Visualizing Low-Abundance Proteins and Post-Translational Modifications in Living Drosophila Embryos via Fluorescent Antibody Injection

Summary

This protocol describes the customized antibody-based fluorescence labeling and injection into early Drosophila embryos to enable live imaging of low-abundance proteins or post-translational modifications that are challenging to detect using traditional GFP/mCherry-tag approaches.

Abstract

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

Introduction

Immunofluorescence is a cornerstone technique of modern cell biology originally developed by Albert Coons, which enables the detection of molecules at their native cellular compartments and characterization of the molecular compositions of subcellular organelles or machineries¹. Coupled with genetic manipulations, immunofluorescence helps establish the now well-accepted concept that protein localization is essential for its function². Aside from specific primary antibodies and bright fluorescent dyes, the success of this technique relies on a preliminary process named fixation and permeabilization, which preserves cellular morphologies, immobilizes antigens, and increases the accessibility of antibodies into intracellular compartments. Inevitably, the fixation and permeabilization process would kill cells and terminate all biological processes³. Therefore, immunofluorescence only provides snapshots of the life journey of proteins. However, many biological processes such as cell migration and divisions are dynamic in nature, requiring investigation of protein behaviors in a spatial-temporally resolved manner^4,5.

To examine protein dynamics in living organisms, live imaging methods based on genetically encoded fluorescent proteins such as green fluorescent protein (GFP)⁶ and high-speed confocal microscopes have been developed. Briefly, the protein of interest can be genetically manipulated to be fused with GFP⁷, and then ectopically expressed from viral or yeast promoters such as cytomegalovirus (CMV)⁸ or upstream activation sequence (UAS)⁹. Because GFP is autofluorescent in nature, no fluorophore-coupled antibodies are required to reveal the localization of target proteins, which bypasses the necessity of preliminary processes of fixation or permeabilization. Over the last two decades, fluorescent tags spanning the whole spectrum of wavelength have been developed¹⁰, enabling multi-color live imaging of several target proteins at the same time. However, compared to chemically engineered fluorescent dyes such as AlexaFluor or ATTO, the autofluorescence of these genetically encoded fluorescent proteins is relatively weak and unstable when expressed from endogenous promoters, especially during live imaging over longer time scales¹⁰. While this shortfall can be mitigated by over-expressing fluorescently tagged target proteins, many with enzymatic activities such as kinases and phosphatases severely disrupt normal biological processes if not expressed at physiological levels.

This protocol presents a method that enables photostable antibody-based target illumination in a live image setup, essentially allowing immunofluorescence without the process of fixation or permeabilization (Figure 1). Through a simple NHS-based primary amine reaction¹¹, one can conjugate fluorescent dyes such as AlexaFluor 488 or 594 with essentially any primary antibody or GFP/HA/Myc nanobody¹². Taking advantage of a developmental feature that all Drosophila embryonic cells share a common cytoplasm during the syncytium stage¹³, one can achieve antigen binding and illumination across entire embryos after the injection of dye-conjugated antibodies. With expanding libraries of endogenously tagged proteins available in Drosophila and other model systems¹⁴, this method can potentially broaden applications of these libraries by revealing dynamics of low-abundance fluorescently tagged proteins and other non-fluorescently tagged (HA/Myc-tagged) proteins in living tissues.

Protocol

The experiments were conducted in accordance with the guidelines and approval of the School of Life Sciences, SUSTech University. The organism used is Drosophila melanogaster, and the genotypes are Notch-Knockin-GFP (Chromosome X) and Sqh-sqh-GFP (Chromosome II), generously provided by the labs of Dr. Francois Schweisguth (Institute Pasteur) and Dr. Jennifer Zallen (Sloan Kettering Institute), respectively. While this protocol mainly focuses on aspects of antibody labeling and live imaging, please refer to published reports for more detailed descriptions of Drosophila embryo collection and injection^15,16.

1. Fluorescent labeling of antibodies

1. Preferably, use monoclonal antibodies or nanobodies for the protein of interest. Prepare the stock concentration of antibodies at 1 mg/mL or higher.
   ​NOTE: In this study, GFP nanobody (see Table of Materials) is used as an example. The commercially available GFP nanobody is packaged at a concentration of 1.0 mg/mL and a volume of 250 µL. Additionally, a commercially available antibody labeling kit (see Table of Materials) is utilized, which conjugates Alexa Fluor 594 to primary amines of proteins through a succinimidyl ester reaction¹¹. Importantly, this labeling process does not significantly alter the antibody concentration.
2. Prepare a 1 M sodium bicarbonate solution by resuspending Component B (provided in the antibody labeling kit) in 1 mL of deionized water.
3. Adjust the antibody concentration to 1.0 mg/mL, then add 1/10^th volume (10 µL for 100 µL GFP nanobody) of the 1 M sodium bicarbonate solution.
4. Add 110 µL of the antibody mix (from step 1.3) directly to the tube containing Alexa Fluor 594 dye. Invert to mix (do not vortex) and incubate on a rotator for 1 h at room temperature.
5. Assemble the purification column from the conjugation kit (see Table of Materials). Prepare a 1.5 mL resin bed (provided in the antibody labeling kit) and centrifuge the column at 1100 x g for 3 min at room temperature (RT) to remove excess liquid from the resin.
6. Add the reaction mix (from step 1.4) dropwise to the top of the resin column and centrifuge at 1100 x g for 5 min at 4 °C to collect the labeled antibody. The collected antibody should appear pink in color, and the volume should be slightly less than 100 µL. Store in foil-wrapped or dark-colored tubes at 4 °C.

