The authors gratefully acknowledge the work of Liuliu Zheng and Zenghui Xue, who helped pioneer this technique in the Sokac lab, as well as Hasan Seede who helped with the analysis. The work for this study is funded by a grant from NIH (R01 GM115111).

1. Prepare embryo collection cups and apple juice agar plates
<ol>
	<li>Five days prior to the injection experiment, construct28 or procure at least two small&#160;embryo collection cups. Make fresh 60 mm apple juice agar plates to be used with small collection cups28. Store plates in plastic boxes covered with damp paper towels at 4 &#176;C. 
	NOTE: Small embryo collection cups, populated with fly numbers as described in step 1.3, will provide sufficient embryo numbers per experiment, while also ensuring that embryo handling and injection can be done in a short enough time to allow imaging of early developmental stages.</li>
	<li>Warm apple juice plates to 18 &#176;C and add a dab of yeast paste to the center of the plate. Yeast paste is a simple paste of active yeast and distilled water.</li>
	<li>To promote the most generous egg laying, set up collection cups with flies 2 days prior to the experiment. Add at least 100 females and 50 male flies to the collection cups, and top with a prepared apple juice plate (Figure 1, step 1). On the days leading up to the injection experiment change the apple juice plates at least twice each day, once in the morning and once in the evening. 
	NOTE: The best injection and imaging results are obtained when embryo collection cups are kept at 18 &#176;C with a 12 h light on/light off cycle.</li>
</ol>
2. Prepare a working stock solution of G-actinRed for microinjection
NOTE: This preparation only makes 2 &#956;L of a 5 mg/mL working stock of G-actinRed, so if users are unaccustomed to the microinjection technique, it is advantageous to skip to step 3 and practice the microinjections with a neutral pH buffer to conserve precious working stock. The 10 &#956;g stock of G-actinRed from the vendor can be stored in its original packaging in a 16 oz screw top jar with ~500 g of desiccant at 4 &#176;C for up to 6 months.
<ol>
	<li>Prepare a G-buffer stock solution in advance: 5 mM Tris-HCl, 0.2 mM CaCl2, pH 8.0. Filter and store at room temperature.</li>
	<li>On the day of injections, prepare 1 mL of a G-buffer working solution in a fresh snap cap microcentrifuge tube on ice using the G-buffer stock from step 2.1, supplemented with the following at the indicated final concentrations: 1 mM dithiothreitol (DTT) and 0.2 mM ATP, pH 8.0. 
	NOTE: The 1 mL volume of G-buffer working solution is more than what is needed for an experiment but simplifies the preparation. Excess can be discarded after the experiment or users can scale down according to their preference.
	<ol>
		<li>Keeping the 10 &#956;g G-actinRed stock on ice, first add 1 &#956;L of filtered, distilled water to the top of the pink droplet of G-actinRed inside the tube.</li>
		<li>Next, add 1 &#956;L of the cold, freshly prepared G-buffer working solution from step 2.2. Pipet up and down ~20 times to mix well with the pipette volume set to 1 &#956;L. The G-actinRed stock will now be at the final dilution of 5 mg/mL.</li>
	</ol>
	</li>
	<li>Incubate the prepared G-actinRed for 30 min on ice undisturbed.</li>
	<li>Centrifuge the prepared G-actinRed at 16,000 x g for 20 min at 4 &#176;C in a microcentrifuge to remove any precipitate.</li>
	<li>Carefully pipet 1.5 &#956;L of the supernatant into a fresh snap cap microcentrifuge tube on ice, avoiding the dark pink pellet.</li>
	<li>Store the prepared G-actinRed supernatant on ice for up to 6 h until ready to load into a microneedle. 
	NOTE: Microneedles can be pulled from capillary tubes in advance on a micropipette puller, then stored at room temperature on a strip of modeling clay in a 100 x 20 mm Petri dish. Suggested parameters for microneedle pulling can be found in the Pipette Cookbook29.</li>
</ol>
3. Collect embryos and mount for injection
<ol>
	<li>Allow flies to lay embryos for 30 min on apple juice plates with yeast paste at 18 &#176;C.</li>
	<li>While flies are laying, pre-warm an apple juice agar plate without yeast paste to room temperature, cut out a 4 cm x 1 cm rectangular wedge of apple juice agar with a razor blade, and place on a 25 mm x 75 mm glass slide.
	<ol>
		<li>Harvest plate from the 30 minute collection and dechorionate embryos by pouring fresh bleach, diluted 1:1 with distilled water, onto the plate and swirling the plate for 1 min, as described in28 (Figure 1, step 2). 
		NOTE: Different brands of bleach are sold at different concentrations. The bleach used here is 6% sodium hypochlorite from the bottle and is diluted to a final concentration of 3% sodium hypochlorite. Other bleach brands at slightly lower concentrations will work equally well.</li>
		<li>Pour the bleach and dechorionated embryos into a collection basket (a 70 &#956;m cell strainer) and rinse the plate twice with distilled water from a squirt bottle, adding these washes to the collection basket.</li>
		<li>Vigorously rinse the dechorionated embryos in the collection basket with distilled water until no yeast clumps are visible and the basket leaves no pink marks from excess bleach when blotted on a paper towel.</li>
	</ol>
	</li>
	<li>Using a paintbrush with bristles dampened with distilled water, transfer dechorionated and washed embryos from the collection basket onto the prepared apple juice agar wedge on the glass slide.</li>
	<li>Use a pair of fine tip tweezers or a dissecting needle to arrange ten embryos in a straight line along the long axis of the rectangular agar wedge (Figure 1, step 3). Arrange the embryos head-to-tail, such that their anterior pole is facing to the right and dorsal side is facing the researcher (Figure 1, step 3, magnified).</li>
