We would like to thank Jacob Solis for his assistance in video editing.
This work was funded in part by an NSF CAREER Award DBI-0953902 to MJ Levene.

1. Animal Perfusion5 and Whole Mouse Brain Clearing
<ol>
	<li>The entire length of the procedure can vary depending on the length of time used per dehydration step, but in total the whole process can be conducted in two days.</li>
	<li>Weigh YFP mice and then deeply anesthetize with an intraperitoneal injection of ketamine/xylazine (100 mg/kg:10 mg/kg).</li>
	<li>Confirm a surgical plane of deep anesthesia before proceeding to surgery. Check the animal every 5 min to see if it reacts to a firm toe or tail pinch. If the animal reacts, a supplemental dose (1/3 of original dose) of ketamine/xylazine is required.</li>
	<li>Once deeply anesthetized, restrain the mouse by adhering each limb to a surgical bed using lab tape so that the mouse is in a supine position (dorsal recumbancy), exposing its chest for surgery. The surgical bed is usually made of a metal or plastic mesh and placed into a sink or on top of a lipped-pan so that blood and fixative waste can easily be collected.</li>
	<li>To begin, make an incision below the xyphoid process. Cut along the base of the rib cage using scissors and tweezers and pull back skin as the cut is made. Make two cuts along either side of the mouse sternum (through the ribs) to create a flap of tissue that is held away from the chest cavity using a hemostat to leave the heart exposed.</li>
	<li>Insert a 23 g needle into the left ventricle of the heart and make a small incision in the muscle wall of the right atrium to allow blood to escape. Please refer to JoVE article 2497 for a video of this procedure.5</li>
	<li>Immediately after the right atrium is cut, begin a perfusion with 4 &deg;C phosphate buffered saline, PBS, (pH 7.2) until blood is no longer observed draining from the right atrium of the heart (30 - 40 ml at a rate of about 5 ml/min).</li>
	<li>Once all the blood has been drained (fluid exiting the right atrium is clear), switch the perfusion medium to a chilled 4% PFA solution. Perfuse until the mouse's body becomes noticeably rigid and cold to the touch (approx. 30 - 40 ml at a rate of about 5 ml/min). (Concentrated 16% PFA solution is diluted according to manufacturer's instructions and NaOH is added until pH 7.2 is reached. Wear gloves and a lab coat and handle and mix chemicals inside a fume hood.)</li>
	<li>After perfusion, remove the mouse from the surgical bed and decapitate to begin the excision of the brain.</li>
	<li>Using forceps and iris scissors, remove the skull in small sections starting from the back of the skull and moving forward. Make small cuts every 2 - 4 mm with the scissors across the skull while using the forceps to carefully pull the bone away from the brain in small sections. Do this until the entire top surface of the brain is exposed.</li>
	<li>Excise the brain from the skull using a 5 mm wide, flat spatula and place into a glass vial. Submerge in 4% PFA for 6 hr at 4 &deg;C for post fixation.</li>
	<li>After post fixation, wash the brain twice in room temperature PBS by pouring the PFA out of the glass vial and replacing it with PBS. Swirl the brain in PBS solution before pouring out and replacing with PBS for a second washing.</li>
	<li>Dehydrate the brain at room temperature by a graded series of ethanol incubations (once in 50%, 70%, 95%, and twice in 100%) at 2 hr per incubation, and then 12 hr for the second 100% ethanol incubation, to extract the water from the fixed tissue. For each incubation, pour out the previous ethanol solution from the glass vial and replace it with the subsequent solution until the brain is completely submerged.</li>
	<li>After the second incubation in 100% ethanol, pour out the solution and replace with equal parts ethanol and the clearing solution containing benzyl alcohol and benzyl benzoate (1:2 vol:vol ratio). After 2 hr of incubation, decant this solution and replace with a 100% solution of benzyl alcohol and benzyl benzoate (1:2 vol:vol ratio). The refractive index of the clearing solution is n = 1.54.</li>
	<li>Once in BABB, the brain will become noticeably transparent, within 4 - 5 hr. For best clearing results, leave the brain to clear for 6 days at room temperature while shielded from bright light.</li>
</ol>
2. Microscope Setup
<ol>
	<li>Once the brain is cleared and ready for imaging, affix it to the bottom of a Petri dish using cyanoacrylate. Allow the adhesive to dry before proceeding.</li>
	<li>After drying, submerge the brain in BABB and place the Petri dish under the microscope objective for imaging.</li>
	<li>We use a multiphoton microscope that incorporates a Mai Tai titanium sapphire laser adjustable between a 710 nm to 990 nm excitation wavelength. The excitation wavelength we use to generate YFP signals is 886 nm. Laser power varies from 30 - 100 mW depending on imaging depth.</li>
	<li>Capture the reflected fluorescent signal using a Nikon 5X objective (NA, 0.5) that allows for large field-of-view imaging (2 x 2 mm).</li>
