Name: free-floating immunostaining of mouse brains
Uploaded: 2021-10-07T13:30:00.000+00:00
Duration: 7 min 58 s
Description: Watch this Scientific Journal Video about Free-floating Immunostaining of Mouse Brains at JoVE.com

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>

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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;positive&#39; staining10. We infer the presence of a brownish-gray liver to indicate a high quality of perfusion, which usually results in blood-free brains. The automated perfusion pump used in this protocol helps to perfuse one animal successfully within a short period. Insufficient fixation will generate soft and fragile brain sections in the subsequent procedures, while over fixation will reduce the sensitivity of antigen reactions due to the enhanced formaldehyde cross-linking of proteins. Different conditions were tested, and overnight fixation at 4 &#176;C was sufficient for post-fixation of mouse brains. In addition to 10% NBF used in the present protocol as a fixative buffer, 4% (w/v) of freshly prepared paraformaldehyde (PFA) in PBS has also been extensively used for tissue fixation11.
Regarding cryosectioning, the thickness of sections needs to be decided depending on the specific needs. For instance, RNAscope studies require a thickness of 14 &#181;m instead of 25 &#181;m, commonly used in free-floating staining. Meanwhile, RNAscope studies require that all procedures are performed in RNAase-free solutions to preserve the integrity of the target mRNAs. Some researchers also use a section thickness of 30-40 &#181;m for a variety of staining procedures. Conventional cryosectioning (i.e., optimal cutting temperature (O.C.T.) compound-embedded samples) allows for much thinner (e.g., 10 &#181;m) brain sections that might be crucial for intracellular structures or other applications. The cryosectioning strategy presented here does not necessarily involve O.C.T. compound-embedding of brain samples and allows for 14-40 &#181;m sections. There may be no significant difference for 3,3-diaminobenzidine (DAB) staining using 25 or 40 &#181;m thick brain sections. However, thinner sections offer better-quality fluorescence images.
The benefit of the strategy presented here is that multiple brain samples (up to 5 brains) can be cut at one time. However, the limitation of this method is that brain samples need to be cut within 1 week after dehydration because submersion in 30% sucrose for too long is more likely to cause protein degradation and other issues. To avoid this potential issue, these brain sections can be transferred into the cryoprotectant buffer and stored at -20 &#176;C. For free-floating staining, the duration of incubation and concentration of both primary and secondary antibodies should be optimized in pilot studies. Generally, overnight incubation at 4 &#176;C or room temperature with mild shaking is appropriate for most primary antibodies, if not instructed otherwise by manufacturers. For secondary antibodies, incubation at room temperature for 1-3 h works well in most situations. However, these details must be optimized for various circumstances. For example, for c-fos staining, we typically incubate the brain sections with a concentration of 1:1,000 overnight at 4 &#176;C for immunofluorescence staining. However, using the same antibody for c-fos DAB staining, we prefer to incubate brain sections with a concentration of 1:5,000 for 48 h at 4 &#176;C.
A cocktail of primary antibodies and secondary antibodies might be used for double-staining to speed up the procedure. More specifically, two different primary antibodies from different species (e.g., one is from rabbit, the other one is from guinea pig, chicken, or mouse) are mixed before incubation, as are the corresponding secondary antibodies. The choice of secondary antibody is dependent on the primary antibody. If the primary antibody is from rabbit, the secondary antibody must be anti-rabbit, for example, donkey anti-rabbit or goat anti-rabbit. The selection of blocking serum depends on the secondary antibody, for example, normal donkey serum will be used if the secondary antibody is from donkey. Antigen retrieval is suggested if the immunostaining still does not work even if all guidelines have been followed strictly.
Mounting and coverslipping of brain sections must be performed in a very delicate manner. The whole process requires no wrinkles, folds, or air bubbles. It will take several trials to determine the optimal exposure time for imaging. We recommend the same exposure time for the same antibody across different sections, which is essential for comparing the signal intensities among different animals or groups. It is reasonable that exposure time might not be the same for different antibodies, even in the same section. For example, the exposure time for DAPI might be shorter than the c-fos signal in most cases.
A few procedures presented in the protocol are helpful to improve both reliability and efficiency throughout the whole process. 1) Using an automated perfusion pump for perfusion can considerably shorten perfusion time and significantly improve tissue quality. 2) This cryosectioning strategy enables slicing multiple brain samples simultaneously, which is much more efficient than conventional practice. This method is also easy for new researchers to learn and master. 3) For free-floating staining, as brain samples are stained in suspension, antibodies can penetrate the sections from both sides. We optimized the incubation strategy by placing all sections from one sample/series into a 1.5 mL microcentrifuge tube for primary and secondary antibodies, which saves antibodies, particularly when we need to stain brain samples in bulk. Another benefit of the free-floating approach is that it can be modified and applied to other histochemical staining methods (e.g., chromogenic IHC, hematoxylin and eosin, cresyl violet) in addition to immunofluorescence staining12.
However, one limitation of free-floating staining is that very thin sections can be difficult to handle. An on-slide staining method might be considered if only a few sections need to be collected and stained immediately, as is frequently the case in clinical pathology. We also tested the on-slide staining method using brain sections generated from this protocol, and it worked well. To do this, mount the brain sections onto slides, wait for the sections to dry, and follow a traditional frozen section on-slide staining protocol. 4) Mounting free-floating sections on the slides can be tedious for certain researchers, especially for beginners. We use a fine paintbrush to gently coax sections onto the slide at the air-buffer interface and then use a transfer pipette to gently remove the buffer to lower the air-buffer interface as mounting advances from the top to the bottom of the slide. Although time-consuming, this strategy is friendly to beginners. Experienced experimenters can mount all the brains sections onto the slides in PBS at one time and only remove the buffer to bring the slide out after the last section is mounted. 5) Finally, we use a scanner for imaging, which is more efficient than fluorescence microscopy, especially when there are a large number of slides for imaging. The scanner enables scanning up to 12 slides in one batch with a 20x magnification. Alternatively, standard fluorescence microscopy can be employed in certain circumstances, for example, when a specific cluster of neurons in the brain must be showcased with a higher magnification (e.g., 40x or even 60x)13,14.
In conclusion, this paper presents an established methodology for histological studies of mouse brains that has been proven to be reproducible, reliable, and efficient. The protocol will help generate optimal and consistent histological results among different researchers and laboratories and serve as a reference for beginners to learn this technique.

