1. You don't need alot of embryos. Assemble a cage and let it mature for at least 2 days before collecting embryos for injection.

2. Collection Schedule is temperature dependent, obviously.

For me the following schedule is one I like to follow:

Change plates at 2:00 P.M.

Change plates again at 6:00 P.M.

Change plates again, so now you've got the earlier collection ready to begin working with and a new collection that you can age while you work on the first one.

Injections take about 2 hours (for about 100 embryos). This means you'll be done around midnite. (Work at 16 degrees. Wear warm sweaters!) If it's impossible to work at the colder temperature, you can get good results at the warmer temperatures as well, but your viability will decline and also, the timing is more difficult. I store the embryos at 16 degrees overnight.

Eleven hours after finishing your injections, your embryos will be at mid-stage 11. At this point you identify NBs, once again, in the dark and in the cold!

24-27 hours after identifying NBs (i.e., about 36 hours after injections, at 16 degrees), your embryos are at stage 17 and ready to dissect, fix and confocal.

1. Prior to injection, set up the following:

Glue-covered cover slips (see below).

Halocarbon oil 600 cStokes (see below) and transfer pipette

Large (150 mm diameter) agar plates containing apple juice agar.

eye-lash brush

double-sided scotch tape

tungsten needle for dechorionating (see below)

paintbrushes

dessication chamber (see below)

2. At your dissecting microscope, set up two glass microscope slides, one with a 1-inch by 0.5-inch rectangular plug of apple juice agar on it (you've sliced this plug out of the middle of one of your 150 mm plates) and the other with a piece of double-sided scotch tape on it, about 3/4 the length of the slide (i.e., about 2 inches).

Using the paint brush, carefully and gently transfer a swath of embryos on to the tape. They should be a monolayer on the tape---not clumped into bunches. Using your tungsten needle, gently dechorionate embryos. I like to work with my fiber-optic tilted so that illumination is coming from the side. Embryos that are clearly at cellularization (stage 7) are scooped up onto the tungsten needle and gently transferred to the agar plug. The agar plug is sufficiently hydrated to keep embryos on it from dessicating. This allows uniform dessication later.

3. Line the embryos up head to tail, in a column on your plug. It is crucial that they be lined up in as straight a line as possible. When learning to do this, I found that making a slight indentation in the agar plug with a razor blade provided a nice template into which to place the embryos for a really straight line. The ventral curved side should face you, the dorsal concave side should be on the agar.

4. Pick a rubber-cement coated cover-slip up and with the sticky side down; gently pick up the embryos by touching the sticky side to the embryos. Sometimes, the plug is too wide and makes it difficult to get close to the embryos, so I usually trim it to a smaller rectangle before picking the embryos up. I immediately place the embryos into a dessication chamber and begin timing. How long to dessicate is going to be a purely empirical thing. The ideal way to determine this is to touch the embryos after dessication with an eye-lash brush. If they deflect with the consistency of a raw bratwurst (sorry---it's the best analogy I can think of), that's perfect. At approximately 60% room humidity (which is approximately the natural humidity in a 16 degree room), I find that 5 minutes in a dessication chamber is perfect. When the room humidity goes up to 70% (which requires a humidifier), the dessication time goes up to 14 minutes! The dessication is extremely important. Too dry and the embryos won't develop. Too wet and they'll explode if you poke them. Generally, though, it's better to err a little on the side of too wet, than too dry.

5. After dessicating, remove the cover slip and look at them under a dissecting scope. Use your eye-lash brush to gently push them down onto the rubber cement. This dramatically increases the size of the "footprint", the area you will be injecting into. As soon as you've pushed them all down, cover them with a single line of 600 cSoke halocarbon oil and set them aside.
6. I have measured the effectiveness of halocarbon oils mixed to produce various densities. The manufacturers of halocarbon oil are happy to advise you on how to achieve accurate halocarbon oil densities by mixing heavy (700 cStoke oil) and light (27 cStoke oil) oils. Halocarbon oil allows oxygen to permeate through to the embryo, but keeps water from evaporating out. So the optimum oil density allows the you to achieve the maximum oxygen in while minimizing the water out. I found that this happened with 600 cStoke oil, and that mixture is achieved by 95% (by volume) heavy oil, 5% by volume light.