	<li>Cut off 0.5 cm of the end of a P200 pipette tip with a razor blade and dip into the &#8220;embryo glue&#8221; (described in28). Generously coat a region 5 mm in width (Figure 1, step 4) along the long edge of a 24 mm x 50 mm rectangular coverslip, and let dry, glue side up. Drying will take ~30 s and is complete once the entire glue-coated region appears matte rather than wet or shiny. 
	NOTE: Prepare &#8220;embryo glue&#8221; at least 48 h in advance. Add n-Heptane to strips of double-sided tape in a scintillation vial as described in28.</li>
	<li>Once the &#8220;embryo glue&#8221; has dried, gently place the coverslip glue side down on top of the row of aligned embryos on the agar, leaving 2-3 mm of space between the edge of the coverslip and the row of embryos. 
	NOTE: Sticking the embryos too close to the edge of the coverslip may lead to the embryos drying out too much during the course of the experiment.</li>
	<li>Flip the coverslip over so that the embryos are now facing up. They should be stuck in a line along one long edge of the coverslip, and their ventral region facing the closest edge of the coverslip (Figure 1, step 4, magnified).</li>
	<li>Desiccate the embryos by placing the coverslip with embryos gently on top of 150 g of fresh blue desiccant stored in a 16 oz screw top jar. Tightly screw on the lid and incubate for 8-10 min (Figure 1, step 5).</li>
	<li>After desiccation, remove the coverslip from the desiccant jar and tape each short side of the coverslip to a microscope slide, embryo side up, with two 4 cm2 pieces of double-sided tape so that the embryo coverslip will fit onto the injection stage (Figure 1, step 6).</li>
	<li>Add 2-3 drops of Halocarbon 27 oil with a Pasteur pipette to cover the aligned embryos and protect them from further dehydration (Figure 1, step 6).</li>
</ol>
4. Inject and heat stress embryos to promote actin rod formation
NOTE: All injections are done in a temperature-controlled room at 18 &#176;C.
<ol>
	<li>Prepare humid incubation chambers from a glass Petri dish at least 100 mm x 20 mm in size and line the chamber with twists of lab tissue wipers dampened with distilled water (Figure 1, step 8). Pre-warm the incubation chambers at 32 &#176;C or the desired incubation temperature prior to injecting embryos.</li>
	<li>Open the airflow valve for the microinjector and turn on the microinjector (compressed air or house air with a pressure of at least 90 psi is suitable).</li>
	<li>While embryos are desiccating, backload the previously prepared G-actinRed supernatant into the microneedle using a micro loader tip. Set the pipette to draw up 1-1.5 &#956;L. 
	NOTE: Because of the viscosity of the actin, loading volumes may not be accurate and there may be enough actin left to load at least one to two more microneedles. Up to 60 embryos can be injected per loaded microneedle if the microneedle is calibrated properly and does not become clogged during the course of the experiment.</li>
	<li>Attach the microneedle to the needle holder and tighten the screw. Connect the air tube to the microinjector and ensure that the backflow pressure on the microneedle equilibrates to 30 hPa.</li>
	<li>Calibrate the microinjector settings to expel a 100 &#956;m diameter bubble of G-actinRed (~500 pL) on a slide micrometer. Rotate the pressure knob (500-1500 hPa) and injection pulse time knob (0.1-0.5 s) on the microinjector to get the right bubble size. Adjust these settings each time a new microneedle is loaded to account for variability in actin viscosity and microneedle tip size. 
	NOTE: The prepared G-actinRed is viscous and there may be air in the tip of the microneedle that should be expelled before injecting embryos. If the G-actinRed does not readily expel from the tip of the microneedle, gently break the microneedle tip against the edge of the slide micrometer.</li>
	<li>Place the slide with mounted embryos onto the microscope stage. 
	NOTE: Every injection set up will be different, so researchers will have to adjust their injection method accordingly. Here the embryos are moved with respect to a stationary microneedle, injecting each embryo by running the embryo into the microneedle.</li>
	<li>Adjust the micromanipulator stage and focus of the 10x objective on the light microscope so that the embryos are visible. The embryos are in the correct focal plane when the outlines of the vitelline membrane are sharpest and the embryo appears largest. Choose embryos to inject that are in the correct developmental stage, so that by the time the post-injection incubation is complete, most of the clutch reaches the desired developmental stage (e.g., inject embryos at Bownes&#8217; stage 2-330 in order to observe rods at cellularization after heat stress at 32 &#176;C).</li>
	<li>Use the microneedle controls to bring the needle into the same focal plane as the embryos. 
	NOTE: If the microneedle catches on the coverslip while moving the stage or microneedle, then the microneedle is too close to the slide and is not in the correct focal plane. The microneedle should be parallel to the coverslip, and not at a significant angle (Figure 1, step 7).</li>
	<li>Insert the microneedle into the embryo so that it hits the embryo in the middle of its ventral region, at the embryo &#8220;equator&#8221;. Trigger injection with the foot pedal or &#8220;inject&#8221; button when the microneedle tip is visible inside the middle of the embryo (Figure 1, step 7).</li>
	<li>Inject the G-actinRed once and slowly remove the microneedle. Move the stage and repeat for each embryo of the appropriate developmental stage. 
