We thank Pakinee Phromsiri, Brian Jencik, Jinghong (James) Tang, and Roger White for their comments on the manuscript. We thank Patrick Oakes, Stefano Di Talia, and Victoria Deneke for sharing their expertise on how to perform PIV analysis. This work was supported by National Institutes of Health grants F31 HD100127 (to M. D. K.) and R01 GM102155 (to M. A. W).

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The authors have no conflicts of interest to disclose.

The Drosophila embryo is a powerful and convenient model to study fundamental questions in cellular and organismal biology. Its relative simplicity, powerful genetics, and small size make it an excellent system for imaging both cellular processes and development. Here, a standard microinjection protocol is adapted to enable FP usage in embryos. This approach allows for fluorescent imaging of specific cellular structures without the need for genetically encoded fluorophores, opening many genetic backgrounds to imaging. Combining multiple dyes plus strategically chosen fluorescently tagged proteins can open multichannel live imaging spanning the whole spectrum of visible light.
Critical steps in the protocol: 
This protocol uses BODIPY 493/503 to label LDs. This approach can easily be adapted to mark other cellular structures. For subsequent image analysis, one of the most important factors is the signal-to-noise ratio, i.e., the brightness of the dye compared to the background signal. Lysosomes have been successfully imaged (LysoTracker Red, 1 mM), as well as mitochondria (Mitoview 633, 200 &#181;M), the ER (ER tracker Green, 10 &#181;M), and microtubules (SiR tubulin, 200 nM in DMSO), as shown in Figure&#160;2, Figure 3, Figure 4, Figure&#160;5. In addition, yolk vesicles are autofluorescent and give off blue light upon UV excitation (image using 405 nm as the excitation wavelength (Figure 6)). For other dyes, aim for the dye concentration to be 100-1,000x of what would be needed for staining cultured cells live; this is similar to the concentration of a stock solution that would be diluted into cell culture media. As this protocol calls for an injection of 100 fL and the Drosophila embryo is roughly 9 nL in volume18, these dye concentrations will average out to an internal embryonic concentration of under 1/100th of what is present in the cell culture media. Temporarily, the local concentration will be higher at the site of injection, which is most relevant for FPs that do not diffuse well (i.e., mitochondrial dyes and SiR-tubulin). For these FPs, start at the recommended high concentrations; if unexpected death is observed, successively dilute two-fold until an acceptable compromise between survival and signal strength is reached.
When co-injecting multiple dyes, both dyes should either be in the same solvent, or both the solvents and dyes should be compatible with the mixture (alcohol concentrations exceeding ~10% are not recommended).
The quality of the needles is essential for the success of this procedure, as the tip needs to be as fine as possible. Otherwise, damage from the injection wound can compromise the subsequent development of the embryo. As commercial needle pullers differ, it is important to follow the suggestions of the manufacturer and try out multiple pulling parameters until the desired shape is achieved. It is critical to perform the quality control step 1.1.3 as working with a cracked, jagged, or large bore needle tip will make the successful injection more difficult or even impossible.
The embryo needs to be partially desiccated so that additional volumes of liquid can be added during the injection. If the embryo is under-desiccated, the needle will not enter easily, and cytoplasm will squirt out as the needle penetrates or as the solution is injected. If the embryo is over-desiccated, it will look deflated and will not develop properly. The exact drying time depends on local conditions, e.g., air humidity, and can change from day to day. It has to be determined empirically for each session.
The ability of the embryo to survive after microinjection depends critically on the quality of the needle, proper desiccation, and limiting injection volume to less than 1 pL (ideally 100 fL). As long as these parameters are optimized, no significant toxicity is apparent when the described dyes are injected at the recommended concentrations. If embryos survive the desiccation and injection steps, they typically develop successfully well into germ-band extension, an exception being the microtubule and ER probes which caused cellularization defects at high levels (100 fL injection of the stock concentration of each). Testing found no obvious developmental defects when DMSO, water, and mixtures of the two were injected at the recommended volumes, with an embryo survival rate through germ-band extension of ~75% or more. Injection volumes more than over 1 pL caused defects and embryos injected with ~4 pL volumes developed for less than 1 h. Therefore, injection volumes need to be kept low, which means that dye concentrations have to be high.
