The investigators were supported by grants from the NIH (K01DK119471 to CW; P01DK113954, R01DK115761, R01DK117281, R01DK125480, and R01DK120858 to YX), USDA/CRIS (51000-064-01S to YX), and American Heart Association Postdoctoral Fellowship (#829565) to LT.

C57BL/6J mice of both sexes (8-16 weeks of age) were used in the present study. Care of all animals and all procedures were approved by Baylor College of Medicine&#39;s Institutional Animal Care and Use Committees.
1. Perfusion
NOTE: Steps 1.1 - 1.6 are performed in a fume hood.
<ol>
	<li>Anesthesia
	<ol>
		<li>Pour 5 mL of isoflurane (see the Table of Materials) onto a paper towel placed at the bottom of a desiccator. Introduce the mouse&#160;onto the holed barrier inside&#160;the desiccator and wait until signs of respiration have disappeared. 
		NOTE: Without respiration, the mouse&#160;still has heartbeat for a short period of time, and it will be perfused before the disappearance of the heartbeat.</li>
		<li>Before proceeding, confirm there is no reflex to a toe-pinch.</li>
		<li>Fix the mouse to a foam board by driving a pin through each foot. Ensure that the foam board is placed in a tray to collect liquid spillover.</li>
	</ol>
	</li>
	<li>Exposing the heart
	<ol>
		<li>Make a longitudinal superficial incision along the midline over the thorax and abdomen, then move the skin aside to expose the muscle wall of the thorax and abdomen. Next, make an incision in the muscle layer to expose the liver and the intestine. Finally, cut the rib cage with scissors to open the thorax and expose the heart and lungs. Use hemostatic forceps to pull the ribcage aside to widen the work area.</li>
	</ol>
	</li>
	<li>Collection of terminal blood (optional)
	<ol>
		<li>Insert a 1 mL syringe (see the Table of Materials) carefully into the right atrium of the heart until the tip is completely embedded. Hold the syringe steady and draw blood slowly until the desired volume is reached. 
		NOTE: Take care not to penetrate past the right atrium; Collection of 300-400 &#181;L of blood is achievable per adult mouse. Additives such as a clot accelerator or anticoagulant may be used, depending on the purpose of blood collection.</li>
	</ol>
	</li>
	<li>Placement of the perfusion cannula
	<ol>
		<li>For beginners, cut a small hole (&#60;1 mm) in the left ventricle of the heart with scissors and insert a blunt cannula (18 G needle for young mice and 21 G needle for aged mice) through the left ventricle into the ascending aorta. Alternatively, for experienced experimenters, penetrate the left heart ventricle with a blunt cannula directly and insert it into the ascending aorta carefully.</li>
		<li>Place pins around the conjunction of the cannula and coupled tubing on the foam board to prevent movement during the perfusion. Alternatively, use hemostatic forceps to fix the cannula in place. Proceed to cut the right atrium to allow the outflow of blood from circulation. 
		NOTE: The perfusion cannula and coupled tubing are connected to the perfusion pump.</li>
	</ol>
	</li>
	<li>Perfusion with saline
	<ol>
		<li>Turn on the saline pressure switch, and perfuse the mouse transcardially with 40-60 mL of saline (0.9% NaCl: 25 &#176;C, pH 7.4). Observe the outflow from the right atrium and the color of the liver closely. 
		&#8203;NOTE: An automated perfusion pump (see the Table of Materials) is used to flush the blood within 1-2 min. As the pressure range of the pump is 1-300 mmHg, the speed can be adjusted accordingly. If saline is leaking from the nose and/or the lung is inflated, decrease the pressure and adjust the cannula. Once the outflow no longer contains blood, the brain is sufficiently flushed with saline. Meanwhile, the liver is devoid of blood and becomes brownish-gray in color.</li>
	</ol>
	</li>
	<li>Perfusion with formalin
	<ol>
		<li>Turn off the saline pressure switch and turn on the formalin pressure switch to perfuse the mouse with 40 mL of 10% neutral buffered formalin (10% NBF: 25 &#176;C, pH 6.8-7.2, see the Table of Materials). Observe the animal&#39;s limbs for evidence of tremors. 
		&#8203;NOTE: Tail movement may also be observed after infusion of 10% NBF. CAUTION: As NBF is hazardous, it is suggested that researchers wear personal protective equipment (e.g., appropriate respirator, face shield) throughout the procedure.</li>
	</ol>
	</li>
	<li>Brain isolation9
	<ol>
		<li>Use scissors to remove the head. Make a middle line incision along the integument to expose the skull. Trim off the skin and muscle attachment with scissors.</li>