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><tbody><tr><td>Alexa Flour 594 donkey anti-rabbit IgG (H+L)</td><td>Invitrogen</td><td>A21207</td><td></td></tr><tr><td>&amp;nbsp;30% Sucrose</td><td>VWR</td><td>470302</td><td>30 g Sucrose dissoved into 100 mL of PBS</td></tr><tr><td>&amp;nbsp;Neutral Buffered Formalin</td><td>VWR</td><td>16004-128</td><td>10%, 25 &amp;deg;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 &amp;deg;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 &amp;deg;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 immunohistochemistry images of a mouse brain section with co-staining of WFA and AVP at lower and higher magnification at Bregma -0.82 mm. AVP signals were observed in the paraventricular nucleus and the supraoptic nucleus. WFA signals were observed in the perifornical area of the anterior hypothalamus and the reticular nucleus.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62876/62876fig01.jpg" /> 
Figure 1: Flow chart of fluorescence immunohistochemistry with mouse brains. Following complete anesthesia, a mouse is perfused with saline and then 10% NBF. The brain is carefully removed and cut into sections after fixation and dehydration. The sections were incubated with WFA followed by Streptavidin-Dylight&#160;488 after three washes with PBS. The brain sections were blocked and then incubated with a primary anti-AVP antibody. Then, the sections were washed 3 times with PBS followed by incubation with the secondary antibody, Alexa Flour 594 donkey anti-rabbit IgG (H+L). The brain sections were mounted on slides and coverslips placed on the slides with antifading mounting medium with DAPI before imaging. Abbreviations: NBF = neutral buffered formalin; WFA = Wisteria florbunda agglutinin; PBS = phosphate-buffered saline; AVP = arginine vasopressin; DAPI = 4&#8242;,6-diamidino-2-phenylindole. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62876/62876fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62876/62876fig02.jpg" /> 
Figure 2: Photos illustrating practical cryosectioning and mounting in the laboratory. (A)&#160;Build up a base on top of the plate with 30% sucrose to hold all samples horizontally, cut the tissues into sections (25 &#181;m/section) and collect the sections into a 48-well cell culture plate filled with phosphate-buffered saline. (B) Submerge a slide in the dish with one end slightly tilted with a stand. Place each row of sections under the air-buffer interface and lower the buffer to bring the sections out of the buffer until the bottom of the slide. The white line indicates the interface of buffer and air. (C) A fully mounted slide with brain sections. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62876/62876fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62876/62876fig03.jpg" /> 
Figure 3:&#160;An example of double immunofluorescence&#160;staining. (A-D) Microscopic images showing the distribution of WFA (red, A), AVP (green, B), DAPI (blue, C), and merged (D) in coronal mouse brain sections at Bregma -0.82 mm. Scale bars = 200 &#181;m. (E-H) Higher magnification microscopic images of the white boxes in A-D, respectively. Scale bars = 100 &#181;m. Abbreviations: 3V = third ventricle; PeFAH = perifornical area of the anterior hypothalamus; PVH = paraventricular nucleus; RT = reticular nucleus; SON = supraoptic nucleus. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62876/62876fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

