Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We have used two slightly different variants of the single neuron labeling method in our laboratories. The differences relate to the embryo collection, dechorionisation, devitellinisation and embryo filleting steps. Figure 1 gives an overview of the common and diverging steps of the variants.

1. Preparation of Micro-needles for Embryo Dissection and Micropipettes for Dye Injection

1. Glass needles for embryo dissection are drawn from glass capillaries (1 mm diameter and 0.1 mm wall thickness) with a Sutter puller (Science Instruments) or comparable equipment. Alternatively, an electrolytically sharpened 0.15 mm tungsten wire mounted in a needle holder is used for dissections. (Instructions for electrolytic sharpening are given in ⁹).
2. Dye Injection micropipettes are drawn from thin-walled glass capillaries with inner filaments (Science Products; GB 100 TF 8P) with a Sutter puller (Science Instruments) or comparable equipment. The micropipette tip needs to be sharp, with a gradual, uniform taper of the shank to aid passage through the tissues of the embryo. The desired micropipette is the “High Resistance Microelectrode, Sharp and Long” variety, as described on p.20 of the Sutter Instrument Pipette Cookbook manual (http://www.sutter.com/contact/faqs/pipette_cookbook.pdf). Generally, micropipettes are pulled shortly before injections are performed to ensure their tips remain sharp and clean.

2. Collection, Mounting and Dissection of Embryos

1. Embryo Collection. We have successfully used the neuron injection method described here in embryos from stages 12 to 17 ³. Embryos progressively develop a cuticle during stage 17 and become increasingly difficult to dissect. Two alternative approaches are available to obtain embryos at a particular developmental stage.
   1. Allow flies to lay eggs overnight on apple juice-agar plates coated with yeast paste. Adjust the temperature of egg collection to suit the planned time of injection the next day. Collect all eggs the next day and select embryos at the desired stage using morphological criteria ³.
   2. Allow flies to lay on agar plates as above, but swap out the agar plate with a fresh one every 1.5 hr at 25 °C. Incubate each plate for a defined time period, until embryos have reached the developmental stage required.
2. 1. Chemical Dechorionation. Use a metal spatula to scrape embryos off the agar and transfer them into a glass vessel (e.g. Petri dish or cavity block) containing approximately 10 ml Drosophila Ringer or Phosphate buffered saline (PBS) solution. Add a few drops of concentrated bleach and agitate for a few minutes on a rotary platform until the chorion sloughs off the embryos. Wash embryos in several changes of Ringer/PBS until there is no smell of residual bleach. During these washes, ensure that the embryos do not come into contact with the liquid surface. Otherwise, they will float and become difficult to submerge for later manipulation. Alternatively, chemical dechorionation may be carried out using the basket technique described by ⁴.
   2. Mechanical dechorionation (see Figure 2, steps 1-4). Transfer embryos from the agar plate to a slide covered with double-sided sticky tape. Avoid transferring agar with the embryos as it will interfere with their adhesion to the tape. Allow embryos to dry for 5 to 10 min until the chorion gets slightly brittle. (While waiting, coat the coverslip needed for later devitellinisation with glue – see 2.3B below). Touch each embryo gently with a needle. The chorion will split open and the embryo will stick to the needle. Transfer embryos immediately to an agar block to prevent drying and align about 10 of them in a row, with their ventral sides facing up.

Devitellinisation and filleting are done manually using either of the two methods described in 2.3A and 2.3B.

