Fixing and Staining L1 LBs for IFM
1. Collect embryos on standard fly food by transferring flies to fresh vials after adding 5-10 granules of live yeast to the food; place vials in the dark to encourage egg-laying. Allow embryos to hatch (~20-22 hr at 22ºC), then maintain ~6-8 hrs to mid-L1 stage. Alternatively, transfer embryos collected on standard fly food to amino acid-depleted agar plates to enrich LBs for qNSCs; wash embryos from food using several changes of DDW, allow embryos to hatch and maintain ~20-24 hrs to mid-L1 stage, as larvae develop more slowly on amino acid-depleted plates.
2. Note the characteristic shape of the brain at the anterior end of the larva (Fig. 1a). Transfer 10-15 L1 larvae into DDW in one well of a two-well dissecting slide. Pipette 25-30 µL of PBS into the other well.
3. Put on appropriate personal protective attire, including a high filtration face mask, safety glasses, gloves and a laboratory coat. Wear until after completing washes following fixation.
4. Prepare a 200 µL PCR tube by firmly affixing it cap-side down onto a small piece of double-sided tape on a microscope slide (Fig. 1b). Use the flat (handle) end of a pair of forceps to push down on the hinge and cap edge to ensure that the cap is firmly affixed to the slide. Use the flat end of the forceps to hold the cap down when opening the tube to ensure that it does not detach from the slide.
5. Place 20 µL of freshly made fixative in the PCR tube cap; close the tube loosely after transferring each LB into the fixative during dissection.
6. Transfer the L1 larvae from the well containing DDW into the well containing PBS using a plastic Pasteur pipette.
7. Dissect each larva by holding it halfway down its length using one pair of fine-tip forceps, then grasp the side of the larva anterior tip with the other pair of forceps and pull the anterior end open. Quickly locate the brain and pull off a small piece of the cuticle with the attached brain, including the mouth hooks, and transfer the tissue into the fixative in the PCR tube cap.
8. Repeat with 10-15 larvae, keeping track of time and stopping dissections after ~5-6 min. Close the PCR tube and continue fixing the LBs for 20-25 min.
9. Using a finely drawn out glass microliter pipette and an improvised pipette bulb (Fig. 1c), remove the fixative from the LBs and discard into a waste receptacle; dispose according to institutional laboratory safety guidelines. Replace the fixative with 20 µL of PBST to wash the fixed LBs; repeat the PBST wash. Place the slide with the PCR tube on a platform rotator for 10 min, then change the PBST and rotate another 10 min.
NB: Glass pipettes are drawn out by holding the pipette at both ends and rotating the pipette while heating in a low flame until soft, then quickly removing from the flame and pulling sharply; excess length is broken off using a pair of blunt-ended forceps to pull on the drawn-out region near the position desired for breaking. Each microliter pipette or capillary tube makes two drawn-out pipettes.
10. After removing the final PBST wash, add 20 µL of blocking buffer to the LBs in the PCR tube cap and place the PCR tube affixed to the microscope slide on a platform rotator for 1-2 hr at RT. After blocking, the LBs can be stained with antibodies or stored in blocking buffer at 4ºC for several weeks.
11. For antibody staining, use a P20 Pipetman with cut-off yellow pipette tip to transfer 3-7 LBs to a new PCR tube cap affixed with double-sided tape to a microscope slide. Replace the blocking buffer with 12-15 µL of primary antibody diluted in blocking buffer. Close the PCR tube tightly and transfer the tube with microscope slide to a platform rotator for several hours to overnight at RT.
12. After incubation, use a drawn-out glass pipette to remove and save the antibody solution for reuse. Primary or secondary antibody solutions can be reused 2-8 times – the reused solutions can show significantly lower background than freshly diluted antibody solutions. Add 20 µL of PBST to the LBs and wash 3-4 hrs with rotation at RT, changing the PBST wash solution 3-4 times during the wash time.
13. React LBs with 12-15 µL of secondary antibody diluted in blocking buffer and rotate for 3-4 hrs. Protect from light as necessary, as secondary antibodies are typically conjugated to a fluorophore. Recover the antibody for reuse, then wash LBs as in step 12.
