Dissection of the Eye-Brain Complex from Fly Pupae: A Method to Isolate Retinal Tissue

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This protocol is an excerpt from DeAngelis and Johnson, Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation, J. Vis. Exp. (2019).

1. Tissue Preparation

1. Set up Drosophila crosses (as described previously in Greenspan, Cold Spring Harbor Laboratory Press, (2004)) or culture specific Drosophila strains to obtain pupae of the desired genotype. To ensure that a large number of pupae emerge coincidently, establish these fly cultures in duplicate on nutrient-rich food media or standard food media generously supplemented with yeast-paste.
2. Maintain Drosophila cultures at 25 °C. For crosses utilizing the UAS-GAL4 system, GMR-GAL4 is an ideal driver expressed in larval eye disc cells posterior to the morphogenetic furrow and throughout pupal development. The cross UAS-lacZ X GMR-GAL4 serves as an ideal control cross as it drives expression of the non-endogenous and inert β-galactosidase protein.
3. Use the tip of a 6" bamboo splint that has been wet with distilled water to gently lift white pre-pupae (Figure 1B) from the side of healthy culture vials (Figure 1A) and place into a 1.5 mL microcentrifuge tube (Figure 1C).
4. Place the tubes into a plastic pipette tip box along with a small piece of tissue soaked in distilled water to maintain the chamber at sufficient humidity to protect the pupae from desiccation (Figure 1D). Incubate the pupae at 25 °C until dissection.
5. Use a dry 6" bamboo splint to gently push the pupae out from the microcentrifuge tube and onto a black dissecting dish. Lay a fresh piece of double-sided tape onto the dissecting dish away from the pupae.
6. Using a pair of forceps, carefully place the pupae dorsal side up (i.e., operculums facing up, Figure 1B) onto the tape (Figure 2A). Ensure that the pupae adhere well to the tape.
   NOTE: Moisture on the pupal case will inhibit secure adhesion to the tape, in which case, allow the pupae to air-dry before placing onto the double-sided tape.

2. Dissection of Eye-brain Complexes

1. Use forceps to remove the operculum of each pupa (Figure 2C).
2. Use microdissection scissors to slice or cut open the pupal case of each pupa. Flap open the pupal case to reveal the head, thorax, and anterior abdominal segment, securing the edges of the pupal case to the double-sided tape (Figure 2C).
   NOTE: It is not necessary to entirely reveal the pupae.
3. Pierce the abdomen of each pupa with sharp forceps, to grasp the animal, and remove it from its pupal case (Figure 2C).
4. Place the pupae onto the black dissection dish, away from the tape, and cover with about 400 µL of ice-cold phosphate-buffered-solution (PBS, pH 7.4).
5. Grasp each pupa by the abdomen with forceps, and with microdissection scissors, make a clean cross-sectional cut through the entire thorax, cutting the pupa in half (Figure 2D).
6. Remove the posterior remnant of each carcass from the PBS and put to one side on the dissecting dish (do not discard, see step 3.2.1).
7. Using two pairs of fine forceps, open the thorax and head of each pupa to reveal the eye-brain complex (Figure 2E). To do this, grasp the cut-edges of the thorax epithelium and gradually tear to open the thorax and then head capsule, exposing the eye-brain complex and surrounding fat tissue.
8. Without grasping the tissue, use forceps to guide the eye-brain complex away from the remnants of the head capsule or scoop the eye-brain complex from the torn-open head capsule.
   NOTE: The eye-brain complex is dumbbell-shaped, off-white, and more translucent than the surrounding cream-colored fat (Figure 2B,E).
9. With forceps, carefully remove most fat associated with the eye-brain complexes without touching the eye tissue.

