the stacks together (Figure 5b).</li>
	<li>Select to save the stitched volume in single-resolution or in multi-resolution format. The latter enables for multi-resolution visualization with the TeraManager plugin. In both cases, if the higher resolution image is larger than a few gigabytes, also select the multi-stack save modality and specify the size of individual substacks. This will enable efficient access to the stored data.</li>
	<li>Launch the second part of the stitching. The software will merge the aligned stacks, and save them in either single- or multi-stack mode (Figures 5c and 5d). At the end close the TeraStitcher plugin.</li>
	<li>Load the TeraManager plugin. Select the folder with the multi-stacked multi-resolution volume, and indicate voxel size and axes orientations. The volume will be loaded at the minimum resolution.</li>
	<li>To zoom to a higher resolution, select a landmark using the right-click modality of Vaa3D, then zoom in using mouse scroll. Alternatively you can directly select the ROI to zoom in with a right-click, or specify the coordinates of the volume of interest.</li>
	<li>To zoom back to lower resolution, simply use the mouse scroll.</li>
</ol>

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Microsc. 231, 134-143 (2008).</li><li>Li, 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=Micro-optical+sectioning+tomography+to+obtain+a+high-resolution+atlas+of+the+mouse+brain.">Micro-optical sectioning tomography to obtain a high-resolution atlas of the mouse brain.</a> Science. 330, 1404-1408 (2010).</li><li>Gong, H., 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=Continuously+tracing+brain-wide+long-distance+axonal+projections+in+mice+at+a+one-micron+voxel+resolution.">Continuously tracing brain-wide long-distance axonal projections in mice at a one-micron voxel resolution.</a> Neuroimage. 74, 87-98 (2013).</li><li>Ragan, T., 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=Serial+two-photon+tomography+for+automated+ex+vivo+mouse+brain+imaging.">Serial two-photon tomography for automated ex vivo mouse brain imaging.</a> Nat. Methods. 9, 255-258 (2012).</li><li>Keller, P. J., Dodt, H. U. <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+microscopy+of+living+or+cleared+specimens.">Light sheet microscopy of living or cleared specimens.</a> Curr. Opin. Neurobiol. 22, 138-143 (2012).</li><li>Erturk, 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=Three-dimensional+imaging+of+solvent-cleared+organs+using+3DISCO.">Three-dimensional imaging of solvent-cleared organs using 3DISCO.</a> Nat. Protoc. 7, 1983-1995 (2012).</li><li>Dodt, H. U., 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=Ultramicroscopy:+three-dimensional+visualization+of+neuronal+networks+in+the+whole+mouse+brain.">Ultramicroscopy: three-dimensional visualization of neuronal networks in the whole mouse brain.</a> Nat. Methods. 4, 331-336 (2007).</li><li>Silvestri, L., Bria, A., Sacconi, L., Iannello, G., Pavone, F. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Confocal+light+sheet+microscopy:+micron-scale+neuroanatomy+of+the+entire+mouse+brain.">Confocal light sheet microscopy: micron-scale neuroanatomy of the entire mouse brain.</a> Opt. Express. 20, 20582-20598 (2012).</li><li>Becker, K., Jahrling, N., Saghafi, S., Weiler, R., Dodt, H. 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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=V3D+enables+real-time+3D+visualization+and+quantitative+analysis+of+large-scale+biological+image+data+sets.">V3D enables real-time 3D visualization and quantitative analysis of large-scale biological image data sets.</a> Nat. Biotechnol. 28, 348-353 (2010).</li><li>Gage, G. J., Kipke, D. R., Shain, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+animal+perfusion+fixation+for+rodents.">Whole animal perfusion fixation for rodents.</a> J. Vis. Exp. (65), e3564 (2012).</li><li>Hama, H., 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=Scale:+a+chemical+approach+for+fluorescence+imaging+and+reconstruction+of+transparent+mouse+brain.">Scale: a chemical approach for fluorescence imaging and reconstruction of transparent mouse brain.</a> Nat. Neurosci. 14, 1481-1488 (2011).</li><li>Booth, M. J., Neil, M. A. A., Wilson, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aberration+correction+for+confocal+imaging+in+refractive-index-mismatched+media.">Aberration correction for confocal imaging in refractive-index-mismatched media.</a> J. Microsc. 192, 90-98 (1998).</li><li>Huisken, J., Swoger, J., Del Bene, F., Wittbrodt, J., Stelzer, E. 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The authors declare that they have no competing financial interests.

