This material is from the 4th edition of The Zebrafish Book. The 5th edition is available in print and within the ZFIN Protocol Wiki.


Labeling Single Cells With Lineage Tracers

(Source: B. Melby and D. Raible)

This is a technique for intracellular labeling of small cells at gastrula stages or later.


The electronics set-up is used as a means to get the dye out of the electrode and into the cell. Use an agar bridge (see Agar bridged ground wires) for the bath ground.


Success in single cell labeling lies primarily with the ability to make good pipettes. Pipettes should be pulled the day of the experiment. Allow at least 10 min for the tips to fill with dye. Electrodes are pulled from glass capillaries (e.g. Sutter thick wall, O.D. 1.2 mm, I.D. .69 mm) with an inner filament for easy dye filling.

The ideal pipette tip can be attained only by fine-tuning and testing. The pipettes must be sufficiently sharp to pierce through the outer layer of the embryo, yet have a large enough tip diameter to pass dye. They should have a long, sharp shank but a blunt tip. An important feature of the pipettes is that they pass dye, either by current injection or by "ringing" the capacitance, and that they do not continuously leak dye. The resistance of the pipettes provides a measure of reproducibility from one experiment to another. Good pipettes usually have a resistance in the range of 100-300 M[[Omega]] when filled with 0.2 M KCl. If the resistance is too high, it may indicate a problem with the dye, or with the bath ground, as well as with the pipette shape.

The dye should be dissolved in 0.2 M KCl and filtered (e.g. 0.2 µm spin filters for the microcentrifuge). A 3% solution of rhodamine dextran (Molecular Probes) works quite well. Dye can be made in advance and stored at 4°C in a sealed tube. To fill pipette tips with dye, place a droplet of dye (about 0.5 µl) at the back of the pipette.

Labeling gastrula stage cells:

Embryo preparation and mounting:

Embryos should be removed from their chorions in agar-coated dishes using fine forceps. After removing the chorions, transfer the embryos with a fire-polished pipet. Experimental slides are made by painting several coats of nail polish in a ring of about 3 cm diameter, on a glass slide. Mount embryos in methyl cellulose (see Methyl Cellulose Mounting, Chapter 4) to permit easy manipulation and reorientation. Chilled methyl cellulose is stiffer, but tends to get bubbly as it warms up, therefore it is best to use methyl cellulose at room temperature.

1. Spread a layer of methyl cellulose in the center of the experimental slide.

2. Cover with embryo medium, being careful not to spill over the nail polish ring.

3. Transfer an embryo to the center of the slide. Orient the embryo under the dissecting microscope by gently nudging it with a hair loop.

4. When the embryo is in the preferred orientation, gently tamp it down into the methyl cellulose. Over time, the methyl cellulose will become diluted with embryo medium and will become too fluid, so that the embryo may roll. If this happens, remove the embryo and remount.

5. Be sure to loosen the embryo from the methyl cellulose before trying to move it with a pipette.

Labeling cells:

It is easiest to label cells if the embryo is oriented such that the surface to be labeled is at the top, lying flat. The ideal situation is to have the embryo oriented so that the cells to be labeled are easily visible in reference to landmarks such as the embryonic shield or the margin.

1. Check the orientation of the embryo at low and then high magnification, to be sure that it is appropriate.

2. Place a hanging droplet of embryo medium on the water immersion lens (e.g. 40x) and gently slide the lens into place over the embryo, to avoid rolling the embryo.

3. Position the bath ground at the back of the slide, being sure that there is continuity between the agar and the embryo medium.

4. Raise the microscope objective so that there is plenty of space between the objective and the embryo.

5. Backfill a pipette with 0.5 M KCl, place it in the pipette holder, and attach the holder to the amplifier probe which should be mounted on the micromanipulator. There should be no large bubbles in the pipette or holder; they will cause electrical discontinuity.

6. Advance the pipette into the embryo medium and position it above the embryo using the white light and by viewing it from the side of the microscope.