3. Processing of Tissue for Immunofluorescence

1. Dissect at least 6-10 pupae as described (steps 2.1-2.9).
   NOTE: Three to four independent replicates of each dissection tend to yield sufficient data to test a hypothesis.
2. Washing and Fixation
   1. Cut the tip of a P20 or P100 pipette tip with a clean razor blade to increase the tip-opening to ~1 mm in radius and lubricate by pipetting a mix of PBS and fat up and down. This fat can be obtained from the carcass remnants removed in step 2.6.
   2. Transfer the eye-brain complexes into ~400 µL of PBS in a 9-well glass dish, on ice. Use the lubricated tip and a P20 or P100 air displacement micropipette to transfer.
      NOTE: Eye-brain complexes will adhere to unlubricated tips during pipetting and be damaged or lost.
   3. Remove the remaining fat associated with the eye tissue by pipetting the PBS up and down to gently swirl the eye-brain complexes. In this and all subsequent washing steps, do not directly pipette eye-brain complexes up and down as this will damage the fragile eye tissue.
   4. Transfer the eye-brain complexes in a minimal volume (<20 µL) of PBS into at least 250 µL of fixative. Mix by pipetting the solution up and down. Incubate for 35 min, on ice.
      NOTE: The same lubricated tip prepared in step 3.1.1 can be used for this transfer.
   5. With the same pipette tip, transfer the eye-brain complexes into a well containing ~400 µL of PBS. Mix by pipetting the solution up and down and incubate for 5-10 min on ice, to wash. Repeat the washing step at least twice.
      NOTE: Eye-brain complexes can be maintained for several hours in the final PBS wash, on ice, or at 4 °C, before proceeding to Step 3.3.1.
3. Antibody Staining
   1. To block the tissue prior to exposure to antibody solutions, transfer the eye-brain complexes to ~400 µL of PBT. Use the lubricated tip and a P20 or P100 air displacement micropipette for the transfer. Incubate for 10 to 60 min, on ice.
   2. Prepare the primary antibody solution by diluting appropriate antibodies in PBT. Since eye-brain complexes are incubated in 10 µL aliquots of antibody solution, the volume prepared will be a replicate of 10.
   3. Aliquot 10 µL of antibody solution into a clean well of a 72-well microwell tray.
   4. Cut the tip of a P10 pipette tip with a clean razor blade to increase the tip-opening to ~0.5 mm in radius and lubricate by pipetting PBT up and down.
   5. Transfer no more than 5 eye-brain complexes, in a volume of <3 µL of PBS, into each 10 µL well of antibody solution using a P10 air displacement micropipette and the lubricated tip.
   6. Homogenize the antibody solution by pipetting the solution up and down. Do not pipette the eye-brain complexes up and down as this will damage the tissue.
   7. To minimize evaporation of the antibody solution, place a small piece of tissue soaked in distilled water into the microwell tray and seal the tray (e.g., with the lid provided). Incubate overnight at 4 °C.
   8. Transfer the eye-brain complexes from the microwell tray into ~400 µL of PBT in a 9-welled glass dish, to wash. Use a P10 air displacement micropipette and PBT-lubricated tip (prepare as described in step 3.3.4). Mix by pipetting the solution up and down. Incubate for 5-10 min on ice.
   9. Repeat the washing step at least twice. Use a P20 or P100 air displacement micropipette and PBT-lubricated tip to transfer the eye-brain complexes between wash solutions.
   10. Prepare the secondary antibody solution by diluting appropriate fluorophore-tagged secondary antibodies in PBT as required. 100 µL of secondary antibody solution is sufficient per batch of 6-10 pupae. Aliquot the secondary antibody solution into a 9-welled glass dissection dish.
   11. Transfer the eye-brain complexes into secondary antibody solution. Use a P20 or P100 air displacement micropipette and PBT-lubricated tip for the transfer.
   12. Incubate the eye-brain complexes in secondary antibody solution for 1-2 h at room temperature, or 3-5 h at 4 °C. To prevent bleaching of fluorophores by light, cover the preparation with foil.
       NOTE: Optimal duration of incubation in secondary antibody solution may differ according to the primary and secondary antibodies used.
   13. Transfer the eye-brain complexes into ~400 µL of PBT in a 9-well glass dish, to wash. Mix by pipetting the solution up and down. Incubate for 5-10 min on ice. This washing step should be repeated at least twice.
4. Secondary Fixation
   1. For a secondary fixation, transfer the eye-brain complexes to at least 200 µL of fixative and incubate for 20 min at room temperature or 35 min at 4 °C.
      NOTE: Secondary fixation stiffens the eye tissue, which aids mounting (step 3.5).
   2. Use a P20 or P100 air displacement micropipette and PBT-lubricated tip to transfer eye-brain complexes into ~400 µL of PBT in a 9-well glass dish, on ice, to wash for 5 min.
   3. Repeat the washing step at least once.
   4. Use a P20 or P100 air displacement micropipette and PBT-lubricated tip to transfer the eye-brain complexes into ~400 µL of PBS in a 9-well glass dish, on ice, to rinse for 1-2 min. Keep the tissue in motion by pipetting PBS, but not the tissue, up and down to prevent eye-brain complexes from settling and adhering to the glass dish during PBS-rinse.
5. Mounting
   1. Transfer the eye-brain complexes into ~200 µL of mounting media. Use a P20 or P100 air displacement micropipette and PBT-lubricated tip to transfer the eye-brain complexes. Allow the tissue to equilibrate in mounting media for 1-2 h.
   2. Place a 5-8 µL drop of fresh mounting media onto a clean microscope slide.
   3. Cut a P10 tip with a clean razor blade to increase the tip-opening to ~0.5 mm in radius and use this and a P10 air displacement micropipette to transfer the eye-brain complexes in 5-8 µL of mounting media into the drop of mounting media on the slide.
   4. Use two tungsten needles to separate the eyes from optic lobes. Pin an eye-brain complex to the slide with a sturdy tungsten needle and slice each eye away from its associated optic lobe with a fine tungsten needle (Figure 2F).
   5. Use the fine tungsten needle to maneuver each eye to the surface of the microscope slide with the basal surface of each eye adjacent to the slide. Gently lower a clean cover-slip over the tissue and secure with nail polish.
      NOTE: Arranging the eyes on the slide so that they are close to each other or in a line will facilitate subsequent microscopy.
   6. Image the retinas using fluorescent or confocal microscopy.
      NOTE: Slides should be stored at 4 °C if not imaged immediately.