Generally, injections along the lateral edge of the embryo are recommended as those result in the least damage. However, the injection site may need to be adjusted depending on the diffusive properties of the FPs employed. BODIPY 493/503 and LysoTracker diffuse faster across the entire embryo than Lipid Spot 610 (another dye to mark LDs), while SiR-Tubulin and Mitoview 633 never diffuse fully across the embryo (imaging as late as 7 h post-injection). Thus, injection in or near the site of interest may be necessary. When injecting in the anterior or posterior regions, a particularly fine needle is recommended.
Image acquisition relies on confocal microscopy to optically section and resolve small organelles and all cytoskeletal components. Techniques requiring analysis of many images (e.g., STORM or PALM) will not work because the embryonic contents are in motion and the fluorophores are not optimized for photoswitching. Epifluorescence microscopy lacks the lateral and axial resolution to make out most organelles and smaller cellular structures. For these reasons, it is strongly recommended to use a confocal microscope or employ light sheet technology.
The reproducibility of the image analysis relies greatly on consistent imaging data. For the greatest chance of success, optimization of the injection technique and image acquisition is required. Establishing and practicing a technique where the dye(s) of interest, site of injection, age of the embryo, injection volume, and acquisition setup are all consistent will generate the most robust data for image analysis.
Modifications and troubleshooting of the method 
This protocol demonstrates a method for analyzing the bulk flow of LDs in cleavage stage embryos using particle image velocimetry. The same approach can be used for other organelles, other developmental stages, and other analysis methods. For example, Figure 2 shows analysis of LDs and acidic organelles flowing in the syncytial stages of embryogenesis, visualized by co-injecting BODIPY 493/503 and LysoTracker Red. Further successful imaging of LD motion in embryos up to 7 h post-fertilization has been achieved; these embryos do retain the injection wound but are able to develop for several hours.
Data gathered using this protocol has been used for particle image velocimetry, but many other analysis techniques are available. For example, particle tracking programs like those found in ImageJ, Imaris, or manual tracking can be used to obtain velocities and directionalities of moving structures. Note that most such tracking software are built to work with data from planar cell culture systems and do not always adapt well to 3D structures like the Drosophila embryo. Further, for the generation of the best quality particle tracking data, multiple Z planes would need to be imaged; this should be feasible if image stack acquisition times are under ~2 s. This benchmark should be reachable on spinning disc confocal, lattice light sheet, and recent laser scanning confocal systems. However, the feasibility of particle tracking for abundant organelles such as LDs, mitochondria, and lysosomes is low as the amount of positive signal in a field of view is too high for the current tracking methods. Tracking of less abundant structures like nuclei or yolk vesicles may be possible. PIV for flow analysis works well for LDs and acidic organelles in cleavage stages because both organelles move freely. Organelles like nuclei, ER, and mitochondria are tethered to other cellular structures and thus do not move freely and are not suited to software analysis that assumes free motion. The investigator should pick the techniques best suited to the organelle of interest.
During the syncytial and cellular blastoderm stages, LDs (as well as some other organelles) move along radially oriented microtubules5. It is therefore possible to find cross-sectional views (like in Figure 5) where single microtubules are in focus for long distances, allowing particle tracking in 2D. Since these optical planes are deep within the embryo, overall signal strength is diminished, and the signal-to-noise ratio is reduced.
For tracking analysis, imaging as fast as possible can reveal crucial details of the motion and thus of the motile machinery. For example, lipid-droplet motion is a mixture of two motile states, slow-short motion (~200 nm/s; average travel distance ~100 nm) and fast-long motion (~450 nm/s; average travel distance ~1,000 nm)19; thus, if images are taken every second or even less frequently, the slow-short state becomes undetectable. However, frequent imaging also induces fluorophore bleaching and phototoxicity. Imaging conditions, therefore, have to be adjusted depending on the exact question to be addressed.