CLSM coupled with chemical clearing represent a powerful tool to study the neuroanatomy of entire mouse brains labeled with fluorescent probes, either proteins or synthetic dyes. Since the light emitted outside the focal plane is blocked by a physical spatial filter, high-contrast imaging is possible also in thick specimens, while maintaining the high frame rates typical of light sheet architecture.
The main issue to deal with when using the described methods is the maximization of the contrast. In fact, the available signal is reduced both by chemical and optical factors. From the chemical point of view, the clearing procedures based on organic solvent may quench emission from fluorescent proteins and other fluorescent dyes7. Filtration of the solvent to remove peroxides, as described by Becker et al.10, allows maintaining a good level of fluorescence also in solvent-cleared tissues. Published water-based clearing methods14 do not reduce fluorescence efficiency, but result in poorer transparency when dealing with entire murine brains, at least with linear optical techniques.
On the other hand, at the moment there are no commercial objectives with long working distance corrected for the high refractive index (n &asymp; 1.56) clearing solutions used. Thus, the refractive index mismatch between the medium in which the sample is immersed and the design one of the objective gives rise to strong spherical aberrations. Besides reducing the resolution of the microscope, such aberrations result in a drop of image intensity and contrast, since the light is spread on a wider area. Possible workarounds to this issue include the use of adaptive optics15 and/or static aspheric correctors.
Any improvement in image contrast and intensity readily affects also imaging speed, since a shorter integration time per image can be chosen. At the moment CLSM can afford an imaging speed of about 106 &mu;m3/sec, with a camera exposure time of 100 msec 9. At this speed it takes from 1-3 days to image an entire mouse brain, depending on sample size. Although no significant degradation in image quality is observed throughout such long imaging session, mostly because of the intrinsic low photobleaching of light sheet illumination6, the long time needed to image a single mouse brain could in practice reduce the applicability of CLSM. A 10-fold increase in system efficiency, coupled with the use of a high-speed camera for detection, would reduce imaging time to several hours, allowing a routine use of the described protocol.
Despite all the abovementioned limitations, CLSM imaging coupled to chemical clearing of tissues permits to reconstruct entire mouse brains with micron-scale resolution in less than a week. Other optical methods for whole brain imaging afford higher resolution than CLSM, but at the price of longer preparation and/or imaging time2-4, or of a sparse z sampling5.
The methods described in this article can be used in combination with different strategies for fluorescent labeling: transgenic animals expressing fluorescent proteins in selected neuronal subsets, viral transfection, intraparenchymal dye injection, etc. In practice, all staining strategies preceding animal sacrifice are compatible with CLSM. In fact, post-mortem fluorescent labeling of entire mouse brains has not been demonstrated up to now; anyway, if such kind of staining would be possible in the near future, the number of specimens that could be reconstructed with CLSM will become much larger, potentially including human brain tissue.
In conclusion, confocal light sheet microscopy used in combination with tissue clearing and multi-resolution data management may allow researchers to reconstruct and analyze macroscopic specimens with resolution in the micron scale, with technological efforts that are well at the hand of most laboratories. Potential applications of CLSM are of course not limited to neuroscience, but spans all the research fields in which light sheet microscopy has already been used, as embryogenesis16,17 and anatomy of small animals18.