	NOTE: Expansion of the embryo is normal as the G-actinRed is injected and a bit of cytoplasm may leak out of the embryo.</li>
	<li>After all the embryos have been injected, place the slide with the embryos in the prepared humid incubation chamber and close the lid (Figure 1, step 8). If trying to obtain embryos that reach cellularization, heat stress the embryos at 32 &#176;C for 60-75 min in the humid incubation chamber. 
	NOTE: Incubation times are noted that allow visualization of rods in cellularizing heat stressed embryos. The incubation time will be longer for non heat stress control embryos because development will be slower at a lower temperature31. These control embryos can be incubated in humid chambers at temperatures such as 18 &#176;C or 25 &#176;C, depending on design of the specific experiment and the question to be asked. The minimum incubation time necessary for the G-actinRed to diffuse throughout the embryo is 30 min.</li>
</ol>
5. Image actin rods in heat stressed embryos by confocal microscopy
<ol>
	<li>While the embryos are being heat stressed, turn on the confocal microscope and select the 561 nm laser channel.</li>
	<li>Move the objective lens (25x, 40x or 63x recommended) to the working position.</li>
	<li>If imaging heat stressed embryos, then set the heated stage incubator to achieve an internal temperature of 32 &#176;C. A point-and-shoot or infrared thermometer can be used to check the temperature at or near the objective.</li>
	<li>Remove the slide with injected embryos from the humid incubation chamber after incubation is complete (Figure 1, step 9).</li>
	<li>Working quickly, gently pry off the double-sided tape pieces that were used to adhere the coverslip with mounted embryos to the slide (Figure 1, step 9). 
	CAUTION: Be gentle during these steps as coverslips can easily shatter if too much force is applied.</li>
	<li>Stick two 2.5 cm long pieces of double-sided tape together and cut the tape in half lengthwise to make two strips, 2.5 x 0.5 cm long (Figure 1, step 10).</li>
	<li>Stick two-thirds of the length of each tape strip onto the first coverslip, flanking each side of the embryos in Halocarbon 27 oil (Figure 1, step 10, orange), leaving one-third of the tape strips hanging off the edge of the first coverslip where the embryos are stuck. Use gloved hands and be careful not to touch the embryos during this step.</li>
	<li>Gently place a second rectangular coverslip on top of the tape strips to sandwich the embryos between the coverslips (Figure 1, step 10, blue). Align the 25 mm edges but keep the 50 mm edges offset from one another by 1 cm in width. 
	NOTE: This second coverslip&#8217;s full surface will become the new imaging surface that will face the objective lens, so take care not to get fingerprints or Halocarbon 27 oil on this second coverslip surface. The offset is necessary so that extra Halocarbon 27 oil can be added to evenly immerse the embryos. If needed, add Halocarbon 27 oil at the seam where the two coverslips meet at the top of the sandwich and it will coat the embryos by capillary action (see dashed lines in Figure 1, step 10).</li>
	<li>Gently tap down on the areas of the coverslip that are directly on top of the tape strips with the blunt side of a razorblade to get the coverslip to adhere to the tape.</li>
	<li>Flip the coverslip sandwich over and place on a lab tissue wiper to keep the imaging surface clean and carry to the confocal microscope (Figure 1, step 11). Ensure that the tape is completely stuck to both coverslips before imaging.</li>
	<li>Confirm that the heated stage is at temperature and if using an inverted microscope, add immersion liquid onto the selected objective lens.</li>
	<li>Place the coverslip sandwich onto the stage carefully to ensure that the new imaging surface (2nd coverslip) is the one touching the immersion liquid (Figure 1, step 11). 
	NOTE: If needed, adhere the coverslip sandwich to the stage with two small pieces of double-sided tape to prevent unnecessary movement during imaging if the coverslips do not fit well in the heated stage.</li>
	<li>Focus on an embryo that is in cellularization (Bownes stage 4a30) or desired developmental stage using either transmitted light or fluorescence.</li>
	<li>Once an embryo has been brought into focus, switch to the laser acquisition mode on the confocal microscope and adjust laser power and gain, frame size, tiling, and projection settings as desired.</li>
	<li>Take surface-view images through the focal planes of the embryo&#8217;s nuclei to find intranuclear actin rods. Rods should appear in multiple orientations as bright streaks or dots inside the comparatively dark nuclei (Figure 2A, 2C).</li>
</ol>
6. Alternative imaging experiments
<ol>
	<li>Perform FRAP to investigate actin turnover along the length of intranuclear actin rods or in cytoplasmic actin structures, such as the tips of the plasma membrane furrows during cellularization.</li>
	<li>In the imaging software, choose a rectangular region around furrow tips or actin rods in which to acquire the experiment.</li>
	<li>Choose a small square region of the center of an actin rod within the rectangular acquisition region to bleach. Ensure that the tips of the actin rod are visible and do not get bleached during the course of the experiment to allow for accurate tracking of the rod inside the nucleus.</li>
	<li>Set the bleach laser to iterate 50x and set the bleach laser power to maximum.</li>
	<li>Choose a time course to acquire images every second for a total up to 120 s at maximum pixel dwell speed and set the laser to bleach after the first two seconds of image acquisition.</li>
	<li>Quantify the data to determine the half-time of fluorescence recovery1.</li>
</ol>