		<li>Make a cut at the orbital ridge, and then place the sharp end of iris scissors into the foramen magnum. Advance the scissors along the inner surface of the skull, maintaining upward pressure to avoid damage to the brain. Remove the parietal/frontal bones and meninges carefully. Finally, remove the brain from the opened skull gently.</li>
	</ol>
	</li>
	<li>Post-fixation
	<ol>
		<li>Place the brain in a 15 mL tube filled with 10 mL of 5% NBF and 15% sucrose for overnight fixation at 4 &#176;C. 
		&#8203;NOTE: To prepare 10 mL of 5% NBF and 15% sucrose, combine 5 mL of 10% NBF and 5 mL of 30% sucrose (w/v, dissolved into phosphate-buffered saline (PBS), see the Table of Materials). Make sure the volume of the fixative buffer is at least 10x larger than the sample itself. Initially, the brain will float in the buffer and will sink to the bottom after overnight fixation.</li>
	</ol>
	</li>
	<li>Dehydration
	<ol>
		<li>Transfer the brain sample to a 15 mL tube filled with 10 mL of 30% sucrose for dehydration at 4 &#176;C until it sinks.</li>
	</ol>
	</li>
</ol>
2. Cryosectioning (coronal sections)
<ol>
	<li>Preparation
	<ol>
		<li>Place dry ice on top of the height adjustment plate of a sliding microtome, and wait until white frost is visible. Carefully spread 5 mL of 30% sucrose on top of the plate to form a layer of a solid base after the sucrose solution is fully frozen. Place all brain samples (up to 5 brain samples in one batch) horizontally in a line on top of the sucrose, and then add 0.5 mL of 30% sucrose to the bottom of each brain. 
		NOTE: The brain will immediately stick to the frozen sucrose base and gradually freeze from bottom to top. Place additional sucrose around the mounted portion of the brain to form a sturdier base for cutting.</li>
	</ol>
	</li>
	<li>Sectioning
	<ol>
		<li>After 5-10 min of freezing, when the brain has become hard and white, trim the brain until the desired layer/region is reached. 
		NOTE: Adjust the amount of dry ice on the plate to control the temperature. The temperature is too low when ice crystals appear on the surface of the brain. The temperature is too high when the brain becomes soft and is no longer white.</li>
		<li>Switch from the Trim mode to the Feed mode and section brain tissue to 25 &#181;m thickness. Prepare a 48-well plate filled with 1x PBS, and mark five wells for one mouse brain. Use a paintbrush to collect each section from one mouse and place it into one well. Collect the subsequent section of the same mouse and place it into the 2nd well.</li>
		<li>Repeat 2.2.2 until the 5th well is reached, placing sections 6-10 in the 1st-5th well and so on. Repeat the same procedure for the rest of the brains simultaneously. Repeat until all the sections from one mouse are collected into the 5 wells in anatomical order. 
		NOTE: Following this procedure, the brain sections from each mouse are aliquoted into 5 series, which can be used for up to 5 different histological studies.</li>
	</ol>
	</li>
	<li>Storage
	<ol>
		<li>Use a paintbrush to transfer the sections from each well to a 1.5 mL microtube filled with cryoprotectant buffer (20% glycerol, 30% ethylene glycol, and 50% PBS), and store the samples at -20 &#176;C. 
		&#8203;NOTE: These sections stored in small tubes (1.5 mL) can save physical space in the freezer, and the integrity of the sections can be preserved for several months.</li>
	</ol>
	</li>
</ol>
3. Free-floating WFA staining and anti-AVP immunostaining
<ol>
	<li>Place a cell strainer into a well of a 6-well cell culture plate filled with PBS, and use a paintbrush to transfer one series of brain sections into the cell strainer. Rinse the sections in PBS by transferring the cell strainer to another well filled with PBS. Rinse for 6 x 10 min on a shaker for sections stored in cryoprotectant buffer and 3 x 10 min for freshly cut sections. 
	NOTE: All the washing procedures are performed in a 6-well cell culture plate with a cell strainer inside each well (see the Table of Materials). Each strainer holds the brain sections of interest from each mouse. To save reagents, all the incubation procedures described below are performed in a 1.5 mL microcentrifuge tube. Between washing and incubation procedures, use a paintbrush to transfer the brain sections between each cell strainer and each tube. For washing with PBS, 12 mL is adequate for each well.</li>
	<li>Prepare 1 mL of biotin-labeled WFA (1:1,000) solution in PBT (2.5 mL of Triton X-100 dissolved in 1,000 mL of PBS) buffer in 1.5 mL tube. Transfer brain sections from the cell strainer to the tube and incubate at room temperature on a rocking platform ~50 rpm overnight.</li>
