TITLE: Microinjection Techniques in Fly Embryos to Study the Function and Dynamics of SMC Complexes
AUTHORS: Catarina Carmo, Margarida Araújo, Raquel Oliveira

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block">Structural maintenance of chromosomes (SMC) proteins are critical to maintain mitotic fidelity in all organisms. Over the last decades, acute inactivation of these complexes, together with the analysis of their dynamic binding to mitotic chromatin, has provided important insights on the molecular mechanism of these complexes as well as into the consequences of their failure at different stages of mitosis.</div><br/><div class = "text-block"><span>Here, we describe a methodology to study both SMC function and dynamics using </span><span style = "font-style:italic;">Drosophila melanogaster </span><span>syncytial embryos. This system presents several advantages over canonical inactivation or imaging approaches. Efficient and fast inactivation of SMC complexes can be achieved by the use of tobacco etch virus (TEV) protease in vivo to cleave engineered versions of the SMC complexes. In contrast to genetically encoded TEV protease expression, </span><span style = "font-style:italic;">Drosophila </span><span>embryos enable prompt delivery of the protease by microinjection techniques, as detailed here, thereby allowing inactivation of the complexes within few minutes. Such an acute inactivation approach, when coupled with real-time imaging, allows for the analysis of the immediate consequences upon protein inactivation. As described here, this system also presents unique advantages to follow the kinetics of the loading of SMC complexes onto mitotic chromatin. We describe the use of </span><span style = "font-style:italic;">Drosophila </span><span>embryos to study localization and turnover of these molecules through live imaging and fluorescence recovery after photobleaching (FRAP) approaches.</span></div></div>

[STEPS]
?. [Collecting and Preparing Embryos for Live Imaging]
Set up a cage with the fly strain of your choosing. Use an apple juice plate with a smear of fresh yeast paste at the bottom of the cage.
When setting a cage, take into consideration that older flies have a decreased egg-laying capacity. Males that have matured for over 3 days mate more efficiently and females reach their peak of egg laying 4–7 days after eclosion.Take into consideration that flies tend to lay more eggs during the night-time. If more convenient, cages can be kept inside a box with an inverted light–dark cycle (e.g., to become dark at 12 pm and light at 12 am). In this manner, flies will lay more eggs during the afternoon.Set up a cage for at least a full day before the day of experiments, in order for flies to acclimatize to the cage and fully recover from CO2 anesthesia.As flies are attracted by the smell of fruit and eat yeast, they will be attracted to the apple juice plate with yeast paste, and lay a lot of eggs that can be collected.
?. [Collecting and Preparing Embryos for Live Imaging]
Change plates at least once a day, even on days without experiments. To do this, invert the cage, tap it strongly, so as to bring the flies to the bottom, and exchange plates.
?. [Collecting and Preparing Embryos for Live Imaging]
On the day of the experiments, start with a precollection of – to release retained eggs and increase staging accuracy.
Flies tend to hold embryos before depositing them but fresh yeast paste stimulates the egg laying. Changing the plates every morning will ensure that you remove the embryos held overnight in their oviducts. This will also give a good idea of the egg-laying efficiency.
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The time of collection will depend on the desired developmental time—shorter collection times for early divisions and longer collection times for late developmental stages. For example, to collect embryos that are, at most, at nuclear division 10 (blastoderm nuclei), corresponding to  of development, you may want to start with a collection of , counting with ~ for embryo preparation.
?. [Collecting and Preparing Embryos for Live Imaging]
To collect embryos from the agar plate (Fig. 3a), use an artist’s brush (moist with water) and swipe them onto a cell strainer, placed on a container with tap water.
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Briefly remove excess water on a tissue paper and transfer the embryo containing cell strainer to a container with 50%(v/v) bleach. Incubate for  at . This will remove nontransparent chorion (dechorionation), essential for injection and imaging.
4 Room temperature
Time left on bleach solution will depend on embryo’s resistance; more fragile embryos may require less time. Also, by the end of the day, the bleach solution will become weaker and it may be required to leave embryos longer.
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Remove excess bleach solution with a tissue paper. Wash embryos with a squeeze bottle with distilled water. Water pressure from the bottle directly on the embryos will help in the removal of chorion. Rinse the embryo containing cell strainer in the container with tap water.
