Download So, you or your lab is interested in learning primary cultures—Here

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Primary Culture Information Packet
McLaughlin Lab
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
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COVERSLIP CLEANING
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In a 500mL beaker, place 2 boxes of small coverslips, or one large box of
coverslips, and add 250 mL of Milli Q water.
Swirl gently for 2 minutes
Decant the water into the sink, make sure not to let any coverslips pour out
Add 250 mL of new Milli Q water
Sonicate for 15 minutes
Repeat steps 1-5
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
Cover the beaker with parafilm, and place the beaker on the orbital shaker for 1
hour
Decant the 0.1 N HCl into the appropriate hazardous waste drum
Wash 3 times with Milli Q water
Add 250 mL of 95% ethyl alcohol, and place the beaker on the orbital shaker for 1
hour
Decant the 95% ethyl alcohol into the appropriate hazardous waste drum
Wash 2 times with Milli Q water in the hood
Add 250 mL of 95% ethyl alcohol, and place parafilm on the top.
Store in the cell culture room until use
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Dissociation for Neuronal Culture
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.
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NOTE: Dissection time should take less than 30 mins!!!
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.
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Maintance of Primary Cultures
Primary cultures are maintained at 37oC & 5% CO2.
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).
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Borate Buffer (4º)
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4.76 g Boric Acid
2.54 g Borax
1000 mL of ddH2O
pH to 8.4 with NaOH
Sterile Filter
Plating Media (4º)
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40 mL F12
40 mL FBS
420 mL Dulbecco Modified Eagle Medium
(DMEM)
1 mL of PS
200 uL of 200 mM L-Glutamine
D2C (4º)
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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)
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1 bottle (500 mL) Neurobasal Media
1 bottle (10 mL) 50X B27
P/S
Stock Ara-C (500X) (-20º)
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100 mg Ara-C
6 mL sterile H2O
Sterile Filter
Aliquot 0.5 mL
Working Ara-C (4º)
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20µL Stock
10 mL sterile ddH2O
10% Formaldehyde (RT)
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5 mL of 37% Stock Solution
45 mL of PBS
Glucose Free Balanced Salt Solution
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5.26 g NaCl
0.125 g KCl
66.6 mg CaCl2
1.43 g Hepes
600 mL ddH2O
Sterile Filter
pH 7.3
PLO
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1 mg PLO : 2 mL Borate Buffer
Sterile filter
Make up fresh each time it is used
HBSS (4º)
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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)
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10% Benzalkonium Chloride (B1383
Sigma)
1.25% Ethyl Alcohol
Dissolve in alcohol with stirring
Add 88.75% Milli Q water
MEM/BSA/Hepes
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487.5 mL of clear MEM
12.5 mL of 1 M Hepes
50 mg of BSA
Sterile Filter
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Tissue Culture Media &
Supplements
Product
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
F-12 Nutrient Ham
Hepes Buffer
Vendor
Invitrogen
Invitrogen
Invitrogen
Invitrogen
Invitrogen
Invitrogen
Invitrogen
Invitrogen
Cambrex
Hyclone
Catalog #
11960-089
CASE
11995-073
CASE
51200-038
21103-049
11058-021
17504-044
17502-048
15140-122
17-605C
SH30070.03
Sigma
Sigma
Sigma
Sigma
C-6645
P-3655
N4888
H0887