3. 1. Transfer a single dechorionated embryo of the appropriate developmental stage from the dish of Ringer solution to several drops of Ringer in a silicone dam on a pre-prepared microscope slide. (Make a 3 cm square dam by smearing a thin layer of silicone sealant on a microscope slide, then add a drop of 0.01% poly-L-lysine solution to the center of the dam. Allow the silicone to cure for 24 hr.) Transfer the embryo with a glass Pasteur pipette, ensuring that the embryo remains beneath the surface of the Ringer during the transfer.
      Brace the posterior end of the embryo with a pair of fine forceps, while gently squeezing the embryo with a pair of fine iridectomy scissors, about a quarter the way back from its anterior end. Do not cut right through the embryo. The aim is to just crack open the vitelline membrane while causing minimal damage to the embryo. With the forceps still in position, use a tungsten needle to ease the embryo out of the opened membrane and position it ventral side down on the slide. The embryo should stick to the poly-lysine coat.
      Fillet the embryo with a sharpened tungsten needle. Gently perforate, then tear the body wall along the dorsal midline. Using the needle, push down on the body wall so that it sticks to the slide. To prevent damage to the body wall, push the body wall with the gut interposed between it and the needle. Then use the needle to tear through the gut at its anterior and posterior ends and lift it away. If desired, additional embryos may be transferred to the same slide, devitellinised and filleted, taking care not to damage other embryos already on the slide.
   2. Coat a 24 x 60 mm coverslip with heptane glue (see Table 1) by placing a small drop of glue in the center of the slide and spreading it with an 18 x 18 mm coverslip to a very thin film. Place the coverslip on a microscope slide and make a frame of one-sided adhesive tape around its edges. Cut away excess agar from the block holding the row of 10 embryos, gently touch the embryos with the coverslip heptane glue coverslip. They should now adhere to the coverslip with their ventral sides facing down. Add a generous amount of PBS to cover the embryos (see Figure 2, steps 4-6).
      Penetrate the vitelline membrane with a glass or tungsten needle on the dorsal side of each embryo near its posterior end. Tear open the membrane along the dorsal midline and drag the embryo out of the membrane. Orient the embryo ventral side down elsewhere on the heptane glue substrate and fillet it using the same method described in 2.3A) (see Figure 2, steps 7-8). Since several embryos will be filleted on the same slide, it is helpful to either be very accurate with their orientation or make a drawing of their orientation on the slide.
4. After filleting, embryos may be lightly fixed for 10-15 min in 7.4% formaldehyde in PBS, followed by 4 washes in PBS. Extreme care must be taken not to bring the embryo into contact with the surface of the solution during this process, as the surface tension forces will destroy the embryo. This pre-fixation step is not strictly necessary, but is useful to prevent contraction of the body wall muscles in late stage embryos and to help stabilize embryonic tissues at young stages. Keep in mind that fixation does make the tissues of the embryo more opaque and so somewhat compromises image quality under DIC observation.

3. Filling of Injection Micropipettes

Half fill a 0.5 ml Eppendorf microfuge tube with a 0.1% solution of the carbocynanine dye DiI (Molecular Probes, Eugene, OR) in 100% ethanol (be sure to use ‘dry’ absolute EtOH to avoid DiI precipitation). Make a narrow hole in the lid of the tube and insert the blunt end of the micropipette through the hole in the lid. Allow the DiI to ascend up the filament for at least 5 min. (Always cover the hole in the lid when not filling a micropipette to avoid evaporation of ethanol).

Equipment required for neuron dye injection (Figure 3).

Microscope. A fixed stage microscope must be used for neuron dye injection to avoid vertical movement of the embryo during focusing. Any such movement would displace the pipette after it has come into contact with the embryo. We have found both the Zeiss Axioskop FS and the Olympus AX50/BX50 fixed stage models to be well suited for this purpose. The microscope should be equipped with transmitted light DIC optics for visualization of neurons before staining. The microscope should also be an upright rather than an inverted model. With an upright microscope, the tip of the micropipette is on the same side of the embryo as the viewing objective, whereas with an inverted microscope the two are on opposite sides of the embryo. The latter arrangement compromises the quality of DIC optics. The microscope should be set up for fluorescence microscopy, with a suitable filter set for DiI observation. Since DiI has a comparably broad spectrum with excitation and emission maxima at 549 and 565 nm many filter sets in this range will work, e.g. those for Alexa 568, Cy3, Rhodamine, Texas Red or TRITC. Although it is convenient to fit an electronically controlled shutter in the path of the fluorescence light source, to allow operation of the shutter without hand contact with the microscope, we also effectively labeled on a setup which was only equipped with a manual shutter. Finally, a high magnification, high numerical aperture water immersion objective is required for observation during injection. This objective should be designed for use without a coverslip. We have successfully used the Zeiss Achroplan 100x/1.0W, Olympus LUMPlan FL 100x/1.0W, and Olympus LUMPlan FL/IR 60x/0.9 W objectives.

