We would like to thank Dr. Jennifer Zallen for providing the guinea pig anti-Par-3 primary antibody. This work was supported by generous funding (1R15GM143729-01 and 1P20GM139768-01 5743) from the National Institute of General Medical Science (NIGMS), one of the members of National Institutes of Health (NIH), as well as the Arkansas Biosciences Institute (ABI), which provided partial funding for the purchase of our confocal microscope.

This protocol follows the University of Arkansas (UARK) guidelines for research on invertebrate animals, such as Drosophila melanogaster, and was approved by the UARK Institutional Biosafety Committee (protocol #20001).
1. Drosophila embryo fixation and devitellinization
NOTE: Step 1 describes a procedure (hand-peeling) for the manual removal of the vitelline membrane, a transparent impermeable membrane that surrounds the embryo. Importantly, hand-peeling allows for the selection of properly staged embryos at the start of the ExM protocol, thus greatly enhancing the likelihood of obtaining embryos in a useable orientation at the end of the ExM protocol. However, this ExM protocol is completely compatible with bulk embryo collection and standard procedures for methanol-based removal of the vitelline membrane, in which case one can skip directly to step 2 (immunofluorescence labeling).
<ol>
	<li>Prepare or purchase a number of fine glass needles. The actual dimensions of the needle tip are not critical, but ensure the needles are rigid and sharp enough to pierce the vitelline membranes of fixed embryos. Make needles from glass capillary tubes (1 mm outer diameter, 0.75 mm inner diameter) using a micropipet puller, as one would prepare&#160;needles for embryo microinjections18; alternatively, purchase pre-pulled needles.</li>
	<li>Collect embryos using standard Drosophila techniques19 by placing &#62;100 adult Drosophila in a vented plastic cup sealed with a fruit-juice/agar plate20. Use timed collection windows to enrich for embryos of the proper stage16. For example, to enrich for the gastrulation (stage 6) and convergent-extension (stage 7) stages, change the fruit-juice plate, collect the embryos for 2 h at 25 &#176;C, then remove the plate, and age it for a further 2 h at 25 &#176;C to obtain embryos that are ~2-4 h old.</li>
	<li>To remove the eggshell-like&#160;chorion&#160;from the embryos, cover the surface of the fruit-juice plates with 50% bleach (Table 1), release the embryos from the surface of the agar by agitating them with a small paintbrush, and wait 3 min for the chorion to dissolve.</li>
	<li>Transfer the dechorionated embryos using a paintbrush to a 30 mL scintillation vial containing 4 mL of heptane (organic top phase) and 4 mL of fixation buffer (aqueous bottom phase; Table 1). Freshly dilute the formaldehyde from a freshly prepared or recently opened stock, and mix with 10x PBS and deionized water immediately before adding the embryos. 
	NOTE: Prepare the formaldehyde from paraformaldehyde power or 16% EM-grade formaldehyde in glass ampules. Stocks of concentrated formaldehyde (e.g., 37% formaldehyde) can be used, although the results may be less consistent.</li>
	<li>The embryos will accumulate at the interface between the organic and aqueous phases. Add as many embryos as will form a single layer at the interface. If too many embryos are added to a vial, they will not fix as well.</li>
	<li>Using strong tape, immobilize the scintillation vials on their sides on a tabletop shaker, and agitate them for 20 min at 220 rpm. For optimal fixation, maintain a vigorous emulsion between the organic and aqueous phases during the entire fixation.</li>
	<li>During the fixation time, prepare one of the following for each of the samples.
	<ol>
		<li>Take a plastic 6 cm Petri dish base filled halfway with 3% agar and score a ~5 cm x 3 cm rectangle in the agar with a razor blade or scalpel. Fruit juice/agar plates can also be used for this purpose.</li>
		<li>Using a small lab spatula, remove the agar slab. Invert the base of the Petri dish, and set it on the bench. Place the agar slab on top of the inverted dish (Figure 2A).</li>
		<li>Take the lid of the Petri dish and make sure it is dry. Wearing gloves, place a piece of double-sided tape inside of the lid (the piece of tape should be slightly larger than the agar slab; Figure 2B).</li>
	</ol>
	</li>
	<li>Remove the vials from the shaker, set them upright on the bench, and allow the organic and aqueous phases to separate. Properly fixed embryos will remain at the interface between the two phases.</li>
	<li>Transfer the fixed embryos onto the agar slab using a glass Pasteur pipet fitted with a latex bulb. To prevent the embryos from adhering to the inside of the pipet, try to keep the embryos within the narrow neck of the pipet, and transfer the embryos in multiple small batches rather than all at once. Once all the embryos are on the agar slab, remove most of the residual heptane from around the embryos using a P200 pipettor. Perform this step as quickly as possible (&#60;3 min) to avoid drying out the fixed embryos, which can negatively affect the morphology.</li>
	<li>From a height of ~2 cm, drop the lid with the double-sided tape onto the agar slab to adhere the embryos to the tape (Figure 2C). Gently remove the lid from the agar slab, place it upside down on the bench, and then add enough PBS-Tween (Table 1) to cover the embryos in the lid.</li>
	<li>Using a stereo dissecting microscope at approximately 100x magnification with indirect lighting, identify properly staged embryos using morphological markers. For stage 6 embryos, use markers like a visible cephalic furrow and invaginated mesoderm; for stage 7 embryos, use markers like an extended germband; for stage 11 embryos, use markers like a fully extended germband and visible segmentation along the head-to-tail axis16.
	<ol>
		<li>To collect the desired embryos, first prick the vitelline membrane (a transparent oval membrane around the embryo) near the anterior or posterior end of the embryo with a fine glass needle; the membrane will deflate a bit as the pressure releases. Then, using fine forceps or a metal probe, gently push the embryo on the other end through the hole; the vitelline membrane will remain adhered to the double-sided tape. Leave undesired embryos adhered to the tape.</li>
