Live Imaging of Epithelial-Mesenchymal Transition in Mesoderm Cells of Gastrulating Drosophila Embryos

Abstract

Epithelial-mesenchymal transitions (EMTs) drive the generation of cell diversity during both evolution and development. More and more evidence has pointed to a model where EMT is not a binary switch but a reversible process that can be stabilized at intermediate states. Despite our vast knowledge on the signaling pathways that trigger EMT, we know very little about how EMT happens in a step-wise manner. Live imaging of cells that are undergoing EMT in intact, living, animals will provide us valuable insights into how EMT is executed at both the cellular and molecular levels and help us identify and understand the intermediate states. Here, we describe how to image early stages of EMT in the mesoderm cells of live Drosophila melanogaster embryos and how to image contractile myosin that suspends EMT progression.

1. Introduction

The epithelial-mesenchymal transition (EMT) is a process where highly organized epithelial cells lose cell polarity and junctions and leave epithelial sheets to become migratory mesenchymal cells. The emergence of EMT allowed the evolution from animals with epithelial cells only, to animals with diverse tissue types [1]. Accordingly, during embryogenesis, the first EMT occurs upon gastrulation to generate three germ layers, which lay out the foundations of different tissues [1]. Despite EMT being first discovered in development, it has also been found to play a central role during tumorigenesis [2, 3]. Recent advances in EMT studies suggest that cells that activate EMT programs often only progress partially along the epithelial-mesenchymal axis, expressing a mixture of epithelial and mesenchymal markers. In addition, EMT can be followed by its reverse process MET, such as in the case of endoderm development, epithelial wound healing, and tumor metastasis. The concept of discrete cell states along the epithelial-mesenchymal spectrum of phenotypes has been raised [4, 5]. Therefore, observing EMT progression of living animals is a powerful tool to understand what the intermediate cell states are, and how cells reach these intermediate states.

One of the central steps during EMT is the loss of cell-cell junctions, so that the cells restricted within the epithelial sheet can dissociate from each other [6]. As the basic structure to connect cells and support cell shapes, cell-cell junctions not only respond to EMT signaling, but also are regulated by many other intrinsic and extrinsic cues, including other signals or physical tension. Therefore, cell-cell junctions such as adherens junctions can act as a platform where other inputs modulate EMT progression. Indeed, EMT during development often happens simultaneously with other morphogenic events. During gastrulation in Drosophila melanogaster (Drosophila), the mesoderm primordium undergoes both EMT and mechanical tension-driven epithelial folding. While the EMT program aims to disassemble junctions, epithelial folding requires strong junctions to maintain tissue integrity and effect tissue-wide shape changes. It was shown that mechanical tension not only drives epithelial folding but also overrides the EMT program to strengthen adherens junctions [7, 8]. This provides an excellent system to study EMT in the context of epithelial morphogenesis.

Drosophila embryos have several characteristics that facilitate high-quality live imaging and in order to understand early embryo morphogenesis, it is important to do live imaging properly. The adherens junctions in early Drosophila embryos are very close to the embryo surface and therefore can be steadily imaged with laser scanning confocal microscopy. Before gastrulation, Drosophila embryos consist of a single layer of epithelial cells surrounding the yolk with apical surfaces facing outside (Fig. 1a). Cells are about 30 μm tall and 6–7 μm wide, with spot adherens junctions localized in a relatively narrow zone within 6–7 μm below the apical surface (Fig. 1b). This single layer of epithelial cells is generated during a process called cellularization, where cell membranes of the syncytial embryo gradually ingress between nuclei and grow deeper to eventually enclose the nuclei and cytoplasm surrounding the yolk. Cellularization happens 2 h after egg laying and it takes 50 min to 1 h for the cellularization membrane front to reach the end of the cytoplasm. Therefore, the depth of cellularization front from the embryo surfaces can be used to estimate the embryo’s age for live imaging (Fig. 2). Spot adherens junctions are formed in all the cells during early cellularization and appear as clusters of cadherin-catenin complexes around the subapical region and only become more belt-like much later in the development. No other junctions, such as septate junctions and focal adhesions, have formed during this early stage of development [9]. Immediately following cellularization, gastrulation starts with the epithelial folding of the mesoderm to form the ventral furrow. This ventral furrow formation is driven by mechanical force generated from myosin contraction, on the apical surface of the mesoderm. This apical myosin contraction leads to apical constriction (Fig. 1b, c). The bulk of contractile or activated myosin is on the very apical surface of the cells and the fibers extend to the adherens junctions. In response to the myosin contraction, adherens junctions in the mesoderm migrate toward apical direction and are strengthened—eventually become highly concentrated at the apical edge of the cells [7].