	<li>Rinse the brain sections with PBS for 3 x 10 min as described in 3.1.</li>
	<li>Prepare 1 mL of streptavidin-Dylight 488 (1:500) solution in PBT in a 1.5 mL tube. Transfer brain sections into the tube and incubate at room temperature for 2 h on a rocking platform at ~50 rpm. 
	NOTE: Cover the plate with foil to avoid light exposure from this step forward.</li>
	<li>Rinse the brain sections with PBS for 3 x 10 min. Incubate the brain sections in blocking buffer (3% normal donkey serum diluted in PBT) for 2 h at room temperature. 
	NOTE: The choice of blocking serum is determined by the species from which the secondary antibody is generated. e.g., if the secondary antibody is from goat, normal goat serum should be used.</li>
	<li>Incubate the brain sections in primary antibody (rabbit anti-AVP, 1:500) at room temperature on a rocking platform at ~50 rpm overnight. 
	NOTE: Prepare both primary and secondary antibodies in blocking buffer. The concentration, duration of incubation, and temperature of incubation need to be optimized in pilot studies.</li>
	<li>Rinse the brain sections with PBS for 3 x 10 min. Incubate the brain sections in secondary antibody (Alexa Flour 594 donkey anti-rabbit IgG (H+L), 1:500) at room temperature for 2 h on a rocking platform at ~50 rpm. Rinse the brain sections with PBS for 3 x 10 min.</li>
</ol>
4. Mounting
<ol>
	<li>Fill two Petri dishes (a diameter of 150 mm) with 100 mL of 1x PBS each. Transfer all the brain sections from one strainer into the first dish, and align the brain sections in neuroanatomical order from caudal to rostral. 
	NOTE: To avoid mixing sections from different animals, mount one series of brain sections from one animal at a time.</li>
	<li>After all the brain sections are aligned in the first dish, submerge one slide into the second dish with one end slightly tilted with a stand (see Figure 1 and Figure 2). Use a fine paintbrush to gently place a brain section just below the air-buffer interface onto the tilted slide. Repeat the same procedure with another brain section and mount it side by side with the first section.</li>
	<li>Use a transfer pipette to slowly and gently remove the buffer to lower its level until both brain sections are entirely above the air-buffer interface. 
	NOTE: Take care to minimize the disturbance of the buffer surface, or the brain sections may float away. When dry, this row of brain sections will adhere to the slide firmly. If necessary, gently adjust the sections on the slide to ensure there are no wrinkles or folds in the tissue when the sections are still wet.</li>
	<li>Repeat this process until the bottom of the slide is reached. Continue to repeat until all sections are mounted onto the slide(s).</li>
</ol>
5. Coverslipping
<ol>
	<li>After all sections have dried (1-24 h at room temperature), place 80-100 &#181;L of anti-fading mounting medium with DAPI (see the Table of Materials) on the slide, and gently apply a glass coverslip to cover the samples. 
	NOTE: Apply the glass coverslip carefully and slowly to avoid bubbles.</li>
	<li>Place the slides in a microscope slide box (see the Table of Materials) and store them at 4 &#176;C. 
	&#8203;NOTE: Image all sections within 1-3 days to avoid fading of fluorescence and the development of autofluorescence.</li>
</ol>
6. Imaging
<ol>
	<li>Turn on the scanner and the computer (see Table of Materials). Position the slides in the slide holder with the loading device and insert the holder in the scanner. 
	NOTE: The scanner enables scanning of multiple channels (e.g., 4&#8242;,6-diamidino-2-phenylindole (DAPI), green, and red channels) simultaneously. The scanner scans slides only with a 20x magnification, and a traditional fluorescent microscope (see Table of Materials) can be used when a higher magnification (e.g., 40x) is needed.</li>
	<li>Open the software for the scanner (see the Table of Materials). Choose the appropriate storage location and scanning profile.</li>
	<li>Start the preview scan by clicking on the Start Preview Scan button. After the preview scan, open the tissue detection wizard and circle the regions of interest for imaging.</li>
	<li>Click on the Start Scan button after choosing the regions for imaging. Wait for the machine to finish scanning. Check the result file and export the images. 
	NOTE: If the profile is unsuitable for the slide, adjust by refocusing and changing the exposure time or light strength to adapt the profile to the tissues. Refer to the instructions for the scanner for more technical details (Table of Materials).</li>
</ol>

