Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Summary

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Abstract

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

Introduction

Drosophila embryos and larvae provide an excellent model to study developmental processes in many organs and tissues. Imaging of individual cells is often necessary in these studies in order to ascertain the complex environments in which cells develop. Visualization of cells in tissues can be accomplished through immunostaining. Well-described immunostaining protocols exist for embryonic Drosophila tissues <17 hr after egg laying (AEL)^1-3. However, a protective cuticle forms toward the end of embryogenesis, preventing effective antibody permeation. Thus, these immunostaining protocols are inefficient in the analysis of tissues in late-stage embryos and in subsequent stages of larval development (1^st instar (L1), 2^nd instar (L2), and 3^rd instar (L3)). This inefficiency imposes a barrier to our understanding of dynamic processes that occur during this extended developmental period ⁴. Tissue dissection is a widely employed technique to circumvent this barrier ^5-7. However, dissection can prove inefficient. Extraction may be encumbered by difficulty in locating or isolating embryonic and larval tissues. Furthermore, the physical removal of target tissues may cause damage by rupturing them or by failing to extract them in their entirety.

Sonication is a method that employs sound waves to disturb intermolecular interactions. It has been used to disrupt the integrity of the Drosophila larval cuticle in order to immunostain developing neural cell types ⁶. This protocol has been adapted to immunostain late-stage embryonic and larval gonads, which can be as small as 50μm in diameter ^8-10. Through such studies, the process of male germline stem cell (GSC) niche formation has been characterized in late stage Drosophila embryos^8-10 and mechanisms regulating stem cell development and differentiation in late stage embryonic gonads and larvae have been elucidated ^9-12. Thus, sonication provides an efficient alternative to tissue dissection that may be difficult because of tissue size. Furthermore, it enables immunostaining of Drosophila tissues in situ, leaving cells within the context of the entire organism and maintaining in situ morphology. Here, we describe a step-by-step protocol for fluorescence immunostaining of late-stage embryonic through early/mid-L3 tissues in situ. Analysis of Drosophila gonadal and neural tissue is shown in the Representative Results to demonstrate the efficacy of this protocol. Furthermore, this immunostaining protocol may be adapted to analyze other Drosophila tissues as well as tissues in other organisms with an outer cuticle.

Protocol

1. Preparation of a Collection Cage

1. Anesthetize young, fertile flies with CO₂. Transfer anesthetized flies to a cage. To obtain optimal yield, use 100-120 adult flies ranging from 2-7 days of age at a 4:1 ratio of females to males. Allow flies an appropriate acclimation period, ~24 hr prior to obtaining sample for fixation. If the cage was set up with virgin females mated to males, a use a 36 - 48 hr acclimation period.
2. On the open end of the cage, place a pre-prepared apple juice agar plate with a dime-sized drop of yeast paste in the center over the opening of the cage. Then, secure with tape. Seal the junction between the apple juice agar plate and the cage with Parafilm.
3. Place the cage in a secondary container with apple juice agar plate as the base and allow the flies to reproduce at a defined temperature for an appropriate period of time as determined by the experiment.
   Note: Sample collection times will vary. For example, when collecting late stage embryos/early L1 larvae (17 - 24 hr), allow flies to lay eggs on apple juice agar plates for 7 hr at 25 °C prior to aging of sample for 17 hr at 25 °C. Similarly, for a collection of mid-late first instar larvae (36 - 48 hr) allow flies to lay eggs for 12 hr at 25 °C prior to aging of sample for 36 hr at 25 °C.
4. Once the time period is complete, tap the cage on a table so that flies drop away from the apple juice agar plate without anesthesia. Quickly replace the used plate with a fresh one containing a drop of yeast paste. Secure the fresh plate with tape and parafilm and store the cage with apple juice agar plate as base.
5. Carefully remove the yeast drop from the used plate with a metal spatula. Place lid on used apple juice plate and allow laid embryos to age if experimentally necessary.
   Note: While aging samples, plates may be stored at 25 °C unless higher temperatures are experimentally required. Examples of how to age embryos for collection are described above (see note for step 1.3). Removal of yeast paste prior to sample aging is performed to prevent larvae from crawling into the yeast and breaking up the agar underneath. The remaining apple juice agar and yeast paste residue provide nutrients for larval growth. While embryos laid directly in the yeast paste are lost through yeast paste removal, this loss is preferable to yeast and agar residue in the sample that can reduce staining efficiency. Should one choose not to remove the yeast paste, yeast may be liquefied with Phosphate Buffer Triton X-100 (PBTx), mixed with a paint brush, and then removed from the sample with a cell strainer prior to fixation.