Fig. 1.

Schematics of an early Drosophila embryo. (a) Before gastrulation, Drosophila embryos have a single layer of epithelial cells with the apical surface facing outside. (b) A mesodermal epithelial cell before apical constriction with spot adherens junctions concentrated at the subapical position. Also shown is the schematic of the en face view of cells from their apical surfaces/embryo surface. Below the schematic is a confocal image of such a view, E-cadherin::GFP in green, and myosin::mCherry in magenta. (c) A mesodermal epithelial cell during apical constriction with reinforced adherens junctions at the very apical edges of the cell and contractile actomyosin network on the apical cortex. The en face views in a schematic and a confocal image

Fig. 2.

Bright-field views of a Drosophila embryo from cellularization to gastrulation. In all images, ventral side is left and the anterior side points up. Yolk appears dark while the cytoplasm appears as a transparent circle. (a) The end of syncytial nuclear cycle 12. The bump-like objects at the periphery of the egg are nuclei. Yolk and cytoplasm are separating, but the boundary is unclear. Embryos of this age or younger are more fragile and should be handled with more care. (b) The end of the last syncytial nuclear division (cycle 13). Nuclei appear to be smaller due to smaller spacing and slight elongation. Yolk and cytoplasm have separated further. The cellularization front is hardly visible. (c) Early cellularizing embryo. The cellularization front becomes visible as a straight, sharp, and thin line, largely in parallel to the embryo surface (yellow arrows). Nuclei are further elongated. The yolk/cytoplasm boundary becomes sharper. (d) Cellularization proceeds further and the cellularization front is readily visible (yellow arrows). Pick embryos of this age or younger for imaging junctions loss and reversal during EMT and epithelial folding. (e) Cellularization on the ventral side is almost finished. Nuclei on the ventral side appear to be less aligned, indicating the initiation of apical constriction. Gastrulation begins. (f) Gastrulation proceeds with the ventral furrow already formed and the posterior midgut reaching the dorsal side

Adherens junction levels during the EMT of Drosophila mesoderm primordium do not change unidirectionally. Instead, there is a transient upregulation following the initial downregulation [7]. EMT in those cells is driven by the transcription factor Snail, a conserved master regulator of EMT in many other systems [10]. In the early Drosophila embryo, Snail is zygotically expressed specifically in the mesoderm and gradually accumulates to high levels from before cellularization to gastrulation [7, 11]. This high level of Snail eventually drives the mesoderm to undergo EMT [12]. Although the disassociation of cells only occurs after the folding of the mesoderm, it was shown that the adherens junctions start to be broken down before the epithelial folding event [7]. This is consistent with many of Snail’s transcriptional targets being activated or repressed before ventral furrow formation [13–15]. However, during the epithelial folding, the contractile myosin, which drives the apical constriction and eventually the folding of the mesodermal epithelium, relocates and strengthens adherens junctions despite the continued expression of Snail (Figs. 2 and 3). Such a reverse in junction levels is possible because junctions are downregulated by Snail at the step of assembly and disassembly [8]. Only after the mesoderm is completely internalized and myosin is inactivated, does Snail-dependent junction disassembly resume, and mesodermal cells undergo a full EMT.

Fig. 3.

Examples of confocal images of live embryos expressing E-Cad::GFP and myosin::mCherry (myosin regulatory light chain). (a) A z-projection of ventral cells undergoing apical constriction. E-Cadherin is in green and Myosin in magenta. The arrow indicates the tracked junction cluster shown in (b). (b) An example of a junction cluster tracked for 3 min. The junction cluster is 3D reconstructed from the confocal stacks. Images are pseudocolored to reflect the fluorescence intensities. The upper stripe is E-cadherin::GFP and lower one is myosin::mCherry within the same volume. The temporal resolution is 5 s

The approach described here relies on mounting the embryos directly on the cover glass taking advantage of the fact that both the embryos and the glass are hydrophobic and therefore stick to each other. This method for live imaging embryos avoids the use of glue and allows continued readjustment of the orientation of the embryo. While it creates a small footprint of the embryo on the glass, within this area light goes through only the vitelline membrane, glass, and emersion oil, thus greatly improving image quality. We will discuss how to identify healthy embryos of the right stages and orientation, as this is also critical for the efficient imaging. Finally, how fast imaging can be achieved through the reduction of photobleaching will also be discussed.