	</ol>
	</li>
	<li>Periodically collect the floating devitellinized embryos with a glass Pasteur pipet, and move them to a 1.5 mL microfuge tube.</li>
	<li>At this point, perform one of the following steps.
	<ol>
		<li>Continue directly to the immunofluorescence labeling steps. The devitellinized embryos can go directly into blocking solution (step 2.2). Do not allow the embryos to remain in PBS-Tween or blocking solution (Table 1) for longer than 16 h before proceeding to the next step.</li>
		<li>Move the embryos into methanol for storage. Remove as much of the PBS-Tween as possible, and then add 1 mL of methanol. Once the embryos have settled, remove as much methanol as possible, and add 1 mL of fresh methanol. Store the embryos at &#8722;20 &#176;C indefinitely. Methanol storage also allows for the pooling of embryos from multiple collections.</li>
	</ol>
	</li>
</ol>
2. Immunofluorescence labeling
NOTE: Aside from the antibody incubation steps, exact liquid amounts and times are not critical in this section. To perform a rinse or wash, allow the embryos to settle to the bottom of the tube, remove as much liquid as possible without sucking up embryos, and then add ~1 mL of new liquid; use a glass Pasteur pipet fitted with a latex bulb for optimal clarity and control. For the rinse step, the embryos are not rocked, just allowed to settle; for the wash step, the embryos are rocked on a nutator for the indicated amount of time and then allowed to settle.
<ol>
	<li>If embryos were not stored in methanol, proceed to step 2.2. If the embryos were stored in methanol, rinse twice with PBS-Tween, and then wash for 20 min twice with PBS-Tween.</li>
	<li>Wash the embryos for 30-60 min in 1 mL of blocking solution.</li>
	<li>Incubate the embryos with primary antibodies diluted in antibody solution (Table 1) for 2 h at room temperature or preferably overnight at 4 &#176;C. Perform this step in as small a volume as possible (50-300 &#181;L) to conserve the primary antibodies; rocking on a nutator is not strictly required.
	<ol>
		<li>Increase the amount of primary antibody used in a typical immunofluorescence experiment by at least 50% for ExM. Use the following primary antibody concentrations: 1:200 for anti-Par-3 guinea pig polyclonal21 and 1:100 for anti-GFP rabbit polyclonal.</li>
	</ol>
	</li>
	<li>Remove the primary antibody solution (save at 4 &#176;C if desired), rinse twice with PBS-Tween, and then wash for 15 min four times with PBS-Tween.</li>
	<li>Incubate the embryos with fluorescent secondary antibodies in a final volume of 300 &#181;L (diluted in antibody solution) for 1 h at room temperature on a nutator. Fluorescently labeled streptavidin can be added during this step. From this step onward, protect the embryos from excessive and prolonged light exposure when possible, for example by covering the tubes with an opaque box lid or keeping the samples in a drawer.
	<ol>
		<li>Use the following concentrations: 1:500 for anti-rabbit IgG goat polyclonal fused to Alexa Fluor 488; 1:500 for anti-guinea pig IgG goat polyclonal fused to Alexa Fluor 568; and 1:1000 for streptavidin-Alexa Fluor 488.</li>
	</ol>
	</li>
	<li>Remove and dispose of the secondary antibody solution. Rinse the embryos twice with PBS-Tween, and wash for 15 min four times with PBS-Tween.</li>
	<li>At this point, the embryos can be stored at 4 &#176;C in the dark but process the samples as quickly as possible (&#60;24 h).</li>
</ol>
3. Preparing PDMS wells
NOTE: The PDMS wells can be made up to 2 weeks in advance.
<ol>
	<li>Set an incubator or hot plate to 55 &#176;C, and set a centrifuge that can spin conical tubes to 15 &#176;C.</li>
	<li>To prepare the PDMS solution (Table 1), place a 50 mL conical tube in a secondary container on a scale, and add 10 g of silicone elastomer base to the tube using a syringe. Then, add 1 g of silicone elastomer curing agent, and invert the tube several times to mix.</li>
	<li>Create a balance tube by adding an appropriate amount of water to a second 50 mL conical tube. Centrifuge the PDMS solution at 500 x g for 3 min at 15 &#176;C, and then pour it into a 10 cm Petri dish to a depth of ~1 mm. If necessary, remove bubbles by blowing gently on the solution with an air hose. Allow the PDMS solution to solidify overnight at 55 &#176;C.</li>
	<li>Once the PDMS slab is solidified, using a scalpel, score square areas that are slightly smaller than a 22 mm x 22 mm coverslip. Inside each square, score and remove an ~8 mm-wide square well.</li>
	<li>Transfer each square PDMS well onto a 22 mm x 22 mm coverslip, and firmly adhere it (Figure 2D). Preparing six or more coverslips should yield a good number of expanded embryos to image.</li>
</ol>
4. Adhering the embryos to the coverslips
<ol>
	<li>Apply enough 0.1% poly-L-lysine to cover the coverslip surface inside of each well (~50 &#181;L), and place them in a 55 &#176;C incubator to air dry. Repeat this step to increase adhesiveness.</li>
	<li>Briefly rinse the embryos once in 1x PBS to remove the Tween detergent, and then transfer &#62;10 embryos into each of the poly-L-lysine-coated wells.</li>
	<li>Allow the embryos to settle to the bottom of the wells. Remove the excess liquid from the adhered embryos using a Pasteur pipet. Immediately proceed to the next step.</li>
</ol>
5. Activation and gelation
NOTE: Activation refers to the addition of MA-NHS to the embryos, which will modify the sample proteins and antibodies so they can bind to the hydrogel. Gelation refers to the generation of a hydrogel in and around the embryos in each well. During gelation, the embryos are permeated with a monomer solution and then treated with a gelation solution to form the hydrogel.
<ol>
	<li>Activate the embryos for 1 h at room temperature by filling the wells with activation solution (1 mM MA-NHS freshly diluted in 1x PBS; Table 1). Change this solution approximately every 10 min over the course of 1 h.</li>
	<li>Rinse the embryos with 1x PBS three times. Incubate the embryos in monomer solution (Table 1) for 45 min at 4 &#176;C.</li>
	<li>While the embryos are sitting in the monomer solution, prepare the gelation solution (Table 1). Preparing ~2 mL of gelation solution is enough to cover the PDMS wells from an entire 10 cm Petri dish. Be sure to add the APS last, as it will initiate polymerization and begin the gelation.