2. Fixation

1. Once embryos laid onto the apple juice agar plate have aged to the desired time point, use a small paintbrush wetted with Phosphate Buffer Triton X-100 (PBTx) solution to carefully remove sample from the plate.
   Note: Older larvae are mobile and may crawl onto the inside of the lid. This sample may be collected similarly by removal with a paintbrush.
2. Wipe the sample-laden paintbrush against the interior-facing ridge of a cell strainer. Then, rinse the walls of the strainer and the paintbrush with a squirt bottle containing PBTx solution. Place a petri dish cover underneath the strainer to capture the PBTx solution.
3. After all the sample has been transferred to the strainer, pour enough PBTx into the strainer to raise the sample off the mesh. Then, lift the strainer and pour the contents of the petri dish into a liquid waste container.
4. Repeat Step 2.3 three times to remove yeast and fly waste that may have been transferred along with the sample.
5. Pour enough 50% bleach (NaOCl)/water (ddH₂O) solution into the strainer to raise the larvae off the mesh. Allow larvae to sit in solution for 5 min. Then, lift the strainer and pour out the contents of the petri dish in a liquid waste container.
   Note: Bleach is required only in embryonic samples aged 0 - 22 hr in order to remove the chorionic membrane. The application of bleach to older samples can be omitted, but its inclusion is not detrimental. If omission is desired, replace the bleach in this step with PBTx. Diluted bleach should be used within 24 hr of solution preparation.
6. Wash sample in the strainer with PBTx by pouring enough PBTx into the strainer to raise the sample off the mesh. Allow sample to sit in solution for 3 min. Then, lift the strainer and pour the contents of the petri dish into a liquid waste container.
7. Repeat Step 2.6 five times.
8. Dab the bottom of cell strainer dry, then transfer sample into a scintillation vial containing 1.75 ml of PEMS solution using a paintbrush.
9. Working in a ventilated fume hood, add 250 μl of 37% formaldehyde and 8 ml of reagent grade heptane to the scintillation vial.
   Note: Formaldehyde and heptane are hazardous. For safety purposes, continue to work in a ventilated fume hood and wear appropriate gloves for steps 2.9 – 3.3.
10. Allow the vial to shake at 200 rpm or a similar moderate speed for 20 min.
11. Add 10 ml of methanol to scintillation vial without removing the previous aqueous phase and allow the vial to shake vigorously at 500 rpm for 1 min.
    Note: Methanol is hazardous. Continue to work in a ventilated fume hood and wear appropriate gloves.
12. Remove vial from shaker and immediately pour contents into a cell strainer over a liquid waste container in the hood. Add extra methanol to the vial and run through the cell strainer as necessary to ensure all sample has been removed.
    Note: Cell strainers may drain slowly. To remedy this, touch a laboratory tissue to the bottom of the cell strainer in order to draw the solution through the strainer more quickly. Methanol, formaldehyde, and heptane should be stored in appropriate waste containers until proper disposal as per institutional guidelines.
13. Dry bottom of cell strainer using a laboratory tissue, then use a paintbrush to transfer sample captured in the strainer to a 1.5 ml microcentrifuge tube containing 0.5 ml of methanol.
    Note: If multiple scintillation vials are in use, complete steps 2.12 and 2.13 one vial at a time to prevent sample from drying on cell strainers.
14. Once the sample has been added to the microcentrifuge tube, remove methanol with a pipette. Ensure sample has settled to the bottom of the tube.
15. Rinse sample in the microcentrifuge tube by adding approximately 0.5 ml of methanol to tube. Wait for sample to settle then remove methanol with a pipette.
16. Repeat Step 15 three times.
17. Add 0.5 ml of methanol to the tube, and then store in a -20 °C freezer for later use.