2. Materials

2.1. Embryo Collection

1. Plastic cups with punctured small holes.

2. 60 mm apple juice agar plates (to make 500 ml: Drosophila Agar type II 9 g, sucrose 12 g, apple juice 250 ml, H₂O 250 ml).

3. Yeast paste.

4. 60 mm petri dish lid.

2.2. Embryo Preparation and Mounting

1. Stereomicroscope with a transmitted light source (e.g., ZeissStemi 508).

2. 200 μm pipettor or glass Pasteur pipette.

3. Fine forceps.

4. Paint brushes: one with most bristles removed and another onewith only one bristle.

5. 35 mm glass bottom Petri dish.

6. Absorbent C-fold paper towels: regular size and cut into about2 cm × 2 cm pieces.

7. Air permeable membrane.

8. Water in a wash bottle.

9. Halocarbon oil 27.

10. 4% sodium hypochlorite/bleach: dilute household bleach with water (household bleach from grocery stores usually contains 6% or 8% sodium hypochlorite).

2.3. Live Imaging

1. Laser scanning confocal microscope.

2. Plan Apochromatic 63× oil immersion objective.

3. Methods

3.1. Embryo Collection

1. Place 40 to 60 young flies expressing desired fluorescent proteins in the egg collection cup and cover the opening with an apple juice plate with a drop of yeast paste. Keep the cup upside down with a rubber band securing the plate to the cup. Change the plate every day to feed the flies for about 2 days so the flies start to lay abundant amounts of eggs.

2. On the day of imaging, change to a room-temperature apple juice plate with yeast paste on it and collect the eggs for desired length of time. Keep the cup in a dark and undisturbed place to increase egg laying. To image embryos from early cellularization to gastrulation, collect eggs for at least 2 h. As flies lay more eggs at this temperature, keeping the egg collection cup in 25 °C is preferred, unless the experiment requires a different temperature.

3. After at least 2 h of collection, change the egg collection cup to a new apple juice plate. Pour Halocarbon oil 27 on the plate with collected embryos to identify embryos with the desired stages.

3.2. Embryo Staging and Dechorionation

1. Submerged in Halocarbon oil, the interior of the embryos become visible for recognizing developmental stages. Embryo preparation alone routinely takes 20 min. Therefore, picking embryos of sufficiently early stages is necessary. To image mesoderm-EMT before apical constriction, identify the embryos just starting cellularization or earlier (Fig. 2). Go through the plate, pick up around 10–15 embryos of the desired stages with forceps. If there are too many embryos to handle while searching around the plate, collect them to a marked place on the agar plate.

2. Sort through the embryos collected to find the 7–10 best ones and pick them up together with forceps. Slowly open the forceps so the embryos are clustered on one leg of the forceps. Dry the oil on a small piece of paper towel. Leave the cluster of embryos on the edge of the paper towel in order to clean the oil on the forceps. Dry the oil as much as possible as it prevents the bleach from reaching the chorion.

3. Put several drops of 4% sodium hypochlorite/bleach on a small piece of paper towel. Just enough to cover the texture of the paper towel. This allows the embryos to completely submerge in the liquid without running off the paper due to liquid overflowing.

4. Place the cluster of embryos in the bleach on the paper towel. Embryos usually get loosened and fall into the bleach once the cluster touches the bleach.

5. Use the paint brush with a few bristles to separate embryos by gently sweeping them around and against the paper towel. This also removes the residual oil on the embryos. From this point one, the embryos will be kept on this small paper towel until they are mounted on the glass (see Note 1).

6. While the embryos are being dechorionated, prepare the water drops to wash embryos. Evenly distribute three large drops of water around the inside rim of a 60 mm petri dish lid. Make three such lids with a total of 9 drops of water. Do not leave the embryos in bleach for more than 1 min.