	<li>Filter the reflected fluorescent signal using a 535/50 bandpass filter and collect it using a GaAsP PMT (H7422PA-40, Hamamatsu, Bridgewater, New Jersey).</li>
	<li>Process images using ScanImage software6 at a resolution of 2048 x 2048 pixels using a scan rate of 2 ms per line to generate high-resolution, YFP images.</li>
	<li>Once imaging is complete, remove the brain from the Petri dish using forceps and store in BABB shielded from light for future imaging. For glued samples, remove the tissue by running a razor blade between the glue and the Petri dish at an angle no greater than 30 degrees. We have stored samples in BABB for up to 1 year with no visible degradation.</li>
</ol>
3. Representative Results
The representative images and videos shown here demonstrate the high-resolution multiphoton imaging capability made possible by optical clearing. Whole brain imaging allows for YFP-labeled neurons in the different layers of the hippocampus and cortex to be clearly visible as deep as 2 mm below the tissue surface. Figure 1 shows a representative image of a coronal view corresponding to the anatomical features of the hippocampus at 2.92 mm caudal to bregma7. The depth of imaging in the sample was 1.94 mm. Some of this difference was due to removal of the cerebellum at the caudal end of the brain and the balance was due to shrinkage from the dehydration process. Another image from the stack shows layer V/VI pyramidal neurons of the neocortex expressing YFP 2 mm beneath the tissue surface (Figure 2). All images were acquired using the Nikon 5X objective and magnification was done using the digital zoom feature for image acquisition in ScanImage software.
By zooming in on the neocortex, the individual axons and neuron cell bodies of pyramidal neurons in Layer V of the neocortex are clearly distinguishable up to 1.02 mm beneath the tissue surface (Figure 3). Using the image stack from Figure 3, a 3D reconstruction of the neuron region was made using ImageJ software8 (Figure 4).
The high-resolution capability of MPM also allows us to present images of whole, individual neurons. Here we show a reconstruction of a Layer V pyramidal neuron of the neocortex in which individual dendritic processes are clearly visible (Figure 5).
<img alt="Figure 1" src="/files/ftp_upload/3848/3848fig1.jpg" /> 
Figure 1. Representative image from a 1.2 mm image stack (0.8 - 2 mm beneath the tissue surface) of whole mouse brain showing neocortex and the different layers of the hippocampus. Features of the hippocampus are visible 1.1 mm below the tissue surface and have been labeled using the following code: DG = dentate gyrus, GrDG = granular layer of DG, Lmol = lacunosum moleculare layer, Mol = molecular layer DG, Or = oriens layer, PoDG = polymorph layer DG, Py = pyramidal cell layer, Rad = stratum radiatum.7 Image is 1.8 x 1.8 mm in size, scale bar = 200 &mu;m. The rostral end of the brain was affixed to a Petri dish with the caudal end facing up toward the objective. This facilitated imaging in the coronal plane.
<img alt="Figure 2" src="/files/ftp_upload/3848/3848fig2.jpg" /> 
Figure 2. Representative frame from the 1.2 mm image stack (0.8 - 2 mm beneath the tissue surface) of whole mouse brain highlighting the neocortex. Pyramidal cells and processes expressing YFP in layer V/VI of cortex are visible 2 mm below the tissue surface. The degree of labeling is consistent with previous reports of the sparse labeling in the neocortex.9 Inset (right) is enlarged from boxed area on left. Image is 1.8 x 1.8 mm in size, scale bar = 200 &mu;m (50 &mu;m inset).
<img alt="Figure 3" src="/files/ftp_upload/3848/3848fig3.jpg" /> 
Figure 3. Representative frame taken 774 &mu;m beneath the tissue surface from the image stack of layer V pyramidal neurons in the neocortex of the brain. Stack goes from 700 &mu;m to 1,020 &mu;m beneath the tissue surface. An 8X digital zoom was used to capture details such as fine processes. Image is 225 x 225 &mu;m in size, scale bar = 20 &mu;m.
<img alt="Figure 4" src="/files/ftp_upload/3848/3848fig4.jpg" /> 
Figure 4. Representative image of 3D reconstruction (225 x 225 x 320 &mu;m) of the image stack in Figure 3. Image is 225 x 225 &mu;m in size, scale bar = 28 &mu;m.
<img alt="Figure 5" src="/files/ftp_upload/3848/3848fig5.jpg" /> 
Figure 5. High-resolution reconstructed image of a layer V pyramidal neuron of the neocortex using a 10X digital zoom. Neuron measures 485 &mu;m in length, scale bar = 25 &mu;m. The field of view for each image tile is 180 &mu;m x 180 &mu;m. Each tile is at a different depth (z-level). The cortical apical dendrites do not, in general, express YFP as well as the soma. To have both in the same field of view, as shown here, the power was lowered to minimize saturation of soma; this makes the fine process look dimmer. Our emphasis here was on demonstrating field-of-view rather than fine detail. To reveal finer details, use a digital zoom, focus on a region without soma, and increase the laser power.

