Small Scale Dissection and Antibody Staining of Eye-Antennal and Wing Imaginal Discs from Drosophila melanogaster

Summary

This article describes an efficient imaginal disc dissection and antibody staining protocol that is economical with reagents and facilitates handling of imaginal discs in small numbers.

Abstract

Drosophila imaginal discs have long been studied as models of epithelial development, patterning, and regeneration. Antibody labeling and imaging of dissected imaginal discs is important for visualizing protein expression patterns as markers of developmental and physiological processes. This method of imaginal disc dissection and labeling was originally developed for immune-electron microscopy of eye-antennal imaginal discs by Tomlinson and Ready. Using 60-microwell plates, and by transferring tissues between solutions rather than transferring solutions, individualized attention to tissue samples is possible with very little wastage. Even single imaginal discs can be processed. The method requires only small volumes of antibodies and other reagents. As an example of the method’s application to other small tissues, the article also demonstrates how wing imaginal discs can be dissected and labeled by the same method. A similar approach can be applied to any small tissue of similar dimensions, and also to other histochemical and labeling procedures besides antibody staining.

Introduction

Drosophila imaginal discs are progenitor tissues that are set aside during embryogenesis and grow within the larva without differentiating and contribute to the adult epidermal structures during the pupal stage¹. Their study has provided many insights into developmental patterning mechanisms, with the eye and wing imaginal discs having been particularly well studied^2,3,4,5,6. Immunohistochemistry is particularly useful to investigate spatial and temporal patterns of gene expression in imaginal discs, as well as to characterize cell behaviors, including cell division and cell death.

To facilitate the additional demands of electron microscopic analysis of antibody-labelled eye imaginal discs, Tomlinson and Ready developed a small-scale protocol that is particularly economical with reagents and provides a very high degree of reliability and customization⁷. Their method is described in print, but no video is available^3,8,9. The method is readily modified for the labelling of other imaginal discs (or other tissues) and can be applied to wing imaginal discs in addition to eye-antennal imaginal discs.

Antibody labeling of Drosophila imaginal discs follows the general outline applicable to any tissue¹⁰ (Figure 1). The tissue must be dissected and fixed before incubation with antibodies specific for proteins of interest. After the unbound antibody has been washed off, a labeled (often fluorescent) secondary antibody is added, which binds to the primary antibody, which is itself bound to the protein of interest in the fixed tissue. After washing to remove unbound secondary antibody, the tissue can be mounted for microscopic examination, and the proteins of interest can be visualized. Many general descriptions of this process are available and can be applied to imaginal discs and other Drosophila tissues¹¹. Here, a protocol is provided for the Tomlinson & Ready method, in which tissue samples are transferred individually between small incubation wells on a microwell plate and are observed by a stereo microscope at every step (Figure 2). This approach works best for staining smaller numbers of tissues at a time (tens of samples) and is not ideal for large batches (i.e., hundreds). This approach requires only small solution volumes of 13 µL in each well and so can be performed with very little antibody. Because there is minimal tissue loss there is no minimum sample size, and even single imaginal discs can be processed reliably.

Protocol

The reagents and the equipment used are listed in the Table of Materials.

1. Making the transfer tool

1. Use pliers to cut ~2.5cm of 0.005” diameter tungsten wire. Use forceps to form one end of the wire into a hook or a loop (Figure 3). Insert the other end into a needle holder. This will be used to transfer the discs with little or no liquid transfer. The loop may be the easier of the two to use, but the hook transfers less liquid between solutions.