	<ol>
		<li>Dilute the catalytic oxidant freshly from powder (e.g., 1% TEMPO w/v in water). Combine 1,960&#160;&#181;L of monomer solution with 30 &#181;L of 10% TEMED and 10 &#181;L of 1% TEMPO.</li>
		<li>To avoid polymerizing the entire batch of gelation solution at once, work in small batches. Split the gelation solution (without APS) into 125 &#181;L aliquots between the eight tubes of a&#160;PCR strip.</li>
		<li>Remove the monomer solution from the three PDMS wells using a vacuum while being careful not to disrupt the embryos. Add 5 &#181;L of APS to one of the PCR tubes containing gelation solution to initiate polymerization. Quickly distribute the polymerizing gelation solution amongst the wells (~40 &#181;L per well). Repeat this until all of the wells and embryos are covered.</li>
	</ol>
	</li>
	<li>Let the samples gel for 1.5-2.5 h at 37 &#176;C. Agitate the hydrogels every so often to monitor the polymerization. Solidified hydrogels will not wiggle. Thicker hydrogels will take longer to complete polymerization and solidify.</li>
</ol>
6. Digestion and expansion
NOTE: Thicker and larger gels will take longer to expand, and the center of the gels may take several hours to completely expand; this can be sped up by trimming the edges of the gel. As the gels expand, their refractive index will become nearly identical to that of water, and they will become very hard to see.
<ol>
	<li>After gelation is complete, peel the PDMS wells from the coverslip while trying not to disturb the hydrogels. Cut away excess hydrogel material, if desired.</li>
	<li>Transfer the hydrogels (still attached to the coverslips) individually into the wells of a six-well plate (Figure 2E). Note that the hydrogels may slightly expand during digestion.</li>
	<li>Cover the gels completely with digestion buffer (Table 1) for 1 h at 37 &#176;C. In general, 30 mL of digestion buffer is sufficient to cover the gels in a 6-well plate.</li>
	<li>After digestion, transfer each hydrogel individually&#160;into a 6&#160;cm Petri dish by sliding them off the coverslip. It may be necessary to use a second coverslip to dislodge the gel. Fill each Petri dish with deionized water to expand the gel. Change the water three to four times over the course of 1-2 h until the gels are fully expanded (expect an approximate&#160;four-fold increase in width).</li>
</ol>
7. Mounting and imaging
NOTE: Expanded hydrogels are composed almost entirely of water, making them nearly transparent and extremely fragile. The gels can be manipulated using long coverslips to move them around and pick them up. Mount and image only one or two gels at a time, as gels will gradually release water and begin to slide around the coverslip.
<ol>
	<li>Using a Pasteur pipet, remove as much excess water from the Petri dish as possible to minimize the gels moving around when handled.</li>
	<li>Maneuver each expanded gel, with the embryos on the bottom surface, onto a large coverslip (e.g., 24 mm x 40 mm) for imaging.</li>
	<li>Mount each coverslip with gel over the objective of an inverted laser-scanning confocal microscope. Locate properly staged and oriented specimens using the epifluorescence or brightfield microscopy modes on the microscope with low-magnification (5x or 10x) or medium-magnification (20x) air objectives.</li>
	<li>To image at high resolution, switch to a high-magnification (60x, 63x, or 100x) oil- or water-immersion objective. The surface of the embryos must be within the focusing range of the objective (&#60;300 &#181;m from the coverslip for a 63x objective) in order to be imaged.</li>
	<li>Collect data from samples using the laser-scanning confocal mode on the microscope. Be sure to collect non-saturated images with good dynamic range, and use an appropriate number of pixels per image to capture the maximal possible information about the sample23.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64662/64662fig02.jpg" /> 
Figure 2: Manual devitellinization and working with hydrogels. (A)&#160;Cutting an agar slab from an agar/fruit-juice plate. (B) Placing double-sided tape inside the lid of a 6 cm Petri dish. (C)&#160;Adhering embryos to the taped lid. (D)&#160;A PDMS slab with a square well adhered to a 22 mm x 22 mm coverslip. (E) Coverslip with a PDMS well inside a 6-well plate. <a href="https://www.jove.com/files/ftp_upload/64662/64662fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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<li>Stelzer, E. H. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Light+sheet+fluorescence+microscopy%5BTitle%5D)+AND+%22Nature+Reviews+Methods+Primers%22%5BJournal%5D)">Light sheet fluorescence microscopy.</a> Nature Reviews Methods Primers. 1, 73(2021).</li>
<li>Ku, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Multiplexed+and+scalable+super-resolution+imaging+of+three-dimensional+protein+localization+in+size-adjustable+tissues%5BTitle%5D)+AND+%22Nature+Biotechnology%22%5BJournal%5D)">Multiplexed and scalable super-resolution imaging of three-dimensional protein localization in size-adjustable tissues.</a> Nature Biotechnology. 34 (9), 973-981 (2016).</li>
<li>Wen, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+Universal+labeling+strategy+for+nucleic+acids+in+expansion+microscopy%5BTitle%5D)+AND+%22Journal+of+the+American+Chemical+Society%22%5BJournal%5D)">A Universal labeling strategy for nucleic acids in expansion microscopy.</a> Journal of the American Chemical Society. 143 (34), 13782-13789 (2021).</li>
<li>M'Saad, O., Bewersdorf, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Light+microscopy+of+proteins+in+their+ultrastructural+context%5BTitle%5D)+AND+%22Nature+Communications%22%5BJournal%5D)">Light microscopy of proteins in their ultrastructural context.</a> Nature Communications. 11 (1), 3850(2020).</li>
<li>Sarkar, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Expansion+revealing%3A+Decrowding+proteins+to+unmask+invisible+brain+nanostructures%5BTitle%5D)+AND+%22bioRxiv%22%5BJournal%5D)">Expansion revealing: Decrowding proteins to unmask invisible brain nanostructures.</a> bioRxiv. , (2020).</li>
</ol>