Replace the water from the tap water container between collections and wash the cell strainer with the help of an artist’s brush in order to get rid of remainders of bleach and embryos (from the cell strainer).
?. [Collecting and Preparing Embryos for Live Imaging]
Cut a small block of a clean apple juice agar with a scalpel and place it on a coverslip, to be viewed under a stereo microscope.
?. [Collecting and Preparing Embryos for Live Imaging]
Using a 24 × 60 mm coverslip, place around – in the middle of the coverslip as a single row and tilt it to make it spread as an even layer. This will be used to mount the embryos for live imaging. For injections: take a smaller coverslip (18 × 18 or 22 × 22 mm) and place it so as to overlay approximately half of the glue layer (Fig. 2 for scheme).
6 µl
[heptane glue]
?. [Collecting and Preparing Embryos for Live Imaging]
Transfer embryos from the cell strainer using a brush and place them on the agar block, with the aid of a steel probe. Dechorionated embryos should have an ovoid shape without the dorsal appendages (Fig. 3b2).
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Align the embryos in a row, making sure all face the same direction (i.e., every embryo has the anterior side, marked by micropyle, oriented to the same direction). Use the edge of the agar block as a reference (Fig. 3b2).
?. [Collecting and Preparing Embryos for Live Imaging]
Once aligned, take the preprepared 24 × 60 mm coverslip (see step 9) and with the glue side facing down lower it until you glue the embryos, by gently pressing on the agar block. Keep it parallel to the smaller coverslip, with the micropyle facing this side (for posterior end injections).
Proper attachment of the embryos onto the coverslip is a critical step for successful injections to avoid that the embryo “escapes” the glue once the needle approaches. Embryos must, therefore, be sufficiently dry, as water can compromise their attachment.
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For injections only: leave the preparation to dry for –.
An extended drying step is critical for injections to decrease the osmotic pressure of the embryo and thereby prevent bursting and cytoplasmic ejection once pierced.
?. [Collecting and Preparing Embryos for Live Imaging]
Using a 20–200 μl tip, take halocarbon oil and place it on top the row of aligned embryos and part of the smaller coverslip. This will keep embryos moist and oxygenated.
?. [Collecting and Preparing Embryos for Live Imaging]
Samples are ready for the following processes, including live cell imaging of unperturbed embryos (see next section).
?. [Microinjection Techniques in Fly Embryos]
For microinjection experiments, 1–1.5 h old embryos (or 0–30 min for mRNA injections) must be collected and processed according to protocol described above. Embryos should preferentially be injected (up to three consecutive injections) at the posterior side—owing to a more uniform surface while maintaining the shape and preventing extensive loss of cytoplasm. Here we describe the use of prepulled needles to ensure repeatability.
?. [Microinjection Techniques in Fly Embryos]
Load the needles using Microloader Tips. Take care not to leave air bubbles during loading, as it will make injection impossible. Prepare all needles needed before starting the experiment, to minimize time between injections.
?. [Microinjection Techniques in Fly Embryos]
Once at the microscope, first turn on the injector controller and the micromanipulator. Then, place the first needle in the holder—must be tight to maintain correct pressure—and connect the capillary to the pressure pump afterward.
?. [Microinjection Techniques in Fly Embryos]
Using the lower magnification objective (10× or 20×), put the needle down slowly in the focal plane of the smaller coverslip. When the needle is close to the coverslip you will start seeing a shadow through the lens (Fig. 4a).
?. [Microinjection Techniques in Fly Embryos]
Prepulled needles need to be further opened prior to injections. The smaller coverslip next to the row of aligned embryos will serve as a barrier to break the tip of the needle and thus to open it. Press gently the needle against the edge of the coverslip until it breaks slightly and try several injections until the correct droplet size is achieved. Press the injection button and evaluate the size of the drop (Fig. 4b), where 6 shows an appropriate drop, that should range from 30 to 50 μm in diameter (up to one-tenth of embryo length). The size of the droplet can be controlled by regulating the amount of pressure and the injection time, for example, when using an Eppendorf FemtoJet Microinjector controller. If the needle gets clogged, it can be opened further using the same strategy as before.