2. Dissecting and fixing eye-antennal imaginal discs

1. Rinse larvae with ~7.5 mL 0.1 M sodium phosphate buffer (pH7.2) in a Petri dish (60 mm diameter) to remove any fly media or other debris. See step 6 for the dissection of wing imaginal discs rather than eye-antennal discs. Both these imaginal discs are located within the anterior portion of the body (Figure 4).
   NOTE: PBS can also be used instead of phosphate buffer in steps 2.1–2.3.
2. Transfer larvae to a Petri Dish containing fresh buffer.
   NOTE: Some workers prefer to dissect individual larvae in single buffer drops in a 60 mm Petri dish; others use depression well slides.
3. While observing under a dissecting microscope, hold the larvae ~50% of the way down the body using forceps. Use a second pair of fine forceps to grasp the mouth hooks and pull. The eye-antennal discs will come out attached to the mouth parts. The brain may also remain attached to the eye discs.
4. Use the wire tool to transfer the dissected tissue into a small Petri dish (35 mm diameter) containing ~3 mL fixative, e.g., 3.7% formaldehyde in 0.1 M sodium phosphate buffer (pH7.2), made by diluting 37% formaldehyde stock solution in buffer.
   NOTE: Follow locally approved safety procedures for handling formaldehyde. 3.7% formaldehyde is a simple fixative that will be satisfactory in many cases, but other fixatives may be optimal for certain antigens. PBS or PEM (0.1 M PIPES pH 7.0, 2 mM MgSO_4, 1 mM EGTA) may be used as the fixative buffer in place of 0.1 M sodium phosphate, or formaldehyde solution may be obtained by dissolving solid paraformaldehyde, rather than from a concentrated stock solution. The fixative paraformaldehyde-lysine-periodate (PLP) combines milder protein fixation with carbohydrate crosslinking and better preserves membrane structures¹². The Drosophila glycoprotein Scabrous, for example, will remain antigenic after PLP fixation but not after formaldehyde fixation. Prepare PLP freshly as follows: (1) Add 2 g of paraformaldehyde powder to 25 mL of deionized water in a conical flask in a fume hood and stir on low heat; (2) Dip a pipette into a 50% sodium hydroxide stock solution and transfer a trace of the liquid (much less than one drop) to the mix. The paraformaldehyde should dissolve within a few minutes without the solution coming to a boil. While cooling the 8% formaldehyde solution, dissolve 0.36 g lysine in 10 mL deionized water, 7.5 mL 0.1 M sodium phosphate pH 7.2, 2.5 mL 0.1 M Na₂HPO₄ and keep on ice. Mix 15 mL of this cold buffered lysine solution with 5 mL of the fresh 8% formaldehyde and 50 mg sodium periodate in a 50 mL screw-cap tube, mix by vortexing and use when fresh.
5. After 15–20 min at room temperature, transfer the dissected tissue into another Petri dish containing 0.1 M sodium phosphate buffer (pH 7.2).
   NOTE: PBS can also be used instead of phosphate buffer. Fixation on ice may be preferable for some antigens, or when it is desired to image the endogenous fluorescence of GFP or RFP expressed in the tissue. PLP fixation is always for 45 min on ice.

