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    • CommentAuthorjuruble
    • CommentTimeJun 26th 2007 edited
     

    This protocol explains how to: culture, immunostain, image, and analyze Xenopus laevis eyebuds.

    Protocol revised: 6/26/2007

    Protocol written and revised by: Rebecca Thomason, Sheena Bossie, Sarah Tyndall, Sherry Messersmith, Barbara Lom, Ian Willoughby, William Wood, Courtney DeBruin, Julie Ruble

    THIS PROTOCOL IS IN PROGRESS

    TABLE OF CONTENTS

    General Notes and Precautions

    General Notes on the Preparation of Hood and Work Space

    How to Prepare Dishes for Cultures
    Acid Cleaned Coverslips
    Sylgard Treatment
    Polyornithine Treatment

    How to Treat the Laminin dishes and mix the Media Culture
    Making Culture Media
    Making Embryo Extract
    Laminin Treatment

    Embryonic Xenopus Explant Cultures

    Creating whole eyebud cultures
    Creating dissociated eyebud cultures


    Taking movies of eyebud cultures
    Using PictureFrame™
    Important tips if Immunostaining
    Addition of FGF/DMBI
    Making a movie in Quicktime

    Immunostaining

    Mounting coverslips on slides

    Analysis using Fluorescent Microscopy
    Taking Pictures
    Organizing Data

    Movie Analysis with Photoshop and ImageJ
    Creating actions
    How to use actions
    Batch Processing
    ImageJ analysis
    Understanding macros

    Closing thoughts


    GENERAL NOTES
    (back to Table of Contents)

    Important things to note before you begin:

    1. Be patient. There is a lot of room for error and problems, just work through them and learn from your mistakes.
    2. Sterilization. It is extremely easy to contaminate your cultures, so be compulsive about cleaning and sterile techniques.
    3. Scheduling and Organization. Everything is very time sensitive. Figure out what can be completed early, and what can be done while you are sitting around waiting for something to finish.
    4. Write down EVERYTHING! It is better to be safe than sorry. This includes the “dumb” stuff such as “the tadpole was at stage 38 when we started and it was really wiggly.”
    5. Read this whole protocol through before you begin. There are so many things that require thawing and setting up the night before you begin, it would be helpful to jump on that stuff right away!


    About this Protocol:

    This will be your bible while you are working on Xenopus laevis embryo explant cultures. At the beginning of each section, we have broken down the important steps into a few words, highlighted in orange.


    Important Pipette Info:

    This information applies to the whole protocol: don’t assume materials will magically appear under the hood. Throughout the culturing process, you will need individually packaged stripettes in two sizes: 10 ml and 25 ml. You will use these a lot. Don’t reuse them, except when you are rinsing with PBS (you’ll learn about this later). Use the Costar® Stipettor™ automatic pipetter located next to the refrigerator to attach to the end of these pipettes. Do not remove the cotton from the end. This automatic pipetter gadget is used to suck automatically the liquid up into the pipette. After you are finished using this gadget, replace it back on the lab cart and plug it back in the wall for the battery to recharge.

    Glass Pipettes are autoclaved and there are three different kinds. One is found in a brown-tinted plastic container. Check to see if there is some masking-type tape with brown stripes: this means they have been autoclaved. The unmodified glass pipettes are used just for rinsing (suctioning out PBS). When you are finished using these, throw them away in the glass trash container. Again, use as many as you need (within reason) because they are easily replaced, as well. Don’t resterlize them (if you run out, tell the lab technician or a work study student). The second kind of pipette is a fire-polished pipette, which is used to transfer the embryos from dish to dish. The endings of these are wider in diameter than the other glass pipettes. Finally, for dissociated eyebud cultures, there is a third type of pipette: a pulled pipette, which has a longer, finer tip and is used to strain the eyebuds into individual cells.

    There are also pipetteman. There should be 2 or 3 located in the hood (you will definitely find a 1000 µl and a 250 µl for sure). These pipettemen take the plastic tips located in the blue and yellow boxes.


