Cortical neuron cultures

9/19/05

 

 

 

Plate and coverslip coating

 

  1. PDL cat #P6407 from Sigma. Stock is 1mg/mL, dilute 1mL in 50mL tissue culture sterile ddH2O for 20ug/mL final. See Sigma cell culture catalog for additional details.

 

  1. Coverslips from Bellco, cat#1943-10012 (12mm) are washed once in batch with nitric acid overnight in a 2L flask shaking. The nitric acid can be collected and stored for reuse. Then ddH2O is used to wash the coverslips in the same flask shaking for about a week, changing the water daily. After a final wash in 70% ethanol, the coverslips are poured from the flask into a 13x9 baking dish, and the ethanol is poured off. The coverslips are autoclaved on the dry cycle (20-20) then transferred when cool to a dish for storage. Cleaned coverslips can be stored indefinitely.

 

  1. Coverslips are always coated overnight at 37 in a 24 well plate. Plastic dishes can be coated for as little as 2 hours. PDL can be reused several times for up to 1-2 months, Store at 4 in between uses. Wash coated CS and plates 2X with ddH2O, then they are ready to use. Coated CS and plates can be stored dry in foil at 4 for up to 2 months.

 

  1. Polyornithine at 40ug/mL is an acceptable alternative substrate. We use this all the time for cerebellar granule cells. Preparation is similar.

 

  1. Often we add Laminin to the PDL for cells cultured on coverslips. The addition of laminin helps the cells to stick especially when a low density is desired. Laminin is cat # 23017-015 from Invitrogen. Thaw on ice as per directions. Stock solution is Ò0.5-2mg/mLÓ, we make 100uL aliquots and re-freeze at –80. Dilute one aliquot in 15mL 20ug/mL PDL to coat coverslips. I do not store the coated coverslips for more than a day or two, and I do not reuse the laminin/PDL solution.

 

Dissection

 

  1. We use E16-17, or P0 CD1 mice or E17 or P0 Sprague-Dawly (CD-1) rats for cortical and E19 or P0 rats for hippocampal cultures. Postnatal cultures work best when done within 24 hours of birth. We have also done cortical cultures from C57BL/6 and 129SvJae mice at P0. Our regular animals are from Charles River Labs.

 

  1. On the day of dissection, soak dissecting tools in 70% EtOH. You need big scissors, big forceps, small scissors, two #5 forceps, one #3 forceps, and the tiny scissors for hippocampus.

 

  1. Prepare one 10cm dish and 2-3 6cm dishes with 1X dissociation medium (DM).

 

  1. Begin to prepare dissociation solutions. For one litter you need about 10mL papain solution and 25mL trypsin inhibitor. For single pup cultures or more litters, you may need to scale up. For 10mL papain at 20U/mL you need to add to 10mL DM 3.2mg cysteine-HCl (cat # Sigma C-7477). If more volume or more papain, scale up. This needs to be pHed by color. Takes about 10uL 1N NaOH. For the trypsin inhibitor solution, to 25mL DM add 250mg trypsin inhibitor (Sigma cat # T-9253)and 250mg BSA (Sigma A-9418). Mix until dissolved, then add about 80uL 1N NaOH to pH to color. Warm these solutions at 37 until ready to use.

 

  1. For embryonic cultures, sacrifice the mother with CO2, then remove the uterus to a dish, and cut the embryo heads into a dish with DM. For P0 cultures, cut the heads into DM. If genotyping is required, clip the tails at the same time into labeled tubes.

 

  1. Place the #3 forceps through the eyes, and pull front to back along the sagital suture with the #5 forceps to open the skull. Grab the skull at the rostral end of the suture and pull laterally to expose the brain. Pinch the forceps closed, then scoop under the brain to free from the skull. Transfer to a new dish.

 

  1. From the superior surface, place forceps inbetween the hemispheres and pull laterally to expose the fibers the connect cortex to subcortical regions. Snip this connection to release hemispheres. Turn hemisphere to place lateral side face up. Using two sets of forceps, pull off meninges, flipping over to medial side to finish. Moved cleaned cortex to a new dish.

 

  1. If hippocampus is desired, use small scissors to cut the whiter colored banana-shaped structure off the cortex and move to new dish.

 

  1. If cortex is desired,  cut off the superior part of the hemisphere with equal contribution front to back. Move to new dish.

 

  1. Put tissue to be dissociated into 15mL tube.

 

  1. Finish dissociation solutions. Get papain (Worthington 3126) and warm in your hand. This is a slurry with units calculated for each bottle. Multiply u/mgP times mgP/mL to get units/mL (often 800-1000). You will want 20U/mL for embryonic cultures, 30U/mL for postnatal. After adding to tube of DM with cysteine, return to 37 degree water bath until solution is completely clear (about 5Õ). Use 0.2um sterile syringe filters to sterilize papain and trypsin inhibitor solutions.

 

  1. Pipette off DM from tissue. Add 2-10mL papain (depending on number of pups) and invert tube to fully mix tissue with solution. Incubate 3-7 minutes at 37 degrees, longer for postnatal than embryonic.