3. Blocking and incubating with primary antibody

1. Transfer tissue into a 35 mm Petri dish containing ~3 mL NSG antibody wash buffer for blocking. Incubate the Petri dish on ice for at least 15 min. NSG is 0.1 M sodium phosphate buffer (pH 7.2), 0.1% saponin, 5% Normal Goat Serum. Wash buffer stocks are stored filter-sterilized at 4 °C.
   NOTE: Different wash buffers may be optimal for particular antibodies. For example, whereas saponin is a mild, non-ionic detergent that particularly solubilizes the plasma membrane, other detergents may be preferable for detecting antigens within the nucleus. As a further example, the antibody developed against the nuclear Atonal protein by Jarman et al.¹³ is most immunoreactive in buffers containing NP40 as the detergent. See the Table of Materials for recipes for the alternative wash buffers: PDT, PBT, or NSG. When using normal serum as the blocking protein, ideally, it should be from the species in which the secondary antibody is produced, e.g., normal goat serum if goat secondary antibodies are to be used, or normal donkey serum if donkey secondary antibodies are to be used. In practice, this makes little difference.
2. Clear any unwanted tissues from the eye-antennal imaginal discs during blocking using fine forceps, except for the mouth hooks. Leave the mouthparts attached for now to facilitate transfers.
   NOTE: Extraneous tissue, e.g, brain, may also be left until slide mounting (step 5.4), so long as it does not interfere with staining or visibility.
3. While blocking, prepare a 1:100 dilution of mAb40-1a antibody solution in NSG, e.g., 1 µL of antibody in 100 µL NSG. Mix by pipetting, not vortexing.
   NOTE: mAb40-1a is a monoclonal antibody raised against beta-galactosidase. The optimal antibody dilution and wash buffer will differ depending on the particular antibodies used.
4. Add 13 µL antibody solutions to each of up to six wells in the first column of a 60-well microwell minitray. Conical microwells with straight sides are important for optical clarity. If more than six wells are necessary, start a second minitray.
5. Transfer eye-antennal imaginal discs, preferably still attached to mouthparts, to the primary antibody solution one at a time using the wire loop or hook.
   NOTE: The number of imaginal discs that can be added to each well depends on the antibody. Too much tissue can deplete some antibodies to the point that labeling diminishes. 5 pairs per well is usually satisfactory. Thus, ~60 discs can readily be labeled using the 6 rows of 60-well microtiter plates. Different antibodies can be used for each of the rows. Whether antibody solutions can be reused requires empirical testing for each case; it is not routine in our laboratory.
6. Close the plate with the lid and seal with parafilm to limit evaporation. Incubate on ice or at 4 °C for 4 h to overnight. Rocking is not necessary.
   NOTE: The optimal time and temperature of antibody incubation depend on the antibody. In many cases, 1–2 h at room temperature will be sufficient, but some antibodies will give improved results after incubation at 4 °C, and it may often be convenient to incubate at 4 °C overnight. In very few cases, primary antibody incubations of 2–3 days may be optimal. Sealing the microwell lid is only necessary for incubations longer than ~4 h.

4. Washing and incubating with secondary antibody

1. Add 13 µL of NSG buffer to the next three columns of wells after each antibody well, as the samples will be washed three times. Transfer the samples to wash for 5 min in each well successively.
2. While washing, prepare a 1:200–400 dilution of fluorescently conjugated secondary antibodies in NSG buffer, e.g., 0.5 µL of antibody in 100 µL NSG. Mix by pipetting, not vortexing. Centrifuge the secondary antibody stock solution for 60 s (at maximum speed) before dilution, as antibody aggregates can bind nonspecifically to the tissue surface.
   NOTE: The optimal secondary antibody dilution may vary by manufacturer. Alexa- or Cyanine-conjugated antibodies are available subtracted for other species to facilitate multiple labeling.
3. Add 13 µL of the secondary antibody solution to the next (5^th) column of wells.
4. Transfer samples into the secondary antibody using the loop or a hook.
5. Close the plate with the lid and seal with parafilm to limit evaporation. Incubate on ice for 4 h overnight.
   NOTE: The optimal time and temperature of antibody incubation depend on the antibody. In many cases, 1–2 h at room temperature will be sufficient, but some antibodies will give improved results after incubation at 4 °C, and it may often be convenient to incubate at 4 °C overnight. Sealing the microwell lid is only necessary for incubations longer than ~4 h.

5. Washing and mounting tissue for microscopy

1. Prepare the next three columns of wells after the secondary antibody well with 13 µL NSG. Transfer the samples through each well for 5 min.
2. Transfer the samples into a final wash in the 9^th well using 13 µL of 0.1M sodium phosphate buffer (pH 7.2). This buffer wash is necessary when wash buffers contain components that precipitate in the mounting medium.
3. . Add 13 µL of mounting medium to the final (10^th) column of wells. The mounting medium is made by dissolving 2% n-propyl gallate in 75% glycerol and 25% 0.1 M sodium phosphate buffer, pH 7.2. Prepare fresh mounting medium every week by mixing the components on a rotating platform. Allow any air bubbles to clear before use.
   NOTE: Mounting media are also available commercially.
4. Pipette 20 µL of mounting medium on a clean microscope slide. Transfer samples into the medium on the slide. This is a final opportunity to remove unwanted tissue. Cuticular mouthparts must be removed from eye-antennal discs.
5. To arrange samples so that they will not drift when the cover glass is overlaid, use the wire tool to drag samples out of the main drop of mountant. Gently lower a clean #1.5 thickness 22 mm x 40 mm cover glass onto the slide using forceps, without introducing air bubbles.
6. Let the slides sit for 1 h in a cardboard slide holder to let the mounting media settle. Avoid disturbing the slides as the fragile samples can be damaged if the cover glass moves.
7. Once the cover glass has settled, seal the edges using nail polish. Do not move or press the cover glass while doing this. Allow nail polish to dry for at least 1 h before imaging.
   NOTE: Unfortunately, most cosmetic nail polish now uses solvents that do not spread evenly on glass slides and are not useful, so that scientific nail polish must be obtained.
8. Store the slides at 4 °C. Samples can normally be imaged and re-imaged for months.
   NOTE: Slides can also be stored at -20 °C.