    ALL PIPETTES AND ATTACHMENTS SHOULD BE STERILIZED

    hood setup

    GENERAL NOTES ON THE PREPARATION OF HOOD AND WORK SPACE (back to Table of Contents)

    Blower — Ethanol — UV — hands

    1. Have a look at your hood space. Are there stray items placed haphazardly around (did someone else who is using the hood leave his or her tools lying around)? Clean up the area, but remember to place all of your tools (such as your microscope, forceps, pipetteman) inside the hood to sterilize.
    2. Turn on the blower (see below for button symbols on the hood). Spray the inside of hood (paying special attention to the space you will be working in) with 70% ethanol (you will find a spray bottle in the hood area). Turn on the UV light for 20 or more minutes. Do not use the UV without the plastic shield in place; without proper protection it can burn skin and eyes (i.e. don’t look directly at the light).
    3. When ready to use the hood, switch off the UV light and turn on the normal light. Leave the blower on (NOTE: if you leave the blower on all the time, anytime you open the shield, the UV light will shut off automatically).
    4. Wash your hands with soap and then let your hands dry. Using gloves will provide additional protection, but isn’t required. Either way, MAKE SURE TO SPRAY YOUR HANDS/GLOVES with the EtOH. NOTE: anytime you take your hands out of the hood, there is the risk of dragging bacteria back. To be on the safe side, always spray your hands prior to placing them back in the hood.

    hood buttons


    HOW TO PREPARE DISHES FOR CULTURES (back to Table of Contents)

    clean dishes — acid clean coverslips
    glue coverslips (for movie dishes) — get sterile — pipette poly-O — wait overnight — rinse PBS (x4) — UV PBS culture dishes and lids for 30+ minutes


    To make culture dishes, you will put acid cleaned coverslips into sterile/new 35 mm dishes under sterile conditions and then do the Poly-O treatment. For movie dishes, you will use 35 mm dishes with holes (20 mm diameter) drilled in the center (using an electric drill), do the sylgard treatment, and then the Poly-O treatment.


    You have TWO options: 1. You can make a bunch of 35 mm culture/movie dishes with the Poly-O treatment, place them in big 150 mm petri dishes, and put them in the fridge, or 2. You can place acid-cleaned coverslips individually in 35 mm petri dishes and put them back in their sleeve to keep in the hood to be poly-O treated later. Either way, it makes life so much easier if you already have a batch of dishes (stuffed with coverslips and Poly-O treated or just stuffed with coverslips) made up.


    Acid cleaned coverslips (back to Table of Contents)

    You probably only need to do this once a semester or two, and usually a lab technician or work study student can make these. Here is the recipe:

    1. Drop coverslips one by one into a beaker of chromic acid. One or two boxes per beaker is enough. Let the coverslips sit over night. NOTE: The chromic acid is extremely caustic; IT BURNS! Wear gloves, goggles and a lab coat to protect yourself. Also, use forceps when moving coverslips that have been exposed to Chromic acid.
    2. Return the chromic acid to its original container. It can be reused many times.
    3. Wash the coverslips for 30 minutes under running double distilled 18 Mohm water (ddH2O). Use a spatula or a pipette to make sure the coverslips are separated. When you are done, there should be no traces of yellow residue on the slips.
    4. You can (for added cleaning) rinse the coverslips three times in a 95% EtOH solution, using a swirling motion.
    5. Using clean forceps, spread out the coverslips on a large (150 mm) Petri dish or on some filter paper to dry in the hood with the blower ON and the UV light. Alternatively, you can autoclave them. After they are completely dry, label the large Petri dish with “Acid Cleaned Coverslips [date].”


    You can now use these whenever you need to make culture dishes! They can be stored on a shelf for weeks; just make sure they are covered and sealed with tape to ensure sterility.

    Sylgard treatment (back to Table of Contents)

    This is the protocol for the movie dish technique. It is very important to mix the agents correctly, or you will end up with glue that won’t dry.