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F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+mechanical+properties+of+early+Drosophila+embryos+measured+by+high-speed+video+microrheology.">The mechanical properties of early Drosophila embryos measured by high-speed video microrheology.</a> Biophysical Journal. 108 (8), 1899-1907 (2015).</li><li>Mollinari, C., Gonz&#225;lez, A., Cid-Arregui, A., Garc&#237;a-Carranc&#225;, <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microinjection+and+transgenesis.">Microinjection and transgenesis.</a> Microinjection and Transgenesis: Strategies and Protocols. , 587-603 (1998).</li><li>Figard, L., Sokac, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+cell+shape+change+in+living+Drosophila+embryos.">Imaging cell shape change in living Drosophila embryos.</a> Journal of Visualized Experiments. (49), 2503 (2011).</li><li>Oesterle, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Pipette Cookbook 2018: P-97 and P-1000 Micropipette Pullers. , (2018).</li><li>Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+photographic+study+of+development+in+the+living+embryo+of+Drosophila+melanogaster.">A photographic study of development in the living embryo of Drosophila melanogaster.</a> Journal of Embryology and Experimental Morphology. 33 (3), 789-801 (1975).</li><li>Powsner, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effects+of+temperature+on+the+durations+of+the+developmental+stages+of+Drosophila+melanogaster.">The effects of temperature on the durations of the developmental stages of Drosophila melanogaster.</a> Physiological Zoology. 8 (4), 474-520 (1935).</li><li>Hunter, M. V., Willoughby, P. M., Bruce, A. E. E., Fernandez-Gonzalez, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxidative+Stress+Orchestrates+Cell+Polarity+to+Promote+Embryonic+Wound+Healing.">Oxidative Stress Orchestrates Cell Polarity to Promote Embryonic Wound Healing.</a> Developmental Cell. 47 (3), 377-387 (2018).</li></ol>