14. Remove final PBST wash using a drawn-out glass pipette, then add 10-15 µL of 50% glycerol in PBS to the LBs in the PCR tube cap. Use a P20 Pipetman and cut-off yellow tip to transfer the LBs to a two-well slide for final dissections. Place a 5 µL drop of 90% glycerol in PBS in the other well.
15. Prepare a clean microscope slide by positioning double-sided tape spacers and vacuum grease supports as shown in Fig. 1d. Place a 5 µL drop of 90% glycerol in PBS in the center.
16. For final dissections, use a fine-tipped pair of forceps to gently hold each LB and remove the attached cuticle and mouth hooks with a scalpel made from a 26 G hypodermic needle fit on a 1 mL hypodermic syringe. The attached cuticle, mouth hooks and other tissue should be cleanly dissected away from the LB. Transfer each LB to the 90% glycerol in PBS in the second well of the dissecting slide using a P20 Pipetman fit with a 10 µL pipette tip and the Pipetman set to 2-3 µL. The LBs will be translucent, but can be visualized by careful observation under incident illumination. After completing the final dissections, transfer the LBs to the drop of 90% glycerol in PBS on the microscope slide using a P20 Pipetman and 10 µL pipette tip. Orient the LBs by gently moving the solution adjacent to the LBs with the tips of a pair of forceps to position as desired.
Note that the final dissections can also be performed by placing 3-5 µL of 50% glycerol on a microscope slide, cleanly dissecting the LBs and transferring the LBs to an adjacent 3-5 µL drop of 90% glycerol, then removing the 50% glycerol and positioning the spacers around the LBs in the drop of 90% glycerol.
17. Carefully place a clean coverslip onto the LBs in the 90% glycerol in PBS, tilting the coverslip to avoid trapping air bubbles. Note that other mounting media can be used instead of 90% glycerol. The 90% glycerol suppresses photobleaching, but does not prevent it. DAPI or other DNA dyes can be added to the 90% glycerol mounting medium (or to the secondary antibody washes) to stain nuclear DNA.
18. The LBs mounted on the slide can then be observed by confocal microscopy using settings appropriate for secondary antibody detection and imaging.
Modifications for Live Imaging
1. 3D printed slides are created on an Ultimaker 2+ 3D printer with PLA filament, printing at 200°C extruder temperature, 70°C bed temperature, 5% infill, 0.8 mm wall thickness, 0.2 mm layer height. Each slide requires ~13 g of PLA filament.
2. Assemble the metal or 3D slide with a Teflon membrane support by cutting a circle from the membrane to fit the opening with sufficient overlap to allow the membrane to be secured by an O-ring. Each membrane sheet can be used to make 4 circles. Using the hypodermic syringe filled with vacuum grease with attached cut-off 18 G needle, make three short lines of vacuum grease around the circumference of the opening on the membrane-covered metal or plastic edge, leaving large gaps to allow air trapped by the coverslip to escape.
3. After collecting embryos, larval hatching, and aging larvae to mid-L1 stage, transfer larvae to DDW in a shallow two-well dissecting slide as in step 2 above. The remaining well can contain PBS or Schneider’s Drosophila medium with or without supplements.
4. Dissect LBs in PBS or Schneider’s Drosophila medium with or without supplements, taking care to remove attached tissue without damaging the brains, then transfer LBs to a 9-10 µL drop of PBS or Schneider’s medium on a Teflon-covered metal or 3D printed slide for live imaging. Fat bodies from L1-L3 larvae can be placed around the LBs to induce qNSC activation and increase the number of actively dividing neuroblasts. After orienting the LBs and fat bodies (if added) as desired, carefully lower a clean round coverslip onto the vacuum grease on the membrane-covered metal or plastic edge; use gentle pressure to push the coverslip down until the LBs are just immobilized between the membrane and coverslip surface.
5. Image the LBs on the slide by confocal or spinning disk microscopy using settings appropriate for live imaging.