    1. To mix Sylgard, you will combine a 10:1 ratio of silicone elastomer to curing agent. From the large silicone elastomer tub, pour as much Sylgard goo as you will need (if you cut near the base of a plastic pipette, you can use that to take up the amount of Sylgard goo that you want) to put a thin layer on the bottom of each movie dish into a thin plastic weighing pan. After doing this look at your weight before hitting the tare button. Now add to the weighing pan 1/10th as much curing agent. Stir this really well for about two minutes using a disposable plastic pipette.
    2. Paint on a circle of Sylgard (using a small paint brush, a disposable pipette, or your gloved finger) around the holes you have drilled on the bottom side of the 35 mm dishes.
    3. Now use forceps to pick up an acid cleaned coverslip and place it on top of the Sylgard you just applied to the 35 mm dish.
    4. Let the glue on the movie dishes harden overnight so that coverslips will be permanently attached over the holes in your dishes. (The Sylgard directions say you can place the dishes in a warm oven (up to 37° C) to accelerate the curing, but we never tried that, and found that overnight drying, about 24 hours, works best). Note: Any excess Sylgard you have mixed up can be poured into a medium sized Petri dish that other students can use as a pin cushion or to line up tadpoles.


    sylgard

    Polyornithine treatment (back to Table of Contents)

    NOTE: Polyornithine treatment (the “Poly-O” treatment) can be done anytime. It is okay to store sterile, labeled polylysine or polyotnithine treated culture/movie dishes in sterile PBS in the fridge in the tissue culture room for up to two weeks (this is an estimate -- they have been used successfully up to a month after making them).

    1. Turn on the blower in the sterile hood, spray the hood with EtOH, sterilize the hood and tools with UV light for at least 20 minutes. Remember to spray your hands with the 70% EtOH to remain sterile. Follow the instructions from the General Notes on Preparation section.

    2. Place about eight 35 mm culture or movie dishes in a 150 mm petri dish under the hood. Remember that you can make more than just 1 big dish at a time; if you made 4 big dishes of 8 smaller dishes each, you'd already have 32 dishes made up for cultures.

      NOTE: When you are working under the hood try to avoid reaching over and across things (e.g., you might spin the large dish to reach the small dishes that are in the back). Also, try to avoid leaving lids open for longer than necessary (i.e. instead of taking off all the lids of your 35 mm dishes, lift one lid at a time, as needed, just briefly. You can stack the small dishes if that makes it quicker to lift one lid at a time).

    3. Pipette approximately 250 µl (0.25 ml) polylysine or polyornithine (1 mg/ml in PBS) onto the coverslip inside each movie or culture dish. Try to spread the drop out to cover most of the coverslip.

    Poly-O

    1. Cover the big dish and allow the dishes to sit overnight (10-24 hours).

    2. Turn on the blower in the sterile hood, spray the hood with EtOH, sterilize the hood and tools with UV light for at least 20 minutes. Remember to spray your hands with the 70% ethanol to remain sterile. Follow the instructions from the General Notes on Preparation section.

    3. Aspirate off polyornithine or polylysine. To do this you will need a flask (usually plastic) with a tube connecting the hood suction (valve labeled “VAC” in hood) to the side of the flask. Remove the cotton from the top of a sterile plastic 10 ml pipette and push the pipette into the flask through the hole in the rubber stopper. Now connect another tube from the top of this pipette to an autoclaved glass pipette. This dentist-like device will be your suction system; usually there's already one sitting around (see picture below of aspirator/"vacuum").

    4. After sucking off the remains of the polylysine or polyortnithine, each dish needs to be rinsed four times with sterile PBS. Use the Stripettor™ automatic pipetter device and a sterile plastic stripette to fill each dish about ½ full with sterile PBS. Use the suction device to remove the PBS you just added to each dish. (Remember to open only one small dish at a time, briefly, and to rotate the bigger dish so you aren't reaching over anything.) Repeat this rinse three more times, changing the glass pipettes with each round of rinsing. When changing the glass pipettes, do NOT touch the end that is sucking. Use your middle and ring finger to carefully remove the pipette from the container and carefully stick it into the end of the rubber tube in the sucker device. The goal here is to BE STERILE.

      NOTE: At the end of the day (or when you are done with cultures), the vacuum flask should be emptied.

    5. Label the big dish. Close the hood (leave the blower on) and turn on the UV light to sterilize the culture/movie dishes for at least 30 minutes. Please do not look into the UV light; you are smarter than that and enjoy vision.

    vacuum


    HOW TO TREAT THE LAMININ DISHES AND MIX THE CULTURE MEDIA (back to Table of Contents)

    Thaw laminin — mix and filter culture media — apply laminin — cover dishes — allow to set for 2+ hours — aspirate — PBS rinse (x3)

    NOTE: The laminin is expensive, so don't waste it and never put it under or near a UV light. This prep should occur in sterile conditions, e.g. under the hood, with the blower on, a few hours before you are ready to do explants.