<ol>
	<li>Must, 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=The+Disease+burden+associated+with+overweight+and+obesity.">The Disease burden associated with overweight and obesity.</a> Journal of the American Medical Association. 282 (16), 1523-1529 (1999).</li><li>Apovian, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Obesity:+definition,+comorbidities,+causes,+and+burden.">Obesity: definition, comorbidities, causes, and burden.</a> The American Journal of Managed Care. 22 (7), 176-185 (2016).</li><li>Wong, S. K., Chin, K. Y., Suhaimi, F. H., Fairus, A., Ima-Nirwana, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+metabolic+syndrome:+a+review.">Animal models of metabolic syndrome: a review.</a> Nutrition &amp; Metabolism. 13 (1), 1-12 (2016).</li><li>Kennedy, A. J., Ellacott, K. L., King, V. L., Hasty, A. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mouse+models+of+the+metabolic+syndrome.">Mouse models of the metabolic syndrome.</a> Disease Models &amp; Mechanisms. 3 (3-4), 156-166 (2010).</li><li>Schacht, V., Kern, J. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Basics+of+immunohistochemistry.">Basics of immunohistochemistry.</a> The Journal of Investigative Dermatology. 135 (3), 1-4 (2015).</li><li>Mepham, B. L., Britten, K. J. M., Jones, D. B., Wright, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Immunostaining+methods+for+frozen+and+paraffin+sections.">Immunostaining methods for frozen and paraffin sections.</a> Lymphoproliferative Diseases. 15, 187-211 (1990).</li><li>Liu, 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=TPH2+in+the+dorsal+raphe+nuclei+regulates+energy+balance+in+a+sex-dependent+manner.">TPH2 in the dorsal raphe nuclei regulates energy balance in a sex-dependent manner.</a> Endocrinology. 162 (1), 1-16 (2021).</li><li>Liu, 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=Defining+vitamin+D+receptor+expression+in+the+brain+using+a+novel+VDR(Cre)+mouse.">Defining vitamin D receptor expression in the brain using a novel VDR(Cre) mouse.</a> Journal of Comparative Neurology. 529 (9), 2362-2375 (2020).</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> Journal of Visualized Experiments: JoVE. (65), e3564 (2012).</li><li>Whittington, N. 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(162), e61622 (2020).</li><li>He, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+small+potassium+current+in+AgRP/NPY+neurons+regulates+feeding+behavior+and+energy+metabolism.">A small potassium current in AgRP/NPY neurons regulates feeding behavior and energy metabolism.</a> Cell Reports. 17 (7), 1807-1818 (2016).</li><li>Xu, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+serotonin+2C+receptors+in+dopamine+neurons+inhibits+binge-like+eating+in+mice.">Activation of serotonin 2C receptors in dopamine neurons inhibits binge-like eating in mice.</a> Biololgical Psychiatry. 81 (9), 737-747 (2017).</li></ol>