free-floating-immunostaining-of-mouse-brains

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.8K Views.  Baylor College of Medicine. Immuno-histochemische kleuring van muizenhersenen is een veelgebruikte techniek in neurowetenschappelijk onderzoek. De kwaliteit, betrouwbaarheid en reproduceerbaarheid van de histologische resultaten van de hersenen kunnen echter variëren tussen verschillende onderzoekers en laboratoria. We presenteren een gevestigde methodologie voor histologische studies van muizenhersenen waarvan is bewezen dat ze reproduceerbaar, betrouwbaar en efficiënt zijn.Helpen om de procedure aan te tonen ...

Video: Free-floating Immunostaining of Mouse Brains

Batch Immunostaining for Large-Scale Protein Detection in the Whole Monkey Brain

Immunohistochemistry (IHC) is one of the most widely used laboratory techniques for the detection of target proteins in situ. Questions concerning the expression pattern of a target protein across the entire brain are relatively easy to answer when using IHC in small brains, such as those of rodents. However, answering the same questions in large and convoluted brains, such as those of primates presents a number of challenges. Here we present a systematic approach for immunodetection of target proteins in an adult monkey brain. This approach relies on the tissue embedding and sectioning methodology of NeuroScience Associates (NSA) as well as tools developed specifically for batch-staining of free-floating sections. It results in uniform staining of a set of sections which, at a particular interval, represents the entire brain. The resulting stained sections can be subjected to a wide variety of analytical procedures in order to measure protein levels, the population of neurons expressing a certain protein.

Large-scale immunodetection of target proteins across the entire primate brain is possible by employing novel tissue embedding and sectioning methods combined with the use of creative apparatus for batch staining of multiple free-floating sections at a given time.

Immunohistochemistry (IHC) is one of the most widely used laboratory techniques for the detection of target proteins in situ. Questions concerning the ...