<ol>
	<li>Parra, S. G., Chia, T. H., Zinter, J. P., Levene, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiphoton+microscopy+of+cleared+mouse+organs.">Multiphoton microscopy of cleared mouse organs.</a> J. Biomed. Opt. 15, 036017 (2010).</li><li>Vesuna, S., Torres, R., Levene, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiphoton+fluorescence,+second+harmonic+generation,+and+fluorescence+lifetime+imaging+of+whole+cleared+mouse+organs.">Multiphoton fluorescence, second harmonic generation, and fluorescence lifetime imaging of whole cleared mouse organs.</a> J. Biomed. Opt. 16, 106009 (2011).</li><li>Zucker, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+insect+and+mammalian+embryo+imaging+with+confocal+microscopy:+Morphology+and+apoptosis.">Whole insect and mammalian embryo imaging with confocal microscopy: Morphology and apoptosis.</a> Cytometry. A69, 1143-1152 (2006).</li><li>Sakhalkar, H. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+imaging+in+bulk+tissue+specimens+using+optical+emission+tomography:+Fluorescence+preservation+during+optical+clearing.">Functional imaging in bulk tissue specimens using optical emission tomography: Fluorescence preservation during optical clearing.</a> Phys. Med. Biol. 52, 2035-2054 (2007).</li><li>Dazai, J., Spring, S., Cahill, L. S., Henkelman, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple-mouse+Neuroanatomical+Magnetic+Resonance+Imaging.">Multiple-mouse Neuroanatomical Magnetic Resonance Imaging.</a> J. Vis. Exp. (48), e2497 (2011).</li><li>Pologruto, T. A., Sabatini, B. L., Svoboda, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ScanImage:+flexible+software+for+operating+laser+scanning+microscopes.">ScanImage: flexible software for operating laser scanning microscopes.</a> Biomed. Eng. Online. 2, (2003).</li><li>Paxinos, G., Franklin, K. B. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Mouse Brain in Stereotaxic Coordinates. , (2001).</li><li>Abramoff, M. D., Magelhaes, P. J., Ram, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Image+Processing+with+ImageJ.">Image Processing with ImageJ.</a> Biophotonics International. 11, 36-42 (2004).</li><li>Feng, G., Mellor, R. H., Bernstein, M., Keller-Peck, C., Nguyen, Q. T., Wallace, M., Nerbonne, J. M., Lichtman, J. W., Sanes, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+Neuronal+Subsets+in+Transgenic+Mice+Expressing+Multiple+Spectral+Variants+of+GFP.">Imaging Neuronal Subsets in Transgenic Mice Expressing Multiple Spectral Variants of GFP.</a> Neuron. 28, 41-51 (2000).</li><li>Hama, H. <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. Neuro. 14, 1481-1488 (2011).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by Yale University's Institutional Animal Care &amp; Use Committee.