Результаты

Figure 1: Selection and culture of pupae for dissection. (A) Wandering third larval instar (L3) larvae and pupae locate along the sides of healthy Drosophila cultures. (B) Pre-pupae can be identified by their translucent white color as pigment has yet to be generated in the protective pupal case. Anterior-posterior and dorsal-ventral axis of the pupa are shown in blue. A dam...

Материалы

Name	Company	Catalog Number	Comments
Bamboo splints, 6"	Ted Pella Inc	116
Black dissecting dish			Glass petri dish filled to rim with SYLG170 or SYLG184 (colored black with finely ground charcoal powder). Leave at room temperature for 24-48 h to polymerize.
Blade holder	Fine Science Tools	10053
Bovine serum albumin	Sigma-Aldrich	A7906
Double-sided tape	3M	665
Drosophila& food media, nutrient-rich			7.5% sucrose, 15% glucose, 2.5% agar, 20% brewers yeast, 5% peptone, 0.125% MgSO4.7H2O, 0.125% CaCl2.2H20
Drosophila& food media, standard			Bloomington Drosophila Stock center cornmeal recipe. (https://bdsc.indiana.edu/information/recipes/bloomfood.html)
Fixative solution			4% formadehyde in PBS, pH 7.4.
Forceps	Fine Science Tools	91150-20	Forceps should be sharpened frequently.
Formaldehyde	Thermo Scientific	28908
Glass 9-well dishes	Corning	7220-85	Also known as 9-well dishes
Glass coverslips (22 x 22 mm)	Fisher Scientific	12-542-B
Glass microscope slides (25 x 75 x 1 mm)	Fisher Scientific	12-550-413
Glass petri dish	Corning	3160-100BO
Glycerol	Sigma-Aldrich	G5516
Microcentrigure tubes	Axygen	MCT-175-C
Microdissection scissors	Fine Science Tools	15000-03
Microwell trays (72 x 10 µL wells)	Nunc	438733
Mounting media			0.5% N-propylgallate and 80% glycerol in PBS
N-propylgallate	Sigma-Aldrich	P3130
PBS (phosphate buffered saline pH 7.4)	Sigma-Aldrich	P5368	Prepare according to manufacturer's instructions
PBT			0.15% TritonX and 0.5% bovine serum albumin in PBS, pH 7.4
Pin holder	Fine Science Tools	26016-12
Primary antibody: goat anti-GAPDH	Imgenex	IMG-3073	For Western blotting. Used at 1:3000
Primary antibody: rabbit anti-cleaved Dcp-1	Cell signaling	9578S	For immunofluorescence. Used at 1:100
Primary antibody: rat anti-DEcad	Developmental Studies Hybridoma Bank	DCAD2	For immunofluorescence. Used at 1:20
Primary antibody: rat anti-DEcad	DOI: 10.1006/dbio.1994.1287	DCAD1	Gift from Tadashi Uemura. Used at 1:100.
Scalpel blades	Fine Science Tools	10050	Break off small piece of scapel blade and secure in blade holder.
Secondary antibody: 488-conjugated donkey anti-rat IgG (H+L)	Jackson ImmunoResearch	712-545-153	For immunofluorescence. Used at 1:200
Secondary antibody: cy3-conjugated goat anti-rabbit IgG (H+L)	Jackson ImmunoResearch	111-165-144	For immunofluorescence. Used at 1:100
Secondary antibody: HRP-conjugated goat anti-rat IgG (H+L)	Cell Signaling Technology	7077	For Western blotting. Used at 1:3000
Secondary antibody: HRP-conjugated rabbit anti-goat IgG (H+L)	Jackson ImmunoResearch	305-035-003	For Western blotting. Used at 1:3000
Stereo dissecting microscope (M60 or M80)	Leica Microsystems		or similar microscope
Sylgard (black)	Dow Corning	SYLG170
Sylgard (transparent)	Dow Corning	SYLG184	Color black with finely ground charcol powder
Tissue: Kimwipes	KIMTECH	34120
TritonX	Sigma-Aldrich	T8787
Tungsten needle, fine	Fine Science Tools	10130-10	Insert into pin holder
Tungsten needle, sturdy	Fine Science Tools	10130-20	Insert into pin holder
Yeast paste	(local supermarket)		Approximately 2 tablespoons Fleischmann's ActiveDry Yeast (or similar) dissolved in ~20 mL distilled H2O