When compared to our understanding of the molecular and cellular mechanisms underlying brain function, our knowledge of fine neuroanatomy looks still in its infancy1. In fact, we still lack comprehensive maps of the position of neuronal somata or of long-range projections throughout the brain. Actually, many technological efforts are devoted to reconstruct the mouse brain with microscopic resolution. Optical methods based on serial sectioning of resin-embedded samples, as Knife-Edge Scanning Microscopy (KESM)2, Micro-Optical Sectioning Tomography (MOST)3 and fluorescence-MOST (fMOST)4, can provide sub-micron three-dimensional resolution imaging of entire mouse brains. However, the total time needed for the preparation and imaging of a single sample is of the order of several weeks, limiting the practical application of such methods. Another technique based on serial sectioning (but without resin embedding), Serial Two-Photon tomography (STP)5, allows high three-dimensional resolution imaging. However, this technique has been demonstrated in whole mouse brains only with a very sparse sampling, collecting 1 section every 50 &#956;m 5, and to our knowledge STP have not produced yet a full-sampling reconstruction of a murine brain.
A cutting-edge optical method to reconstruct whole specimens in three dimensions at high speed is light sheet microscopy6. In this optical scheme the sample is illuminated with a thin sheet of light and the fluorescence emission is collected along an axis perpendicular to the illumination plane. In this way only in-focus fluorophores are excited and it is possible to achieve optical sectioning in wide field configuration, guaranteeing high frame rates. When coupled to clearing protocols based on the substitution of water with a refractive index matching liquid7, light sheet microscopy has been applied to macroscopic specimens, as whole mouse brains8. However, specimen-induced light scattering, which affects even cleared specimens, degrades the light sheet leading to the excitation of out-of-focus fluorophores which adds an overall blurring to the acquired images.
To selectively collect only in-focus photons, we recently coupled light sheet illumination to a confocal detection scheme9. In confocal light sheet microscopy (CLSM), out-of-focus and scattered photons are rejected by a linear spatial filter (slit) prior to detection (Figure 1). To achieve confocal operation in light sheet architecture, the illumination is produced by means of a scanning line, and a de-scanning system in the detection path creates a static image of the excitation scanning line at the location of the slit. A third scanning system reconstructs a two-dimensional image on the camera sensor. CLSM affords a 100% contrast enhancement in cleared mouse brains with respect to conventional light sheet microscopy, while maintaining a 10 Hz frame rate. With CLSM it is possible to reconstruct entire mouse brains with resolution of 2 &#956;m in XY and 9 &#956;m in Z, and with sufficient contrast to discern single fluorescently labeled neurons.
In this Article we describe the experimental pipeline to reconstruct a mouse brain with CLSM. Aldehyde-fixed mouse brains are dehydrated in a graded series of peroxide-free tetrahydrofuran and subsequently cleared in peroxide-free dibenzyl ether (Figure 2), following the protocol developed by Becker et al.10 The cleared specimen is then carefully mounted on a tipped plate and positioned in the imaging chamber of a custom-made confocal light sheet microscope. The sample can be directly mounted by plunging it on a tip of the plate (Figure 3a); alternatively it can be glued on a disc of cleared agarose gel (Figure 3b) or placed in the cavity of a beaker made of cleared agarose gel (Figure 3c). An optical tomography of the entire brain is then performed, as many parallel adjacent stacks are acquired to cover all the brain volume (Figure 4). A pre-tomography sampling, which produces a rough map of the specimen position and shape, guarantees that imaging is performed only in the volume occupied by the sample. The tomography stacks are subsequently stitched together with custom-made software and saved in a multi-resolution hierarchical scheme11. Mouse brains can eventually be explored with prototypal multi-resolution Google Maps-like visualization software. Both these software instruments are integrated as plugins in the open-source platform Vaa3D12.
We finally show representative images of mouse brains to demonstrate the capabilities of the described experimental pipeline in studying fine murine neuroanatomy.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Phosphate Buffered Saline Tablets</td>
 <td>Sigma-Aldrich</td>
 <td>P4417_100TAB</td>
 <td></td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>Agar Scientific</td>
 <td>R1018</td>
 <td></td>
 </tr>
 <tr>
 <td>Peroxide-free Tetrahydrofuran</td>
 <td>Sigma-Aldrich</td>
 <td>186562</td>
 <td></td>
 </tr>
 <tr>
 <td>Aluminum oxide activated, basic, Brockmann I</td>
 <td>Sigma-Aldrich</td>
 <td>199443</td>
 <td></td>
 </tr>
 <tr>
 <td>Dibenzyl ether</td>
 <td>Sigma-Aldrich</td>
 <td>108014</td>
 <td></td>
 </tr>
 <tr>
 <td>Butylated hydroxytoluene</td>
 <td>Sigma-Aldrich</td>
 <td>W218405 </td>
 <td></td>
 </tr>
 <tr>
 <td>Agarose Type III-A, High EEO</td>
 <td>Sigma-Aldrich</td>
 <td>A9793</td>
 <td></td>
 </tr>
 <tr>
 <td>Acrylic glue</td>
 <td>Bostik</td>
 <td>D2498</td>
 <td></td>
 </tr>
 <tr>
 <td>Vaa3D software</td>
 <td>open source</td>
 <td></td>
 <td>freely downloadable from <a href="http://www.vaa3d.org/" target="_blank">http://www.vaa3d.org/</a> </td>
 </tr>
 </tbody>
</table>