6. Dissecting and fixing wing imaginal discs

1. To dissect wing imaginal discs rather than eye-antennal imaginal discs, follow the steps below in place of steps 2.3–2.5 above.
2. Under a dissecting microscope, grip the larval mouthparts with fine forceps. Using microdissection scissors, bisect the larvae 30%–40% of the way down from the anterior.
3. Invert the anterior portion of the larva by spreading the open, cut end of the larvae using a second pair of forceps and pushing the head and mouthparts through, similar to inverting a sock
4. Use forceps to transfer the inverted anterior larvae into fixative into a small Petri dish (35 mm diameter) containing ~2.5 mL fixative, e.g., 3.7% formaldehyde in 0.1 M sodium phosphate buffer (pH 7.2), made by diluting 37% formaldehyde stock solution in PBS.
   NOTE: If desired, inverted carcasses can be transferred to 400 µL of fixative in a microfuge tube, which facilitates fixation in a fume hood. The same fixative options available for eye-imaginal disc fixation also apply to wing disc fixation (see NOTE on step 2.4). The optimal fixation and for labeling eye-antennal imaginal discs will usually be the same for wing discs.
5. After 15–20 min at room temperature, transfer the dissected tissue into another Petri dish containing 0.1 M sodium phosphate buffer (pH 7.2).
   NOTE: The same wash buffer options available for eye-imaginal disc fixation also apply to wing disc fixation (see NOTE on step 2.4). The optimal wash buffer for labeling eye-antennal imaginal discs will usually be the same for wing discs.
6. Remove wing discs from the larvae using fine forceps. If still present, the smaller haltere disc and leg disc can either be removed now or remain attached to the wing discs to facilitate transfers.
   NOTE: The wing discs may not be readily visible in the carcass until fat body or other tissue is pulled away (always identify a tissue before removing it). The wing discs are the largest imaginal discs and are attached to the body wall, closely associated with the trachea. They have a teardrop shape, like the outline of South America¹ (Figure 4).
7. Continue with steps 3–5 as above.

Results

The Tomlinson and Ready procedure regularly yields excellent antibody staining results. Here, triply labeled eye, antennal, and wing imaginal discs are illustrated. In Figure 5A, a third-instar eye-antennal imaginal disc has been triple-labeled to detect the transcription factor Senseless (blue), neural progenitor marker Elav (red), and anti-GFP labeling of cell clones expressing RNAi for the Ocho transcript (green). In Figure 5B, a third instar wing im...