While standard organic dyes are compatible with a range of organic solvents, and therefore do not pose a particular challenge for clearing protocols, fluorescent proteins are often less tolerant of changes in solvent.4 The goal of the present work was to overcome a serious limitation of previous optical clearing protocols where fluorescence of XFPs was lost or severely degraded. The images that are presented here demonstrate that fluorescence from YFP was preserved. The optical clearing technique described ...

<table border="1">
<tbody>
 <tr>
 <td>Name 			of the reagent</td>
 <td>Company</td>
 <td>Catalog 			number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Phosphate 			Buffered Saline </td>
 <td>Sigma-Aldrich, 			Inc.</td>
 <td>D8537</td>
 <td>500 ml, pH 7.2</td>
 </tr>
 <tr>
 <td>Paraformaldehyde </td>
 <td>Electron 			Microscopy Sciences</td>
 <td>15710</td>
 <td>10 x 10 ml, 			16% paraformaldehyde</td>
 </tr>
 <tr>
 <td>Ethyl alcohol</td>
 <td>American 			Bioanalytical</td>
 <td>AB00515-00500</td>
 <td>500 ml, 200 			proof</td>
 </tr>
 <tr>
 <td>Ethyl alcohol</td>
 <td>Pharmco 			Products, Inc.</td>
 <td>111000190</td>
 <td>1 gal., 190 			proof</td>
 </tr>
 <tr>
 <td>Benzyl alcohol</td>
 <td>Sigma-Aldrich, 			Inc.</td>
 <td>402834</td>
 <td>500 ml, 99+%</td>
 </tr>
 <tr>
 <td>Benzyl 			benzoate</td>
 <td>Sigma-Aldrich, 			Inc.</td>
 <td>B6630-IL</td>
 <td>500 ml, &ge; 99%</td>
 </tr>
 <tr>
 <td>5X/0.5 NA 			objective</td>
 <td>Nikon</td>
 <td>AZ Plan Flour 			5X</td>
 <td>15 WD</td>
 </tr>
 </tbody>
</table>

multiphoton microscopy of cleared mouse brain expressing yfp

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Watch this Scientific Journal Video about Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP at JoVE.com

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

multiphoton-microscopy-of-cleared-mouse-brain-expressing-yfp

Research

JoVE Journal

Neuroscience

13.7K Views.  Yale University. Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Video: Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Preparation of Mouse Brain Tissue for Immunoelectron Microscopy

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments (dendrite, dendritic spine, axon, axon terminal, perikaryon), as well as their intracellular organelles and cytoskeleton, in the central nervous system at high spatial resolution. Combined with TEM, pre-embedding immunocytochemistry allows the discrimination of cellular elements with few distinctive features and identification criteria (e.g., microglial perikarya and processes, when using an antibody against the microglia-specific marker Iba1 (ionized calcium binding adaptor molecule 1; as presented here)), identifying the neurotransmitter contents of cellular elements (e.g., serotonergic) and their ultrastructural localization of soluble or membrane-bound proteins (e.g., 5 HT1A and EphA4 receptors). Here, we describe a protocol for transcardiac perfusion of mice with acrolein fixative, removal and sectioning of the brain, as well as immunoperoxidase-diaminobenzidine (DAB) staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with TEM. When rigorously performed, this technique provides an excellent compromise between optimal ultrastructural preservation and immunocytochemical detection.