7. Locate the tip under magnification, and check to make sure that it isn't broken.

8. Switch on the amplifier, this should only be turned on when there is a complete circuit (i.e. from probe, through the bath and bath ground, back to the amplifier).

9. A good pipette will release a small puff of dye when the capacity compensation (usually a button, sometimes a dial) is turned on (i.e. "ringing the capacitance"), or when the pipette check is pressed (this injects a small amount of current). A bad pipette will leak, or sometimes not release dye at all. Throw it away and get a new one, but be sure to turn off the amplifier before you pull the pipette out of the bath.

10. Carefully lower the pipette to the level of the embryo, keeping it in focus. Superficial cells can be labeled by positioning the pipette above the cell and lowering the tip through the EVL layer until the cell membrane dimples.

11. Monitor the fluorescence, and label the cell by ringing the capacitance, or by injecting current. Ringing the capacitance causes the pipette tip to vibrate, enabling penetration and it will also cause a little bit of dye to be released.

12. After a cell has been labeled, rapidly remove the pipette using the stage control or the micromanipulator, and turn off the amplifier.

13. To label cells in deeper layers, it is best to bring the pipette in from the side. Focus on the cell layer of interest. Bring the pipette into the same focal plane as this cell layer. Advance the pipette up to the cell using a microdrive, or fine micromanipulator adjustment.

Sometimes cells along the pipette track tend to be labeled by dye leaking from the tip. The tip may appear to have passed through these cells, but there is still continuity with their membranes. Single cell labels can be achieved by stopping dye injection as soon as one cell starts to label. Because of this problem, it is important to check the fluorescence without Nomarski optics (it will be much brighter) in order to see all of the cells that have been labeled.

Localizing the labeled cell:

For mapping purposes, it is best to check the cell's location using three different orientations of the embryo. In the orientation in which the embryo was labeled, check the cell's distance from the margin under the 40x water immersion lens. Then turn the embryo onto it's side and check the cell's radial depth at 40x. It is important to have a fairly precise side orientation, or else the cell may appear deeper or shallower than it is. The depth can also be assessed by focussing up and down in the original orientation, although this method gives no information about the relationship of the cell to the hypoblast. Finally, assess the position with respect to the dorsal midline by orienting the embryo animal pole up and looking at it under low power (10x objective).

Labeling single cells in older embryos

1. Mount embryos in agar as described in the section on Agar Mounting (Chapter 4). Orient the embryo so that the cell of interest is visible with Nomarski optics and accessible with the pipette.

2. Focus on the cell of interest, and bring the pipette along side the embryo in the same focal plane.

3. Using the stage controls, force the mounted embryo against the pipette until the skin springs back and the pipette tip enters the embryo.

4. Move the embryo away with the stage controls until there is no compression of the embryo by the pipette.

5. Then, using the fine control of the micromanipulator, move the pipette into position against the cell to be labeled. For cells close to the surface, the pipette can be inserted into the embryo at a different focal plane.

6. Label the cell as described above, and quickly remove the pipette from the embryo by moving the embryo away with the stage controls.


Labeling rig:

  • vibration-free table
  • oscilloscope
  • micromanipulator
  • stimulator (optional)
  • amplifier
  • bath ground
  • probe
  • microscope with fluorescence light source
  • electrode holder
  • 10x and 40x water immersion objectives
  • Pipettes:

  • Sutter thick wall glass with capillary
  • electrode puller
  • dye (fluorescent conjugated dextran)
  • electrode holder
  • Hamilton syringe needles
  • 0.2 and 0.5 M KCl solutions
  • Miscellaneous:

  • squirt bottle of embryo medium
  • squirt bottle of distilled water for rinsing
  • Gastrulae mounting:

  • embryo medium
  • 3% methyl cellulose (in embryo medium)
  • agar-coated dishes
  • experimental slides
  • fire polished pipets
  • dissecting microscope
  • hair loop
  • forceps
  • Older embryos:

  • 1.2% agar in Ringer's
  • water bath at 42-44°C
  • experimental slides
  • fine probes

  • The Zebrafish Book