The authors have no conflicts of interest to disclose.

This protocol provides an established method for neuroanatomical studies of the mouse brain, including perfusion, tissue sectioning, free-floating immunostaining, tissue mounting, and imaging. However, a few key details essential for consistent and reliable results must be optimized.
The quality of perfusion is critical for successful staining. Staining results might be affected if blood remains in the brain, given that blood cells (e.g., red blood cells) can generate an artificial &#39;positi...

The prevalence of obesity and associated comorbidities has reached epidemic levels, causing a tremendous socioeconomic burden1,2. Various mouse models have been developed to better understand the biological processes responsible for obesity3,4. Because central mechanisms are important for the regulation of energy homeostasis in these animal models, neuroanatomical studies of mouse brains have become a necessary technique in this field. However, the quality, reliability, and reproducibility of brain histology techniques vary considerably among laboratories and even researchers within the same laboratory for various reasons (e.g., antibodies, tissues, treatments, species, research objectives). Therefore, it is necessary to establish a general protocol for histological studies of the mouse brain, including perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging. Meanwhile, beginners can quickly learn, master, and adjust this protocol to satisfy their individual needs.
Immunohistochemical staining is an established method that has been used extensively to visualize specific cell types, mRNAs, and proteins in a variety of tissues (e.g., brain and peripheral tissues)5,6. More specifically, an antigen of interest can be labeled by a specific primary antibody and a corresponding secondary antibody linked to an enzyme (e.g., chromogenic immunohistochemistry) or a fluorescent dye (fluorescein isothiocyanate)6. As an example of the utility of these techniques, &#946;-endorphin [one peptide encoded by pro-opiomelanocortin (POMC)] and c-fos (a marker of neuronal activity) were stained in the arcuate nucleus. Deletion of tryptophan hydroxylase 2 (an enzyme integral to serotonin synthesis) in the dorsal raphe nucleus was shown to decrease c-fos expression in POMC neurons in the arcuate nucleus7. In addition, the distribution of vitamin D receptor mRNA was mapped in the mouse brain via in situ hybridization (RNAscope)8. This paper presents a reliable and efficient method with a step-by-step workflow for free-floating immunostaining, aiming to improve the quality and reproducibility of histological studies of the mouse brain.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alexa Flour 594 donkey anti-rabbit IgG (H+L)</td>
			<td>Invitrogen</td>
			<td>A21207</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;30% Sucrose</td>
			<td>VWR</td>
			<td>470302</td>
			<td>30 g Sucrose dissoved into 100 mL of PBS</td>
		</tr>
		<tr>
			<td>&#160;Neutral Buffered Formalin</td>
			<td>VWR</td>
			<td>16004-128</td>
			<td>10%, 25 &#176;C, pH 6.8-7.2</td>
		</tr>
		<tr>
			<td>1 mL Sub-Q Syringe</td>
			<td>BD</td>
			<td>309597</td>
			<td></td>
		</tr>
		<tr>
			<td>48 Well Cell Culture Plate</td>
			<td>Corning</td>
			<td>3548</td>
			<td></td>
		</tr>
		<tr>
			<td>6 Well Cell Culture Plate</td>
			<td>Corning</td>
			<td>3516</td>
			<td></td>
		</tr>
		<tr>
			<td>Antifading mounting media with DAPI</td>
			<td>Vector Laboratories</td>
			<td>H-1200</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclavable plastic desiccator</td>
			<td>Thermo Scientific Nalgene</td>
			<td>5315-0150</td>
			<td></td>
		</tr>
		<tr>
			<td>AVP antibody</td>
			<td>Phoenix Pharmaceuticals</td>
			<td>H-065-07</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell Strainer</td>
			<td>Corning</td>
			<td>431752</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryoprotectant buffer</td>
			<td>User preference</td>
			<td>Not applicable</td>
			<td>20% glycerol, 30% ethylene glycol, and 50% PBS</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Covetrus</td>
			<td>11695-6777-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica DFC310FX microscope</td>
			<td>Leica</td>
			<td>Not applicable</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slide Boxes (50-place)</td>
			<td>VWR</td>
			<td>Not applicable</td>
			<td></td>
		</tr>
		<tr>
			<td>PBT</td>
			<td>User preference</td>
			<td>Not applicable</td>
			<td>2.5 mL of Triton X-100 dissolved into 1000 mL of PBS</td>
		</tr>
		<tr>
			<td>Perfusion two automated Perfusion System</td>
			<td>Leica</td>
			<td>39471005</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS) 20x</td>
			<td>VWR</td>
			<td>VWRVE703-1L</td>
			<td>25 &#176;C, pH 7.3-7.5, 1x composition:137 mM NaCl, 2.7 mM KCl, 9.8 mM Phosphate buffer</td>
		</tr>
		<tr>
			<td>Slideing Microtome Microm HM450</td>
			<td>ThermoFisher</td>
			<td>Microm HM450</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride</td>
			<td>RICCA Chemical</td>
			<td>7220-32</td>
			<td>0.9%, 25 &#176;C, pH 7.4</td>
		</tr>
		<tr>
			<td>Streptavidin Protein, DyLight 488</td>
			<td>ThermoFisher</td>
			<td>#21832</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>089k01921</td>
			<td></td>
		</tr>
		<tr>
			<td>WFA antibody</td>
			<td>Sigma-Aldrich</td>
			<td>L1516</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss Axio Z1 Scanner</td>
			<td>Zeiss</td>
			<td>Not applicable</td>
			<td></td>
		</tr>
		<tr>
			<td>Zen 3.1 software</td>
			<td></td>
			<td></td>
			<td>scanner software</td>
		</tr>
	</tbody>
</table>

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.