We describe a protocol for transcardiac perfusion of mice, removal and sectioning of the brain, as well as immunoperoxidase staining, resin embedding, and ultrathin sectioning of the brain sections. Upon completion of these procedures, the immunostained material is ready for examination with transmission electron microscopy.

Transmission electron microscopy (TEM) is extremely useful for visualizing microglial, oligodendrocytic, astrocytic, and neuronal subcellular compartments ...

Intravital Microscopy of the Mouse Brain Microcirculation using a Closed Cranial Window

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, inflammatory and metabolic conditions, using in vivo fluorescence microscopy. A closed cranial window is placed over the left parieto-occipital cortex of the mice. Local microcirculation is recorded in real time through the window using epi and fluorescence illumination, and measurements of vessels diameters and red blood cell (RBC) velocities are performed. RBC velocity is measured using real-time cross-correlation and/or fluorescent-labeled erythrocytes. Leukocyte and platelet adherence to pial vessels and assessment of perfusion and vascular leakage are made with the help of fluorescence-labeled markers such as Albumin-FITC and anti-CD45-TxR antibodies. Microcirculation can be repeatedly video-recorded over several days. We used for the first time the close window brain intravital microscopy to study the pial microcirculation to follow dynamic changes during the course of Plasmodium berghei ANKA infection in mice and show that expression of CM is associated with microcirculatory dysfunctions characterized by vasoconstriction, profound decrease in blood flow and eventually vascular collapse.

Intravital microscopy to follow temporal and spatial hemodynamic and inflammatory events in the pial microcirculation.

This experimental model was designed to assess the mouse pial microcirculation during acute and chronic, physiological and pathophysiological hemodynamic, ...

Focussed Ion Beam Milling and Scanning Electron Microscopy of Brain Tissue

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron microscope (FIB/SEM). The samples are fixed with aldehydes, heavy metal stained using osmium tetroxide and uranyl acetate. They are then dehydrated with alcohol and infiltrated with resin, which is then hardened. Using a light microscope and ultramicrotome with glass knives, a small block containing the region interest close to the surface is made. The block is then placed inside the FIB/SEM, and the ion beam used to roughly mill a vertical face along one side of the block, close to this region. Using backscattered electrons to image the underlying structures, a smaller face is then milled with a finer ion beam and the surface scrutinised more closely to determine the exact area of the face to be imaged and milled. The parameters of the microscope are then set so that the face is repeatedly milled and imaged so that serial images are collected through a volume of the block. The image stack will typically contain isotropic voxels with dimenions as small a 4 nm in each direction. This image quality in any imaging plane enables the user to analyse cell ultrastructure at any viewing angle within the image stack.

This protocol describes how resin embedded brain tissue can be prepared and imaged in the three dimensions in the focussed ion beam, scanning electron microscope.

This protocol describes how biological samples, like brain tissue, can be imaged in three dimensions using the focussed ion beam/scanning electron ...

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

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 ...

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow for an in-depth characterization to identify pathways involved in these events. Cerebellar granule neurons (CGNs) that are derived from the developing cerebellum are an ideal model system that allows for morphological analyses. Here, we describe a method of how to genetically manipulate CGNs and how to study axono- and dendritogenesis of individual neurons. With this method the effects of RNA interference, overexpression or small molecules can be compared to control neurons. In addition, the rodent cerebellar cortex is an easily accessible in vivo system owing to its predominant postnatal development. We also present an in vivo electroporation technique to genetically manipulate the developing cerebella and describe subsequent cerebellar analyses to assess neuronal morphology and migration.