Limitations of the method 
Depending on the dye desired, the method can be limited by the compatibility between the dye solubility and the toxicity of the injection solution. Alcohols like isopropanol and ethanol are difficult to handle within a needle due to their lower viscosity and appear to damage cellular components and kill the embryo.
The method is also not well suited for visualizing the earliest steps in embryogenesis because it takes 30+ min to prepare the embryos for injection. At room temperature, the initial cell cycles of the embryo are just ~10 min long each; so, even if one were to pick a newly fertilized egg in step 5.2.3, the first few cell cycles would already be completed by the time the embryo is ready for imaging.
Illumination with light in the UV/blue range is considerably more phototoxic than for longer wavelengths. Under such conditions (e.g., to follow autofluorescent yolk vesicles; Figure 6), one has to limit imaging time (leading to shorter time series) or use lower laser power (resulting in a reduced signal-to-noise ratio).
After cellularization, dyes injected in a specific location tend to diffuse poorly, as they must traverse many cell membranes. This limits the region of observation in later developmental stages.
The significance of the method with respect to existing/alternative methods 
The motion of LDs and other lipid-containing organelles in early embryos can be visualized with genetically encoded fluorophores, label-free techniques, and by the introduction of FPs. The latter can be achieved by permeabilization of the vitelline membrane12 or the microinjection approach discussed here.
Genetically encoded fluorophores are versatile markers whose levels are typically highly reproducible from embryo to embryo. However, they have lower quantum yields and bleach more easily than FPs. Typically, they are only available in one or two tagged version(s) (e.g., GFP or mCherry), limiting the choice of which structures can be imaged simultaneously. FPs, on the other hand, often exist in a large variety; for example, various lipid-droplet specific dyes are available with emission spectra from Autodot in the UV/blue spectrum to Lipidtox and LipidSpot 610 in the far-red spectrum. FPs can also be directly applied to any strain of interest, and thus do not require strain construction to, for example, introduce the desired organelle marker into a mutant strain of interest. This advantage is particularly pronounced when multiple structures are to be labeled simultaneously; instead of time-consuming crosses spanning multiple generations, this can be achieved in a single day by mixing the relevant dyes and introducing them at the same time. Finally, if cellular processes are to be probed with pharmacological inhibition, drugs and dyes can be introduced together.
Label-free methods are very powerful approaches for detecting specific cellular structures. For example, LDs can be specifically detected in early embryos by third-harmonic generation microscopy20 or by femtosecond Stimulated Raman Loss microscopy21. Like FPs, these approaches can be applied in any genetic background, and because they do not cause bleaching, they potentially allow for faster image acquisition. However, they are typically limited to specific organelles and thus do not by themselves support multiplex imaging; they also require specialized microscopes.
There are two general strategies for introducing small molecules into embryos. One is the microinjection approach employed here; the other is chemical (terpene) treatment to permeabilize the vitelline membrane. The latter approach12 is less involved than microinjection, but also more variable from embryo to embryo. In addition, after permeabilization, the protection provided by the vitelline membrane is compromised and the embryo proper is accessible to the external medium, making it more challenging to keep it alive. Microinjection is much less likely to derail embryonic development than permeabilization. However, permeabilization is recommended if many embryos need to be monitored simultaneously, e.g., for drug screening purposes. To follow the movement of cellular structures and obtain reproducible image series suitable for image analysis, microinjection is the method of choice.
Importance and potential applications of the method in specific research areas 
The Drosophila embryo is an important model system for studying many cell-biological and developmental processes1,5,6. Tagging organelles with fluorescent proteins has made major contributions to the understanding of how the early embryo develops, how various organelles traffic, and how such trafficking is modulated developmentally and genetically. However, their propensity to bleach and the challenges of generating strains in which multiple organelles are labeled with different colors limit the application of this approach. The use of FPs introduced by microinjection solves many of these challenges and can even be combined with fluorescently tagged proteins. This technique allows for the imaging of multiple organelles, cell structures, and cytoskeletal components in any genetic background. As a result, several genotypes can be compared via live imaging, making it possible to determine the effect of mutations on the trafficking of multiple organelles.