The authors have no conflicts of interest to declare.

Manual devitellinization 
Most Drosophila embryo fixation protocols involve removing the vitelline membrane by shaking fixed embryos in an emulsion of methanol and heptane, which causes the membranes to burst off via osmotic rupture26. While methanol-based devitellinization (methanol popping) is effective and appropriate for many applications, manual devitellinization (hand-peeling) offers some significant advantages. First, hand-peeling allows one to choose precisely staged embryos to devitellinize and collect, greatly increasing the likelihood of obtaining expanded embryos in a useable orientation at the end of the experiment.&#160;This enrichment is critical when studying specific aspects of rapid developmental processes (e.g., mesoderm invagination or convergent extension), for which appropriately staged embryos may represent only a few percent of all embryos, even within a tightly timed collection window. Of course, for many applications, the more traditional bulk methanol popping of embryos from a timed collection window will be sufficient, and hand-peeling may not be worth the extra effort. Second, the binding of certain primary antibodies and dyes is negatively affected by previous exposure of the sample to methanol. For this reason, hand-peeling can yield significant increases in the immunofluorescence signal quality compared with methanol-popped samples, making it a useful general technique for Drosophila developmental biologists.
High-resolution confocal microscopy in expanded whole-mount Drosophila embryos 
While performing high-resolution confocal microscopy on expanded samples is conceptually the same as on unexpanded samples, ExM does introduce some technical hurdles. Notably, embryo orientation, which is random, becomes even more important as the sample size increases, because high-magnification, high-NA objectives are only able to focus light from sample regions that are very close to the coverslip27. Therefore, it is usually only possible to focus on the cells at or near the surface of the embryo that ended up adjacent to the coverslip when the gel was formed. The best way to ensure there are specimens of the correct orientation at the end is to start the ExM protocol with a tightly staged collection of fixed embryos (e.g., by using hand-peeling) and to seed many embryos in each well (&#62;10). To visualize cells deep in the interior of the embryo, it may be necessary to utilize more specialized imaging setups, such as light-sheet microscopy28. Additionally, we find that image quality can be improved by opening the confocal pinhole to a size greater than one airy unit. Of course, an increased pinhole size will come at the cost of decreased maximal resolution, but in practice, even small increases in pinhole size can significantly boost the signal intensity (data not shown). Future studies should systematically address pinhole size and effective resolution in ExM samples.
Variations on basic ExM 
The protocol described here is a relatively simple example of ExM that should work for many applications and be easy to implement in most developmental biology labs. However, there are numerous variations on the basic concept of ExM4,5,7 that can be used to increase the signal intensity, achieve even further degrees of expansion, and detect nucleic acid molecules as well as proteins. In this protocol, the embryos are incubated with antibodies prior to gelation and expansion. Alternatively, the samples can be treated with antibodies after they are expanded6,30, which can increase signal intensity due to increased epitope accessibility and decreased loss of bound antibodies during the expansion steps. In addition, specific crosslinker molecules can be used to attach RNA molecules to the hydrogel to allow the detection of RNA in expanded gels using the hybridization chain reaction method30. Finally, the samples can be subjected to multiple rounds of expansion, as in iterative expansion microscopy (iExM)31, pan-ExM32, and expansion revealing (ExR)31, to achieve even higher degrees of increased resolution.

