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     collagenase solution (see below) 
              Locate two Corning 50 ml tubes 
              Measure 40 mg (2 mg/ml) of Sigma Collagenase stored at -20 deg C into each Corning 50 ml tube 
              Add 20 ml of Ca-free OR2 solution into each tube 
         Fill a few Petri dishes with Ca-free OR2 solution 
          We will place the eggs initially to rinse out the excess blood etc. 
         Rinse surgical tools (forceps and scissors) in 95% ethanol. 
         Get out a 23 or 21 gauge needle and a packet of chromic gut sutures. 
         Put ice in a silver-color aluminum tray. 
         Remove frog and place on ice. Cover head with ice. Poke feet with needle to be sure frog is out (20 minutes
     should be enough time). 
         Insert needle under skin layer on either side (if frog has had a previous surgery, go to opposite side, if frog has
     had multiple surgeries choose side that is the best healed). Make an incision. Grab muscle layer with tissue forceps
     and make another incision. 
         Tease oocyte lobes carefully out. Cut and place in Petri dishes with OR2. Take many cells as it is better to be
     choosy about which oocytes to inject. 
     Post Op Care of the frog 
              Put two to three sutures in the muscle layer and two to three sutures in skin layer. 
              Put frog in Rubbermaid tub with lid (holes punched in) with a small amount of ddH2O and allow to recover
          2-3 hours in lab. 
              Using forceps, carefully tease apart lobes, releasing clumps of oocytes then place oocytes in the collagenase
          solution. 
         Place the cells in the two Corning tubes with collagenase and close the lids tightly. 
         Place on orbiter at almost 45 to 50 RPMs for 1 hour (or more) 
          It is best to check every 15 minutes and shake dish to see if cells are dispersing. 
         Wash cells with OR2. 
          Fill the tube with OR2 to the top and gently swirl, and then decant the solution. 
          Repeat this at least 4 or 5 times until the solution is very clear. It is very important to wash the cells very well
          to rinse away collagenase. 
         Wash cells with ND96 for a few times. 
         Fill a few petri dishes with ND96 and transfer the cells to the petri dishes using a fire-polished blunt, large
     diameter pasture pipette. 
          You want a single layer of cells. You may need to use more petri dishes if you have many cells. 
         Fill a 60 mm petri dish with ND96. 
          You will collect "GOOD" cells in this dish 
         Using a dissection microscope, select good oocytes using a fire-polished glass pasture pipette with an
     approximately 2 mm diameter. 
         Record the surgery and the egg appearance in the frog database. 
         PUT THE FROG BACK IN THE ORIGINAL TANK 
          This has been neglected a few times before </protocol>
<protocol xml:space="preserve"
  title="Establishment of a Primary Culture, Chick Embryo"
  type="growth"
>
Materials 

     Chick embryo (approximately 8 days old)
     70% (v/v) ethanol for swabbing
     Sterile scissors, forceps and probes
     Sterile petri plates
     Phosphate buffered saline (PBS)
     Trypsin, cold sterilized in a 125 ml sterile erlenmeyer containing a magnetic stirring bar
     Minimum Essential Medium
     Fetal Calf Serum
     Clinical centrifuge with sterile capped centrifuge tubes
     Culture flasks
     Inverted phase contrast microscope (Optional) 

Procedure 8 

   1.Candle an 8 day old egg to ensure that it is alive. This is easily accomplished by holding the egg in front of a bright light source; the embryo can
     be seen as a shadow. Circle the embryo with a pencil. 

   2.Place the egg in a beaker with the blunt end up, and wash the top with a mild detergent, followed by swabbing with ethanol. 

   3.Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a circle of shell, thus exposing the underlying
     membrane (the chorioallantois). 

   4.With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic membrane, exposing the embryo. 

   5.Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass rod, and place the embryo in a 100mm petri dish
     containing phosphate buffered saline (PBS). Wash several times with PBS by transferring the embryo to fresh petri plates. After removal of all
     yolk and/or blood, move the embryo to a clean dish with PBS. 

   6.Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb by pulling at the proximal end. Move the
     remaining tissues of the embryo to yet another dish and wash with PBS. 

   7.Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS. Allow the tissue pieces to settle. 

   8.Remove the PBS with a sterile pipette and add 25 ml of trypsin, a
proteolytic enzyme. Stir the solution gently at 37 deg. C for 15-20 minutes. 

   9.Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal volume of Minimal Essential Medium (MEM) + 10%
     Fetal Calf Serum (FCS). FCS contains protease inhibitors which will inactivate the trypsin. 

  10.Centrifuge the cells in MEM at 1000 rpm for 10 minutes in a standard clinical centrifuge. Remove the supernatant and resuspend the pellet in 25
     ml of fresh MEM + 10% FCS. 

  11.Remove 0.1 ml of the culture and determine cell concentration and viability as directed in the previous section. 

  12.Seed two 25 cm plastic culture flasks containing 25 ml of MEM + 10% FCS to a final concentration of 10 cells/ml. 

  13.Label and place your cultures in the tissue culture incubator at 37 deg.  C and examine daily for cell density and morphology. 

  14.Note any changes in the color of the media. Tissue Culture media has a pH indicator (Phenol Red) added in order to check on the growth of cells.
     The media initially is a cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby reducing the media. Should
     this color change occur within 24 hours, the culture is most likely contaminated and should be disposed of. 

  15.Examine the cultures using an inverted phase contrast microscope. This will allow observation of the cells without opening or disturbing the
     growth. 

  16.Make cell density determinations at 10 X magnification using a square ocular grid, as explained in Chapter One for the determination of area. 

  17.Plot the cell density on a log scale vs. time of culture. 

  18.Diagram the shape of the cells at each phase. 

Notes 

The cultures will develop differently than the suspension cultures. The viable cells will grow out of the trypsinized pieces of tissue and will remain in
contact with the bottom of the culture flask. They will continue to divide and migrate until the entire bottom of the flask is covered with a single layer of
cells (contact inhibition and the formation of a monolayer). 
</protocol>
<protocol xml:space="preserve"
  title="Protocol: Growth of PC12 cells"
  type="growth"
>

     Medium:    

          DME, high glucose (4500 mg/L)