Labeling single cells in ovo and then studying their fate during subsequent development can be a powerful tool in the investigation of how embryos develop. It can be used to construct fate maps at the single-cell level, analyze morphogenetic movements, address issues of tissue specification, and assess the importance of lineage in the determination of regional or cellular identity. One way of labeling single cells is by infection with replication-deficient retroviral vectors. This method allows cells and their descendants to be permanently labeled with a genetic tag, thus allowing for the long-term analysis of clonally related cells. A second method is to inject single cells with tracer dyes using intracellular microelectrodes. The advantages of using the single-cell injection technique are that you can accurately target particular areas of the embryo, there are no doubts about clonality because each injection can be checked visually the time, and by using fluorescent tracers, the sequential analysis of the same clone at different times of development becomes possible. The disadvantage of the single-cell injection technique is that the marker is diluted by cell division and increased cell volume. This effectively limits its usefulness to between about 6 and 9 rounds of cell division, depending on the quality of the initial injection.
Labeling single cells in ovo by intracellular injection is not the sort of technique that can easily be learned by simply reading about it. It is much better to see the technique in action. This account will assume a basic understanding of intracellular recording technology and electrophysiological technique. If you do not have this, then make friends with a pharmacologist or neurophysiologist.
1. Make microelectrodes—we use 1.2-mm diameter, thin-walled aluminosilicate glass with internal filament (A-M Systems, Everett, WA). Electrode tip should have a nice constant taper and be fine but not wispy. When back-filled with dextran and 1 M potassium chloride, they should have a resistance of between 50 and 150 MQ. In practice, the precise electrode resistance does not matter; if the electrode penetrates a cell, records a stable membrane potential, and passes sufficient dye, then it is a good electrode (see Notes 7 and 8). Electrodes should first be back-filled with approx 0.5 |L of fluorescent dextran (100 mg/mL in distilled water, Molecular Probes cat. no. D-3308 and D-3306 for tetramethylrhodamine and fluorescein fluorescence, respectively) and then with a little 1 M potassium chloride. Don't worry about air bubbles. The capillary action of the internal filament will deal with them.
2. Window egg, inject ink subblastodermally, make a small hole in vitelline membrane over target area, and carefully drop a little saline solution onto the embryo to decrease the risk of drying out and to improve visibility. Do all this under a stereomicroscope.
3. Transfer egg to platform on fixed-stage injection microscope and insert silver/ silver chloride reference electrode into albumin via air hole in shell.
4. Illuminate embryo with fiber-optic directed into the windowed egg. Find and focus embryo under x20 objective.
5. Manipulate microelectrode tip down into the saline covering the embryo.
6. Switch on amplifier, and check that you are recording a stable baseline potential. You need to work with a "gain" of 10 mV/division and a time base set to 1 s/division. The oscilloscope must be set to monitor DC potentials.
7. Gently manipulate electrode tip down onto the surface of the cells (see Notes 9 and 10). You probably will not be able to see individual cells. As the electrode tip touches a cell membrane, there will be small change in the appearance of the trace on the oscilloscope. This may be an increase in the thickness of the trace or more usually a small positive deflection. This change is the best indication that the electrode has just touched a cell membrane; it is better than trying to see it happen down the microscope.
8. "Ring," "buzz," or "zap" the capacitance-compensation button or knob on the amplifier to penetrate the cell membrane. If successful, a small but stable negative deflection should be seen on the oscilloscope (anywhere between -5 and -30 mV is common for neural plate cells). This is the most efficient method of getting the electrode tip inside a cell, but it does have one big disadvantage. When you "ring," "buzz," or "zap" an electrode, you simultaneously squirt dye out of the end of the electrode. With a successful penetration, most if not all of this dye will go straight into the cell to be labeled. However, quite often some dye will also be sprayed out onto adjacent cells, and if these have been damaged (by the electrode scraping past them for instance), they will often take up significant amounts of the dye. If this happens, this is the end of your lineage analysis, because you will have labeled more than one cell. It is therefore essential to check briefly each injection visually with epifluorescence in order to ensure only one cell has been labeled. Do not admire your cell for too long, since phototoxicity can kill it. Cell penetration in the absence of simultaneous dye injection can be effected with a piezo-electrode stepper device, but these are not as efficient at getting into small cells as "ringing." No matter how you penetrate your cells, you should then iontophorese more dye into them by using the amplifier's current injection facility (for lysinated rhodamine dextran use positive current pulses of about 4 nA and 250-ms duration for about 30 s).
9. Rapidly withdraw the microelectrode using the axial drive of the manipulator. The oscilloscope trace should spring back up to its original baseline level.
10. Carefully drop a little more saline onto the embryo, and then reseal the egg with tape and reincubate to the appropriate stage.
11. Fix embryo in a solution containing 3.5% paraformaldehyde.
12. Observe results in either sectioned material or whole mounts using conventional epifluorescence or confocal microscopy.
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