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.

The described protocol can be used to reconstruct with micron-scale resolution either entire mouse brains or excised parts, without the need for physical sectioning. As a representative result, in Figure 6 the whole cerebellum of an L7-GFP mouse (post natal day 10) is shown. In this animal all Purkinje neurons are labeled with EGFP. If we zoom in, the typical lamellar structure of the cerebellar cortex can be seen (Figure 7). Further zooming in allows clearly distinguishing each Purkinje cell soma (Figure 8). The described protocol can thus be used to screen neuronal spatial organization in various neurodevelopment studies.
As a second representative result, we present images from the unsectioned brain of an adult thy1-GFP-M mouse. In this transgenic animal EGFP is expressed in a random sparse neuronal subset. The right half of the brain is shown at in Figure 9. As we zoom-in further and further (Figures 10 and 11), neuronal processes become distinguishable. This result demonstrates that the described protocol allows micron-scale resolution imaging in entire adult mouse brains, opening the possibility of studying whole-brain anatomy at cellular resolution in mouse models of neurodegenerative disease.
<img alt="Figure 1" fo:content-width="6in" src="/files/ftp_upload/50696/50696fig1.jpg" /> 
Figure 1. Optical scheme of confocal light sheet microscopy (CLSM). (a) Top view of the apparatus, showing the excitation pathway. Laser emission from a 488 nm diode-pumped solid state (DPSS) laser, after expansion and collimation by a first telescope, enters an acousto-optic tunable filter (AOTF) which regulates beam intensity. Then, a second telescope further expands the beam. A galvo mirror vertically (along y) scans the beam, which is focused by a lens inside the specimen chamber. (b) Lateral view of the apparatus, showing the detection pathway. Light collected by the detection objective is vertically descanned by a galvo mirror and focused by a lens, producing a fixed image of the moving excitation line. At the position of the fixed image a linear spatial filter (slit) is placed. After the slit, a galvo mirror placed inside a 4f lens system reconstructs a 2-dimentional image on the chip of an electron-multiplying charge-coupled device (EM-CCD). A fluorescence bandpass filter remove all excitation stray light before detection. Representative rays of excitation, detection and bandpass-filtered detection light are depicted by blue, light blue and green rays, respectively. The focal lengths of all lenses actually used in the microscope are indicated. <a href="https://www.jove.com/files/ftp_upload/50696/50696fig1large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/50696/50696fig2.jpg" /> 
Figure 2. Specimen clearing. Formaldehyde-fixed mouse brains, which are stored in PBS 0.1 M at 4 &#176;C (a), are dehydrated in a graded series of tetrahydrofuran (THF, b) and subsequently cleared in 100% dibenzyl ether (DBE, c). The dehydration step causes also some tissue shrinkage.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/50696/50696fig3.jpg" /> 
Figure 3. Specimen mounting for CLSM imaging. The cleared brain can be directly plunged on the tipped mounting plate (a), glued on a cleared agarose disc (b), or inserted in a cleared agarose beaker (c).
<img alt="Figure 4" fo:content-width="6in" src="/files/ftp_upload/50696/50696fig4.jpg" /> 
Figure 4. Tomography acquisition. The pre-tomography routine samples a parallelepiped containing the brain (a) to define the number and length of parallel adjacent stacks needed to image all the specimen (b). After tomography, black images are generated to complete the virtual acquired volume to a parallelepiped (c).
<img alt="Figure 5" fo:content-width="4in" src="/files/ftp_upload/50696/50696fig5.jpg" /> 
Figure 5. Volume stitching. The collected image stacks partially overlap between each other (a), allowing their precise alignment with the TeraStitcher plugin (b). The plugin then merges the aligned stacks in either a single-stacked (c) or multi-stacked (d) multi-resolution representation.
<img alt="Figure 6" fo:conteint-width="5in" src="/files/ftp_upload/50696/50696fig6.jpg" /> 
Figure 6. Whole cerebellum of a P10 L7-GFP mouse, in which all the Purkinje neurons are labeled with EGFP. Scale bar, 1 mm.
<img alt="Figure 7" fo:content-width="4in" src="/files/ftp_upload/50696/50696fig7.jpg" /> 
Figure 7. Zoom-in of a portion of the cerebellum shown in Figure 6, showing the typical lamellar structure of the cerebellar cortex. Scale bar, 500 &#956;m.
<img alt="Figure 8" fo:content-width="4in" src="/files/ftp_upload/50696/50696fig8.jpg" /> 
Figure 8. Further zoom-in of a portion of the cerebellum shown in Figure 6. Purkinje cells soma can be clearly distinguished. Scale bar, 200 &#956;m.
<img alt="Figure 9" fo:content-width="5in" src="/files/ftp_upload/50696/50696fig9.jpg" /> 
Figure 9. Right half of the brain from an adult thy1-GFP-M mouse, characterized by a sparse unspecific neuronal EGFP labeling. Scale bar, 1 mm.
<img alt="Figure 10" fo:content-width="4in" src="/files/ftp_upload/50696/50696fig10.jpg" /> 
Figure 10. Zoom-in of a portion of the half brain shown in Figure 9, showing part of the hippocampus and of cortex. Scale bar, 200 &#956;m.
<img alt="Figure 11" fo:content-width="4in" src="/files/ftp_upload/50696/50696fig11.jpg" /> 
Figure 11. Further zoom-in of a portion of the half brain shown in Figure 9. Several neurites in the cortex can be clearly distinguished. Scale bar, 50 &#956;m.