 

  1. Pipette off papain and add 10mL trypsin inhibitor. Invert to mix and let tissue settle. Pipette off and repeat.

 

  1. Pipette off trypsin inhibitor and replace with 5 mL serum-containing cortical medium. Pipette up and down 10-15X with 5mL pipette until tissue is fully dissociated. It should fall apart easily and should not become gooey. If it looks like snot, the cells are all lysed and all is lost. For hippocampal neurons dissociation is optimal with a final few passes through a fire polished pasteur pipette. Optimally this can be siliconized to prevent cell sticking and loss.

 

  1. Count the cells on a hemacytometer. You should see little debris in the background (you always see some with postnatal, but hope to minimize) and you should see some cells with processes still attached.  You can expect about 500,000 per animal for hippocampus, 5-10x10 6 per animal for cortex.

 

  1. Plate at an appropriate density for application. For cell staining, ideal is 10,000-100,000 per well of 24 well plate. 50,000 is a nice easy density that works well. For biochemistry, plate anywhere from 1-3 million per well of a 6 well. For luciferase assays we do 3 million in serum, for westerns you can get away with 1 in NB-B27.

 

  1. Best is to plate the cells in serum-containing medium so they stick well. Then if you want the cells in serum-free, at least 4 hours later or after overnight,  remove the medium and replace with Neurobasal-B27. If you want to block glia, treat with AraC to 10uM on DIV2. For long-term cultures, feed every 3-4 days by removing 1/2 the medium and replacing with fresh. Mouse neurons seem to be especially sensistive to the feeding schedule, and need to it start by 4-5DIV. Rat neurons can wait longer, up to 7DIV before feeding is required.

 

  1. We use at 5DIV for gene expression assays (transfecting as early as 3DIV), we are infecting with virus at 1DIV for assays at 8DIV, for synapses transfect at 4DIV for staining at 16-20DIV, see spines at 21-28DIV. 

 

 

Glia

 

  1. For long-term cultures (21-28 days), for good low density cultures (10-50,000 cells/12mm coverslip) or for optimal synapse development, the best strategy is to plate the neurons on top of a confluent layer of glial cells.

 

  1. Both mouse and rat neurons can be plated on rat glia if that happens to be what you have. In fact, for some reason the rat glia seem to be easier to maintain, though we havenÕt tried especially hard. Rat neurons do not appear to survive on mouse glia, however.

 

  1. To get glia, you need about 3 P1 pups. Chop up the brain and dissociate as described for the neurons. Plate on uncoated plastic dishes in DMEM with 10% FBS (glial medium, see below). Plate 0.5-2 million cells per 10cm plate. Next day change the medium to get rid of unstuck cells. Allow to grow to confluence (this will take 5-7 days) then split with trypsin onto coverslips. Do not replate neurons after trypsinization. However, you can hold them at confluence for a week or two (feed every 3-4 days by replacing 1/2 the medium with fresh) and do not add AraC.

 

  1. Plate glia at 10K/well if you are going to let them grow for at least 5 days or 30K/well if you need to use in the next few days. 100K/ well if you need to use them next day. Once the glia are confluent, add AraC at 10uM final to inhibit further glial growth.

 

Reagents

 

1. Cortical plating medium (with serum – gene expression studies):

 

500ml BME (Sigma cat#B1522 or Gibco/Invitrogen cat#21010-046)

6.67 mL 45% glucose (XM final)

10% calf serum

5ml pen/strep (100X stock Sigma cat#P0781)

5ml glutamine (100X stock Sigma cat#G7513)

 

 

2. Serum-free medium:

 

500ml Neurobasal (Gibco cat#21103-049)

10ml B27 supplement (Gibco Cat#17504-044)

5ml pen/strep (100X stock Sigma cat#P0781)

5ml glutamine (100X stock Sigma cat#G7513)

 

 

 

 

3. Glial Medium

500mL DMEM (Gibco 11960-069)

50mL FBS

5mL pen/strep

5mL gluamine

 

 

4.  1M Hepes (pH 7.4)

23.8g Hepes (Sigma)

80mL ddH2O

pH to 7.4 with NaOH

ddH2O to 100mL and sterile filter. Store 4 degrees for 6 months

 

 

5 .1M MgCl2

20.3 g MgCl2.6H2O

to 100mL with ddH2O

Stir into solution and autoclave or sterile filter

 

 

6. 10XK= Ky/Mg stock (10mM Kynurenic acid/100mM MgCl2):

1X HBSS (Ca/Mg free Gibco 14170-112)     400 mL

kynurenic acid (Sigma K3375)                       945 mg

1M Hepes (pH 7.4)                                         50 mL

1M MgCl2                                                      50 mL

 

Takes all day shaking and heating at 37 to get into solution. Adjust pH by color as you go along by adding 1N NaOH. May need to leave overnight. Break up last clumps of KA by smashing with a pipette. Aliquot in 50mL and store –20 for a year.

 

 

7.  Dissociation Medium (DM)

500mL 1X HBSS (Ca/Mg free Gibco 14170-112)

6.67mL 45% glucose (2.5M stock)

50mL 10X Ky-Mg stock

Store 4 degrees 6 months

 

8. AraC 1mM (100X)

 MW=243.3g/mol

cytosine-b-d-arabino-furanoside (Sigma C-1768)       2.43mg

ddH2O to 10mL

Store –20 for