Biology

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brain tissue, using markers for microglia and neuronal elements as an example. Specifically, this paper is a step-by-step protocol for fluorescent visualization of microglia and neurons via immunohistochemistry for Iba1 and Pan-neuronal, respectively. Fluorescence double-labeling is particularly useful for the localization of multiple proteins within the same sample, providing the opportunity to accurately observe interactions between cell types, receptors, ligands, and/or the extracellular matrix in relation to one another as well as protein co-localization within a single cell. Unlike other visualization techniques, fluorescence immunohistochemistry staining intensity may decrease in the weeks to months following staining, unless appropriate precautions are taken. Despite this limitation, in many applications fluorescence double-labeling is preferred over alternatives such as 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) or alkaline phosphatase (AP), as fluorescence is more time efficient and allows for more precise differentiation between two or more markers. The discussion includes troubleshooting tips and advice to promote success.

This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brains, using markers for microglia and neuronal elements as an example.

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level ...

Immunofluorescence Staining Using IBA1 and TMEM119 for Microglial Density, Morphology and Peripheral Myeloid Cell Infiltration Analysis in Mouse Brain

This is a protocol for the dual visualization of microglia and infiltrating macrophages in mouse brain tissue. TMEM119 (which labels microglia selectively), when combined with IBA1 (which provides an exceptional visualization of their morphology), allows investigation of changes in density, distribution, and morphology. Quantifying these parameters is important in providing insights into the roles exerted by microglia, the resident macrophages of the brain. Under normal physiological conditions, microglia are regularly distributed in a mosaic-like pattern and present a small soma with ramified processes. Nevertheless, as a response to environmental factors (i.e., trauma, infection, disease, or injury), microglial density, distribution, and morphology are altered in various manners, depending on the insult. Additionally, the described double-staining method allows visualization of infiltrating macrophages in the brain based on their expression of IBA1 and without colocalization with TMEM119. This approach thus allows discrimination between microglia and infiltrating macrophages, which is required to provide functional insights into their distinct involvement in brain homeostasis across various contexts of health and disease. This protocol integrates the latest findings in neuroimmunology that pertain to the identification of selective markers. It also serves as a useful tool for both experienced neuroimmunologists and researchers seeking to integrate neuroimmunology into projects.

This protocol describes a step-by-step workflow for immunofluorescent costaining of IBA1 and TMEM119, in addition to analysis of microglial density, distribution, and morphology, as well as peripheral myeloid cell infiltration in mouse brain tissue.

This is a protocol for the dual visualization of microglia and infiltrating macrophages in mouse brain tissue. TMEM119 (which labels microglia ...

Immunology and Infection

Quantifying the Heterogeneous Distribution of a Synaptic Protein in the Mouse Brain Using Immunofluorescence

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a protein, it is vital to also determine its distribution. Here, we describe a protocol employing immunofluorescence, confocal microscopy, and computer-based analysis to determine the distribution of the synaptic protein Mover (also called TPRGL or SVAP30). We compare the distribution of Mover to that of the synaptic vesicle protein synaptophysin, thereby determining the distribution of Mover in a quantitative manner relative to the abundance of synaptic vesicles. Notably, this method could potentially be implemented to allow for comparison of the distribution of proteins using different antibodies or microscopes or across different studies. Our method circumvents the inherent variability of immunofluorescent stainings by yielding a ratio rather than absolute fluorescence levels. Additionally, the method we describe enables the researcher to analyze the distribution of a protein on different levels: from whole brain slices to brain regions to different subregions in one brain area, such as the different layers of the hippocampus or sensory cortices. Mover is a vertebrate-specific protein that is associated with synaptic vesicles. With this method, we show that Mover is heterogeneously distributed across brain areas, with high levels in the ventral pallidum, the septal nuclei, and the amygdala, and also within single brain areas, such as the different layers of the hippocampus.

Here, we describe a quantitative approach to determining the distribution of a synaptic protein relative to a marker protein using immunofluorescence staining, confocal microscopy, and computer-based analysis.

The presence, absence, or levels of specific synaptic proteins can severely influence synaptic transmission. In addition to elucidating the function of a ...