A micromanipulator is required to position and move the micropipette during neuron dye injection. We have used both the Leica micromanipulator and a stage-mounted Narashige 3-axis hydraulic micromanipulator.

An intracellular DC amplifier, with the facility for current injection, is required for iontophoretic injection of DiI from the micropipette.

4. Procedure for Dye Injection

1. Place the slide with the mounted embryo(s) on the stage of the injection microscope (Figure 3). Use a 10x objective to locate an embryo and bring it into the center of the field of view.
2. Swing the high power objective into the viewing position and bring the embryo into focus. Locate the cell of interest under DIC optics (or using fluorescence if the target cell expresses GFP) and bring it into the center of the field of view. Ensure that the field diaphragm of the microscope is opened just to the edge of the field of view, not beyond. A narrow illumination beam will aid positioning of the micropipette (see step 4.4 below). Raise the objective until it is as high as possible above the embryo while still remaining in contact with the Ringer/PBS solution.
3. Place the bath electrode for the DC amplifier into the PBS well clear of the embryo and the path of the micropipette.
4. Insert the injection micropipette into its holder, which has been filled with a 0.1 M LiCl solution. Attach the micropipette holder to the micromanipulator. Using the micromanipulator coarse controls, bring the micropipette into the space between the tip of the objective and the embryo. Ensure that it is well above the level of the embryo. Because of the short working distance of the high power objective, the micropipette will have to be held at a shallow angle in order to bring it into focus (see Figure 3). You will have to establish the appropriate angle by trial and error in the first instance. If the angle is too steep, you will not be able to get the micropipette tip into focus. If it is too shallow, the micropipette shaft will foul against the wall of the dam holding the bathing solution in place.
5. Center the tip of the micropipette in the field of view. To achieve this, bring your eyes to the level of the embryo and move the micropipette back and forth in the Y-axis of the microscope stage. When its tip crosses the light path, you will see the bright reflection of the light rays from it. Move the tip of the micropipette forward in the X-axis so that it overshoots the light beam. It will be easier to locate the micropipette in the small field of view of the high power objective if you are looking for its shaft rather than its tip. Now look through the microscope eyepieces and gradually lower the high power objective until the shaft of the micropipette comes into focus. Moving the micropipette back and forth in the Y-axis with the micromanipulator controls helps to locate it, as it gradually comes into focus. When the shaft is in focus, move the micropipette in the X-axis until its tip lies in the center of the field of view. At this stage the micropipette tip should be well above the level of the embryo. Switch to fluorescence illumination and examine the tip to check for leakage of DiI. Adjust the DC current to prevent any DiI crystal from forming at the tip.
6. Using the micromanipulator controls, lower the micropipette in the Z-axis. Bring it back into focus with the microscope focus control. Progressively lower the micropipette down towards the embryo in this way. Initially, you can do this with the coarse Z-axis controls of the micromanipulator and microscope. However, as the embryo comes into focus you should switch to the fine control knobs.
7. When the surface of the embryo comes into focus, move the tip of the micropipette towards the edge of the field of view, in the direction of the micromanipulator. Focus on the cell to be injected and check that it lies at the center of the field of view. Refocus on the micropipette tip and move it in the Y-axis until it lies at the same level in the Y-axis as the cell. Move the tip of the micropipette towards the cell in the X-axis as you lower it in the Z-axis. If using a Leica micromanipulator, you will probably find that the microscope stage controls give you finer control of movement in the X-axis than the micromanipulator controls, so switch to the stage control as the tip gets close to the cell.
8. Bring the micropipette tip into contact with the cell and make a depression on its surface. Pass several nanoamps of depolarizing current for a few seconds. You should see a small crystal of DiI forming on the cell. To confirm that the cell has been labeled, briefly open the shutter to illuminate the embryo with fluorescent light, then switch back to DIC. If the body of the cell of interest shows signs of labeling, apply current for several more seconds. Turn off the current and quickly pull the embryo away from the micropipette in the X-axis using the microscope stage control. Then withdraw the micropipette from the solution. If no DiI labeling is evident, the micropipette may be blocked and will require replacement with a fresh micropipette. You may find that the tip of the micropipette has snagged other tissues en route to the cell of interest and this tissue is stained rather than the cell. In this case, try withdrawing the micropipette slightly, and re-approach the cell. It may be necessary to replace the micropipette and try again.