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

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 ...

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Research

JoVE Journal

Neuroscience

16.6K vistas.  European Laboratory for Non-linear Spectroscopy (LENS).  En este artículo se describe el procedimiento experimental completo para reconstruir, con alta resolución, la anatomía del cerebro multa de cerebros de ratón marcados con fluorescencia. El protocolo descrito incluye la preparación de muestras y el desmonte, la muestra de montaje para imágenes, datos post-procesamiento y visualización multi-escala. 

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

Optical Recording of Suprathreshold Neural Activity with Single-cell and Single-spike Resolution

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also propagation of suprathreshold spiking activity involves populations of neurons. Empirical studies addressing cortical function directly thus require recordings from populations of neurons with high resolution. Here we describe an optical method and a deconvolution algorithm to record neural activity from up to 100 neurons with single-cell and single-spike resolution. This method relies on detection of the transient increases in intracellular somatic calcium concentration associated with suprathreshold electrical spikes (action potentials) in cortical neurons. High temporal resolution of the optical recordings is achieved by a fast random-access scanning technique using acousto-optical deflectors (AODs)1. Two-photon excitation of the calcium-sensitive dye results in high spatial resolution in opaque brain tissue2. Reconstruction of spikes from the fluorescence calcium recordings is achieved by a maximum-likelihood method. Simultaneous electrophysiological and optical recordings indicate that our method reliably detects spikes (&gt;97% spike detection efficiency), has a low rate of false positive spike detection (&lt; 0.003 spikes/sec), and a high temporal precision (about 3 msec) 3. This optical method of spike detection can be used to record neural activity in vitro and in anesthetized animals in vivo3,4.