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within cells. Neurochemical phenotyping of functionally identified neurons usually requires concurrent labelling with multiple antibodies (targeting protein) using immunohistochemistry (IHC) and optimization of in situ hybridization (targeting RNA), in tandem. A &#34;neurochemical signature&#34; to characterize particular neurons may be achieved however complicating factors include the need to verify FISH and IHC targets before combining the methods, and the limited number of RNAs and proteins that may be targeted simultaneously within the same tissue section.
Here we describe a protocol, using both fresh frozen and fixed mouse brain preparations, which detects multiple mRNAs and proteins in the same brain section using RNAscope FISH followed by fluorescence immunostaining, respectively. We use the combined method to describe the expression pattern of low abundance mRNAs (e.g., galanin receptor 1) and high abundance mRNAs (e.g., glycine transporter 2), in immunohistochemically identified brainstem nuclei.
Key considerations for protein labelling downstream of the FISH assay extend beyond tissue preparation and optimization of FISH probe labelling. For example, we found that antibody binding and labelling specificity can be detrimentally affected by the protease step within the FISH probe assay. Proteases catalyze hydrolytic cleavage of peptide bonds, facilitating FISH probe entry into cells, however they may also digest the protein targeted by the subsequent IHC assay, producing off target binding. The subcellular location of the targeted protein is another factor contributing to IHC success following FISH probe assay. We observed IHC specificity to be retained when the targeted protein is membrane bound, whereas IHC targeting cytoplasmic protein required extensive troubleshooting. Finally, we found handling of slide-mounted fixed frozen tissue more challenging than fresh frozen tissue, however IHC quality was overall better with fixed frozen tissue, when combined with RNAscope.

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

This protocol describes a method for combining fluorescence in situ hybridization (FISH) and fluorescence immunohistochemistry (IHC) in both fresh frozen and fixed mouse brain sections, with the goal of achieving multilabel FISH and fluorescence IHC signal. IHC targeted cytoplasmic and membrane attached proteins.

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within ...

Lineage Tracing of Inducible Fluorescently-Labeled Stem Cells in the Adult Mouse Brain

A telomerase reverse transcriptase (Tert) lineage-tracing mouse line was developed to investigate the behavior and fate of adult tissue stem cells, by crossing the &#39;Tet-On&#39; system oTet-Cre mouse with a novel reverse tetracycline transactivator (rtTA) transgene linked to the Tert promoter, which we have demonstrated marks a novel population of adult brain stem cells. Here, administration of the tetracycline derivative doxycycline to mTert-rtTA::oTet-Cre mice will indelibly mark a population of cells that express a 4.4 kb fragment of the promoter region of the gene Tert. When combined the Rosa-mTmG reporter, mTert-rtTA::oTet-Cre::Rosa-mTmG mice will express membrane tdTomato (mTomato) until doxycycline treatment induces the replacement of mTomato expression with membrane EGFP (mGFP) in cells that also express Tert. Therefore, when these triple-transgenic lineage tracing mice receive doxycycline (the &#34;pulse&#34; period during which TERT expressing cells are marked), these cells will become indelibly marked mGFP+ cells, which can be tracked for any desirable amount of time after doxycycline removal (the &#34;chase&#34; period), even if Tert expression is subsequently lost. Brains are then perfusion-fixed and processed for immunofluorescence and other downstream applications in order to interpret changes to stem cell activation, proliferation, lineage commitment, migration to various brain niches, and differentiation to mature cell types. Using this system, any rtTA mouse can be mated to oTet-Cre and a Rosa reporter to conduct doxycycline-inducible &#34;pulse-chase&#34; lineage&#160;tracing experiments using markers of stem cells.

The ability to permanently mark stem cells and their progeny with a fluorophore using an inducible transgenic lineage tracing mouse line allows for spatial and temporal analysis of activation, proliferation, migration, and/or differentiation in vivo. Lineage tracing can reveal novel information about lineage commitment, response to intervention(s), and multipotency.

A telomerase reverse transcriptase (Tert) lineage-tracing mouse line was developed to investigate the behavior and fate of adult tissue stem cells, by ...