I generally dissect 6 groups of 15-20 embryos (keeping track of phenotypes on the agar plugs......make sure you cut off a corner of the plug and make an accurate note to yourself as to which embryos are where on the plug!), and then line all six groups up. Then rapidly pick them up, beginning the dessication timing with the deposit of the first coverslip into the dessication chamber. I take the first slide out after 5 minutes, push embryos down with my eyelash brush (GENTLY!!!!), cover them with oil and move on to the next slide.....until all 6 slides are done. In this way, I can plan to inject about 120 embryos/injection session.

As soon as I've finished the first group, I can begin on the second set of embryos that have now aged appropriately. I can comfortably inject 6 coverslips in about an hour. I can rush myself and do 8 cover slips, but I prefer simply to do the 6 and then have another set of embryos to begin again. Rushing is never good when microinjecting.

Figure 1: How I cut out agar plugs and then use the de-plugged plates as storage chambers for injected embryos.

6. Injection techniques also vary from one person to the next. Generally, I position the cover-slip onto the stage with blobs of clay. I have adapted the stage using time tape--(see diagram) and attach the cover slip to lab (time) tape using children's modeling clay. See Figure 2.

Figure 2: How the microscope stage is set up for DiI injections.

7. I position the needle to the appropriate height at low magnification. I locate the top embryo in the line, and then switch to 100X magnification (using high density (type B) immersion oil). I lower the condenser lens carefully. It is crucial that you have positioned the needle correctly, because the working distance is small---it is terrible to have gotten to this point, only to break your needle by lowering the condenser lens!

Figure 3: The injection procedure:

8. The footprint should be obvious to you if the embryo is properly dessicated. See Figure 3. The big heavy line to the left of the purple line (the edge of the embryo) and to the right of the ventral furrow is the "footprint". If the slides are old (i.e., were coated with the rubber cement mixture more than 2 days ago) or if the embryos are not properly dessicated, there will be no footprint. Without a footprint, embryos can still be injected, but it's much more difficult to do.

Injection should be easy. Push the needle gently into the peri-vitelline space; there should be something of the sensation of popping in. Squeeze out a small drop. It is here that the the quality of the needle comes in to play. If your needle's tip is too large, there will be great resistence to the initial outflow of DiI and when the DiI finally exits the tip of the needle, the drop will be huge. The needle should look beveled to you under the mricoscope--i.e., the tip should look diamond-shaped. And it should be approximately 1/5 the diameter of a neurectodermal cell. When squeezing the drop of DiI out, it should flow nicely and you ought to be able to control the size of the drop----you should have no trouble leaving a too-small sized drop.

It is crucial that you do not actually touch the surface of these neurectodermal cells---they are fragile and will die if you touch them. The idea here is to squeeze the droplet onto their surface from the peri-vitelline space, without ever actually touching the cells. For this reason it is to your advantage to place the embryo on the coverslip, so that the footprint falls at the spot you want labeled. Thus if you are planning to label medially, the embryo should be tilted when you place it on the coverslip so that the footprint falls close to the midline. Conversely, when you want to label laterally, the embryo must be tilted in the opposite direction.

It is also to your advantage to label neurectodermal cells four rows apart. This corresponds to the antero-posterior length of one segment---frequently you will find that you are labeling the identical clones only one segment apart. This is extremely convenient when using paired-GAL4 drivers, as one segment will express paired early on and the next won't express it until much later----this allows you to examine within a single embryo, the differences in affected vs. relatively unaffected segments. If you are injecting any segmental background (such as en-GFP), this is also convenient, in that it allows you to examine intersegmental fibers more accuartely and whether or not identical clones connect to one another, from one segment to the next. And finally, it gives a better idea of motoneuronal arrangement, if for example, 3-2 clones are labeled in adjacent segments, one sees that SNa anteriorly is tightly fasciculated with ISN fibers from homologous lineage in the adjacent segment.

I generally try to inject a single embryo at least four times, but obviously, you can label as many as 8 or ten times in a single embryo at stage 8.

When finished, remove the cover-slip to a prepared apple-juice agar plate with a moistened kimwipe in the middle (see fig 1). This leaves the immersion oil intacton the under-side of the coverslip (so you can use it again the next day) and in general disturbs everything to a minimal degree while also keeping the chamber optimally hydrated. It is not a bad idea to put a little more halocarbon oil over your embryos as you put them away for the night.