9. If desired, multiple cells can be individually labeled in the same embryo.
10. Remove most of the solution in the injection chamber, then fix the embryos by adding 7.4% formaldehyde/PBS for 10 min followed by 4 washes with PBS. The embryo is then left at room temperature for at least 4 hr (or overnight at 4 °C) in a dark, humid chamber (to avoid bleaching or dry falling) to allow the DiI to diffuse throughout neuronal processes. At this stage, the DiI-labeled cells may be either photoconverted or viewed directly in the confocal microscope.

5. Photoconversion for Examination Using DIC Optics

The principle of photoconversion is the (photo-)oxidation of DAB by the fluorescent light emitted by the stained cell during illumination with the appropriate wavelength. This means bright cells are photoconverted in shorter time compared to weakly stained cells. If multiple cells labeled in one specimen differ too much regarding their brightness this can lead to difficulties in photoconverting all of them in same quality (see Figure 4C): in this case one may have to decide on a compromise between neglecting the weaker cells (using short illumination) or accepting that the brighter cells begin to swell (using longer illumination). If all cells are too weakly labeled (thus needing very long illumination), background caused by endogenous peroxidases can become a problem. Since DAB is toxic it should be handled with care. Waste can be ‘deactivated’ with 7% chlorine bleaching.

1. Photoconversion must be carried out for each embryo individually. In cases where only one embryo is injected per slide, the PBS solution covering the embryo is replaced with a DAB solution (3 mg DAB/ml Tris buffer), the slide placed on a fluorescence microscope and the filled cell viewed with a 100x objective. (Using an upright microscope the objective dips into the DAB solution and should be cleaned several times with water after the photoconversion procedure is finished; this can be avoided if an inverted microscope is used). A Cy3 filter set and a 100W Hg lamp are used and the cell exposed to the red excitation wavelengths until a brown DAB reaction product becomes evident in the labeled neuron (check periodically by switching to bright field illumination). The time taken for optimal DAB staining is variable, but requires exposure to the excitation light beyond the stage where the fluorescence has completely faded. After photoconversion is complete, wash the preparation several times with PBS. Replace the PBS with a drop of 70% glycerol, scrape away the silicone with a scalpel blade, replace with a ring of Vaseline and add a coverslip.
2. When multiple embryos have been filled on the one slide, transfer each embryo to a small drop of PBS on a separate 24 x 60 mm coverslip with the ventral side of the embryo facing the glass. Place a frame of adhesive tape around each embryo to create a well and cover the preparation with PBS. Keep the slides in a moist chamber before photoconversion to prevent them from drying out. Exchange the PBS covering the embryo with DAB solution. Immediately place the slide onto an inverted microscope equipped with a 100W Hg lamp and use a Cy3 filter set to excite the DiI, using a x 50 objective. After photoconversion, transfer the embryo to a fresh slide in a drop of 70% glycerol, add a coverslip and seal with nail polish.

6. Examination by Confocal Microscopy

1. After step 4.10, transfer embryos to a small drop of PBS on a fresh 24 x 60 mm coverslip. We achieve optimal optics by mounting the flattened embryos with the dorsal side towards the coverslip (leaving only a small amount of PBS between the coverslip and embryo).
2. Place a frame of adhesive tape around the embryo and enlarge the PBS drop to fill the frame when covered with a slide. Place a slide on it carefully and fix it with nail polish. Filled neurons should be examined on a confocal (or fluorescence) microscope as soon as possible.