No conflicts of interests declared.

The significance of this method is that it utilizes the well-established protocol of microinjection in Drosophila embryos21,22,23,24,25,26,27 to enable new research regarding the ASR and accompanying actin rod assembly. A major advantage of injecting G-actinRed into live ...

This protocol describes how to inject G-actinRed to visualize the assembly of intranuclear actin rods in heat-stressed embryos that are undergoing an inducible Actin Stress Response (ASR)1. We developed this protocol to aid studies of the ASR, which in embryos leads to disrupted morphogenesis and reduced viability, and in adult human cell types is associated with pathologies including renal failure2, muscle myopathies3, and Alzheimer&#8217;s and Huntington&#8217;s Disease4,5,6,7,8. This ASR is induced by numerous cellular stresses, including heat shock9,10,11, oxidative stress4,6, reduced ATP synthesis12, and abnormal Huntingtin or &#946;-amyloid oligomerization4,5,6,7,9,13,14,15,16. A hallmark of the ASR is the assembly of aberrant actin rods in either the cytoplasm or nucleus of affected cells, which is driven by stress-induced hyperactivation of an actin interacting protein, Cofilin1,5,6,10. Unfortunately, key knowledge gaps remain regarding the ASR. For example, the function of the actin rods is not known. We do not understand why rods form in the cytoplasm of some cell types, but the nucleus of others. Nor is it clear whether the ASR is protective or maladaptive for cells or embryos undergoing stress. Finally, we still do not know the detailed mechanisms underlying Cofilin hyperactivation or actin rod assembly. Thus, this protocol provides a rapid and versatile assay to probe the ASR by visualizing actin rod formation and dynamics in the highly tractable experimental system of the living fruit fly embryo.
The protocol to microinject G-actinRed into living Drosophila embryos was initially developed to study the dynamics of normal cytoplasmic actin structures17 during tissue building events. In those studies, we found that G-actinRed injection did not adversely affect early developmental processes in the embryo, including cytokinesis or gastrulation17,18. We then modified the protocol, adapting the embryo handling and G-actinRed injection to allow imaging of actin rods in heat stressed embryos undergoing the ASR1. Other methods besides G-actinRed injection can be used to visualize actin in embryos. These methods rely on expressing fluorescent proteins (FPs) tagged to actin or to domains of actin binding proteins, such as Utrophin-mCherry, Lifeact, F-tractin-GFP, and Moesin-GFP (reviewed in19). However, using these FP probes requires caution because they can stabilize or disrupt some actin structures, do not equally label all actin structures20, and in the case of actin-GFP, are highly overexpressed &#8211; problematic for the analysis of rod assembly which is not only stress dependent but also actin concentration dependent1. Thus, G-actinRed is the preferred probe for rod studies in fly embryos, and the large size of the embryo allows its easy injection.
The workflow of this protocol is similar to other well-established microinjection techniques that have been used for injecting proteins, nucleic acids, drugs, and fluorescent indicators into Drosophila embryos21,22,23,24,25,26,27. However, following the microinjection of G-actinRed here, embryos are exposed to mild heat stress to induce the ASR and intranuclear actin rod assembly. For labs with access to flies and an injection rig, this method should be readily implementable and adaptable for specific lines of study in regard to the ASR, including its induction by different stresses or modulation in distinct genetic backgrounds.