We have had Chroma Technologies design a filter set for our UV lights that filter out a maximum of the very strong DiI fluorescence while allowing optimal GFP illumination. It has a very narrow band-pass, so the illumination is clear and specific, but very dim. The dimness is compounded by high index neutral density filters. We therefore us a CCD camera to amplify the low-light optics and find this system allows us to use Mercury lamp illumination on our microscope. The Technau lab has used a different illumination system to conquer the problem of DiI associated phototoxicity.

We generally find the oil drop using transmitted light optics and very low levels of light. We then briefly illumine the droplet with the UV light. If a NB appears to be labeled, we circle it on the screen of our TV monitor with a water marker and also mark the edges of the engrailed stripe, (which demarcates rows 6 and 7), and then immediately switch the UV irradiation off. We then return to low level transmitted light optics to definitely identify the NB.

In general, assigning an identification to the NB cannot be described----it requires having someone like Chris Doe teach you in person. There are several morphological landmarks that can be used early on to aid in NB identification although nothing beats an in vivo marking system, such as the en-GFP system we used in this study. In general, the ventral furrow and the segmental and parasegmental grooves are the most obvious things to observe. These are the landmarks that the Technau lab studies used to make their NB identifications. In addition, several NBs are easy to see and can be used to identify the other less obvious ones. Examples include NB 6-1, 7-1 and 1-2 which line up in a nice diagnonal row; the row 5 NBs which are just above the parasegmental groove, the guitar-pick shaped NB 3-1. Once you can be confident of these the rest become easier to score.

It is essential that you not only find a way to keep track of which embryo on your converslip has which NB labeled in it, but which of the many labelings you''ve done in a single embryo correspond to what you are identifying the NB as. Thus, I have developed a worksheet that works well for me----with a map of the embryo on it, and 6 lines for each embryo, on which I note whether the injection/clone has died at the stage of identification, or whether it has formed an epidermal clone (which is the most frequent case) or whether a NB has been labeled and if so, which one.

Develop a short-hand! The embryo will be at stage 11 for only a very short time, even at 16 degrees, so you have to be able to do this all very very quickly.

37 hours AEL (at 16 degrees C), the embryos should be entering mid-stage 17, and this is the last stage at which you will be able to do simple filleted dissections.

~20 mls. (methanol-free) paraformaldehyde in PBS, pH=7.6

PBS

capillary pipettes pulled for DiI labeling, but unbeveled.

razor blades

diamond pencil

Kimwipes

welled slides (see below).

Check cover-sleips on a fluorescent scope (I use the same one I inject on) and note (preferably on the same worksheets that have NB identifications) which embryos have produced useable clones. Make note of cover-slip number genotype. You will be cutting that information off of the coverslip, so it is crucial that you have that written down somewhere.

On a kimwipe, use a razor blade to wipe off as much of the immersion oil as possible. I get right in close. After removing as much oil as possible this way, tear a Kimwipe and use the long sides to blot off as much of the remaining oil as possible. I routinely spend as much as 5 minutes on a cover slip blotting off oil. The embryos should be visible under the dissecting scope as though there were almost no oil on them at all. Use the diamond pen to cut the sides off of the cover-slip. Score a line and pull apart the cover slip. Be very careful when doing this---you do not want to crack the coverslip at a point where the embryos are. The cover-slip fragment should be about the size as shown in Figure 4.

Slip the edges of this trimmed cover slip under the silicon edges around the superfrost plus slide you have made a well on. Cover with PBS or insect saline. Using your notes to remind you, remove the embryos with clones in them from their vitelline membranes and move them over to the glass surface in the center of your slide. At mis stage 17, there is only one tiny little spot left on their ventral surfaces that will still stick to the glass. Push this spot down, FIRMLY. By this stage, embryos should be fairly robust. If not, maybe because their mutant phenotypes include general sickliness, you uually don't have a problem getting them to stick to the glass---their cuticles are usually not intact. Holding the embryo down at that spot with a tungsten needle in your left hand, use a BEVELED capillary pipette to slice the embryo open along its dorsal surface. This may take time and patience! If it is especially late nad mature, the cuticle will always want to slam back shut. But once open, the body wall can be attached to the glass jsut by repeatedly "smearing it out."

I generally apply a brief light fix to the well and then confocal the clone.