Understanding the function of the vertebrate central nervous system requires recordings from many neurons because cortical function arises on the level of populations of neurons. Here we describe an optical method to record suprathreshold neural activity with single-cell and single-spike resolution, dithered random-access scanning. This method records somatic fluorescence calcium signals from up to 100 neurons with high temporal resolution. A maximum-likelihood algorithm deconvolves the underlying suprathreshold neural activity from the somatic fluorescence calcium signals. This method reliably detects spikes with high detection efficiency and a low rate of false positives and can be used to study neural populations in vitro and in vivo.

Signaling of information in the vertebrate central nervous system is often carried by populations of neurons rather than individual neurons. Also ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

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 ...

Free-floating Immunostaining of Mouse Brains

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the regulation of energy metabolism and other neurobiological processes. However, the quality, reliability, and reproducibility of brain histology results may vary among laboratories. For each staining experiment, it is necessary to optimize the key procedures based on differences in species, tissues, targeted proteins, and the working conditions of the reagents. This paper demonstrates a reliable workflow in detail, including intra-aortic perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging, which can be followed easily by researchers in this field.
Also discussed are how to modify these procedures to satisfy the individual needs of researchers. To illustrate the reliability and efficiency of this protocol, perineuronal nets were stained with biotin-labeled Wisteria florbunda agglutinin (WFA) and arginine vasopressin (AVP) with an anti-AVP antibody in the mouse brain. Finally, the critical details for the entire procedure have been addressed, and the advantages of this protocol compared to those of other protocols. Taken together, this paper presents an optimized protocol for free-floating immunostaining of mouse brain tissue. Following this protocol makes this process easier for both junior and senior scientists to improve the quality, reliability, and reproducibility of immunostaining studies.

This protocol describes an efficient and reproducible approach for mouse brain histological studies, including perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging.

Immunohistochemical staining of mouse brains is a routine technique commonly used in neuroscience to investigate central mechanisms underlying the ...

Characterization of Neuromuscular Junctions in Mice by Combined Confocal and Super-Resolution Microscopy

Neuromuscular junctions (NMJs) are highly specialized synapses between lower motor neurons and skeletal muscle fibers that play an essential role in the transmission of molecules from the nervous system to voluntary muscles, leading to contraction. They are affected in many human diseases, including inherited neuromuscular disorders such as Duchenne muscular dystrophy (DMD), congenital myasthenic syndromes (CMS), spinal muscular atrophy (SMA), and amyotrophic lateral sclerosis (ALS). Therefore, monitoring the morphology of neuromuscular junctions and their alterations in disease mouse models represents a valuable tool for pathological studies and preclinical assessment of therapeutic approaches. Here, methods for labeling and analyzing the three-dimensional (3D) morphology of the pre- and postsynaptic parts of motor endplates from murine teased muscle fibers are described. The procedures to prepare samples and measure NMJ volume, area, tortuosity and axon terminal morphology/occupancy by confocal imaging, and the distance between postsynaptic junctional folds and acetylcholine receptor (AChR) stripe width by super-resolution stimulated emission depletion (STED) microscopy are detailed. Alterations in these NMJ parameters are illustrated in mutant mice affected by SMA and CMS.

This protocol describes a method for morphometric analysis of neuromuscular junctions by combined confocal and STED microscopy that is used to quantify pathological changes in mouse models of SMA and ColQ-related CMS.

Neuromuscular junctions (NMJs) are highly specialized synapses between lower motor neurons and skeletal muscle fibers that play an essential role in the ...