The flow chart of this protocol is briefly illustrated in Figure 1. This laboratory&#39;s cryosectioning procedure is demonstrated in Figure 2A, in which 5 brain samples were sectioned simultaneously. The mounting of brain sections is shown in Figure 2B, and a fully mounted slide with brain sections is illustrated in Figure 2C. In Figure 3, representative fluorescence immun...

Watch this Scientific Journal Video about Free-floating Immunostaining of Mouse Brains at JoVE.com

Free-floating Immunostaining of Mouse Brains

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 ...

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Neuroscience

12.5K Views.  Baylor College of Medicine. This protocol describes an efficient and reproducible approach for mouse brain histological studies, including perfusion, brain sectioning, free-floating immunostaining, tissue mounting, and imaging.

Video: Free-floating Immunostaining of Mouse Brains

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

Ex vivo Culturing of Whole, Developing Drosophila Brains

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for investigating the cell biology and development of central brain structures by allowing acute pharmacological interventions and live imaging of cellular processes. As an example of the technique, prior work from our lab1 has shown that a previously unrecognized subcellular compartment lies between the axonal and somatodendritic compartments of axons of the Drosophila central brain. The development of this compartment, referred to as the axon initial segment (AIS)2, was shown genetically to depend on the neuron-specific cyclin-dependent kinase, Cdk5. We show here that ex vivo treatment of wild-type Drosophila larval brains with the Cdk5-specific pharmacological inhibitors roscovitine and olomoucine3 causes acute changes in actin organization, and in localization of the cell-surface protein Fasciclin 2, that mimic the changes seen in mutants that lack Cdk5 activity genetically.
A second example of the ex vivo culture technique is provided for remodeling of the connections of embryonic mushroom body (MB) gamma neurons during metamorphosis from larva to adult. The mushroom body is the center of olfactory learning and memory in the fly4, and these gamma neurons prune their axonal and dendritic branches during pupal development and then re-extend branches at a later timepoint to establish the adult innervation pattern5. Pruning of these neurons of the MB has been shown to occur via local degeneration of neurite branches6, by a mechanism that is triggered by ecdysone, a steroid hormone, acting at the ecdysone receptor B17, and that is dependent on the activity of the ubiquitin-proteasome system6. Our method of ex vivo culturing can be used to interrogate further the mechanism of developmental remodeling. We found that in the ex vivo culture setting, gamma neurons of the MB recapitulated the process of developmental pruning with a time course similar to that in vivo. It was essential, however, to wait until 1.5 hours after puparium formation before explanting the tissue in order for the cells to commit irreversibly to metamorphosis; dissection of animals at the onset of pupariation led to little or no metamorphosis in culture. Thus, with appropriate modification, the ex vivo culture approach can be applied to study dynamic as well as steady state aspects of central brain biology.

Ex vivo Culturing of Whole, Developing Drosophila Brains

This article describes a method by which one can mimic in vivo development of the Drosophila mushroom body in an ex vivo culture system.

We describe a method for ex vivo culturing of whole Drosophila brains. This can be used as a counterpoint to chronic genetic manipulations for ...

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 ...

Stereotaxic Surgery for Genetic Manipulation in Striatal Cells of Neonatal Mouse Brains

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, they may function to regulate the developmental process and/or physiological function in neonatal brains. To investigate neurobiological functions of specific genes in the brain, it is essential to inactivate genes in the brain. Here, we describe a simple stereotaxic method to inactivate gene expression in the striatum of transgenic mice at neonatal time windows. AAV-eGFP-Cre viruses were microinjected into the striatum of Ai14 reporter gene mice at postnatal day (P) 2 by stereotaxic brain surgery. The tdTomato reporter gene expression was detected in P14 striatum, suggesting a successful Cre-loxP mediated DNA recombination in AAV-transduced striatal cells. We further validated this technique by microinjecting AAV-eGFP-Cre viruses into P2Foxp2fl/fl mice. Double labeling of GFP and Foxp2 showed that GFP-positive cells lacked Foxp2 immunoreactivity in P9 striatum, suggesting the loss of Foxp2 protein in AAV-eGFP-Cre transduced striatal cells. Taken together, these results demonstrate an effective genetic deletion by stereotaxically microinjected AAV-eGFP-Cre viruses in specific neuronal populations in the neonatal brains of floxed transgenic mice. In conclusion, our stereotaxic technique provides an easy and simple platform for genetic manipulation in neonatal mouse brains. The technique can not only be used to delete genes in specific regions of neonatal brains, but it also can be used to inject pharmacological drugs, neuronal tracers, genetically modified optogenetics and chemogenetics proteins, neuronal activity indicators and other reagents into the striatum of neonatal mouse brains.

