This neural cell dissociation protocol is intended for samples with a low amount of starting material and yields a highly viable single-cell suspension for downstream analysis, with optional fixation and staining steps.
Successfully dissociating small quantities of neural tissue can equip labs to gain structure-specific insight into treatment efficacy, cellular function, as well as disease and treatment mechanisms of action. This neural dissociation protocol consistently yields a highly viable and actual single-cell suspension. In addition, we can process samples that are only a fraction of those that the commercial kits are intended for.
Proper planning and preparation are key to a successful outcome for this technique. Running several practice rounds to familiarize yourself with the protocol would be beneficial. After anesthetizing a six-month-old female C57BL/6J mouse, pinch the lower abdomen and lift the skin using forceps.
Then using scissors, cut through the fur and skin to the bottom of the ribcage. Make two diagonal incisions beginning below the ribcage and moving toward each shoulder. Then carefully resect the diaphragm and ribcage to expose the heart.
After carefully removing any connective tissue around the heart, use scissors to clip the right atrium. Then turn off the isoflurane flow to the breather. Holding the heart steady with forceps and with the bevel of the butterfly needle facing up, pierce the left ventricle while keeping the needle level and parallel to the animal.
Next, holding the needle in place, turn on the pump and perfuse at least 30 milliliters of the saline or heparin solution until the fluid leaving the heart is opaque and the liver and lungs pale in color. After perfusion, turn off the pump, remove the needle, and transfer the mouse to the dissection area. After decapitation, cut the fur from the back of the head up to the eyes and peel the skin back to expose the skull.
Next, clip the skull between the eyes and make two cuts at the back of the skull at 10 and 2 o'clock positions. Then make one long cut along the midsagittal line of the skull to the original cut between the eyes. Next, using forceps, peel the two halves of the skull away to the sides.
Then use a spatula to remove the brain and place it in a 60 millimeter glass Petri dish filled with cold DPBS and kept on ice. Using a scalpel or razor, separate each hemisphere and remove the olfactory bulbs and cerebellum. Then remove the midbrain until the hippocampus is exposed.
Next, secure the brain with forceps. Then using a second set of forceps, gently tease the hippocampus out of each hemisphere. Transfer both hippocampi to a labeled 1.5 milliliter tube containing cold DPBS and place the tube on ice.
Using forceps, transfer the hippocampi tissue pieces to a C tube and add 30 microliters of enzyme mix 2 into the tube. After twisting the cap and tightening until it clicks, place the tube in a dissociator and run the appropriate program. While the program is running, pre-wet a 70 micron cell strainer placed on a 50 milliliter conical tube with two milliliters of BSA buffer.
After the dissociation program is complete, add four milliliters of BSA buffer to the dissociated tissue and filter the mixture through the cell strainer on the 50 milliliter conical tube. Next, add 10 milliliters of DPBS to the C tube. Then close the tube, swirl the solution gently, and filter it through the cell strainer on the 50 milliliter conical tube.
Centrifuge the filtered cell suspension, discard the supernatant without disturbing the pellet and store the pellet. For debris removal, resuspend the pellet with 1, 550 microliters of cold DPBS and transfer the suspension to a labeled 50 milliliter conical tube. Then add 450 microliters of cold debris removal solution and pipette up and down.
Gently overlay the cell suspension with one milliliter of cold DPBS, keeping the tip against the wall of the conical tube. Repeat the procedure until the total overlay is two milliliters. Next, centrifuge the suspension at 3, 000 times G for 10 minutes with full acceleration and full break.
Aspirate the topmost layer, then sweep the pipette tip back and forth to aspirate the white middle layer. Remove as much of the middle layer as possible without disturbing the bottommost layer. Next, add two milliliters of cold DPBS and pipette up and down to mix.
Then centrifuge the suspension at 1, 000 times G for 10 minutes with full acceleration and full break. After centrifugation, discard the supernatant and resuspend the pellet in one milliliter BSA buffer. After cell counting, centrifuge the remaining cell suspension and resuspend the pellet in 50 microliters of diluted live dead stain.
Then transfer the sample to a labeled flow tube and incubate at room temperature for 8 to 10 minutes in the dark. After incubation, add 500 microliters of BSA buffer and repeat the centrifugation step. Then discard the supernatant, leaving a small amount of buffer in the tube.
The primary gate excluded debris in the forward scatter versus side scatter plots and the dead cells were subsequently excluded. The second gate excluded cells positive for myelin basic protein. And of the remaining cells, density plots of cells positive for each fluorochrome were created.
The frequency of each neuronal cell population was calculated out of the third gate. Samples processed with manual mechanical dissociation and enzymatic digestion yielded a substantially lower population of cells of interest, whereas both fresh and fixed samples processed with automated mechanical dissociation and enzymatic digestion showed a several-fold higher population of cells of interest. While the perfusion and debris removal steps can be challenging initially, maintaining a smooth and steady hand can help ensure success.