?. [Microinjection Techniques in Fly Embryos]
To perform an injection, as the needle comes in contact with the embryo’s posterior pole, notice how the membranes retract with it (Fig. 4c2, 3 and ESM Movie S1). Move the needle further until it goes through the embryo (Fig. 4c4) and inject.
?. [Microinjection Techniques in Fly Embryos]
For multiple injections, change the needle and inject the second/third solution through the same hole. The small opening from the first injection facilitates the entry of the second needle inside the embryo, without membrane retraction. Figure 4c6 displays a second injection using the same injection site.
Sequential injections should be performed exactly at the same site to avoid cytoplasm release. To facilitate this process, ensure a wide opening during the first injection. This can be achieved either by breaking slightly more the first needle to be used or by introducing it further inside the embryo (the wider part of the needle helps to introduce an opening which is then easier to find in subsequent injections).If it is not possible to spot the opening, the needle can be used to probe where the injection site is by scrolling up and down slowly through the posterior side of the embryo until it gets in by itself.
?. [Inactivation of SMC Complexes by TEV Cleavage]
TEV-mediated inactivation requires prior establishment of Drosophila strains surviving solely on the TEV-cleavable version of the protein. Strains should also contain the desired fluorescent markers (e.g., H2Av- or H2B-fluorescently tagged proteins to monitor chromatin behavior). This strategy allows full and acute inactivation of targeted proteins in a time-resolved manner and thus can be applied to investigate both the establishment and maintenance of the intricate mitotic chromosome morphology. As an example of a time-restricted inactivation protocol, we detail the steps for cohesin inactivation in metaphase-arrested embryos, as originally described in [15].
The experimental layout described here focuses on the use of TEV protease to study the role of SMC complexes in the maintenance of metaphase chromosome structure. Canonical studies on the role of these complexes for the establishment of chromosome architecture can be achieved by injection of TEV during interphase, leading to precocious sister chromatid disjunction (for cohesin) or impaired sister chromatid resolution (condensin) [9, 15], depending on the question to be addressed.
?. [Inactivation of SMC Complexes by TEV Cleavage]
Switch to a lower magnification lens for injections.
?. [Inactivation of SMC Complexes by TEV Cleavage]
Induce a metaphase arrest through the injections of – into the embryo. Imaging acquisition can be performed (using a 63× or 100× lens). After –, every nucleus should have their chromosomes aligned forming the metaphase plate (Fig. 5a).
[UbcH10C114S]
?. [Inactivation of SMC Complexes by TEV Cleavage]
Subsequently, perform a second injection with TEV protease at –. If the protease is at this concentration, sister chromatid separation should be observed within – after TEV protease injection in flies carrying TEV-sensitive cohesin complexes (Fig. 5b).
?. [Fluorescence Recovery After Photobleaching (FRAP)]
FRAP studies on chromatin-binding proteins revealed that many have a very dynamic behavior, with turnover within seconds (e.g., transcription factors [29]). In contrast, both cohesin and condensin complexes were shown to display a slow turnover or be stably bound to mitotic chromatin [19, 20, 22]. More importantly, turnover rates may also vary for specific time points of cell cycle. Thus, microinjection techniques can be used to arrest the fast embryonic cycles at specific stages, thereby enabling long-term FRAP experiments. What follows below is an example of analysis of condensin I turnover on mitotic chromosomes, similar to the one previously published [22]. For this analysis, strains expressing fluorescent-tagged versions of the protein of interest are required.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Switch to a lower magnification lens for injections.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Inject – UbcH10C114S (intact spindle forces) or   into the embryo to induce a metaphase arrest, if required. After –, every nucleus should be arrested in prometaphase or metaphase.
[colchicine (microtubule poison)]
Depending on the anticipated time of recovery, FRAP analysis can be performed in cycling embryos instead of inducing a metaphase arrest.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Select a field for imaging, preferably including the nuclei closer to the coverslip.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Image for a short period a time (e.g., ) before inducing a bleaching pulse. This will provide a reference for basal fluorescent intensity before bleaching and recovery occur.
For FRAP analysis it is crucial to minimize photodamage stimulated by laser/light power. Time between frames should also be short enough to detect fast exchange events but without further enhancing phototoxicity (e.g., 30 s/frame).
