Procedures and precautions for the preparation of mouse spleen single-cell suspensions
Procedure for preparation of mouse spleen single cell suspensions
a) Grinding method
1. Mice were killed by dislocation of the cervical vertebrae, soaked in 75% alcohol for 5 min, then taken out and placed on a sterile operating table with the left ventral side facing upwards.
2. A small cut was made in the middle of the left ventral side of the mouse, and the skin was torn open to expose the abdominal wall, revealing a long red spleen.
3. The peritoneum was lifted on the lower side of the spleen, cut and turned upwards to expose the spleen. Lift the spleen with forceps and ophthalmic scissors to separate the connective tissue underneath the spleen, remove the spleen, and immerse it in clean PBS solution.
4. Place the spleen in a 200-mesh sieve and gently grind it with a tissue grinder stick until there are no visible red clumps.
5. Rinse the sieve with 15 mL of PBS and collect the rinsate in a 15 mL centrifuge tube and centrifuge at 300 g for 5 min, discarding the supernatant.
6. Resuspend the cells by adding 2 mL of 1× erythrocyte lysate. After lysis for 2~3 min at room temperature, immediately add 10 mL of PBS and centrifuge at 300 g for 5 min, discard the supernatant.
7. Resuspend splenocytes with cell staining buffer, filter the cell suspension through a 200-mesh sieve again and count the cells, and adjust the cell concentration to 1 × 107/m.
b) Blowing method
1. Mice were dislocated from the cervical vertebrae, soaked in 75% ethanol for 5 min, and then removed and placed on a sterile operating table with the left ventral side facing up.
2. A small cut was made in the middle of the left ventral side of the mouse, and the skin was torn open to expose the abdominal wall, revealing a long red spleen.
3. Lift up the peritoneum on the lower side of the spleen, cut it open and turn it upward to expose the spleen. Lift up the spleen with forceps and ophthalmic scissors to separate the connective tissue underneath the spleen, take out the spleen and soak it in clean PBS solution.
4. Take a sterile 2.5 mL syringe to aspirate PBS, hold the spleen with forceps in the left hand and the syringe in the right hand, and carefully insert it into the spleen and blow it out until the spleen cells are completely blown out and observe that only white connective and adipose tissues are left, and gently rinse the remaining white tissues in PBS with forceps.
5. The blown cells were filtered through a 200-mesh sieve, collected in a 15 mL centrifuge tube and centrifuged at 300 g for 5 min, and the supernatant was discarded.
6. Resuspend the cells by adding 2 mL of 1× erythrocyte lysate, lysed for 2~3 min at room temperature, immediately added 10 mL of PBS, centrifuged at 300 g for 5 min, and discarded the supernatant.
7. Resuspend spleen cells with cell staining buffer, count, and adjust the cell concentration to 1×107/mL.
FSC/SSC map of mouse spleen
Notes:
1. A normal-sized spleen will yield approximately 4×107 cells based on experience, and the actual cell count will be based on the results of the count.
2. Lymphocytes account for about 60%~70% of the total cell volume after lysing erythrocytes in the mouse spleen.
3. Unstained spleen cell samples can be seen in the fluorescence channel with a small amount (about zero percent) of non-specific signal.
4. If you do not have a tissue grinding rod, you can also use the pusher inside the syringe instead. Use the rubber pad on the front of the pusher to grind the spleen.
5. If the collected spleen cells need to be cultured further, place the mouse spleen on a sterile bench when removing it. If you are just doing a normal flow-through experiment, you can disregard the sterile environment.
6. PBS can be replaced with cell staining buffer.
7. For the step of lysing red blood cells, the lysis time should be determined by the effect of lysing red blood cells during the experiment.
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