We describe a protocol of stereotaxic surgery with a homemade head-fixed device for microinjecting reagents into the striatum of neonatal mouse brains. This technique allows genetic manipulation in neuronal cells of specific regions of neonatal mouse brains.

Many genes are expressed in embryonic brains, and some of them are continuously expressed in the brain after birth. For such persistently expressed genes, ...

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate tangentially from their place of origin in the ventral telencephalon (including from the medial and caudal ganglionic eminences (MGE, CGE)) to the dorsal cortical plate in response to a variety of intrinsic and extrinsic cues. Different methodologies have been developed over the years to genetically manipulate specific pathways and investigate how they regulate the dynamic cytoskeletal changes required for proper IN migration. In utero electroporation has been extensively used to study the effect of gene repression or overexpression in specific IN subtypes while assessing the impact on morphology and final position. However, while this approach is readily used to modify radially migrating pyramidal cells, it is more technically challenging when targeting INs. In utero electroporation generates a low yield given the decreased survival rates of pups when electroporation is conducted before e14.5, as is customary when studying MGE-derived INs. In an alternative approach, MGE explants provide easy access to the MGE and facilitate the imaging of genetically modified INs. However, in these explants, INs migrate into an artificial matrix, devoid of endogenous guidance cues and thalamic inputs. This prompted us to optimize a method where INs can migrate in a more naturalistic environment, while circumventing the technical challenges of in utero approaches. In this paper, we describe the combination of ex utero electroporation of embryonic mouse brains followed by organotypic slice cultures to readily track, image and reconstruct genetically modified INs migrating along their natural paths in response to endogenous cues. This approach allows for both the quantification of the dynamic aspects of IN migration with time-lapse confocal imaging, as well as the detailed analysis of various morphological parameters using neuronal reconstructions on fixed immunolabeled tissue.

Ex Utero Electroporation and Organotypic Slice Cultures of Embryonic Mouse Brains for Live-Imaging of Migrating GABAergic Interneurons

Here, we provide a low-cost and reliable method to generate electroporated brain organotypic slice cultures from mouse embryos suitable for confocal microscopy and live-imaging techniques.

GABAergic interneurons (INs) are critical components of neuronal networks that drive cognition and behavior. INs destined to populate the cortex migrate ...

Homochronic Transplantation of Interneuron Precursors into Early Postnatal Mouse Brains

Neuronal fate determination and maturation requires an intricate interplay between genetic programs and environmental signals. However, disentangling the roles of intrinsic vs. extrinsic mechanisms that regulate this differentiation process is a conundrum for all developmental neurobiologists. This issue is magnified for GABAergic interneurons, an incredibly heterogeneous cell population that is born from transient embryonic structures and undergo a protracted migratory phase to disperse throughout the telencephalon. To explore how different brain environments affect interneuron fate and maturation, we developed a protocol for harvesting fluorescently labeled immature interneuron precursors from specific brain regions in newborn mice (P0-P2). At this age, interneuron migration is nearly complete and these cells are residing in their final resting environments with relatively little synaptic integration. Following collection of single cell solutions via flow cytometry, these interneuron precursors are transplanted into P0-P2 wildtype postnatal pups. By performing both homotopic (e.g., cortex-to-cortex) or heterotopic (e.g., cortex-to-hippocampus) transplantations, one can assess how challenging immature interneurons in new brain environments affects their fate, maturation, and circuit integration. Brains can be harvested in adult mice and assayed with a wide variety of posthoc analysis on grafted cells, including immunohistochemical, electrophysiological and transcriptional profiling. This general approach provides investigators with a strategy to assay how distinct brain environments can influence numerous aspects of neuron development and identify if specific neuronal characteristics are primarily driven by hardwired genetic programs or environmental cues.

Challenging young neurons in new brain regions can reveal important insights into how the environment sculpts neuronal fate and maturation. This protocol describes a procedure to harvest interneuron precursors from specific brain regions and transplant them either homotopically or heterotopically into the brain of postnatal pups.

Neuronal fate determination and maturation requires an intricate interplay between genetic programs and environmental signals. However, disentangling the ...