Registration of Calcium Transients in Mouse Neuromuscular Junction with High Temporal Resolution using Confocal Microscopy

Estimation of the presynaptic calcium level is a key task in studying synaptic transmission since calcium entry into the presynaptic cell triggers a cascade of events leading to neurotransmitter release. Moreover, changes in presynaptic calcium levels mediate the activity of many intracellular proteins and play an important role in synaptic plasticity. Studying calcium signaling is also important for finding ways to treat neurodegenerative diseases. The neuromuscular junction is a suitable model for studying synaptic plasticity, as it has only one type of neurotransmitter. This article describes the method for loading a calcium-sensitive dye through the cut nerve bundle into the mice's motor nerve endings. This method allows the estimation of all parameters related to intracellular calcium changes, such as basal calcium level and calcium transient. Since the influx of calcium from the cell exterior into the nerve terminals and its binding/unbinding to the calcium-sensitive dye occur within the range of a few milliseconds, a speedy imaging system is required to record these events. Indeed, high-speed cameras are commonly used for the registration of fast calcium changes, but they have low image resolution parameters. The protocol presented here for recording calcium transient allows extremely good spatial-temporal resolution provided by confocal microscopy.

The protocol describes the method of loading a fluorescent calcium dye through the cut nerve into mouse motor nerve terminals. In addition, a unique method for recording fast calcium transients in the peripheral nerve endings using confocal microscopy is presented.

Estimation of the presynaptic calcium level is a key task in studying synaptic transmission since calcium entry into the presynaptic cell triggers a ...

Coherent Anti-Stokes Raman Spectroscopy (CARS) Application for Imaging Myelination in Brain Slices

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature vibrations of molecules. However, these vibrational signatures are also characteristic of molecules within anatomical tissue such as the brain, making it increasingly useful and applicable for Neuroscience applications. For example, CARS can measure lipids by specifically exciting chemical bonds within these molecules, allowing for quantification of different aspects of tissue, such as myelin involved in neurotransmission. In addition, compared to other techniques typically used to quantify myelin, CARS can also be set up to be compatible with immunofluorescent techniques, allowing for co-labeling with other markers such as sodium channels or other components of synaptic transmission. Myelination changes are an inherently important mechanism in demyelinating diseases such as multiple sclerosis or other neurological conditions such as Fragile X Syndrome or autism spectrum disorders is an emerging area of research. In conclusion, CARS can be utilized in innovative ways to answer pressing questions in Neuroscience and provide evidence for underlying mechanisms related to many different neurological conditions.

Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices

Visualizing myelination is an important goal for many researchers studying the nervous system. CARS is a technique that is compatible with immunofluorescence that can natively image lipids within tissue such as the brain illuminating specialized structures such as myelin.

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature ...

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 ...

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.

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 ...

Mouse Retina OCT Fibergram Alignment with Confocal Images

Alignment of Visible-Light Optical Coherence Tomography Fibergrams with Confocal Images of the Same Mouse Retina

In recent years, in vivo retinal imaging, which provides non-invasive, real-time, and longitudinal information about biological systems and processes, has been increasingly applied to obtain an objective assessment of neural damage in eye diseases. Ex vivo confocal imaging of the same retina is often necessary to validate the in vivo findings especially in animal research. In this study, we demonstrated a method for aligning an ex vivo confocal image of the mouse retina with its in vivo images. A new clinical-ready imaging technology called visible light optical coherence tomography fibergraphy (vis-OCTF) was applied to acquire in vivo images of the mouse retina. We then performed the confocal imaging of the same retina as the &#34;gold standard&#34; to validate the in vivo vis-OCTF images. This study not only enables further investigation of the molecular and cellular mechanisms but also establishes a foundation for a sensitive and objective evaluation of neural damage in vivo.

Author Spotlight: Advancements in In Vivo and Ex Vivo Retinal Imaging for Improved Glaucoma Diagnosis and Treatment 

The present protocol outlines the steps for aligning in vivo visible-light optical coherence tomography fibergraphy (vis-OCTF) images with ex vivo confocal images of the same mouse retina for the purpose of verifying the observed retinal ganglion cell axon bundle morphology in the in vivo images.

In recent years, in vivo retinal imaging, which provides non-invasive, real-time, and longitudinal information about biological systems and processes, has ...