3. Rehydration and Preparation of Sample for Immunostaining

1. Remove methanol from the microcentrifuge tube with a pipette, leaving the sample in the tube. Store methanol waste to the appropriate waste container until time of proper disposal.
   Note: To enhance sonication efficiency, use no more than 3 mm of sample settled at the bottom of the 1.5 ml microcentrifuge tube. Excess sample may be transferred to a separate tube with a pipette before beginning the rehydration step above. Cutting the pipette tip with a clean razor blade can be used to prevent clogging of the pipette tip.
2. Add 1.0 ml of a 50% methanol/Phosphate Buffer Tween (PBTw) solution to the tube and rock for 3 min.
3. Remove methanol solution to the proper waste container and rinse with 1.0 μl of PBTw twice. After each rinse, allow the sample to settle before drawing off the PBTw with a pipette.
4. Add 1.0 ml of Bovine Serum Albumin/Phosphate Buffered Tween (BBTw) mix to the vial and rock. After 3 min on a rocker, allow the sample to settle and remove the BBTw with pipette.
5. Repeat Step 3.4 two times taking care to remove as much BBTw as possible without losing sample after the last wash.
6. Add 0.5 ml of BBTw to the tube and place the tube on ice.

4. Sonication of Sample

1. Set the sonicator to 10% maximum amplitude and designate a run time of 2 sec constant sonication.
   Note: These settings are specific to the sonicator indicated in the Materials and Equipment Table and may require optimization for different sonicators.
2. Prior to sonication of sample, clean the sonicator probe by placing the probe tip into a 50 ml conical tube filled with deionized water and run sonicator to clean the probe. Dry sonicator probe with laboratory tissue after run.
3. Submerge the probe into the microcentrifuge tube containing sample so that it is positioned approximately 3 to 4 mm above the sample and start sonication.
4. Remove the microcentrifuge tube and place it on ice for at least 30 sec to dissipate heat from sonication and allow sample to settle at the bottom of the microcentrifuge tube.
5. Repeat steps 4.3 and 4.4 as needed based on age of sample being processed.
   Note: For optimum sonication sample volume, embryos younger than 17 hr do not require sonication, but those between 17 and 24 hr (stage 17/early-L1) require two sonications. Larvae between 24 - 36 (early/mid-L1), 36 - 48 (mid/late-L1), 48 - 60 (early/mid-L2), 60 - 72 (mid/late-L2), 72 - 84 (early-L3), 84 - 96 (early/mid-L3), 96 - 108 (mid/late-L3) hr require 5, 9, 11, 13, 15, 18, and 22 sonications respectively. See discussion for further elaboration on sonication times.
6. Rinse with 1.0 ml BBTw twice. After each rinse allow the sample to settle before drawing off BBTw with a pipette.
7. Add 1.0 ml of BBTw to the vial and rock. After 3 min on the rocker, allow the sample to settle and remove the BBTw with pipette.
8. Repeat Step 4.7 two times.

5. Immunostaining

1. Block sample by adding 5% normal serum diluted in BBTw to the microcentrifuge tube. Remove block after rocking for 1 hr.
   Note: Use normal serum from the species in which the secondary antibodies are generated.
2. Dilute primary antibodies to appropriate working concentration in 5% normal serum/BBTw solution.
3. Rock sample in primary antibody solution O/N at 4 °C or for at least 3 hr at RT (approximately 22 °C.
   Note: Antibody incubation times and temperatures may vary. O/N incubation at 4 °C or 3 hr at RT is typically sufficient. However, a two-day incubation period may enhance staining quality, especially with older samples or when low-affinity primary antibodies are used. Longer incubations are typically performed at 4 °C. Similar considerations must be made when using secondary antibodies (see 5.10).
4. Allow sample to settle to bottom of microfuge tube. Draw off primary antibodies with pipette. Save antibodies for reuse if desired.
5. Rinse with 1.0 ml of BBTw twice. After each rinse allow sample to settle before drawing off BBTw with a pipette.
6. Add 1.0 ml of BBTw to the vial and rock. After 3 min on the rocker, allow sample to settle and remove BBTw with pipette.
7. Repeat Step 5.6 five times.
8. Block sample by adding 5% normal serum/BBTw solution to the microcentrifuge tube. Remove block after rocking for 30 min to 1 hr.
   Note: This additional blocking step is not required, but may improve staining quality for specific antibodies.
9. Dilute secondary antibodies to appropriate working concentration in 5% normal serum/BBTw solution.
10. Wrap microcentrifuge tube in aluminum foil to reduce fading due to light exposure. Rock sample in secondary antibody solution for 12 to 48 hr at 4 °C or for at least 3 hr at RT (approximately 22 °C.
11. Allow sample to settle to bottom of microfuge tube. Draw off secondary antibodies with pipette.
12. Rinse with 1.0 ml of PBTw twice. After each rinse allow sample to settle before drawing off the PBTw with a pipette.
13. Add 1.0 ml of PBTw to the vial and rock. After 5 min on the rocker, allow the sample to settle and remove the PBTw with pipette.
14. Repeat Step 5.13 three times.
15. OPTIONAL: Add DAPI diluted 1:1,000 in PBTw. Rock sample wrapped in aluminum foil for 3 min; then remove DAPI solution with a pipette. Rinse five times with 1.0 ml of PBTw, allowing sample to settle before removing PBTw with pipette.
16. Add 80 - 100 μl of 1,4-diazabicyclo[2.2.2]octane (DABCO) solution to microcentrifuge tube and store at -20 °C.
    Note: Another glycerol-based anti-fade agent may be substituted for .