2. Preparation of Drosophila embryos

1. Place 200 newly hatched adult Drosophila in a plastic cylinder-shaped cage (diameter = 5 cm, height = 8 cm) with a male-to-female ratio of 1: 10. Seal one side with porous metal mesh for ventilation and the other side with a fruit-juice/yeast-paste agar plate.
2. Change the plate every 12 h for three days before embryo collection. This allows synchronization of Drosophila egg laying and improves collection efficiency.
3. On the day of injection, change the cage to a fruit-juice plate with minimal yeast paste and collect embryos at 25 °C for 1 h. If the first round of collection does not yield sufficient embryos (<50 embryos), discard the first plate and repeat this step for a second round of collection until more than 100 embryos are laid within 1 h.
4. Remove the plate from the cage to stop the egg laying and incubate at 25 °C for another 2 h to obtain embryos that are 2-3 h old. The correct stage of embryos is critical for the success of antibody injection.
5. To remove the eggshell of embryos, add 2 mL of 50% bleach to the fruit-juice plate and gently dislodge embryos from the plate using a paintbrush (Figure 2A-4). Often, embryos will be laid directly on top of the yeast paste. In this case, it is acceptable to brush the yeast paste into the bleach solution to collect all the embryos.
6. Incubate the floating embryos in 50% bleach for 2 min and swirl the fruit-juice plate every 30 s to improve the efficiency of eggshell removal.
7. Transfer the embryo/bleach mix by pouring it into a 70 µm nylon cell strainer (Figure 2A-7). Wash the embryos thoroughly using a water bottle to remove any residual bleach and yeast paste. Dry the cell strainer completely with paper towels, ensuring that any excess water around the embryos is removed.

3. Embryo alignment and desiccation

1. Prepare a 5 cm x 5 cm x 0.5 cm (width, length, height) 4% agarose gel (Figure 2A-9) and allow the gel to cool down to room temperature. Cut a 1 cm x 3 cm x 0.5 cm (width, length, height) agarose block using a clean razor blade and place the block on a glass slide (Figure 2A-2). Examine the gel block under the dissecting scope to ensure that the edges are cut straight, and the surface is flat and dry.
2. Transfer embryos from the strainer onto the gel block using a paintbrush. Spread the embryos evenly along the midline of the block by brushing gently. Ideally, clusters of embryos are separated into single ones or doublets without touching each other.
3. Prepare a tweezer with two legs taped tightly together to create a single fine tip (Figure 2A-5). Under a dissecting scope, pick up individual embryos using the tweezer and place them along the long edge of the gel block. Align 20-30 embryos, ensuring their anterior-posterior axis is parallel to the edge (Figure 2C).
4. Pipette 10 µL heptane glue (see Table of Materials) at the center of a 24 mm x 50 mm coverslip (Figure 2A-1) and spread the glue into a 0.5 cm x 3 cm rectangular area using a pipette tip. Wait for the glue to completely dry before use.
5. Place the glass slide with the gel block at the edge of the desk, with embryos facing outward. Lift the coverslip with glue using a flat-tip tweezer (Figure 2A-6) and hold it still on top of the embryos, with the glue side facing toward the embryos at a tilted angle of around 45 degrees.
   NOTE: Gently press the coverslip against the gel block so that the embryos will be in contact with the glue, and then quickly release the pressure and lift the coverslip. The amount of tension applied is critical so that embryos can be stably attached to the glue without being pressed too hard and bursting.
6. Pipette 10 µL water at the center of a new glass slide and place the coverslip with the embryos on top of it, with the side of the embryo facing upwards (Figure 2B). Ensure to use water instead of nail polish or other adhesive glue for attaching the coverslip to the glass slide, as this side of the coverslip will be placed directly on top of the confocal lens for live imaging.
7. Dry the embryos in a desiccation chamber (Figure 2A-3) (see Table of Materials) for 10-15 min until the vitelline membrane wrinkles (Figure 2E). The required time may vary with the humidity of the room and should be experimentally tested for each lab condition.
8. Pipette 40 µL of halocarbon oil (a mix of type 27 and type 700 at a 1: 1 ratio in volume, see Table of Materials) at one end of the embryo strip. Tilt the slide until the halocarbon oil covers the entire surface of the embryos.