    First, get enough microfuge tubes of the 10µg laminin out of the freezer and move it to the fridge to thaw for 30 minutes before using.
    There are two different tubes in the purple plastic box in the small Lom lab freezer, the "7" tube and the "14" tube. Each "7" tube is enough for 4 dishes, and each "14" tube is enough for 8 dishes, so calculate how many tubes you need based on how many dishes you're making.


    Making the Culture Media
    (back to Table of Contents)

    NOTE: Once you make the culture media, it lasts for a while, so you don’t need to make it every time. When you run out, you can make it while you are waiting for the laminin to thaw.

    1. While the laminin is thawing you can prepare your culture media under the hood. You will need a 0.2 micron Nalgene filter, L15 (phenol red is the pH indicator), FBS (fetal bovine serum, orangish-brown), antibiotic/antimycotic solution (in brown glass bottle), EE (embryo extract, pink: see recipe for making this below) and double distilled 18 Mohm water (ddH2O). You can find all of these things in the refrigerator in the tissue culture room or in Dr. Lom’s lab. Ask her if you cannot find something.

      NOTE: The FBS takes a while to thaw, so although you can thaw it that day, it might be beneficial to get it out of the freezer (in Dr. Lom’s lab) and place it in the fridge to thaw over night.

    2. To prepare the culture media combine these ingredients: (making 100 ml is easiest and most of these solutions are already measured out to the correct percentage, so you can just pour them together
    3. in your 0.2 micron Nalgene filter, which has an attachment that can be connected to the aspirator. This speeds the filtration process up).

      60% L15 + 10% FBS + 1% FB + 1%EE + 28% double distilled 18M? filtered water = 100%

      DO NOT poke sterile plastic pipette on the bottom as it will ruin the filter.

    4. Allow media to flow through the filter and then carefully (being sterile), quickly remove the top half of the filter and replace with the cap. Label this bottle of culture media with the name, date, contents, and creator.

     

    Making Embryo Extract (recipe also available in the recipe section, here) (back to Table of Contents)

    NOTE: This is another solution that can be made once and then stored at 4ºC for a few months.
    Use 1 embryo per ml finished media

    1. Select tadpoles around stage ~24-28 and then remove the vitelline envelopes with forceps. Place embryos in a tissue homogenizer and aspirate the extra Steinberg’s off of the top.
    2. Add in one squirt of the following solution: 60% L15 with 10% FBS : 1% FB into the tissue homogenizer (15 ml wheaton) with the embryos (no Holtfaeter’s).
    3. Crush the embryo 50 times (push down slowly, pull up quickly)
    4. Transfer the embryo “mush” into Oakridge centrifuge tube full (or ¾ full) of the 60% L15 with 10% FBS : 1% FB.
    5. Spin 90’ at 15K with a SS-34 rotor.
    6. You will end up with three layers: top liquid, middle liquid, and bottom solid
    7. Carefully pipette out the middle layer and place into a 115 ml filter unit (try to avoid pouring the bottom solid layer in because it can clog the filter). Filter to sterilize.
    8. Add more 60% L15 with 10% FBS : 1% FB to make the total volume equal to the number of embryos used (i.e. 100 embryos = 100 ml).

    Adding laminin to coverslips (back to Table of Contents)

    1. Now that your laminin has thawed you can apply it to coverslips. Mix the 10 µl laminin and sterile PBS in the original laminin microfuge tube.

    2. There are two different tubes in the purple plastic box in the small Lom lab freezer:

      "7" tube: Mix in 903 µl PBS (use Pipetteman) into the tube. Swish and turn tube to mix the solutions.

      "14" tube: Mix 1806 µl PBS (use Pipetteman) into the tube. Swish and turn tube to mix the solutions.