<table><tbody><tr><td>Adenosine triphosphate (ATP)</td><td>Millipore-Sigma</td><td>A23835G</td><td>Component of G buffer</td></tr><tr><td>Apple juice, Mott&#039;s, 64 fl oz</td><td>Mott&#039;s</td><td>014800000344</td><td>Component of apple juice plates</td></tr><tr><td>Bacto Agar</td><td>BD</td><td>214010</td><td>Component of apple juice plates</td></tr><tr><td>Bleach, PureBright Germicidal, 6.0% sodium hypochlorite</td><td>KIK International</td><td>059647210020</td><td>For dechorionating embryos</td></tr><tr><td>Calcium chloride</td><td>Millipore-Sigma</td><td>C1016500G</td><td>Component of G buffer</td></tr><tr><td>Cell strainer, 70 &amp;mu;m</td><td>Falcon</td><td>352350</td><td>For collecting dechorionated embryos</td></tr><tr><td>Confocal microscope, LSM 880 34-channel with Airyscan</td><td>Zeiss</td><td>0000001994956</td><td>For imaging intranuclear actin rods</td></tr><tr><td>Desiccant</td><td>Drierite</td><td>24001</td><td>For desiccating embryos</td></tr><tr><td>Dissecting microscope, Stemi 508 Stereoscope with 8:1 zoom</td><td>Zeiss</td><td>4350649000000</td><td>For arranging embryos on agar wedge</td></tr><tr><td>Dissecting needle, 5 in</td><td>Fisher Scientific</td><td>08965A</td><td>For arranging embryos on agar wedge</td></tr><tr><td>Dithiothreitol (DTT)</td><td>Fisher Scientific</td><td>BP1725</td><td>Component of G buffer</td></tr><tr><td>Double-sided Tape, Scotch Permanent, 0.5 in x 250 in</td><td>3M</td><td>021200010323</td><td>For making embryo glue</td></tr><tr><td>Embryo collection cage</td><td>Genessee Scientific</td><td>59100</td><td>For housing adult flies and collecting embryos</td></tr><tr><td>Fine tip tweezers, Dumont Tweezer, Style 5</td><td>Electron Microscopy Sciences</td><td>72701D</td><td>For arranging embryos on agar wedge</td></tr><tr><td>Glass capillaries, Borosillicate glass, thin 1 mm x 0.75 mm</td><td>World Precision Instruments, Inc.</td><td>TW1004</td><td>For microneedles</td></tr><tr><td>Halocarbon oil 27</td><td>Millipore-Sigma</td><td>H8773100ML</td><td>For hydration of embryos</td></tr><tr><td>Heated stage incubator</td><td>Zeiss</td><td>4118579020000, 4118609020000, 4118609010000</td><td>For confocal imaging</td></tr><tr><td>Lab Tissue Wipers, KimWipes</td><td>Kimberly-Clark</td><td>34155</td><td>Lab tissue wipers</td></tr><tr><td>Light microscope, Invertoskop 40C Inverted Phase contrast microscope, refurbished</td><td>Zeiss</td><td>Discontinued</td><td>Injection microscope</td></tr><tr><td>Methyl-4-hydroxybenzoate</td><td>Millipore-Sigma</td><td>H36471KG</td><td>Component of apple juice plates</td></tr><tr><td>Microinjector, FemtoJet4x</td><td>Eppendorf</td><td>5253000025</td><td>Microinjector</td></tr><tr><td>Micro loader tips, epT.I.P.S. 20 &amp;mu;L</td><td>Eppendorf</td><td>5242956003</td><td>For loading microneedles</td></tr><tr><td>Micromanipulator and injection stage with x,y,z dials for needle adjustment</td><td>Bernard Instruments, Inc (Houston, TX)</td><td>Custom</td><td>For performing microinjections</td></tr><tr><td>Micropipette puller, Model P-97, Flaming/Brown</td><td>Sutter Instruments</td><td>P97</td><td>For pulling capillary tubes to make microneedles</td></tr><tr><td>Microscope cover glass 24x50-1.5</td><td>Fisher Scientific</td><td>12544E</td><td>For mounting embryos</td></tr><tr><td>Microscope slides, Lilac Colorfrost, Precleaned, 25 x 75 x 1mm</td><td>Fisher Scientific</td><td>22037081</td><td>For mounting embryos for injection</td></tr><tr><td>n-Heptane</td><td>Fisher Scientific</td><td>H3601</td><td>Component of embryo glue</td></tr><tr><td>Objective, 10x</td><td>Zeiss</td><td>Discontinued</td><td>10x objective for injection microscope</td></tr><tr><td>Objective, C-Apochromat 40x/1,2 W Korr. FCS</td><td>Zeiss</td><td>4217679971711</td><td>40x water objective for confocal</td></tr><tr><td>Objective, LD LCI Plan-Apochromat 25x/0.8 Imm Cor DIC M27 for oil, water, silicone oil or glycerine immersion (D=0-0.17mm) (WD=0.57mm at D=0.17mm)</td><td>Zeiss</td><td>4208529871000</td><td>25x mixed immersion objective for confocal</td></tr><tr><td>Objective, Plan-Apocrhomat 63x/1.40 Oil DIC f/ELYRA</td><td>Zeiss</td><td>4207829900799</td><td>63x oil objective for confocal</td></tr><tr><td>Paintbrush, Robert Simmons Expression E85 Pointed Round size 2</td><td>Daler-Rowney</td><td>038372016954</td><td>For transferring embryos</td></tr><tr><td>Paper towels, Kleenex C-fold paper towels, white</td><td>Kimberly-Clark</td><td>884266344845</td><td>For blotting cell strainer</td></tr><tr><td>Pasteur pipette, 5 3/4 in</td><td>Fisher Scientific</td><td>1367820A</td><td>For covering embryos with oil</td></tr><tr><td>Petri dish, glass, 100 x 20 mm</td><td>Corning</td><td>3160102</td><td>For humid incubation chamber</td></tr><tr><td>Petri dish, plastic, 60 x 15 mm</td><td>VWR</td><td>25384092</td><td>For apple juice plates</td></tr><tr><td>Pipette, Eppendorf Reference 0.5-10 &amp;mu;L</td><td>Eppendorf</td><td>2231000604</td><td>For loading the microneedle</td></tr><tr><td>Pipette tip, xTIP4 250 &amp;mu;L</td><td>Biotix</td><td>63300006</td><td>For adding embryo glue to coverslip</td></tr><tr><td>Razor blade</td><td>VWR</td><td>55411050</td><td>For cutting agar wedge, tape, pipette tips</td></tr><tr><td>Rhodamine-conjugated globular actin, human platelet (non-muscle; 4x10 &amp;mu;g)</td><td>Cytoskeleton, Inc.</td><td>APHR-A</td><td>G-actin^Red</td></tr><tr><td>Scintillation vial, 20 mL Glass borosillicate with polyethylene liner and urea caps</td><td>Fisher Scientific</td><td>033377</td><td>For making embryo glue</td></tr><tr><td>Screw top jar, 16 oz</td><td>Nalgene</td><td>000194414195</td><td>For desiccating embryos</td></tr><tr><td>Stage micrometer</td><td>Electron Microscopy Sciences</td><td>602104PG</td><td>For calibrating volume of G-actin injection</td></tr><tr><td>Sucrose</td><td>Millipore-Sigma</td><td>840971KG</td><td>Component of apple juice plates</td></tr><tr><td>Trizma base</td><td>Millipore-Sigma</td><td>T15031KG</td><td>Component of G buffer</td></tr><tr><td>Yeast, Lesaffre Yeast Corporation Yeast, Red Star Active Dry, 32 oz</td><td>Lesaffre Yeast Corporation</td><td>117929157002</td><td>Component of yeast paste</td></tr></tbody></table>