4. Antibody Injection and imaging

1. Prepare a few injection glass needles using a micropipette puller (see Table of Materials for parameter settings) and load each needle with 5 µL of AlexaFluor-labeled antibody solution (from step 1.6).
2. Place a 25 mm x 25 mm coverslip on top of a glass slide. Pipette 40 µL of halocarbon oil and spread it along the edge of the coverslip.
3. Under the injection scope, align the tip of the needle against the edge of the coverslip under oil. Adjust the injection air pressure of the picopump (see Table of Materials) so that one pump generates one bubble filled with antibodies. The bubble diameter should be limited to 20-50 µm (Figure 2F).
4. Replace the empty glass slide with one containing embryos under oil. Align the injection needle tip perpendicularly against the anterior-posterior axis of the embryos (Figure 2D,G).
5. Check the stage of the embryos to ensure that most have initiated cellularization but have not yet started gastrulation (stage 4 and stage 5), remaining as a syncytium (Figure 2F,G).
   NOTE: The hallmark of this stage is the absence of any grooves or folds and the presence of an oval-shaped, dark-colored yolk sac visible at the center of the embryo. Morphological characterization of the embryo stage under oil is based on the book chapter "Stages of Drosophila Embryogenesis" by Volker Hartenstein¹⁷.
6. Use the X-Y stage manipulator to move embryos toward the needle until the tip arrives at the center of the yolk. Pump two to three times to inject antibodies into the yolk. Successful injection is indicated by the quick disappearance of the vitelline membrane wrinkle as embryos gain volume from the antibody solution (Figure 2G).
7. Use the X-Y stage manipulator to move embryos away from the needle after injection and move downward to the next embryo. Repeat step 6 until all embryos on the slide are injected.
8. Transfer the glass slide with injected embryos to a humidity chamber (Figure 2A-8). Incubate at 25 °C until the desired stage of embryo development is achieved.
9. Lift the 24 mm x 50 mm coverslip off the glass slide and place it directly into the slide holder of the microscope. The side without embryos will be facing the objectives. Locate the embryos using a low-magnification lens under bright field illumination and switch to a 40x or 63x lens for high-resolution fluorescent live imaging.

Results

To demonstrate the advantages of the antibody injection method over fluorescent-tag-based live imaging or immunofluorescence, two case studies are provided that characterize the dynamic localization of a low-abundance transmembrane receptor, Notch, and a type of post-translational modification called tyrosine phosphorylation in living embryos.

Notch signaling activity plays a major role in cell fate determination during embryogenesis and adult organ homeostasis^18,<...

Discussion

This presented procedure outlines the specialized method of fluorescence labeling with custom antibodies and subsequent injection into early-stage Drosophila embryos. This technique facilitates real-time visualization of proteins or post-translational modifications that exist in low quantities and are typically difficult to observe through conventional GFP/mCherry tagging methods.

Caution should be exercised when extending this method to make quantitative comparisons between wild-type...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Agarose	Sangon Biotech	A620014
Alexa Fluor 594 Antibody Labeling Kit	Invitrogen	A20185	Purification column from step 1.6 is included in this kit
Biological Microscope	SOPTOP	EX20	Eyepiece lens: PL 10X/20. Objective lens: 10x/0.25
Bleach	Clorox®
Borosilicate Glass Capillaries	World Precision Instruments	TW100F-4
Centrifuge	Eppendorf	5245
Cell Strainer	FALCON	352350
Desiccation chamber	LOCK&LOCK	HSM8200	320ml
Dissecting Microscope	Mshot	MZ62	Eyepiece lens: WF10X/22mm.
Double-sided Tape	Scotch	665
Fine Super Tweezer	VETUS	ST-14
Fisherbrand™ Cover Glasses: Rectangles	Fisherbrand	12-545F
Fisherbrand™ Superfrost™ Plus Microscope Slides	Fisherbrand	12-550-15
Forcep	VETUS	33A-SA
Halocarbon oil 27	Sigma-Aldrich	H8773-100ML
Halocarbon oil 700	Sigma-Aldrich	H8898-100ML
Heptane	Sigma-Aldrich	H2198-1L	Heptane glue is made of double-sided tape immersed in heptane
Dehydration reagent	TOKAI	1-7315-01	Fill to 90% volume of the dessication chamber
Manual Micromanipulator	World Precision Instruments	M3301R
Micropipette puller	World Precision Instruments	PUL-1000	Procedure: step 1, Heat: 290, Force:300, Distance:1.00, Delay:50. Step 2, Heat: 290, Force:300, Distance:2.21, Delay:50
Pneumatic picopump	World Precision Instruments	PV 830	Eject: 20 psi;  Range: 100ms; Duration: timed
PY20	Santa Cruz	SC-508
Square petri dishes	Biosharp	BS-100-SD
GFP nanobody	Chromotek	gt