In cell and developmental biology, seeing is believing, and the ability to accurately determine the localization patterns of proteins is fundamental to many types of experiments. Laser-scanning confocal microscopy is the standard tool for imaging fluorescently labeled proteins in three dimensions within intact samples. Conventional confocal microscopes are incapable of distinguishing (resolving) adjacent fluorescent signals that are separated by less than one-half of the wavelength of the light they emit1. In other words, two point sources must be separated by at least 200-300 nm in the lateral direction (500-700 nm in the axial direction) to resolve them as two distinct signals. This technical barrier is known as the diffraction limit, and it is a fundamental hurdle to studies of complex subcellular structures (e.g., the actomyosin cytoskeletal or mitochondrial networks) with spatial features below the diffraction limit. Therefore, techniques for increasing the resolving power of conventional confocal microscopes are of general interest to the biological community.
To sidestep the diffraction limit, a number of different super-resolution microscopy technologies have been developed that allow for resolution in the order of tens of nanometers or less1,2,3, revealing a world of biological complexity that was previously only accessible via electron microscopy. Despite the obvious advantages of these hardware-based methods, super-resolution microscopes often have specific sample labeling requirements and long acquisition times, limiting their flexibility, or they may simply be too expensive for some labs to access. An alternative to microscope-based super-resolution is expansion microscopy (ExM), which is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules4. The isotropically expanded samples can then be observed at an effectively increased resolution using a traditional fluorescence confocal microscope. ExM was first described by the Boyden group in 20155, and the basic technique has since been adapted for use in a variety of experiments6,7,8. ExM has also been adapted for use in whole-mount embryos, notably in Drosophila9,10,11, C. elegans12, and zebrafish13, making it a powerful tool for developmental biologists.
ExM is based on two different hydrogel chemistries: 1) swellable polyelectrolyte hydrogels, which greatly increase in size when soaked in water14, and 2) polyacrylamide hydrogels, which have extremely small polymer spacing to allow for isotropic sample expansion15. While there are many published ExM protocols, they generally share the following steps: sample fixation, labeling, activation, gelation, digestion, and expansion4. The fixation conditions and fluorescence labeling strategies will of course vary based on the needs of the experiment and system, and in some protocols, labeling occurs after expansion. The target molecules in the sample must be primed (activated) for binding to the hydrogel, which can be achieved using different chemistries4. During the gelation steps, the sample is saturated with monomers of the future hydrogel (sodium acrylate, acrylamide, and the cross-linker bisacrylamide), and the hydrogel is then formed by free-radical polymerization catalyzed by an initiator, such as ammonium persulfate (APS), and an accelerator, such as tetramethylenediamine (TEMED)4. After gelation, the sample is enzymatically digested to homogenize sample resistance to swelling and ensure the isotropic expansion of the hydrogel4. Finally, the digested hydrogel is placed in water, which causes it to expand to approximately four times its original linear size4.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64662/64662fig01v2.jpg" /> 
Figure 1: Overview of expansion microscopy in Drosophila embryos. ExM is a multi-step protocol that takes at least 4 days to complete. Embryo collection, fixation, and devitellinization take 1 day or more depending on whether embryos from multiple collections are pooled. Immunofluorescence labeling takes 1 day or 2 days depending on whether the embryos are incubated overnight with the primary antibodies. Embryo activation, gelation, digestion, and expansion can be performed in a single day. The gels can be mounted and imaged immediately after expansion, although for practical reasons it is often desirable to start imaging the following day. <a href="https://www.jove.com/files/ftp_upload/64662/64662fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
This protocol describes how to perform ExM on whole-mount early- to mid-stage Drosophila embryos16 to visualize subcellular protein localization patterns at super-resolution (Figure 1). This method uses methylacrylic acid N-hydroxysuccinimidyl ester (MA-NHS) chemistry to activate and anchor protein molecules to the hydrogel17, and it is a modification of a previously published ExM protocol for use in late-stage Drosophila embryos and tissues11. This protocol uses polydimethylsiloxane (PDMS) wells to mold the hydrogels and facilitate solution exchange during activation and gelation. An alternative method that does not require the creation of PDMS wells involves lowering embryos attached to coverslips into drops of monomer solution sitting on a piece of laboratory sealing film22. In addition, this protocol describes a method for manually removing the impermeable vitelline membrane that surrounds Drosophila embryos, which is a prerequisite for immunofluorescence staining. Importantly, this method of hand-peeling embryos can be used to select only properly staged Drosophila embryos prior to sample labeling, which greatly increases the likelihood of ending up with expanded samples of the correct stage and orientation and, thus, makes the downstream data collection much more efficient.