imaging intranuclear actin rods in live heat stressed drosophila embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin rods are a hallmark of a conserved, inducible Actin Stress Response (ASR) that accompanies human pathologies, including neurodegenerative disease. Previously, we showed that the ASR contributes to morphogenesis failures and reduced viability of developing embryos. This protocol allows the continued study of mechanisms underlying actin rod assembly and the ASR in a model system that is highly amenable to imaging, genetics and biochemistry. Embryos are collected and mounted on a coverslip to prepare them for injection. Rhodamine-conjugated globular actin (G-actinRed) is diluted and loaded into a microneedle. A single injection is made into the center of each embryo. After injection, embryos are incubated at elevated temperature and intranuclear actin rods are then visualized by confocal microscopy. Fluorescence recovery after photobleaching (FRAP) experiments may be performed on the actin rods; and other actin-rich structures in the cytoplasm can also be imaged. We find that G-actinRed polymerizes like endogenous G-actin and does not, on its own, interfere with normal embryo development. One limitation of this protocol is that care must be taken during injection to avoid serious injury to the embryo. However, with practice, injecting G-actinRed into Drosophila embryos is a fast and reliable way to visualize actin rods and can easily be used with flies of any genotype or with the introduction of other cellular stresses, including hypoxia and oxidative stress.

A schematic workflow of embryo handling is depicted in Figure 1, and a timetable for a typical experiment is presented in Table 1. An estimate for a good experimental outcome is that for every 10 embryos injected, at least half of the embryos viewed will be at the correct developmental stage, undamaged, and exhibit a robust ASR with heat stress at 32 &#176;C. This ASR will be evidenced by the assembly of intranuclear actin rods as shown in the representative surface view ima...

Watch this Scientific Journal Video about Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos at JoVE.com

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The goal of this protocol is to inject Rhodamine-conjugated globular actin into Drosophila embryos and image intranuclear actin rod assembly following heat stress.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin ...

imaging-intranuclear-actin-rods-live-heat-stressed-drosophila

Research

JoVE Journal

Developmental Biology

3.4K Views.  Baylor College of Medicine. Het doel van dit protocol is om Rhodamine-geconjugeerde bolvormige actine te injecteren in Drosophila embryo's en beeld intranucleaire actine staaf assemblage na hitte stress.

Video: Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The Drosophila larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila is well known for the ease of powerful genetic manipulations and the larval nervous system has proven particularly useful in studying not only normal function but also perturbations that accompany some neurological disease (Lloyd and Taylor, 2010). Many key synaptic molecules found in Drosophila are also found in mammals and like most CNS excitatory synapses in mammals, the Drosophila NMJ is glutamatergic and demonstrates activity-dependent remodeling (Kohet al. , 2000). Additionally, Drosophila neurons can be individually identified because their innervation patterns are stereotyped and repetitive making it possible to study identified synaptic terminals, such as those between motor neurons and the body-wall muscle fibers that they innervate (Keshishian and Kim, 2004). The existence of evolutionarily conserved synapse components along with the ease of genetic and physical manipulation make the Drosophila model ideal for investigating the mechanisms underlying synaptic function (Budnik, 1996).
The active zones at synaptic terminals are of particular interest because these are the sites of neurotransmitter release. NC82 is a monoclonal antibody that recognizes the Drosophila protein Bruchpilot (Brp), a CAST1/ERC family member that is an important component of the active zone (Waghet al. , 2006). Brp was shown to directly shape the active zone T-bar and is responsible for effectively clustering Ca2+ channels beneath the T-bar density (Fouquetet al. , 2009). Mutants of Brp have reduced Ca2+ channel density, depressed evoked vesicle release, and altered short-term plasticity (Kittelet al. , 2006). Alterations to active zones have been observed in Drosophila disease models. For example, immunofluorescence using the NC82 antibody showed that the active zone density was decreased in models of amyotrophic lateral sclerosis and Pitt-Hopkins syndrome (Ratnaparkhiet al. , 2008; Zweieret al. , 2009). Thus, evaluation of active zones, or other synaptic proteins, in Drosophila larvae models of disease may provide a valuable initial clue to the presence of a synaptic defect.
Preparing whole-mount dissected Drosophila larvae for immunofluorescence analysis of the NMJ requires some skill, but can be accomplished by most scientists with a little practice. Presented is a method that provides for multiple larvae to be dissected and immunostained in the same dissection dish, limiting environmental differences between each genotype and providing sufficient animals for confidence in reproducibility and statistical analysis.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

The Drosophila  larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila  is well known for ...

