1
So, you or your lab is interested in learning primary cultures—Here is what we can do for you.
1.
Attached are protocols for our plate prep, dissection/dissociation, media formulations, and culture care instructions.
2.
You are welcome to watch 2 dissections in our lab, performed by our expert staff, provided you schedule 2 weeks in advance.
3.
Once our cells are plated, you may look at them twice during their maturation.
We suggest that you already have an IACUC protocol in place for the animals you will be sacrificing. One week after watching our dissection, it would be wise to perform a dissection/dissociation on your own while the information is still fresh.
That way, when watch another dissection in our lab, you can trouble shoot your methods and techniques.
Contact Information
Lab Phone: 936-3872
Location: 8141A MRB III

2
1.
In a 500mL beaker, place 2 boxes of small coverslips, or one large box of coverslips, and add 250 mL of Milli Q water.
2.
Swirl gently for 2 minutes
3.
Decant the water into the sink, make sure not to let any coverslips pour out
4.
Add 250 mL of new Milli Q water
5.
Sonicate for 15 minutes
6.
Repeat steps 1-5
7.
Decant the water, and add 250 mL of 0.1 N HCl a.
248 mL of Milli Q water b.
2 mL of 12 N HCl
8.
Cover the beaker with parafilm, and place the beaker on the orbital shaker for 1 hour
9.
Decant the 0.1 N HCl into the appropriate hazardous waste drum
10.
Wash 3 times with Milli Q water
11.
Add 250 mL of 95% ethyl alcohol, and place the beaker on the orbital shaker for 1 hour
12.
Decant the 95% ethyl alcohol into the appropriate hazardous waste drum
13.
Wash 2 times with Milli Q water in the hood
14.
Add 250 mL of 95% ethyl alcohol, and place parafilm on the top.
15.
Store in the cell culture room until use

3
One Day Before Dissection
1.
UV culture plates for 1 hr.
2.
Make Borate Buffer/Poly-L-Ornithine [sigma, P-3655] (2 mL: 1 mg) to coat plates with and without coverslips.
3.
Add 2 ml of PLO to each well, and leave it on for at least 1 hr.
4.
Wash each well 2X with tissue culture water. Remove lids and allow it dry in the hood for 2-4 hrs.
5.
Add 1.5 ml of plating media to each well, and place plates in the incubator until dissection.
Before Dissection
1.
Weight out 3 mg of Trypsin [sigma, T9935] and dissolve in 5 mL of HBSS. Sterile filter into a
35 mm culture dish & label.
2.
Add 3 mL of HBSS to 5 of the 35 mm culture dishes.
3.
Add 450 µL of Trypan blue to a 1.5 mL eppendorf tube.
4.
Prepare 70% alcohol, scissors, forceps, and absorbent pads for dissection.
5.
Use syringe with an 18 gage needle to remove sedative from the bottle; then, replace the 18 gage needle with a 22 or 25 gage needle for injection into the rat.
6.
Calculate the number of mL that are need to plate all the wells.
7.
Place plating media into water bath at 37ºC.
Dissection
1.
Inject the rat with the sedative, Nebutol, by IP injection, then wait a few minutes for her to relax and lay down
2.
Check her “blink” reaction and squeeze her foot to determine if she is sedated enough.
3.
Place her on the absorbent pad belly up, and spray the belly with 70% alcohol.
4.
Use the large forceps to lift the belly skin. Cut open with the heavy-duty dissecting scissors.
Cut the skin along the midline up to the diaphragm area.
5.
Then cut the muscle along the midline to expose the embryos and the diaphragm.
6.
Cut both sides of the rat’s diaphragm to stop respiration.
7.
Use the large forceps to pull all the embryos. Cut the embryos open with fine-tip dissecting scissors. Cut off the head of the embryo into HBSS culture dish.
8.
Transfer one embryo head into a new dish of HBSS that is now under the microscope.
9.
Hold the head straight up. Puncture the skin and membrane outside of the brain with the dissecting forceps. Peel the skin and membrane toward the front of the head until the brain pops out. Turn the head around and peel toward the back of the head until the brain stem is exposed. Put the dissecting forceps under the brain stem and pull toward the back of the head until the brain is separated with the skull. Pinch-cut the brain stem to separate the brain completely.
10.
Place the brain in a new HBSS dish. With the dorsal of the brain facing up, hold the brain stem gently with forceps. Separate the hemispheres with the dissecting forceps. Pull the cerebral cortex away from the brain stem.
11.
Place one cortex in a new dish of HBSS. Flip the cerebral cortex so that the outside is facing up. Pinch-cut the olfactory bulb with the dissecting forceps. Peel the meninges from the front toward the back. Try to peel it off in one piece.
12.
Pinch cut any mid brain structures underlying the cortex.
13.
Transfer the cortex to a new dish with HBSS. Repeat for the other cortices.