<table><tbody><tr><td>acrylamide</td><td>Milipore Sigma</td><td>1490-100ML</td><td></td></tr><tr><td>ammonium persulfate</td><td>VWR</td><td>BDH9214-500G</td><td></td></tr><tr><td>anti-GFP rabbit polyclonal antibody</td><td>Torrey Pines BioLabs</td><td>TP-401</td><td></td></tr><tr><td>anti-guinea pig IgG goat polyclonal antibody, Alexa Fluor 568</td><td>Thermo Fisher Scientific</td><td>A-11075</td><td></td></tr><tr><td>anti-rabbit IgG goat polyclonal antibody, Alexa Fluor 488</td><td>Thermo Fisher Scientific</td><td>A-11008</td><td></td></tr><tr><td>bisacrylamide</td><td>Research Products International</td><td>A11275</td><td></td></tr><tr><td>bovine serum albumin (30% solution)</td><td>Millipore Sigma</td><td>A7284</td><td></td></tr><tr><td>conical tubes, 50 mL</td><td>fisherscientific&amp;nbsp;</td><td>21008-940</td><td></td></tr><tr><td>coverlip glass, square 22 mm</td><td>VWR</td><td>48366-227</td><td></td></tr><tr><td>coverslip glass, rectangular 40 mm x 24 mm</td><td>VWR</td><td>48393-230</td><td></td></tr><tr><td>glass capillaries for pulling needles</td><td>World Precision Instruments</td><td>TW100F-4</td><td></td></tr><tr><td>glass microinjection needles (pre-pulled)</td><td>World Precision Instruments</td><td>TIP10LT</td><td></td></tr><tr><td>guanidine HCl</td><td>VWR</td><td>101970-606</td><td></td></tr><tr><td>heptane</td><td>VWR</td><td>EM-HX0078-1</td><td></td></tr><tr><td>latex pipet bulbs</td><td>VWR</td><td>82024-554</td><td></td></tr><tr><td>methanol</td><td>VWR</td><td>BDH1135-4LP</td><td></td></tr><tr><td>methylacylic acid N-hydroxysuccinimidyl ester</td><td>VWR</td><td>730300-1G</td><td></td></tr><tr><td>microfuge tube, 1.5 mL</td><td>VWR</td><td>20170-038</td><td></td></tr><tr><td>multi-well plate, 6-well</td><td>Genesee</td><td>25-100</td><td></td></tr><tr><td>paraformaldehyde (16%, EM-grade, methanol-free)</td><td>Electron Microscopy Sciences</td><td>509804487 (Fisher)</td><td></td></tr><tr><td>Pasteur pipet (2 mL, short tip)</td><td>VWR</td><td>14673-010</td><td></td></tr><tr><td>PDMS kit (Sylgard 184 Kit, base and curing agent)</td><td>VWR</td><td>102092-312</td><td></td></tr><tr><td>Petri plates</td><td>Genesee</td><td>32-107</td><td></td></tr><tr><td>phosphate-buffered saline (10x solution)</td><td>VWR</td><td>97063-660</td><td></td></tr><tr><td>Poly-L-lysine solution (0.1% solution)</td><td>VWR</td><td>P8920-1ooML</td><td></td></tr><tr><td>Proteinase K</td><td>Thermo Fisher Scientific</td><td>E00491</td><td></td></tr><tr><td>scintillation vials (30 mL)</td><td>VWR</td><td>66022-128</td><td></td></tr><tr><td>sodium acrylate</td><td>VWR</td><td>101181-226</td><td></td></tr><tr><td>sodium azide (powder)</td><td>Millipore Sigma</td><td>71289</td><td>make a 1% w/v working stock; acute POISON at this concentration!</td></tr><tr><td>Streptavidin, Alexa Fluor 488</td><td>Thermo Fisher Scientific</td><td>S32354</td><td></td></tr><tr><td>TAE (50x)</td><td>VWR</td><td>97063-692</td><td></td></tr><tr><td>tape (double-sided, 1 inch wide)</td><td>Scotch&amp;nbsp; 3M</td><td>665 Scotch double sided 1inch/1296 inches Boxed</td><td></td></tr><tr><td>TEMED</td><td>Thermo Fisher Scientific</td><td>PI17919</td><td></td></tr><tr><td>TEMPO</td><td>VWR</td><td>EM8.14681.0005</td><td>catalytic oxidant</td></tr><tr><td>Tween-20</td><td>VWR</td><td>97063-872</td><td>extremely viscous when pure; make a 10% working stock with water</td></tr><tr><td>Zeiss LSM 900</td><td>Zeiss</td><td></td><td>Laser scanning microscope used without AiryScan</td></tr></tbody></table>

using expansion microscopy to physically enlarge whole-mount drosophila embryos for super-resolution imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged molecules within complex biological samples. While traditional confocal microscopes allow one to resolve two adjacent fluorescent point sources located a few hundred nanometers apart, observing the finer details of subcellular biology requires the ability to resolve signals in the order of tens of nanometers. Numerous hardware-based methods for super-resolution microscopy have been developed to allow researchers to sidestep such resolution limits, although these methods require specialized microscopes that are not available to all researchers. An alternative method for increasing resolving power is to isotropically enlarge the sample itself through a process known as expansion microscopy (ExM), which was first described by the Boyden group in 2015. ExM is not a type of microscopy per se but is rather a method for swelling a sample while preserving the relative spatial organization of its constituent molecules. The expanded sample can then be observed at an effectively increased resolution using a traditional confocal microscope. Here, we describe a protocol for implementing ExM in whole-mount Drosophila embryos, which is used to examine the localization of Par-3, myosin II, and mitochondria within the surface epithelial cells. This protocol yields an approximately four-fold increase in sample size, allowing for the detection of subcellular details that are not visible with conventional confocal microscopy. As proof of principle, an anti-GFP antibody is used to distinguish distinct pools of myosin-GFP between adjacent cell cortices, and fluorescently labeled streptavidin is used to detect endogenous biotinylated molecules to reveal the fine details of the mitochondrial network architecture. This protocol utilizes common antibodies and reagents for fluorescence labeling, and it should be compatible with many existing immunofluorescence protocols.

