Free-floating Immunostaining of Mouse Brains

Immuno-histochemical staining of mouse brains is a commonly-used technique in neuroscience research. However, the quality, reliability, and reproducibility of brain histology results may vary among different researchers and laboratories. We present an established methodology for histological studies of mouse brains that has been proven to be reproducible, reliable, and efficient.

Helping to demonstrate the procedure will be Dr.Chunmei Wang, an assistant professor from my laboratory After confirming successful anesthetization in an eight to 16 week old C57 black/6J mouse, Fix the mouse on a foam board and 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. Then, using scissors, cut the ribcage to open the thorax and expose the heart and lungs.

Using hemostatic forceps, pull the ribcage aside to widen the work area.Next. to place a perfusion cannula, directly penetrate the left heart ventricle with a blunt cannula and carefully insert it into the ascending aorta. Place pins around the conjunction of the cannula and coupled tubing on the foam board to fix the cannula in place during the perfusion, and proceed to cut the right atrium to allow the outflow of blood from circulation.

For perfusion with saline, turn on the saline pressure switch and perfuse the mouse transcardially with 40 to 60 milliliters of saline. Observe the outflow from the right atrium and the color of the liver closely. Next, to perfuse with formalin, turn off the saline pressure switch and turn on the formalin pressure switch to perfuse the mouse with 40 milliliters of 10%neutral-buffered formalin.

Observe the animal’s limbs for evidence of tremors. For brain isolation, use scissors to remove the head and make a middle line incision along the integument to expose the skull, then trim off the skin and muscle attachment. Next, make a cut at the orbital ridge and place the sharp end of iris scissors into the foramen magnum.

Advance the scissors along the skull’s inner surface, maintaining upward pressure to avoid damage to the brain. Then, carefully remove the parietal and frontal bones and meninges before removing the brain from the open skull. Place dry ice on top of the height adjustment plate of a sliding microtome and wait until white frost is visible, then carefully spread five milliliters of 30%sucrose on top of the plate to form a solid base layer after the sucrose has frozen.

Next, place all brain samples horizontally in a line on top of the sucrose base with 500 microliters of 30%sucrose. After five to 10 minutes of freezing, when the brain has become hard and white, trim the brain until the desired layer or region is reached, then, switch from the trim mode to the feed mode and section brain tissue to generate 25-micrometer-thick sections. Next, prepare a 48-well plate filled with PBS and mark five wells for one mouse brain.

Using a paintbrush, collect one section and place it into one well, then collect the subsequent section of the same mouse and place it into the second well. Repeat the procedure until the fifth well is reached placing sections six to 10 in the first to fifth well, and so on. Place a cell strainer into one well of a six-well cell culture plate filled with PBS and, using a paintbrush, transfer one series of brain sections into the cell strainer.

Rinse these sections in PBS by transferring the cell strainer to another well filled with PBS on the shaker. Then, transfer the brain sections from the cell strainer into a 1.5-milliliter tube containing one milliliter of biotin-labeled WFA and incubate on a rocking platform at 50 RPM overnight. The following day, rinse the brain sections with PBS as demonstrated previously, then transfer the sections into a tube containing one milliliter of streptavidin-daylight 488 solution and incubate for two hours on a rocking platform.

After rinsing the brain sections with PBS again, incubate them in blocking buffer for two hours, followed by incubation in primary antibody overnight on a rocking platform. The next day, following PBS rinses, incubate the sections in secondary antibody for two hours. Fill two Petri dishes with 100 milliliters of PBS and transfer all the brain sections from one strainer into the first dish.

Align the brain sections in neuroanatomical order from caudal to rostral. Then, submerge one slide into the second dish with one end slightly tilted with a stand, and using a fine paintbrush, gently place a brain section just below the air buffer interface onto the tilted slide. Next, using a transfer pipette, slowly and gently remove the buffer to lower its level until the brain section is entirely above the air buffer interface.

Continue to repeat this process until the bottom of the slide is reached and all sections are mounted onto the slide. For imaging, turn on the scanner and computer, then position the slides in the slide holder with the loading device, and insert the holder into the scanner. Open the software for the scanner, and choose the appropriate storage location and scanning profile.

Start the preview scan by clicking on the Start Preview Scan button. After the scan, open the tissue detection wizard, and circle the regions of interest for imaging. After choosing the regions for imaging, click on the Start Scan button and wait for the machine to finish scanning.

Check the result file and export the images. Shown here are representative fluorescence immuno-histochemistry images of coronal mouse brain sections at Bregma-negative 0.82 millimeters, displaying the distribution of wisteria floribunda agglutinin, arginine vasopressin, and DAPI. Wisteria floribunda agglutinin signals are observed in the perifornical area of the anterior hypothalamus and reticular nucleus, whereas arginine vasopressin signals are observed in the paraventricular nucleus.

When trying this protocol for the first time, we suggest performing it under the supervision of an experienced researcher. The protocol will help generate optimal and consistent histological results among different researchers and laboratories, and will serve as a reference for beginners learning this technique.