7. Get embryos out of the bleach by gently picking up the small piece of paper towel from a corner with forceps. Dry this small piece of paper towel on regular paper towels. Do not over dry. Wash the small paper by touching a drop of water, previously prepared, with the corner of the paper grabbed by the forceps. Then dry on the regular paper towel. Repeat this with all 9 drops of water so the embryos are washed thoroughly. Keep the damp paper towel with embryos in a lid of a 35 mm dish (see Notes 2 and 3).

3.3. Embryo

Mounting

1. Clean the glass of the 35 mm glass bottom dish with Kimwipes soaked in 70% ethanol. Do not touch the rim of the well to prevent rubbing off debris.

2. Pick up each embryo one by one from the paper towel using the paint brush with a single bristle. Land the embryos on the glass in the dish. Avoid placing embryos on any remaining debris on the glass (see Note 4).

3. After placing all embryos on the glass, put a few drops of water on top of the embryos. Rock the dish so that the whole well is covered by the water. Due to both the glass and vitelline membrane being hydrophobic, embryos stick to the glass underwater without floating. An air bubble will often form attaching to each embryo.

4. Carefully pick up each embryo using the one-bristle paintbrush, within the water, leaving the air bubble on the glass. Do not lift the embryo outside the water, otherwise the embryo will be lost.

5. Choose a clean area, adjust the angles of the paint brush so that the side of the embryo to be imaged is landed on the glass. To image the mesoderm, have the ventral posterior side of the embryo landing first on the glass. Gently lay the embryo down to the middle of its ventral side on the glass. Try to arrange the embryos in one or two rows to facilitate switching between embryos with high magnification objective (see Notes 5 and 6).

6. Once all embryos are orientated correctly, cover the well of water with a small piece of air permeable membrane. Place the hydrophilic side of the membrane down and let the water spread to the entire membrane. Let the excessive water run away from the membrane so that the membrane sticks to the petri dish (see Note 7).

3.4. Confocal Imaging

1. Put a drop of immersion oil on the glass bottom of the dish where the embryos are located on the other side of the glass. Place the dish on the motorized stage of the confocal microscope (see Note 8).

2. Use the 63×/1.4 oil objective (Plan Apochromatic). It can cover about 10 × 20 cell on the ventral side of the embryo. Watch the embryos from above the stage and move them to the center of the objective. Embryos can be seen as white spots by bare eyes when the bright-field light is on. There is no need for epifluorescence mode (see Note 9).

3. Under bright field, adjust the focus and move the stage slightlyto find the shadow of the unfocused embryo and then focus properly. Go through the rows of the embryos to find one of appropriate stages by examining the depth of cellularization. Further inspect the orientation of the embryo: the embryo should appear very symmetric including the pole cells at the posterior end. Damaged embryos can show visible leakage of yolk droplets. Finally move the focus to the surface of the embryo to facilitate focusing under confocal mode.

4. Set the laser power to low levels and visualize the embryo in live mode. Do a final check on the age, orientation, and healthiness of the embryo. Red fluorescent protein is almost always weaker and faster bleached than GFP so use 488 nm laser alone for this purpose if possible (see Note 10).

5. Once the embryo to be imaged is identified, adjust the laser power, gain, zoom factor, and temporal resolution. Find a balance between them to minimize photobleaching and maximize the image quality (see Note 11).

4. Notes

1. Bleach can lose potency over time and the dechorionation becomes slow. It is tempting to push and roll the embryo to facilitate dechorionation. However, doing so often results in embryo damage or embryos rolling inside the vitelline membrane. This will lead to difficulties in properly orienting the embryos when trying to precisely image the mesoderm. Furthermore, embryos rolled inside the vitelline membrane always roll back to the original position exactly during ventral furrow formation. Therefore, the cells being imaged will be moving in or out of the field of view.

2. Usually preparing water drops and paper towels for the washing step is sufficient for dechorionation. While not all embryos will appear to be dechorionated before washing, they will by the first two to three rounds of drying and washing cycle. As the water rushes through the small paper towel from its corner, all embryos should be completely dechorionated, if they are not already. When the bleach is not potent enough, drying the small paper towel and reapplying the bleach can significantly speed up the dechorionation process.

3. Do the washing and picking-up under the stereomicroscope to prevent losing embryos. Pick up different corners of the small paper towel when embryos are washed from one corner to another.