To characterize the general efficacy of ExM in whole-mount Drosophila embryos, embryo length along the head-to-tail axis was measured in unexpanded control embryos versus expanded embryos (Figure 3A-C). The unexpanded control embryos were subjected to the same fixation conditions and immunofluorescence labeling steps as the expanded embryos, except they were mounted using a solidified mounting medium prior to imaging. The individual unexpanded embryos spanned approximately one-half of a field of view when using a 10x objective (Figure 3A). By contrast, the expanded embryos spanned approximately two full fields of view when using the same 10x objective (Figure 3B). To assess how the degree of expansion varied both within and between experiments, the same ExM protocol was performed on three separate occasions, and embryo length was measured in three different gels within each individual experiment. The average head-to-tail length of the unexpanded control embryos was 398.8 &#181;m (standard deviation [SD] = 22.93 &#181;m; n = 74; Figure 3C). For experiment 1, experiment 2, and experiment 3, the average embryo lengths were 1,596 &#181;m (SD = 159.9 &#181;m; n = 57), 1,868 &#181;m (SD = 150.5 &#181;m; n = 51), and 1,954 &#181;m (SD = 120.3 &#181;m; n = 44), respectively, representing expansion factors of 4.0-fold, 4.7-fold, and 4.9-fold, respectively (Figure 3C). The intra-experimental variation between the gels was much less noticeable than the inter-experimental variation, which was approximately 20% (Figure 3C). To assess the effects of ExM on cell and embryo morphology, an antibody against the adherens junction component Par-3 (Bazooka)21 was used to label the apical cell membranes, and we imaged the developing mouth segments of stage 11 Drosophila embryos&#8212;a stage with a complex segmented structure (Figure 3D-F). In the control sample, the cells in the maxillary segment had an average width of 4.76 &#181;m (SD = 1.053 &#181;m, n = 25; Figure 3D,F). In the expanded samples imaged using the same 40x objective and zoom factor (1x), the cells in the maxillary segment had an average width of 19.10 &#181;m (SD = 3.966 &#181;m, n = 18; Figure 3E,F), representing a 4.0-fold expansion. Therefore, consistent with previous reports11, we were able to expand whole-mount Drosophila embryos approximately four-fold in linear dimensions using ExM without sample tearing or obvious distortions in the cellular or tissue morphology.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64662/64662fig03.jpg" /> 
Figure 3: Four-fold expansion of Drosophila embryos. (A) Unexpanded and (B) expanded Drosophila embryos imaged using a 10x objective (0.3 NA) at 1x zoom. Individual fields of view (FOV) are indicated with dashed lines. The embryos expressed a GFP-tagged version of myosin light chain and were stained with an anti-GFP antibody. (C)&#160;Quantification of embryo length (along the head-to-tail axis) in three hydrogels per experiment and from three separate ExM experiments compared with unexpanded controls. (D,E)&#160;Maxillary segments from (D) unexpanded and (E) expanded&#160;stage 11 Drosophila embryos imaged using a 40x objective (1.3 NA) at 1x zoom. The cell outlines (adherens junctions) were detected with an anti Par-3/Bazooka antibody (white). (F)&#160;Quantification of the cell width (long axis) from equivalent groups of cells from (D) and (E). The box plots in (C) and (F) show the 25th, 50th, and 75th percentile ranges; the whiskers indicate the minimum and maximum values; the &#34;+&#34; symbols indicate the mean. <a href="https://www.jove.com/files/ftp_upload/64662/64662fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
To demonstrate that ExM can be used to resolve subcellular details below the typical diffraction limit, the actomyosin cytoskeleton was imaged in unexpanded control versus expanded embryos undergoing convergent extension (stage 7). The tissue remodeling events of gastrulation and convergent extension are largely controlled by changes in the localization of the motor protein myosin II24. However, in the densely packed columnar epithelium of the early Drosophila ectoderm, it is difficult to observe many fine details of the myosin II localization pattern, even when imaged at 158x magnification (63x objective with a 2.5x optical zoom)&#8212;a typical maximal resolving power for a laser-scanning confocal microscope. For example, because myosin II is a cortical protein (located directly beneath the plasma membrane), pools of myosin II25 located on either side of cell-cell contacts were not resolvable in stage 7 embryos, and they appeared as a single line where neighboring cells met (Figure 4A). By contrast, in expanded stage 7 embryos, parallel lines of myosin II could be observed at cell-cell junctions, representing cortical protein pools in adjacent cells (Figure 4B). The distance between parallel myosin II lines in expanded samples was 892.7 nm (SD = 0.171 nm, n = 12); when divided by four, this yields a predicted distance of ~220 nm between the myosin lines in adjacent cells in unexpanded embryos, which is indeed just below the diffraction limit for a signal detected with Alexa 488 (peak emission of ~520 nm/2 = 260 nm).
In addition, we also tested whether ExM could be used to resolve the mitochondrial network architecture in densely packed cells of gastrulating Drosophila embryos (stage 6). Mitochondrial function is closely linked to network structure (i.e., fused vs. fragmented organelles), but the details of mitochondrial network organization are hard to visualize using conventional confocal microscopy in cell types that are not flat and/or thin. Mitochondria are naturally rich in biotinylated molecules, and, thus, mitochondria can be labeled in the early Drosophila embryo using fluorescently labeled streptavidin26. In the unexpanded stage 6 embryos labeled with streptavidin-Alexa 488, the signal appeared as cytoplasmic puncta that were often overlapping and difficult to resolve (Figure 4C). By contrast, in the expanded stage 6 embryos, many more fine details of the mitochondrial network were visible and puncta were more easily resolvable (Figure 4D)26, 27. These results indicate that ExM can be used to study mitochondrial network organization in cell types not traditionally suited for mitochondrial analysis.
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64662/64662fig4v2.jpg" /> 
Figure 4: Details of actomyosin cytoskeleton and mitochondria revealed by expansion microscopy. (A,B)&#160;Myosin II localization in neuroectoderm (germband) cells imaged with a 63x objective (1.4 NA) at 2.5x zoom in stage 7 (A) unexpanded and (B) expanded embryos. Myosin II was detected in the embryos expressing a transgenic GFP-tagged version of the myosin II regulatory light chain (sqh-GFP), which was detected with an anti-GFP antibody (red). Distinct pools of cortical myosin located in adjacent cells can be resolved in the expanded embryo (white arrows). (C,D) Mitochondrial networks in neuroectoderm cells imaged with a 63&#215; objective (1.4 NA) at 2.5x zoom in stage 6 unexpanded (C) and expanded (D) embryos. The mitochondria were detected with streptavidin-Alexa 488 (green), and the cell outlines were detected with an anti-Par-3/Bazooka antibody (magenta). The experiments were performed with a laser-scanning confocal microscope. <a href="https://www.jove.com/files/ftp_upload/64662/64662fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Table 1: Solution recipes. Composition for the solutions used in this protocol in order of appearance. All the stocks are liquids unless otherwise noted. The chemicals were resuspended or diluted in autoclaved filtered water unless otherwise noted. <a href="https://www.jove.com/files/ftp_upload/64662/JOVE_Parveen 2023_Revision_Recipes.zip" target="_blank">Please click here to download this Table.</a>

Watch this Scientific Journal Video about Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging at JoVE.com

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Here, a protocol for the implementation of expansion microscopy in early Drosophila embryos to achieve super-resolution imaging using a conventional laser-scanning confocal microscope is presented.