Genetic Manipulation of Cerebellar Granule Neurons In Vitro and In Vivo to Study Neuronal Morphology and Migration

Neuronal morphogenesis and migration are crucial events underlying proper brain development. Here, we describe methods to genetically manipulate cultured cerebellar granule neurons and the developing cerebellum for the assessment of morphology and migratory characteristics of neurons.

Developmental events in the brain including neuronal morphogenesis and migration are highly orchestrated processes. In vitro and in vivo analyses allow ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

Purification of Mouse Brain Vessels

In the brain, most of the vascular system consists of a selective barrier, the blood-brain barrier (BBB) that regulates the exchange of molecules and immune cells between the brain and the blood. Moreover, the huge neuronal metabolic demand requires a moment-to-moment regulation of blood flow. Notably, abnormalities of these regulations are etiological hallmarks of most brain pathologies; including glioblastoma, stroke, edema, epilepsy, degenerative diseases (ex: Parkinson&#8217;s disease, Alzheimer&#8217;s disease), brain tumors, as well as inflammatory conditions such as multiple sclerosis, meningitis and sepsis-induced brain dysfunctions. Thus, understanding the signaling events modulating the cerebrovascular physiology is a major challenge. Much insight into the cellular and molecular properties of the various cell types that compose the cerebrovascular system can be gained from primary culture or cell sorting from freshly dissociated brain tissue. However, properties such as cell polarity, morphology and intercellular relationships are not maintained in such preparations. The protocol that we describe here is designed to purify brain vessel fragments, whilst maintaining structural integrity. We show that isolated vessels consist of endothelial cells sealed by tight junctions that are surrounded by a continuous basal lamina. Pericytes, smooth muscle cells as well as the perivascular astrocyte endfeet membranes remain attached to the endothelial layer. Finally, we describe how to perform immunostaining experiments on purified brain vessels.

We describe a protocol allowing the purification of the mouse brain's vascular compartment. Isolated brain vessels include endothelial cells linked by tight junctions and surrounded by a continuous basal lamina, pericytes, vascular smooth muscle cells, as well as perivascular astroglial membranes.

In the brain, most of the vascular system consists of a selective barrier, the blood-brain barrier (BBB) that regulates the exchange of molecules and ...

High-Resolution 3D Imaging of Rabies Virus Infection in Solvent-Cleared Brain Tissue

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and study the distribution, tropism, and abundance of pathogens inside of organ tissues provides pivotal data on disease development and progression. Using conventional microscopy methods, immunolabeling is mostly restricted to thin sections obtained from paraffin-embedded or frozen samples. However, the limited 2D image plane of these thin sections may lead to the loss of crucial information on the complex structure of an infected organ and the cellular context of the infection. Modern multicolor, immunostaining-compatible tissue clearing techniques now provide a relatively fast and inexpensive way to study high-volume 3D image stacks of virus-infected organ tissue. By exposing the tissue to organic solvents, it becomes optically transparent. This matches the sample&#8217;s refractive indices and eventually leads to a significant reduction of light scattering. Thus, in combination with long free working distance objectives, large tissue sections up to 1 mm in size can be imaged by conventional confocal laser scanning microscopy (CLSM) at high resolution. Here, we describe a protocol to apply deep-tissue imaging after tissue clearing to visualize rabies virus distribution in infected brains in order to study topics like virus pathogenesis, spread, tropism, and neuroinvasion.

Novel, immunostaining-compatible tissue clearing techniques like the ultimate 3D imaging of solvent-cleared organs allow the 3D visualization of rabies virus brain infection and its complex cellular environment. Thick, antibody-labeled brain tissue slices are made optically transparent to increase imaging depth and to enable 3D analysis by confocal laser scanning microscopy.

The visualization of infection processes in tissues and organs by immunolabeling is a key method in modern infection biology. The ability to observe and ...