6. Analysis

1. Before mounting sample onto slides, add an extra 5 - 10 μl of DABCO/p-phenylenediamine (PPD) antifade solution to microcentrifuge tube.
   Note: Sample can be added to slides without dissection. Sample volume added to a slide varies with cover slip size. When pipetting sample onto slide, the pipette tip may be cut off using a razor blade to prevent sample shearing. It is often useful to line sample up in rows for easy analysis. The addition of PPD as an additional antifade agent may not be required, but can prevent photobleaching if samples are stored long-term.
2. Gently place glass cover slip over sample. Then, secure cover slip in place using nail polish.
3. View mounted sample using fluorescence microscopy.

Results

To demonstrate the efficacy of sonication-based immunostaining in analysis of late-stage embryonic and larval tissues in situ, wild-type embryos and larvae were processed for immunostaining of testes, ovaries, and neural tissue. Samples were imaged via confocal microscopy and representative results are shown (Figure 1 and Figure 2). Results reveal that the described protocol is effective for visualizing morphological features as well as individual cells in situ during l...

Discussion

This protocol provides a method to successfully immunostain target Drosophila embryonic and larval tissues in situ, thus eliminating the need for dissection. As per prior protocols for staining early embryos^1,2,3, the chorionic membrane is removed using 50% bleach (NaOCl). Samples are fixed in formaldehyde and methanol. Because the larval cuticle causes older sample to float, the entire sample is then passed through a cell-strainer to ensure larval retention. Sample is stored, if desired, in ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Table 1: Reagents and Buffers
Phosphate buffer Triton X-100 (PBTx)			For 5 L: 500 ml PBS 10X, 4.45 L ddH2O, 50 ml Triton 10%. Store at RT.
Phosphate buffer saline 10X (PBS 10X)			For 1 L in dH2O: 80 g NaCl, 2 g KCl,14.4g Na2HPO4, 2.4 g KH2PO4. Add components and fill to appropriate volume. Store at 4 °C.
Triton 10%			For 50 ml: 5 ml of Triton, 5 ml of PBS 10X, 45 ml of ddH2O. Rock to mix. Store at RT.
PEMS			0.1 M Pipes (pH 6.9), 2.0 mM MgSO4, 1.0 mM EGTA. Store at RT.
Pipes			For a 400 ml of a 0.25 M solution (pH 6.9): 30.24 g Pipes dH20 NaOH. Dissolve Pipes in 300 ml dH2O and then adjust to pH 6.9 with NaOH. Bring the total volume to 400 ml with dH2O and autoclave. Store at RT.
Formaldehyde			37% formaldehyde by weight in methanol. Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.
Heptane		CAS 142-82-5	n-Heptane. Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.
Methanol		CAS 67-56-1	Store at RT. Store formaldehyde, heptane, and methanol waste mixture in a tightly sealed container in fume hood before disposal as per institutional guidelines.
Phosphate buffer Tween (PBTw)			To make 1 L: 100 ml PBS 10X, 890 ml ddH2O, 10 ml Tween 10%. Filter sterilize after adding all components. Store at 4 °C.
Tween 10%			For 50 ml: 5 ml Tween, 5 ml of PBS 10X, 40 ml of ddH2O. Rock to mix. Store at RT.
Bovine serum albumin/phosphate buffer Tween (BBTw)			To make 1 L: 100 ml PBS 10X, 890 ml ddH2O, 10 ml Tween 10%, 1 g Bovine Serum Albumin (BSA). Add BSA then sterilize using a 0.2 μm vacuum filter unit. Store at 4 °C.