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

The workhorse of developmental biology is the confocal microscope, which allows researchers to determine the three-dimensional localization of tagged ...

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Research

JoVE Journal

Developmental Biology

1.6K Views.  University of Arkansas. Dit protocol kan worden geïmplementeerd in de meeste laboratoria voor ontwikkelingsbiologie van de slokdarm, waardoor onderzoekers die geen toegang hebben tot superresolutiemicroscopen superresolutiebeelden kunnen produceren. Het belangrijkste voordeel van deze techniek is dat het onderzoekers in staat stelt de diffractielimiet van conventionele confocale microscopie te omzeilen zonder dat een superresolutiemicroscoop nodig is door het monster zelf uit te breiden. Dit protocol zal gunstig zi...

Video: Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are expressed in spatial relation to each other in situ. Multi-target chromogenic whole-mount in situ hybridization (MC-WISH) greatly facilitates the instant comparison of gene expression patterns, as it allows distinctive visualization of different mRNA species in contrasting colors in the same sample specimen. This provides the possibility to relate gene expression domains topographically to each other with high accuracy and to define unique and overlapping expression sites. In the presented protocol, we describe a MC-WISH procedure for comparing mRNA expression patterns of different genes in Drosophila embryos. Up to three RNA probes, each specific for another gene and labeled by a different hapten, are simultaneously hybridized to the embryo samples and subsequently detected by alkaline phosphatase-based colorimetric immunohistochemistry. The described procedure is detailed here for Drosophila, but works equally well with zebrafish embryos.

Multi-target Chromogenic Whole-mount In Situ Hybridization for Comparing Gene Expression Domains in Drosophila Embryos

We describe a multi-target chromogenic whole-mount in situ hybridization (MC-WISH) procedure in intact Drosophila embryos allowing simultaneous and specific detection of three different mRNA distribution patterns by contrasting color precipitates.

To analyze gene regulatory networks active during embryonic development and organogenesis it is essential to precisely define how the different genes are ...

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Biology

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility in Drosophila

Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at multiple length scales, such as cell division, cell migration, epithelial folding, and branching morphogenesis. An in-depth understanding of the role of actomyosin contractility in morphogenesis requires approaches that allow the rapid inactivation of actomyosin, which is difficult to achieve using conventional genetic or pharmacological approaches. The presented protocol demonstrates the use of a CRY2-CIBN based optogenetic dimerization system, Opto-Rho1DN, to inhibit actomyosin contractility in Drosophila embryos with precise temporal and spatial controls. In this system, CRY2 is fused to the dominant negative form of Rho1 (Rho1DN), whereas CIBN is anchored to the plasma membrane. Blue light-mediated dimerization of CRY2 and CIBN results in rapid translocation of Rho1DN from the cytoplasm to the plasma membrane, where it inactivates actomyosin by inhibiting endogenous Rho1. In addition, this article presents a detailed protocol for coupling Opto-Rho1DN-mediated inactivation of actomyosin with laser ablation to investigate the role of actomyosin in generating epithelial tension during Drosophila ventral furrow formation. This protocol can be applied to many other morphological processes that involve actomyosin contractility in Drosophila embryos with minimal modifications. Overall, this optogenetic tool is a powerful approach to dissect the function of actomyosin contractility in controlling tissue mechanics during dynamic tissue remodeling.

Author Spotlight: Optogenetic Inhibition of Rho1-Mediated Actomyosin Contractility Coupled with Measurement of Epithelial Tension in Drosophila Embryos  

Actomyosin contractility plays an important role in cell and tissue morphogenesis. However, it is challenging to manipulate actomyosin contractility in vivo acutely. This protocol describes an optogenetic system that rapidly inhibits Rho1-mediated actomyosin contractility in Drosophila embryos, revealing the immediate loss of epithelial tension after the inactivation of actomyosin in vivo.

Contractile forces generated by actin and non-muscle myosin II ("actomyosin contractility") are critical for morphological changes of cells and tissues at ...

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

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

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&#160;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.

Author Spotlight: Unveiling the Dynamics of Mechanical and Biochemical Signals in Animal Morphogenesis Research 

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.

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Multiplex Detection of Gene Expression in the Intact Drosophila Brain Using Expansion-Assisted Iterative Fluorescence In Situ Hybridization

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ within a given tissue remain limited. The EASI-FISH protocol described here has been adapted to detect the expression of dozens of genes in the intact adult Drosophila central nervous system (CNS) using commercially available reagents. This protocol includes a new gel formulation that enhances gel robustness, enabling multiple rounds of hybridization and allowing the embedding of multiple brains per gel. This improvement increases throughput, facilitates optimal comparison of experimental conditions, and reduces reagent costs. Additionally, by employing the GAL4-UAS system for co-detection of green fluorescent protein (GFP), gene expression can be visualized in specific neuronal or glial cell types. Notably, the high resolution achieved through expansion microscopy, combined with the sensitivity of the method, allows for the detection of single RNA transcripts. This approach effectively integrates high image quality with high throughput, making it a powerful tool for studying gene expression throughout the intact fly brain.

Expansion-Assisted Iterative Fluorescence In Situ Hybridization (EASI-FISH) is a robust technique that combines expansion microscopy with fluorescence in situ hybridization (FISH) to detect the expression of multiple genes in thick tissue. This protocol outlines recent advancements in the method and its adaptation for labeling the intact Drosophila central nervous system..

Understanding gene expression is essential for deciphering cellular functions. However, methods for analyzing the expression of numerous genes in situ ...

Neuroscience

Super-Resolution Live Cell Imaging of Subcellular Structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

Using Expansion Microscopy to Physically Enlarge Whole-Mount Drosophila Embryos for Super-Resolution Imaging

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