4. Plastic bristles are hydrophobic like the vitelline membrane so usually the embryo sticks to the bristle without problems. For the one-bristle brush, the thicker it is, the easier it is to pick up embryos as the contact area between the bristle and embryos is larger. Obviously, bristles cannot be too thick as it becomes difficult to manipulate. One way to make such a brush is to cut one bristle off from the root and stick the tip of this bristle back to the other end of the pen brush. This way the root, which is thicker, becomes the tip of the one-bristle brush. Flattening the bristle also increases its area.

5. During this step, once the embryos are on the glass they are exposed to dry air. It is preferable to do this quickly as possible to avoid drying the embryos too much. As the orientation of the embryos will be adjusted later, landing the embryos on its dorsal side is often the safest and easiest way. Due to its geometry, the dorsal side is physically sturdier and requires less care. It is possible to place many embryos down on the glass within a short period of time without damaging them or pushing the embryos to roll inside the vitelline membrane.

6. It is crucial to orient the embryo precisely so that between the desired tissues and objective there is only immersion oil, glass, and vitelline membrane. Again, to decrease the chance of the embryos rolling inside the vitelline membrane, avoid rolling the embryos from side to side. Rocking the embryos from anterior to posterior is more acceptable. If the embryo does not land precisely in the middle of the ventral side, lift the embryo up from anterior and before the posterior leaves the glass adjust the paint brush angle to place the embryo down from posterior to anterior.

7. Placing an air permeable membrane on the well has several functions. It maintains the embryos in a very thin layer of water with sufficient air exchange. Meanwhile it prevents this thin layer of water from evaporation too fast while imaging. Furthermore, as this membrane sticks to the well due to atmosphere pressure, there is no worry that the water may spill out when transported to the microscope and placed on the stage. It also allows imaging through upright microscope, as long as there is sufficient working distance, since the petri dish can be placed upside down with the membrane holding up the water in the well.

8. Placing the oil on the dish where the embryos are rather thanon the objective has several advantages. First it prevents the oil from running down the objective before being held between the objective and the cover glass on the bottom of the dish. It also restricts the oil to the area where the embryos are, so that the oil does not spread over large area, resulting in too little oil between the objective and area to be imaged.

9. A 60× or 63× oil objective is necessary to get sufficient quality images of detailed morphology of adherens junctions and contractile myosin.

10. Determine the exact age of the embryo by the depth of the cellularization, as shown by any proteins localizing around the cellularization front such as E-cadherin or myosin. DIC can be helpful to determine the age if no fluorescent proteins in the embryo associate with cellularization structures.

    The pattern of E-cadherin or myosin around cellularization front in the precisely oriented embryo should appear very symmetric. Since cellularization proceeds faster in the mesoderm, the pattern will appear asymmetric in a slightly tilted embryo.

    Normally the footprint of the ventral side of the embryo on the glass is a smooth oval. When embryos leak content due to damage, the footprint will appear abnormally big and show wrinkles due to loss of internal pressure. The outline of the footprint can also appear wavy. Additionally, avoid embryos that have landed on debris. Damaged embryos often also show defects in cellularization. The cellularization front in a healthy embryo will appear very even on both the XY plane or in the z direction, both in terms of cell sizes and cellularization speeds. Cells in damaged embryos may have shrinking cells surrounded by stretched cells. If the significantly damaged cell is outside field of view, cells in the field of view often appear stretched to a certain direction. Do not image such an embryo.

11. To image embryos from cellularization to gastrulation, a good starting point of imaging format is the following: physical size of each pixel: 0.12 μm; digital dimension of each image: 1024 × 512; Z stack depth: 10–15 μm with 1 μm interval; temporal interval: 1 min. Scanning goes much faster along X than Y, to save time, always use more pixels along X rather than the other way around. For most confocal microscopes, each stack will take much less than 1 min. Therefore, there is no need to use bidirectional imaging as less frequent and spreadout exposures to lasers appears to reduce photobleaching compared to concentrating all the laser dosage in a short period of time. At the end of an imaging session, zoom out to examine if the imaged area is significantly darker than the rest of the embryo.

    To image adherens junction dynamics and track junction clusters during apical constriction, the following format can be tested first: physical size of each pixel: 0.12 μm; digital dimension of each image: 512 × 256; Z stack depth: 7 μm with 0.5 μm interval; temporal interval: 5 s. Z Piezo driver is helpful to image at such a high temporal resolution. Bidirectional mode may be used depending on the speed of the microscope.