Normal goat serum (NGS)	ackson ImmunoResearch Laboratories	005-000-121	To make 10 ml: Normal goat serum 10 ml ddH2O. Add ddH2O to vial of NGS and sterilize using a 0.2 μm syrninge filter. Store aliquots at -20 °C.
1,4-diazabicyclo[2.2.2]octane (DABCO)		CAS: 281-57-9	To make 100 ml: 25 ml ddH2O, 1 ml Tris HCl (1M, pH 7.5), 2.5 g of DABCO solid, 3.5 ml 6N HCl, 250 μl 10N NaOH, 70 ml glycerol. In 250 ml beaker with stir bar, add ddH2O, Tris HCl and DABCO. Stir and then add 6N HCl, 10 N NaOH, and glycerol. Then add 10N NaOH dropwise until solution reaches pH 7.5. Aliquot. Store aliquots at -20 °C.
DABCO + p-phenylenediamine (PPD) solution			1.765 ml NaHCO3, 0.353 Na2CO3, 0.02 g PPD (CAS: 106-50-3). Dissolve PPD in NaHCO3 and NaCO3 solution. Add 60 μl of PPD solution to 500 μl of DABCO. Store aliquots at -20 °C.
Apple juice plates			To make ~200 plates: 45 g agar (CAS#9002-18-0), 45 g granulated sugar (store bought), 500 ml apple juice (store bought), 15 ml Tegosept 10%, 1.5 ml ddH2O. Add agar to ddH2O in 4 L flask then autoclave for 30 min. Mix apple juice and sugar on heated stir plate. Gradually add apple juice mixture to autoclaved agar. Mix on heated stir plate then aliquot 10 ml volumes into 35 mm petri dishes and let stand at RT to solidify. Store at 4 °C.
Tegosept 10%			To make 100 ml: 10 g Tegosept, 100 ml ethanol. Store aliquots at -20 °C.
Yeast paste			~50 g dry active yeast. Gradually add ddH2O to beaker containing yeast while stirring until paste-like consistency reached. Store at 4 °C.
Table 2: Staining Materials
DAPI	Invitrogen	D3571	1:1000, stock at 5 mg/ml.
Rabbit anti-Vasa			1:250, a gift from Ruth Lehmann.
Mouse anti-Fasciclin III	Developmental Studies Hybridoma Bank (DSHB)	7G10	1:10
Mouse anti-1B1	Developmental Studies Hybridoma Bank (DSHB)	1B1	1:4
Guniea pig anti-Traffic Jam			1:2500, a gift from Dorthea Godt (Li et al, 2003).
Mouse anti-Prospero	Developmental Studies Hybridoma Bank (DSHB)	Prospero MR1A	1:10
Rat anti-Elav	Developmental Studies Hybridoma Bank (DSHB)	7EBA10	1:30
mouse anti-Repo	Developmental Studies Hybridoma Bank (DSHB)	8D12	1:10
Goat anti-rabbit Alexa546	Invitrogen	A11010	1:500
Goat anti-mouse Alexa488	Invitrogen	A11029	1:500
Goat anti-guniea pig Alexa633	Invitrogen	A21105	1:500
Goat anti-rat Alexa488	Invitrogen	A11006	1:500
Table 3: Materials and Equipment
Fly Cages	Hand-made; Genesee Scientific Corporation	Not applicable; Bottles: 32-130; Pre-made cage: 59-101	Made by cutting clear cast acrylic tubing (1 3/4 inch in diameter) into 4 inch tall segments with a compound miter saw at 400 rpm. Ultrafine stainless steel screening (was attached to one end of the tub with acrylic compund glue. An alternate method using an empty fly food bottle can be found in Drosophila Protocols ISBN 0-87969-584-4. Cages may also be purchased from the Genesee Scientific Corporation.
Sonicator: Branson 250 Digital Sonifier	Branson	Model: Branson Digital Sonifier 250
Sonicator probe	Branson	Model #: 102C (CE)	EDP: 101-135-066; S/N: OBU06064658
Syringe filter	Nalgene	190-25-20	0.2 μm cellulose, acetate membrane filter
Imaging system: Spinning disc confocal microscope with multichromatic light source, digital CCD camera, and imaging software	Microscope: Olympus, Light source: Lumen Dynamics, Camera: Q-Imaging, Imaging Software: Intelligent Imaging Inc.		Microscope: BX51 equipped with DSU spinning disc, Light source: X-Cite 120Q, Camera: RETIGA-SRV, Imaging Software: Slidebook 5.0
Vacuum filter unit	Nalgene	450-0020	0.2 μm cellulose nitrate membrane filter