Histological-Based Stainings Using Free-Floating Tissue Sections

Immunohistochemistry is a widely used technique to visualize specific tissue structures as well as protein expression and localization. Two alternative approaches are widely used to handle the tissue sections during the staining procedure, one approach consists of mounting the sections directly on glass slides, while a second approach, the free-floating, allows for fixed sections to be maintained and stained while suspended in solution. Although slide-mounted and free-floating approaches may yield similar results, the free-floating technique allows for better antibody penetration and thus should be the method of choice when thicker sections are to be used for 3D reconstruction of the tissues, for example when the focus of the experiment is to gain information on dendritic and axonal projections in brain regions. In addition, since the sections are kept in solution, a single aliquot can easily accommodate 30 to 40 sections, handling of which is less laborious, particularly in large-scale biomedical studies. Here, we illustrate how to apply the free-floating method to fluorescent immunohistochemistry staining, with a major focus on brain sections. We will also discuss how the free-floating technique can easily be modified to fit the individual needs of researchers and adapted to other tissues as well as other histochemical-based stainings, such as hematoxylin and eosin and cresyl violet, as long as tissue samples are properly fixed, typically with paraformaldehyde or formalin.

The free-floating technique allows researchers to perform histological-based stainings including immunohistochemistry on fixed tissue sections to visualize biological structures, cell type, and protein expression and localization. This is an efficient and reliable histochemical technique that can be useful for investigating a multitude of tissues, such as brain, heart, and liver.

Immunohistochemistry is a widely used technique to visualize specific tissue structures as well as protein expression and localization. Two alternative ...

Immunolabeling of Neuronal Membrane Proteins in a Freeze-fractured Specimen of Mouse Brain Tissue

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/53853/combined-optogenetic-freeze-fracture-replica-immunolabeling-to'>Source: Sch&#246;nherr, S., et. al. Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate Receptors in the Mouse Amygdala. J. Vis. Exp. (2016)</a>This video demonstrates the immunolabelling of freeze-fractured replicas of mouse brain tissue for electron microscopy. The fractured and coated replicas are digested to expose receptors. Primary and gold-tagged secondary antibodies are added to label specific neuronal membrane receptors. The replica is mounted on a grid and visualized under an electron microscope.

Source: Sch&#246;nherr, S., et. al. Combined Optogenetic and Freeze-fracture Replica Immunolabeling to Examine Input-specific Arrangement of Glutamate ...

JoVE Encyclopedia of Experiments

Neuropathology

Epilepsy and Seizure Disorders

Immunostaining of Fluorescently Labeled Astrocytes in a Mouse Brain Tissue Section

Source: Eaker, A. R., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/63804">Analysis of Astrocyte Territory Volume and Tiling in Thick Free-Floating Tissue Sections.</a> J. Vis. Exp. (2022)
This video demonstrates the procedure for immunostaining a fixed mouse brain tissue section with fluorescently labeled astrocytes using primary and secondary antibodies and capture the images of the stained astrocytes through confocal microscopy.

Source: Eaker, A. R., et al. Analysis of Astrocyte Territory Volume and Tiling in Thick Free-Floating Tissue Sections. J. Vis. Exp. (2022)
This video ...

Neurodegenerative Diseases

Immunostaining of Neurons in Mouse Brain Slices Following Fluorescence In Situ Hybridization

<a href='https://app-jove-com.remotexs.ntu.edu.sg/v/52799/detection-axonally-localized-mrnas-brain-sections-using-high'>Source: Baleriola, J., et. al., Detection of Axonally Localized mRNAs in Brain Sections Using High-Resolution In Situ Hybridization. J. Vis. Exp.&#160;(2015)</a>This video demonstrates the detection of mRNA within cholinergic axons in fixed mouse hippocampal slices using fluorescent in situ hybridization and antibody staining. Fluorescence microscopy reveals the spatial localization of probe-tagged mRNAs and choline acetyltransferase in cholinergic neurons.

Free-floating Immunostaining of Mouse Brains

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