    3. In the same manner as when you applied the Poly-O to the dishes, use a pipetteman to drop 250/300 µl (¼ ml) of laminin onto the center of each clean, dry, sterile coverslip. Be careful to lift their lids only briefly, and to only touch the outsides of the dishes.
    4. Fill a 35 mm dish with sterile double distilled 18 Mohm water (ddH2O) or PBS and put it in the center of the 150 mm dish to prevent dehydration. Alternatively, you can use wet kimpwipes around/under the small culture or movie dishes. Put the lid on the big dish and slide it to the back of the hood to allow the laminin to set for 2-6 hours at room temperature. If your tads aren’t old enough yet, you can let the laminin sit in the dishes for more than 6 hours, but don’t make this a habit, its not the best idea to let the laminin sit for a long period of time, the laminin does have the potential to go bad. You can also cover the laminined dishes and move them out of the hood if you want to UV the hood while the laminin sets up.
    5. After the laminin has set, aspirate off the excess laminin. Use the Stripettor™ automatic pipetter device and a sterile plastic stripette to fill each dish about ½ full with sterile PBS. Use the suction device to remove the PBS you just added to each dish. (Remember to open only one small dish at a time, briefly, and to rotate the bigger dish so you aren't reaching over anything.) Repeat this rinse two more times, changing the glass pipettes with each round of rinsing. When changing the glass pipettes, do NOT touch the end that is sucking. Use your middle and ring finger to carefully remove the pipette from the container and carefully stick it into the end of the rubber tube in the sucker device. The goal here is to BE STERILE. IMPORTANT: Please do not allow laminin treated dishes to dry or come in contact with UV light.

    EMBRYONIC XENOPUS EXPLANT CULTURES (back to Table of Contents)

    Get embryos — sterilize tools — REAG + anesthesia rinse (5x) — remove eye buds — REAG rinse (5x)

    Eye bud removal is very challenging. It’s hard to do at first, and it can be frustrating at times. But once you get good at it, it's fun! Notice that the eye buds are like a ball of cells. They are delicate, so be gentle. You can’t just rip off a big section of the epithelial layer; it makes it hard to find the eye bud. There are two bumps. The eye bud is the smaller bump located closer to the mouth (the only black oval on the tadpole) -- in fact it’s directly over the mouth. You might want to remove more eye buds than actually needed, because 9.9 out of 10 times you WILL lose some in the process of moving them to the culture media dishes. Read below for more in-depth details of removal and transfer.


    1. Select embryos. Choose healthy embryos at or slightly younger than the stage you want to work with (the best ages to work with are stages 25-30. Stage 28 has produced excellent results). Always record the embryo stage, the explant time (beginning and ending), and general notes about the health of the tadpoles. It is important to keep the embryos in clean dishes before culture (you can also use extra antibiotics — talk to the lab technician — in the 20% Steinberg’s) to reduce the possibility of bacteria contamination.
    2. Turn on the gas valve and light the small burner in the hood with a match or flint. Spray 70% EtOH on the tips of all forceps and other tools you intend to use. Briefly insert the tips into the flame and allow the ethanol to burn off and sterilize your tools. BE CAREFUL not to get the EtOH on yourself (it will catch on fire). Don’t stick your hands near the flame, or burn yourself on the warm tools after they have been flamed.
    3. Fill four 35 mm dishes with ~2 ml of sterile reaggregation solution with anesthesia (REAG with 0.05% MS222, labeled REAG w/anest [containing (in mM) 116.6 NaCl, 0.67 KCl, 1.31 MgSO4 (7H2O), 10.0 CaCl2 (dihydrate), 4.6 Tris Base; 0.002% Phenol Red, pH 7.8, 0.05% tricane methanesulfonate]).
    4. Select the PERFECT embryos (most healthy looking, big eye buds). You're going to want about 5-7 eyes for each culture or movie dish you have (so if you're using 8 culture dishes, you want 40-56 eyes, meaning 20-28 embryos). Using the sterile glass fire-polished pipette (already UVed), place embryos in one of the 35 mm dishes of REAG that you just prepared. If you are using embryos younger than stage 27, it is likely that they will still be in a vitelline envelope. Gently pull off each vitelline envelope by pinching both sides of the bubble with your forceps.
    5. After this is completed, use a different glass fire-polished pipette to transfer the embryos to another sterile REAG dish. Rinse a total of 4 times in the 35 mm dishes of REAG you prepared in step 3, using a new pipette for each transfer and swirling gently in each dish. The embryos are the least controlled, sterile part of the experiment, and these rinses provide at least some sterilization for them. Feel free to rinse them as many times as you would like.
    6. The fourth and final dish of REAG is where you'll dissect out the eyebuds of the embryos. Gently "cut" around the eyebud with your forceps and then slowly peel back the top layer of pigmented skin. Finally, lift out the eyebud itself. Place the eyebuds in a small pile in a "corner" of the dish. When working with a lot of embryos, dissect tissue from the older embryos first, thus allowing younger embryos to reach the appropriate stage while you work.
    7. Prepare 4 small dishes of Serum Free Culture Media. Use the 200 µl pipetteman to rinse the eyebuds 4 times, using a different yellow tip each time. NOTE: Any time you are transferring eyebuds, suck up liquid a few times first to wet the inside of your pipette tip. Otherwise, the eyebuds could get stuck inside the tip.
    8. From this step you can either move on to the next section, Creating whole eyebud cultures, or to the subsequent section, Creating dissociated eyebud cultures, depending on which experiment you're doing.

