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Passaging of Neuronal Stem Cells for Studies of the Effects of Anesthetics
By: Edwin S. Masangkay & Martin Chin
Introduction:
The purpose of this experiment is to study the effects of anesthetics on
stem cells of the brain. Anesthetics are used to allow patients to undergo surgery by
creating a loss of feeling or awareness in the body. A general anesthetic puts the person
to sleep. A local anesthetic causes loss of feeling in a part of the body such as a tooth or
an area of skin without affecting consciousness. Regional anesthesia numbs a larger part
of the body such as a leg or arm, also without affecting consciousness. 1
Possible
anesthetics used in this experiment include midazolam, isoflurane, ketamine, fentanyl,
propofol, etomidate, and lidocaine. Midazolam, isoflurane, propofol, and etomidate act
on the inhibitory neurotransmitter gamma-aminobutyric acid (GABA); while, ketamine
interacts with N-methyl-D-aspartate (NMDA) receptors, fentanyl binds to serotonin (5-
HT1A) receptors, and lidocaine, a local anesthetic, blocks sodium channels in
peripheral sensory neurons.
The procedure for experimentation requires the harvesting of neuronal stem cells
from the hippocampus of a rat brain and the passaging of these cells. This protocol will
present an aseptic technique to culture and passage neuronal stem cells to test for the
effects of anesthetics on them.
Note: This protocol requires the procedures to be done at the same time, three (3)
hours per day, for six (6) days straight.
Materials:
DMEM T-25 flasks containing passage cells
B27 T-75 flasks (for passaging excess cells)
L-glutamine Five (5) 6-well- plates
bFGF Microtubes
Penstrep 15 ml conical tubes
F12 5 ml and 10 ml pipets
Heparin 200 µl pipets
Versene Hemacytometer
Trypan Blue Syringe

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Media Preparation
Base Media:
DMEM + F12 (Premixed 3:1 with Penicillin-streptomycin).
Proliferation Media (Pro-Media):
DMEM + F12 (Premixed 3:1 with Penicillin-streptomycin).
200 µl B27 supplement/10 ml of media.
7.5 units of heparin/10 ml of media.
1 µl of bFGF/1 ml of media.
Note: Media must be fresh.
Culturing of Stem Cells
Procedure:
Day One (1):
1. Prepare pro-media in amount as needed.
2. Transfer cells from each T-25 flask to a 15 ml conical tube.
3. Add 1-2 ml of versene into each flask to detach remaining adherent
cells. Incubate flasks for 5 minutes at 37o
C.
4. Transfer remaining cells to the appropriate 15 ml conical tube.
5. Centrifuge tubes at 300 RCF for six (6) minutes in room temperature
(RT).
6. Aspirate media from the tubes leaving ~ 200 µl of supernatant.
7. Triturate (30 – 35 times) gently using 200 µl pipet.
8. Fill to volume to 2 ml total with pro-media to each of the conical tubes.
Mix well.
9. Prepare three 6-well- plates for control, a total of 15 wells will be used, as follows: On plate
1, label 3 wells as day ‘0C’ for control’ and leave remaining three wells unlabelled. On
plate 2, label 3 wells as day “1C” and 3 wells as day “2C”. On plate 3, label 3 wells as day
‘3C’ and 3 wells as day ‘4C’. See figure 1, following page.
10. Prepare two 6-well- plates for anesthetic exposure. A total of 12 wells will be used. On
plate 1, label 3 wells as day ‘1’ and label 3 wells as day “2”. On plate 2, label 3 wells as
day ‘3’ and 3 wells as day ‘4’.
11. Prepare and count the cells using a hemacytometer. See cell counting protocol below.
12. After the cell count from sample tubes make a 100,000 (105
) cells/well- dilution to each of
day “0” to day “3”, a total of 24 wells including the controls. Make a 50,000 cells/well on
day “4” wells, a total of 6 wells including the controls.
13. Plate a total of 3 ml cell solution in each of the appropriate wells.
14. Any remaining cells in solution should be transferred onto T-75 flasks for passaging. Fill to
volume to 10 ml of each flask with pro-media.
15. Incubate 6-well plates and T-75 flasks at 37o
C with CO2.

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Day Two (2):
1. Count day ‘0C’ wells. Record to lab notebook. Refer to cell counting protocol.
2. Make a dilution of anesthetic solution for each well of cells to be tested.
3. Pipet an appropriate amount of diluted anesthetic into each well of day ‘1’ through day ‘4’.
4. Incubate cells at 37o
C with CO2.
Day Three (3):
Count day ‘1C’ and day ‘1’ wells of cells exposed to the anesthetic. Record cell count to
lab notebook.
Day Four (4):
Count day ‘2C’ and day ‘2’ wells of cells exposed to the anesthetic. Record cell count to
lab notebook.
Day Five (5):
Count day ‘3C’ and day ‘3’ wells of cells exposed to the anesthetic. Record cell count to
lab notebook.
Day Six (6):
Count day ‘4C’ and day ‘4’ wells of cells exposed to the anesthetic. Record cell count to
lab notebook.
Figure 1. Setup of Six-well- Plates for Culturing Stem Cells.
Day 0C Not Used Day 1C Day 2C Day 3C Day 4C Day 1 Day 2 Day 3 Day 4
Exposed to Anesthesia

4. Total number of cells is determined by
the number of cells per ml counted
multiplied by the media dilution factor, by
the trypan blue dilution, and by the the
cell density per section of the
hemacytometer grid.
Since 2 ml of media was used multiply
cells/ml by 2 and because 20 μl of
traypan blue was added to the 20 μl of
cell suspension, it is multiplied by two
again. Finally, the total cell count is
determined by multiplying by 10,000, the
cell density per section of the
hemacytometer.
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Cell Counting Protocol
Procedure:
1. Using an appropriate sized pipet, transfer cells from each well
of the 6-well-plate into a 15 ml conical tube. See figure 2.
2. Treat remaining cells in the well with 1-2 ml of versene.
Incubate for 5 minutes.
3. Pipet remaining cells into the appropriate tube.
4. Spin tube at 300 RCF for 6 minutes at RT.
5. Aspirate supernatant leaving ~ 200 μl total volume.
6. Triturate gently ~ 30-35 times to separate neurospheres.
7. Bring total volume of tube to 2 ml with DMEM media.
Mix samples well.
8. From 2 ml cell solution, take 20 μl of sample and place into a
microtube.
9. Add 20 μl of trypan blue and mix well.
10. Pipet 10 μl of each cell solution on each side of the
hemacytometer making sure to cover each grid.
11. Using only the middle square, on each of the grids of the
hemacytometer, count the cells.
12. Record the average of the cells counted.
13. Calculate the number of cells times 2 (the dilution factor) times
2 (for the trypan blue solution) times 1x104
. {total cells = cell
count * 2 * 2 * 1x104
(the cell density)}. See sidebar.
Note:
• Day ‘4’ total cell count should be multiplied by 2 since they were plated at 50,000 cells/well.
• Results should be graphed for better analysis, using an application such as Excel or Prism.
References:
1. MedicineNet.com, Definition of anesthesia, http://www.medterms.com/script/main/art.asp?articlekey=2246, January 2007.
Count middle square of both grids.
1 2 3 1 2 3 1 2 3
        
Figure 2. Setup for counting, Day 0 Control (Left) to Day 4C (center) and Day 1 to
Day 4 (right).