Neuroscience

Imaging Calcium in Drosophila at Egg Activation

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. One aspect of egg activation, conserved across all organisms examined, is a change in the intracellular concentration of calcium (Ca2+) often termed a 'Ca2+ wave'. While the speed and number of oscillations of the Ca2+ wave varies between species, the change in intracellular Ca2+ is key in bringing about essential events for embryonic development. These changes include resumption of the cell cycle, mRNA regulation, cortical granule exocytosis, and rearrangement of the cytoskeleton.
In the mature Drosophila egg, activation occurs in the female oviduct prior to fertilization, initiating a series of Ca2+-dependent events. Here we present a protocol for imaging the Ca2+ wave in Drosophila. This approach provides a manipulable model system to interrogate the mechanism of the Ca2+ wave and the downstream changes associated with it.

Imaging Calcium in Drosophila at Egg Activation

This article describes an adaptable ex vivo protocol for visualizing Ca2+ during egg activation in Drosophila.

Egg activation is a universal process that includes a series of events to allow the fertilized egg to complete meiosis and initiate embryonic development. ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

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 ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Biology

Real-Time Imaging of Germline Stem Cells in Drosophila Ex Vivo Germaria Culture

Extended Live Imaging of Female Drosophila melanogaster Germline Stem Cell Niches

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to explore the dynamics of a myriad of developmental processes, such as cell division, stemness, differentiation, migration, apoptosis, autophagy, cellular adhesion, etc., over time. Recently, we have implemented an extended ex vivo culture and live imaging of the female Drosophila GSC niche. Using a Drosophila line harboring a GFP::Par-1 transgene as an example, this method allows the visualization of the GSCs&#39; asymmetric division within their niche and the description of the changes in the spectrosome morphology along the cell cycle. Here, we present a detailed protocol for the ex vivo culture of Drosophila germaria, enabling prolonged visualization of the female GSC niche. Importantly, this protocol is broadly applicable to live imaging GSCs with multiple fluorescently tagged proteins of interest that are available in stock centers and/or in the Drosophila research community.

Here, we provide a detailed protocol to perform live imaging of the asymmetric division of germline stem cells (GSCs) in the Drosophila ovarian niche. We use a transgenic line that ubiquitously expresses a green fluorescent protein (GFP) fusion of the spectrosome protein Par-1.

Live imaging methods allow the analysis of dynamic cellular processes in detail and in real-time. The Drosophila ovary represents an excellent model to ...

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Viewing Microtubule Networks in Drosophila Neuromuscular Junctions and Muscle Cells

Using Drosophila Larval Neuromuscular Junction and Muscle Cells to Visualize Microtubule Network

The microtubule network is an essential component of the nervous system. Mutations in many microtubules regulatory proteins are associated with neurodevelopmental disorders and neurological diseases, such as microtubule-associated protein Tau to neurodegenerative diseases, microtubule severing protein Spastin and Katanin 60 cause hereditary spastic paraplegia and neurodevelopmental abnormalities, respectively. Detection of microtubule networks in neurons is advantageous for elucidating the pathogenesis of neurological disorders. However, the small size of neurons and the dense arrangement of axonal microtubule bundles make visualizing the microtubule networks challenging. In this study, we describe a method for dissection of the larval neuromuscular junction and muscle cells, as well as immunostaining of &#945;-tubulin and microtubule-associated protein Futsch to visualize microtubule networks in Drosophila melanogaster. The neuromuscular junction permits us to observe both pre-and post-synaptic microtubules, and the large size of muscle cells in Drosophila larva allows for clear visualization of the microtubule network. Here, by mutating and overexpressing Katanin 60 in Drosophila melanogaster,&#160;and then examining the microtubule networks in the neuromuscular junction and muscle cells, we&#160;accurately reveal the regulatory role of Katanin 60 in neurodevelopment. Therefore, combined with the powerful genetic tools of Drosophila&#160;melanogaster, this protocol greatly facilitates genetic screening and microtubule dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Author Spotlight: Understanding Microtubule Network in Drosophila Neuromuscular Junctions 

Here, we present a detailed protocol to visualize the microtubule networks in neuromuscular junctions and muscle cells. Combined with the powerful genetic tools of Drosophila melanogaster, this protocol greatly facilitates genetic screening and microtubule-dynamics analysis for the role of microtubule network regulatory proteins in the nervous system.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

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