?. [Fluorescence Recovery After Photobleaching (FRAP)]
According to the imaging software available to induce FRAP, draw ROIs of the nuclei to be bleached, bleaching a maximum of one-fourth of the metaphases in the field (see Fig. 6a).
Several options can be used for the shape and size ROI to be bleached: bleaching of entire metaphases, bleaching of half metaphase, or bleaching of a smaller area within the metaphase (e.g., a small circle, rectangle). Be aware that recovery dynamics may be challenged by larger areas simply due to bleaching of a higher number of fluorescent molecules. Also, nuclei, although arrested at metaphase, will not be static, and may hinder FRAP efficiency. We find it best to bleach half of a metaphase plate, where the other unbleached half can be used as an internal control.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Induce the pulse.
When optimizing bleaching pulses, take care that they should be short, typically less than 20 ms, in order to avoid phototoxicity for the sample and localized heating of the sample, generated by high laser intensity [34]. As such, increasing the laser power works better than increasing the timing of the bleaching pulse. This is also useful to minimize the diffusion effect of fluorescent molecules during the pulse [18].
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Image immediately after bleaching for a longer period of time (e.g., ), keeping the same settings as prebleaching imaging.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Analyze the recovery using quantitative imaging software (e.g., Fiji [30]).
?. [Fluorescence Recovery After Photobleaching (FRAP)]
The mean fluorescence intensity can be normalized in several ways (e.g., to the first time point before pulse (t0) or to unbleached half metaphase for each time point).
Several options can be used for the shape and size ROI to be bleached: bleaching of entire metaphases, bleaching of half metaphase, or bleaching of a smaller area within the metaphase (e.g., a small circle, rectangle). Be aware that recovery dynamics may be challenged by larger areas simply due to bleaching of a higher number of fluorescent molecules. Also, nuclei, although arrested at metaphase, will not be static, and may hinder FRAP efficiency. We find it best to bleach half of a metaphase plate, where the other unbleached half can be used as an internal control.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Plot the relative mean fluorescence intensity versus time in a xy  manner.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
For estimation of protein turnover, fit the data to the appropriate function (e.g., the One Phase Association equation y = yo + (Plateau−yo)*(1 − exp(−K × x)) can be used to estimate several dynamic parameters, as indicated in Table 1).   AB1VariableDefinition2Y0Value of y at t = 0 Expressed in the same units of y3PlateauValue that y tends to for infinite of x Expressed in the same units of y4KRate constant Expressed in –t (inverse of x units)5TauTime constant Expressed in the inverse of y units6Half-timeTime of fluorescence recovery after the pulse where the fluorescence intensity is half of the final recovered intensity Expressed in the same units of x7Span (mobile fraction)Difference in intensity between y0 and plateau Expressed in the same units of y
AB1VariableDefinition2Y0Value of y at t = 0 Expressed in the same units of y3PlateauValue that y tends to for infinite of x Expressed in the same units of y4KRate constant Expressed in –t (inverse of x units)5TauTime constant Expressed in the inverse of y units6Half-timeTime of fluorescence recovery after the pulse where the fluorescence intensity is half of the final recovered intensity Expressed in the same units of x7Span (mobile fraction)Difference in intensity between y0 and plateau Expressed in the same units of y
?. [Fluorescence Recovery After Photobleaching (FRAP)]
FRAP will result in three possible scenarios: no recovery of fluorescence intensity, indicating that there was no replacement of fluorescent molecules and hence, the protein is stable and did not turn over; partial recovery of fluorescence intensity or complete recovery of fluorescence intensity, where there was limited or complete exchange of the tagged protein (Fig. 6b) [31].
?. [Microinjection Techniques in Fly Embryos]
Before usage, centrifuge your microinjection sample at high speed for  at , so as to impede precipitates to be extracted and possibly clog the injection needle.
4 °C
?. [Inactivation of SMC Complexes by TEV Cleavage]
Use your reference channel (e.g., fluorescent histones) to select an embryo with the required nuclear density and in late interphase using a 63× or 100× lens.
?. [Fluorescence Recovery After Photobleaching (FRAP)]
Use your reference channel (e.g., fluorescent histones) to select an embryo with the required nuclear density and in late-interphase using a 63× or 100× lens.