I'm looking for some help regarding mounting for zebrafish embryos/larvae for live confocal imaging. It seems most protocols in the literature use low MP agarose; however, our lab does not have this. We can make up 3% methylcellulose. I'm struggling to find a sufficient protocol to describe how to conduct live imaging with zebrafish in methylcellulose so that the larvae (from 2-10 dpf) are positioned correctly/remain anesthetized/don't die.
Currently, we have been using silica gel to encase the larvae in water and Tricaine between two coverslips, but it seems that this method does not provide standardized positioning.
Any help/protocol for this would be much appreciated!
1. Using an embryo loop or a small wooden stick [cold, 4 degree Celsius], place a small dab of 3% methylcellulose (see recipe below) in the center of the depression of a glass depression slide. Don't just drop the methylcellulose on, push it down, otherwise it will float away. If it does float away, then just push it back into the depression of the glass depression slide.
2. Dechorionate and/or anesthetize the embryos as necessary as listed in “How to use anesthetics on zebrafish.”
3. Place the embryo on top of the methylcellulose, and layer fish water on top to fill the depression.
4. Move the slide under the stereomicroscope and gently push the embryo into the methylcellulose with an embryo loop until it is secure. Use the loop to position the embryo in the orientation that you want. Be careful to keep the embryo covered with water during your observations or it will dehydrate and die.
Recipe for Methylcellulose
1. For 3% methylcellulose it is equivalent to 3g methylcellulose in 100mls of fish water. 2. Rock overnight in a rocker until the solution is dissolved. 3. Split the solution into 50 ml tubes and freeze the solution until needed.
4. When the solution is needed, thaw for usage (use heat for time purposes to help methylcellulose go into solution).
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