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    • CommentAuthorjuruble
    • CommentTimeSep 5th 2007 edited

    This protocol explains how to: create Xenopus dissociated eyebud cultures.

    Protocol revised: 4/19/2007

    Protocol written by: Julie Ruble

    NOTE: There is a "master resource" for multiple types of culturing (including dissociated eyebud cultures) and related techniques (immunostaining, movie-capturing, etc) here.  This protocol is a short, concise guide to making dissociated eyebud cultures. Please review both protocols to find which is most helpful for your purpose.


    Days Before Culturing

    • Autoclave at least a dozen fire-polished pipettes and a dozen pulled pipettes the day before you'll be culturing.  You can also autoclave your tools -- forceps, etc.
    • Make sure you will have stage 24-28 tadpoles when you're ready to begin your cultures.
    • Make sure you have enough poly-ornithine dishes (please see this section of the master protocol for how to make poly-o dishes)

    30 Minutes Before Culturing

    • Take out laminin tubes from the blue laminin box in the small freezer in the lab and defrost them in the refrigerator.  You need 1 tube for every 4 culture dishes you plan to make, since each tube has 900 ul and each dish will need 200 ul.
    • Spray forceps with ethanol and place them in the hood.  Spray all hood surfaces with ethanol.
    • UV the hood: pull down the screen, turn on receptacle, germicidal, and blower, and start timer.  These instructions are for the EdgeGARD hood and may vary slightly with other hoods.


    NOTE:  You'll be using sterile technique as you culture.  This means avoid uncapping solutions for longer than absolutely necessary, avoid passing your hands/arms over open containers, avoid touching the inside of containers/foil.  In addition, don't touch things outside of the hood and bring your hands back in without cleaning with ethanol.

    1. Here is a list of the items you should have in your hood when you start to culture:
      • Autoclaved fire-polished pipettes
      • Autoclaved pulled pipettes (for instructions on using the autoclave, click here)
      • 2 autoclaved and/or ethanol'ed forceps
      • Small p35 petri dishes
      • Poly-o dishes
      • Defrosted laminin tubes
      • Sterile PBS (from tissue culture room fridge)
      • REAG (from tissue culture room fridge)
      • DISAG (from tissue culture room fridge)
      • Serum free culture media (from tissue culture room fridge)
      • Pipettemen -- p200, p20, and p1000
    2. Turn off the germicidal switch and turn on the fluorescent light in the hood.  Pull up the screen.  Wash your hands and then ethanol them generously.  From now on, after touching anything outside of the hood, you must re-ethanol your hands. 
    3. Use the p1000 pipetteman to dilute each laminin tube with sterile PBS to a concentration of 10 ug/ml.  With our current laminin tubes (which are 1.3 ug/ml), this means put 903 ul of sterile PBS into each laminin tube.  Always check the concentration of your laminin before you begin (it's written on a card in the laminin box) to calculate how much sterile PBS to add.
    4. Turn on the vacuum knob.  Use the vacuum / aspirator (looks like an Erlenmeyer flask with tubes sticking out of it) to suck the liquid out of the poly-o dishes one at a time, getting as much as you can around the edges.  Don't take all the lids off at once -- remember sterile technique.
    5. Using the p200 pipetteman, put 200 ul of the laminin mixture in a "bubble" right on top of the coverslip. If laminin doesn't stay on coverslip, you can suck it up and "rebubble."  Set these laminin-coated culture dishes aside -- we will come back to them later.
    6. Get out 4 small p35 petri dishes and set the lids aside (we will use them later).  Fill each dish with REAG.  These dishes will be used to rinse the tadpoles.
    7. Using a fire-polished pipette, pick up tads (5-7 for each culture dish you'll be using) and put the total number of tads into the first dish of REAG.  This is the first rinse, and during this, you can remove any vitelline envelopes (clear bubbles that may still be around the tadpole) using your forceps.
    8. Using a fresh fire-polished pipette, put tads into second REAG dish.  Repeat this, using a fresh fire-polished pipette each time, with the last two dishes, rinsing the tads a total of 4 times.  On the last rinse, you'll keep the tads in the REAG and dissect out their eyebuds, gathering 1-2 from each tad and leaving them in a pile in the corner of the REAG dish. To dissect out eyebuds, peel back top layer of skin and pluck out eyebud.
      • NOTE: They can't sit in the REAG too long (no longer than ~40 minutes), so be efficient or do your rinses in batches to ensure you can get the dissections done in a timely manner.
      • Here is a video of eyebud removal (taken by S. Bossie and R. Thomason) to help you understand the process:

    9. Prepare the 4 small p35 petri dish lids that you saved in step 6 with serum free culture media.  We use the lids to save dishes, but you could also use 4 new dishes.
    10.  Using the p200 pipetteman with a yellow tip, pump serum free culture media up and down a few times to coat the inside of the tip (this ensures that the eyebuds won't get stuck in the tip as you transfer them).  Suck up the eyebuds from the REAG and gently pipette them into the first dish of serum free culture media.  This is the first of 4 rinses.
    11. Complete the 4 rinses using a different yellow pipette tip for each transfer (and remembering to coat the inside of each tip first).
    12. Take out 2 small p35 petri dishes.  Fill one with DISAG and in the other, make 10ul "bubbles" of serum free culture media (as shown in the figure below).  Make as many bubbles as you have culture dishes; the figure below is for 8 culture dishes. 
      SFCM bubbles
    13. You'll do the DISAG step that follows in "rounds" so none of the eyebuds stay in DISAG too long.  Take ~18 eyebuds (enough for ~2-3 dishes) and put them in one corner os the DISAG dish.  On the lid above this pile of eyebuds, write "0" and start your timer.  This is your first round of eyebuds.  After ~2 minutes, you'll put in the next batch of ~18 eyebuds in a separate spot and write "2" on the lid over this pile, etc.  This "round" system will prevent you from getting backed up and unable to take the eyebuds out at the correct time (15-25 minutes).
    14. While eyebuds are in DISAG, you will do 3 rinses of the laminin-covered poly-o culture dishes with sterile PBS.  Be sure that by now the laminin has been left on for at least 30 minutes (it can be left on for up to half a day).  For each rinse, pipette the sterile PBS into each dish (lifting only one lid at a time), swirl gently, and then use a vacuum/aspirator to suck out most of the PBS. 
    15. After sucking out the last rinse of sterile PBS, use the electric pipette gun with a stripette attached to suck up a lot of serum free culture media and dispense ~2 ml into each culture dish.
    16. The eyebuds in the DISAG will get fluffy.  As you prepare to remove them, clear off your workspace and place the tips of your pulled pipettes next to the eyebuds to choose a good one.  The ideal tip will be very fine compared to the size of the eyebud.  When you've found a good pipette, squeeze all the air out of a rubber bulb and put it on the pipette. 
    17. Using a p200 pipetteman with a new (and lubricated with serum free culture media) tip, suck up however many eyebuds you're culturing per dish (~5-7) and gently pipette them into one of your serum free culture media "bubbles." 
    18. When all of your eyebuds are in their bubbles, use the fine pulled pipette to pump them up and down 2-3 times, disintegrating them into a very fine speckle (look for "snow," not "snot").
    19. When you have all of your eyebuds disintegrated, you can use the pulled pipette to suck up one bubble at a time and gently spray the "snow" out in a zigzag pattern over the coverslip in one of your poly-o culture dishes.  If you see any of the "snow" that is floating above the slip and not sticking, you can suck it up again and gently re-spray.  NOTE: KEEP YOUR DISH OF SFCM BUBBLES COVERED any time you're not working in it!  Otherwise the air flow in the hood will cause your bubbles to evaporate, ruining the eyebuds in that bubble.
    20. When all dishes are finished, you can put them in a big petri dish and label them.  Clean up the area.
    • CommentAuthorjoiordanou
    • CommentTimeNov 10th 2007
    I tend to really rip my tadpoles apart when removing eyebuds which makes it difficult to keep the actual eyebud tissue separate from the rest of the debris. I have found that transfering eyebuds to serum free culture media after every few tadpoles helps me to not lose as many and not plate lots of exta tissue.
    • CommentAuthorkilang
    • CommentTimeNov 11th 2007
    Before Culturing:
    I put pretty much everything except the tadpoles and laminin (& iPod) in the hood while it was being sterilized.

    Also, I added to the "Things to have in the hood" list to make setup simpler:
    -4 fire-polished pipettes (autoclaved)
    -pulled pipettes (autoclaved)
    -rubber bulbs
    -2 ethanoled forceps (one "good" pair for removing skin/plucking out eyebud and one "bad" pair, with a bent tip that helped immobilize the tadpole's body)
    -pin-on-a-stick (helped lift skin over the eyebud)
    -parafilm pieces (to cover the big petri dish at the end)
    -boxes of pipette tips (yellow and blue)
    -ethanol spray bottle
    -sleeve of small petri dishes
    -1 large petri dish (to hold the smaller ones)
    -electric pipette gun with plenty of stripettes
    -fresh glass tip for vacuum
    -poly-o dishes
    -solutions (REAG,DISAG, etc)

    Dissecting out eyebuds:
    -Older tadpoles have thicker, stickier skin but their eyebuds hold together better and are easier to get out.
    -Having separate areas within the dish for whole tadpoles, eyebuds, the tadpole being dissected, and the 'used' tadpoles helps things go more smoothly (and fewer eyebuds are lost)
    • CommentAuthorjuruble
    • CommentTimeJan 4th 2008

    Originally posted by Courtney Cron:


    When using the EdgeGARD hood, the germicidal switch turns on the UV light. Be sure that the only switches that remain on while you are working in the hood are the RECEPTACLE and the BLOWER, not the germicidal. You risk exposure to UV light if the germicidal switch stays on: there is no automatic safety feature like there are in the other hoods.

    • CommentAuthorrethomason
    • CommentTimeAug 13th 2009
    Awww my old protocol. Glad it's still in use and being updated!

    Laminin should never go into the hood during UV sterilization. Your best bet is to put the laminin in the fridge as you set up your UV sterilization so that when that time period is done, the laminin will be ready for you to culture