Simultaneous Cryosectioning of Multiple Rodent Brains

Histology and immunohistochemistry are routine methods of analysis to visualize microscopic anatomy and localize proteins within biological tissue. In neuroscience, as well as a plethora of other scientific fields, these techniques are used. Immunohistochemistry can be done on slide mounted tissue or free-floating sections. Preparing slide-mounted samples is a time intensive process. The following protocol for a technique, called the Megabrain, reduced the time taken to cryosection and mount brain tissue by up to 90% by combining multiple brains into a single frozen block. Furthermore, this technique reduced variability seen between staining rounds, in a large histochemical study. The current technique has been optimized for using rodent brain tissue in downstream immunohistochemical analyses; however, it can be applied to different scientific fields that use cryosectioning.

Here, we present a protocol to freeze and section brain tissue from multiple animals as a timesaving alternative to processing single brains. This reduces staining variability during immunohistochemistry and reduces time cryosectioning and imaging.

Histology and immunohistochemistry are routine methods of analysis to visualize microscopic anatomy and localize proteins within biological tissue. In ...

Targeting Alpha Synuclein Aggregates in Cutaneous Peripheral Nerve Fibers by Free-floating Immunofluorescence Assay

To date, for most neurodegenerative diseases only a post-mortem histopathological definitive diagnosis is available. For Parkinson&#39;s disease (PD), the diagnosis still relies only on clinical signs of motor involvement that appear later on in the disease course, when most of the dopaminergic neurons are already lost. Hence, there is a strong need for a biomarker that can identify patients at the beginning of disease or at the risk of developing it. Over the last few years, skin biopsy has proved to be an excellent research and diagnostic tool for peripheral nerve diseases such as small fiber neuropathy. Interestingly, a small fiber neuropathy and alpha synuclein (&#945;Syn) neural deposits have been shown by skin biopsy in PD patients. Indeed, skin biopsy has the great advantage of being an easily accessible, minimally invasive and painless procedure that allows the analysis of peripheral nervous tissue prone to the pathology. Moreover, the possibility of repeating the skin biopsy in the course of the follow-up of the same patient allows studying the longitudinal correlation with the disease progression. We set up a standardized reliable protocol to investigate the presence of &#945;Syn aggregates in skin nerve fibers of the PD patient. This protocol involves few short fixation steps, a cryotome sectioning and then a free-floating immunofluorescence double-staining with two specific antibodies: anti Protein Gene Product 9.5 (PGP9.5) to mark the cutaneous nerve fibers and anti 5G4 for detecting &#945;Syn aggregates. It is a versatile, sensitive and easy to perform protocol that can also be applied for targeting other proteins of interest in skin nerves. The ability to mark &#945;Syn aggregates is another step forward to the use of skin biopsy as a tool for establishing a pre-mortem histopathological diagnosis of PD.

Here, we present a protocol for a free-floating indirect immunofluorescence assay on skin biopsy sections that allows for the identification of disease specific conformation variants of alpha synuclein involved in Parkinson disease and multiple proteins of the peripheral nervous system.

To date, for most neurodegenerative diseases only a post-mortem histopathological definitive diagnosis is available. For Parkinson&#39;s disease (PD), the ...

Transplantation of Chemogenetically Engineered Cortical Interneuron Progenitors into Early Postnatal Mouse Brains

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is a complicated task, which is particularly difficult in regards to the development of &#947;-aminobutyric acid (GABA)ergic cortical interneurons (CIs). CIs are the main inhibitory neurons in the cerebral cortex, and they play key roles in neuronal networks, by regulating both the activity of individual pyramidal neurons, as well as the oscillatory behavior of neuronal ensembles. They are generated in transient embryonic structures (medial and caudal ganglionic eminences -&#160;MGE and CGE) that are very difficult to efficiently target using in utero electroporation approaches. Interneuron progenitors migrate long distances during normal embryonic development, before they integrate in the cortical circuit. This remarkable ability to disperse and integrate into a developing network can be hijacked by transplanting embryonic interneuron precursors into early post-natal host cortices. Here, we present a protocol that allows genetic modification of embryonic interneuron progenitors using focal ex vivo electroporation. These engineered interneuron precursors are then transplanted into early post-natal host cortices, where they will mature into easily identifiable CIs. This protocol allows the use of multiple genetically encoded tools, or the ability to regulate the expression of specific genes in interneuron progenitors, in order to investigate the impact of either genetic or environmental variables on the maturation and integration of CIs.

Here we present a protocol, designed to use chemogenetic tools to manipulate the activity of cortical interneuron progenitors transplanted into the cortex of early postnatal mice.

Neuronal development is regulated by a complex combination of environmental and genetic factors. Assessing the relative contribution of each component is ...