    Creating whole eyebud cultures (back to Table of Contents)

    Apply culture
    media — transfer eye buds — allow to set — gently move from hood

    1. After step 7 in the "EMBRYONIC XENOPUS EXPLANT CULTURES" section, use the plastic 10 ml stripette and the Stripettor™ automatic pipetter device to pipette 3 ml of Serum Free Culture Media (pink liquid) into each laminin/polyornithine treated sterile movie or culture dish. These smaller movie or culture dishes should be sitting in a large petri dish to facilitate moving them later on. Suck up eyebuds a few at a time, again being sure to keep the pipette tip under the liquid. Carefully move about four eyebuds into a movie or culture dish filled with Serum Free Culture Media.
    2. Ideally, let the eye buds sit in the movie or culture dish in the hood for an hour to allow time for the eye buds to settle and stick. If this settling time is not possible because other students need to use the hood, then immediately after pipetting the eyebuds into the movie dish, place them in a safe place outside of the hood where they will remain undisturbed. A lot of times, there will be problems with “floaters.” This is when the eye buds do not stick in place, which means that neurites cannot grow.
    3. Label the large Petri dish with your name, date and DO NOT TOUCH sign.
    4. Give the little eye buds at least 24 hours to attach and produce neurites.

     

     

    Creating dissociated eyebud cultures (back to Table of Contents)

    Move eyebuds to DISAG — transfer buds to disag bubbles — pump up and down to dissociate — gently spray "snow" over culture dish coverslip


    1. After step 7 in the "EMBRYONIC XENOPUS EXPLANT CULTURES" section, take out 2 small (35 mm) petri dishes. Fill one with Disaggregation Solution and in the other, make 10 ul "bubbles" of Serum Free Culture Media (make as many bubbles as you have dishes; for example, the dish below would have enough bubbles for 8 culture dishes).

      dish

    2. You will 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 of the DISAG dish. On the lid above this pile of eyebuds, write "0" and start timer. After ~2 minutes, you'll put the next batch of ~18 eyebuds in a different spot in the DISAG dish and write "2" over their pile, etc. This "round" system will prevent you from getting backed up and being unable to get them out at the correct time (15-25 minutes).
    3. Eyebuds in DISAG will get "fluffy." Hold tips of pulled pipettes next to the eyebuds and pick one to use that is extremely fine -- diameter should be about a 10th of the diameter of the eyebud. Squeeze all the air out of a rubber pipette bulb and put it on the pulled pipette you select.
    4. When eyebuds have been in DISAG long enough (15-25 minutes), use the p200 pipetteman to suck up however many eyebuds you're doing per dish (~6-7). Remember to suck up DISAG to wet the tip first. Put these eyebuds in one of your SFCM "bubbles" made in step 1.
    5. When eyebuds are in the bubble, use the pulled pipette to strain them. Pump them up and down 2-3 times to disintegrate them into a fine speckled snow (not a stringy snot!). KEEP LID ON BUBBLE DISH DURING ANY BREAK OR BUBBLES WILL EVAPORATE, RENDERING IT IMPOSSIBLE TO DO YOUR CULTURES.
    6. Repeat step 5 until all of your bubbles are filled with disintegrated eyebuds. Then use the pulled pipette to suck up one bubble at a time and gently spray it in a zigzag motion over the coverslip in one of your culture dishes filled with serum free culture media. If you see any of the material floating above the coverslip, you can suck it back up and re-spray it.
    7. When all the dishes are finished, you can label them and set them in a calm area.