NOTE: Dissection time should take less than 30 mins!!!
4
14.
Transfer the dish with all the cortices to the hood. Transfer the cortices into Trypsin solution with forceps, making sure to take as little HBSS as possible with them.
15.
Let the cortices sit in Trypsin for 30 mins in the hood at RT.
During Trypsinization
1.
Aspirate the coating media from the culture plates.
2.
Add 3 mL of HBSS into two more 35 mm culture dishes for wash.
3.
Add 5 mL of plating media into two 15 mL tubes. Set out 3 more tubes for titration.
After Trypsinization
1.
Transfer the cortices form the Trypsin to a new dish with HBSS with a Pasteur pipette, one at a time. Do not break the cortices. Swish them around to wash off the Trypsin. Repeat
2.
Transfer the cortices with a new Pasteur pipette to the first tube of plating media. Slowly triturate the cortices with the Pasteur pipette to maximize the total cell number & viability. Let the chunks settle down to the bottom of the tube.
3.
Transfer ONLY the cell suspension (not the chunks) to a new 15 mL tube with a Pasteur pipette. Again, slowly triturate with cortices. Continue this process until the suspension looks even.
4.
When Titrations are complete, add 50 µL of the cell suspension to the eppendorf tube with
Trypan blue (a 1:10 dilution). Mix well by flipping the tube upside-down several times.
5.
Remove 50 µL of Trypan blue mixture, and add it to the hemocytometer (it takes approximately 15-20 µL to fill each side).
6.
Count the number of live cells and dead cells in 3 of the 4X4 quadrants (1 on one side, & 2 on the other side).
7.
Use this series of formulas to determine the needed cell to media dilution. a.
(Live Cells)/(3 squares) = y b.
y * 10 = z (for 1:10 Dilution in Trypan blue) c.
z * 10,000 = (hemocytometer volume correction)
8.
Dilute the cell suspension to the desire plating density with proper media a.
335,000 cell/mL
9.
Plate 2 mL/well, and change the pipette after every 2 plates.
10.
Swirl media bottle to insure proper homogeneity of cell suspension in between plates.
11.
Make sure to mark the date on the plate cover.
12.
Carefully return dishes to the incubator, making not to splash media on lids or disrupt cells.

5
Primary cultures are maintained at 37 o
C & 5% CO
2
.
Media is partially replaced every 2 – 3 days.
For Neuronal Cultures : Glial cell proliferation is stopped by adding 1-2µM cytosine arabinoside to the cultures two days after dissociation. On the third day, the media is changed from Plating Media to
Neurobasal. All Neuronal experiments are conducted 3 weeks after dissociation.
For Mixed Cultures: Glial cells are inhibited after two weeks in culture with 1-2

M cytosine arabinoside, after which the cultures were maintained in growth medium containing 2% serum and without F12-nutrients. All experiments are conducted one week following mitotic inhibition (25-29 days in vitro ).

Borate Buffer (4º)

4.76 g Boric Acid

2.54 g Borax

1000 mL of ddH
2
O
 pH to 8.4 with NaOH

Sterile Filter
Plating Media (4º)

40 mL F
12

40 mL FBS

420 mL Dulbecco Modified Eagle Medium
(DMEM)

1 mL of PS

200 uL of 200 mM L-Glutamine
D
2
C (4º)

10 mL FBS

1 mL PS

12.5 mL of 1M hepes

471.5 mL Dulbecco Modified Eagle
Medium (DMEM)

6.25 mL of 200 mM L-Glutamine
Neurobasal (4º)

1 bottle (500 mL) Neurobasal Media

1 bottle (10 mL) 50X B27

P/S
Stock Ara-C (500X) (-20º)

100 mg Ara-C

6 mL sterile H
2
O

Sterile Filter

Aliquot 0.5 mL
Working Ara-C (4º)
 20µL Stock

10 mL sterile ddH
2
O
10% Formaldehyde (RT)

5 mL of 37% Stock Solution

45 mL of PBS
6
Glucose Free Balanced Salt Solution

5.26 g NaCl

0.125 g KCl

66.6 mg CaCl
2

1.43 g Hepes

600 mL ddH
2
O

Sterile Filter
 pH 7.3
PLO

1 mg PLO : 2 mL Borate Buffer

Sterile filter

Make up fresh each time it is used
HBSS (4º)

8.4 g NaCl

224 mg KCl

2.38 g Hepes

1.0 g Glucose

1000 mL Milli Q
 pH to 7.3 with NaOH

Sterile Filter
Roccal II (10%) (RT)

10% Benzalkonium Chloride (B1383
Sigma)

1.25% Ethyl Alcohol

Dissolve in alcohol with stirring

Add 88.75% Milli Q water
MEM/BSA/Hepes

487.5 mL of clear MEM

12.5 mL of 1 M Hepes

50 mg of BSA

Sterile Filter

Product
Vendor Catalog #
DMEM w/o L-glutamine
DMEM w/ L-glutamine
MEM w/o L-glutamine
Neurobasal
Opti-Mem
B27 Supplement
N2 Supplement
Pen/Strep
L-Glutamine
FBS
Cytosine Arabino Furanoside
(Ara-c)
Poly-L-ornithine
Sigma
Sigma
F-12 Nutrient Ham
Hepes Buffer
Sigma
Sigma
Invitrogen 11960-089 CASE
Invitrogen 11995-073 CASE
Invitrogen 51200-038
Invitrogen 21103-049
Invitrogen 11058-021
Invitrogen 17504-044
Invitrogen 17502-048
Invitrogen 15140-122
Cambrex
Hyclone
17-605C
SH30070.03
C-6645
P-3655
N4888
H0887
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