Discussion

Antibody staining of Drosophila tissues using 60-well conical-well plates using the Tomlinson and Ready method results in excellent results and is highly reliable, as each imaginal disc is followed under the dissecting microscope during each manipulation. The method requires only small amounts of antibody reagents, and exchanges solutions more completely with each step since samples are transferred with very little associated liquid. It is particularly suited to handling modest numbers of samples that may easily...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Disodium Phosphate	Fisher	7558-79-4	Na2HPO4 - sodium phosphate dibasic anhydrous. Other hydration states can be used but change the molecular weight
Forceps: Dumont #5 Biologie forceps	Fine science tools	11252-20	Style: #5
			Tip Shape: Straight
			Length: 11cm
			Autoclave Safe: Yes
			Tips: Biology
			Tip Dimensions:
			0.05 x 0.02mm
			Alloy / Material: Inox
Formaldehyde	Fisher	BP531-500	Molecular Biology-grade, 37% Formaldehyde. 500ml
Formaldehyde Fixative	Fisher	BP531-500	3.7% formaldehyde in 0.1M Sodium Phosphate Buffer pH7.2 made by dilution from 37% stock.
Glycerol	Fisher Bioreagents	BP229-1
Ice	User Preference
L-Lysine, 98%	Thermoscientific	J62225.22
mAb40-1	DHSB	40-1a-s	Anti-beta galactosidase
Microcentrifuge tubes	USA scientific	1615-5520	1.5mL tubes to hold samples.
Microdissection Scissors	Fine Science Tools	15003-08	5mm cutting edge, straight tip, 8.5cm
Monosodium Phosphate	Fisher	7558-80-7	NaH2PO4 - Sodium Phosphate Monobasic anhydrous. If using a hydrated form, the weight needed to make a 0.1M solution would change.
Mounting Medium			75% glycerol solution in 0.1M sodium phosphate buffer pH7.2 with 2.5% n-propyl gallate. Mix by inversion to minimize bubbles.
NSG			0.1M sodium phosphate buffer pH7.2, 0.1% saponin, 5% NGS
n-propyl gallate	MP Biomedicals	210274780
Nail Polish	Ted Pella, Purchased from Fisher	NC0381432	For sealing slides
Needle holder	Fisher	NC0099380	For microdissecting needles. Hollow stainless steel; handle length: 4 3/4" long
Nunc Microwell MiniTrays	Thermo Scientific	12565155	MicroWell MiniTray for serological applications. Well count 60
Parafilm	StatLab	PM996	StatLab ParaFilm M Self-Sealing Flexible Film
Paraformaldehyde	Polysciences, Inc.	00380-250
10x PBS	User Preference		Prepare 800 mL of distilled water in a suitable container.
			Add 1.37M of Sodium chloride to the solution.
			Add 26.8mM of Potassium Chloride to the solution.
			Add 101mM of Sodium Phosphate Dibasic to the solution.
			Add 18.0mM of Potassium Phosphate Monobasic to the solution.
			Adjust solution to desired pH (typically pH ≈ 7.4).
			Add distilled water until the volume is 1 L.
PBT	User Preference		0.1M sodium phosphate buffer pH7.2, 0.1% bovine serum albumin, and 0.2% Triton X-100.
PDT	User Preference		0.1M Sodium Phosphate buffer (pH = 7.2), 0.3% deoxycholate, and 0.3% Triton x-100
Petri Dish	Falcon	Corning 351007	Petri Dish; 60mm; non treated
Petri Dish	Falcon	Corning 351008	Petri Dish; 35mm; non treated
Pliers	User Preference
PLP	n/a		2% paraformaldehyde 1.35% lysine in 0.05M sodium phosphate buffer pH7.2
Secondary Antibodies	Jackson ImmunoResearch	715-545-151	0.5mg Alexa Fluor 488 Donkey Anti-Mouse IgG.
Slide Covers	VWR	48393-172	VWR Coverglass #1.5 22x40 (CASE)
Slides	Fisher	125442	Fisher Glass Slides 10bx/CS
Sodium Deoxycholate	Thermo Fisher Sci	89904	Sodium Deoxycholate Detergent; 5mg
Sodium Periodate	Sigma Aldrich	71859-25G	Sodium Periodate – 25 grams
Sodium Phosphate Buffer			0.1M NaPO4 buffer pH 7.2. Mix 28ml of 0.1M anyhdrous monosodium phosphate stock (11.998g NaH2PO4 to 1L of distilled water) with 72ml of 0.1M anyhydrous disodium phosphate stock (14.196g Na2HPO4 to 1L of distilled water) to generate 100ml of 0.1M NaPO4 buffer. Alternatively, PBS may be used.
Tungsten wire	Ted Pella	27-11	.005” dia.
Triton x-100	Thermo Fisher Sci	A16046AE
Vortex	User Preference