    Taking movies of eyebud cultures (back to Table of Contents)

    Select eye bud — set up and turn on camera — create folders for images — click Picture frame program — manipulate for good image quality — start filming — create QuickTime movie

    Making movies is something you can easily set up, leave, and come back to later. The important thing is to make sure the cultures have had a good amount of time to set up. This process cannot be rushed. We have found that neurites are growing at their best between 36-78 hours post explant culture. Once you’ve decided how long to wait, you can put the eye buds up on the inverted microscope (the Axiovert 25). Look at all of them and take notes on them. Which ones are producing the most neurites? How many are stuck? How many are floating?

    1. After viewing all cultures, choose one that appears to show some promising growth.
    2. Set objective set to the best view (we almost always used 20x). Remove the left eyepiece from the microscope and there should be a silver eyepiece you can insert in its place. This is where the camera lens will be inserted into the microscope.
    3. To set up the camera, we have found it better to place the top (with the Olympus® Microfire™) of the camera (it’s heavier than the bottom) upside-down. You may think that the image will be filmed upside-down, but, it will actually be viewed normally on the computer. By placing the camera in this fashion, there is less risk of the camera moving your image while you are recording.
    4. Make sure the computer is on and you are logged in. The camera is going to take a series of images, so create a folder on the desktop in which to save them (we usually named the folder with the date and a, b, c, etc. based on how many movies we took that day. For the rest of the protocol, we made up the folder name "12-10-03a," which would be the first movie taken on December 10, 2003). Turn the camera on (the switch is located on the black box). Open the PictureFrame™ program found on your desktop.
    5. The image that your camera is focused on will pop up on the screen (in the window “MicroFIRE™”). Use the fine focus knob to make it clear on the computer screen. You can also change the exposure time, contrast, balance, etc. We found that it is better to increase exposure time and decrease the microscope light on the neurites (reduces condensation in the movie dish).
    6. When you get this image like you want, click on the background program (“PictureFrame™”), and click “File” and pick “Snap direct to disk.” (Or, you can look at the toolbar below that and choose the icon that looks like an arrow pointing into a diskette that performs the same command). Re-click this every time you want to save images to a different folder. This will put a check by that option. A window should pop-up. First select “Choose directory” and click on the folder that you created on the desktop earlier (i.e. 12-10-03a). Change the “File Base Name” to that same folder name also. Next, to the right of “File base name” is a folder called “file type and bit depth”: select JPEG 24-Bit. This ensures that the save images will be saved in this folder.
    7. Now, go back to “MicroFIRE™” and two windows appear: one with your image and another that displays the image contrast. This window is small and rectangular, and is in the upper left corner, there is a big button that looks colorful and a smaller button to the right that has a film-looking picture. Click the small film-looking button (it’s called the “Sequence Snap Mode” button). Immediately, a “Sequence Snaps” window should pop up.
    8. Now you are ready to set up your sequence! On top of the PictureFrame™ screen (background screen) the “Sequence Snap” window should pop up in the left corner. This is where you create your settings for your movies. There are various commands you can do, so here is what each option does:


    START TIME:
    - Select Now: it will start as soon as you click “Start Seq” at the bottom
    - At: this option allows you to set up the exact time you would like the camera to start the sequence
    STOP TIME:
    - After: ## many of shots you would like to take
    - By: Time you want to set it at to finish
    - After: ## of hours you would like it to run for
    INTER-SNAPSHOT DELAY:
    - No delay: it keeps filming
    - For: Chose how many seconds/minutes/hours you want between each image taken.

    After you’ve decided what you want to do, click “Start Seq” and you can walk away! (We would usually chose: Select: Now; After: 3 hours; For: 1 minute).

    Now you can leave for however long your movie is filming for! Woopeeee! But, plan to come back when they are done. They eye buds are sitting in the light for however long your movie is, and the heat of the light isn’t necessarily good for them. Being prompt in your movie making can be the key to getting a ton of data.

    Okay so your movie is done; (wait, you did check the bottom of the sequence screen, if its still says ## shots taken #### seconds left, then you aren’t done!). All of your images should be saved in the designated folder. Now, open the folder that the images were saved in. A bunch of little images will pop up. Just for your sanity, make sure that there are the number of images you asked it to take. (~174 images for our settings)