This protocol demonstrates the FP injection approach for embryos of Drosophila melanogaster, but in principle, this approach applies to any insect eggs for which microinjection techniques have been established, including other species of Drosophila, crickets22, and aphids23.

Vital dyes and molecular probes are powerful tools to image specific cellular structures and organelles live. In the Drosophila embryo, many different organelles display cytoskeleton-driven localization1,2,3,4, but the application of these small molecules is challenging because the eggshell is impermeable to many of them. This protocol describes a method to use fluorescent probes (FPs) in live embryos via microinjection in order to detect large-scale trafficking of organelles. The procedure covers preparation of the injection solution, egg collection and preparation of embryos, microinjection, imaging, and image analysis.
Dramatic spatial rearrangements of organelles are common in many animal oocytes, eggs, and embryos, in part because of the large size of these cells. In the Drosophila embryo, for example, lipid droplets (LDs) and yolk vesicles move toward the embryo center just before cellularization5. This motion depends on microtubules and leaves a ~40 &#181;m region all around the periphery of the embryo depleted of the two organelles. During earlier cleavage stages, many organelles are transported by cytoplasmic flows that are driven by actin-myosin-based contractions at the embryo surface6. Although the embryos of many species exhibit similar rearrangements, the Drosophila embryo is particularly suited for following these processes by imaging because it develops externally at standard laboratory &#39;room temperature&#39;, is relatively transparent, small enough to fit on most microscope setups, and can be manipulated using powerful genetic tools.
For some organelles, fluorescently tagged proteins are available that specifically label these structures. For example, LSD-2 (also known as dPLIN2) is a protein that in embryos specifically targets LDs7. Fly lines are available that carry either inducible transgenes encoding a fusion between Green Fluorescent Protein (GFP) and LSD-28 or a gene trap in which yellow fluorescent protein (YFP) is inserted into the coding region of the endogenous LSD-2 gene9,10. However, this approach has limitations, including that these fusion proteins have low quantum yields and tend to bleach easily. In addition, labeling multiple different structures simultaneously can be challenging: for many organelles, only one type of fluorescent tag (often GFP or mCherry) is currently available, so imaging two organelles at the same time may require new transgenes or insertions; also, even if compatible tags are available, introducing them into a single strain can require time-consuming crosses. It also makes using the many powerful genetic resources less convenient, e.g., if two organelle markers, a Gal4 driver, and an inducible RNAi construct all have to be present in the same mother.
In principle, these limitations can be overcome with the use of FPs, including vital dyes (e.g., LysoTracker to mark lysosomes), molecular probes (e.g., SiR-tubulin to label microtubules), and fluorescently labeled biological molecules (e.g., C12 BODIPY to probe fatty acid metabolism11). From use in cultured cells, they are typically well-validated as powerful tools for probing cellular biology. FPs are versatile, have superior photo properties, and are compatible with fluorescent proteins. Multiple dyes can be mixed and applied simultaneously, often with the benefits of separation into the violet/blue and far-red areas of the spectrum and small Stokes shifts, preventing channel bleed through. Small Stokes shifts allow for simultaneous capture of multiple imaging channels, enabling the tracking of several organelles at once. Finally, they can equally be applied to labeling organelles in the embryos of other Drosophila species or even other insects where no fluorescently tagged proteins may be available.
However, most of these FPs cannot traverse the elaborate eggshell of the Drosophila embryo. It consists of five layers: three outer chorionic layers (chorion) that prevent mechanical damage plus a waxy layer surrounding the vitelline membrane that creates a chemical barrier12. For simplicity, the combination of the waxy layer and the vitelline membrane will be referred to as the &#34;vitelline membrane&#34; below. To bypass the eggshell, this protocol adapts an established embryo microinjection approach to introduce FPs into the Drosophila embryo. The protocol describes how to monitor the cytoplasmic flow of LDs in cleavage stage embryos. It includes the preparation of injection needles and egg collection cages, the process of egg collection, and mechanical removal of the chorion. It goes over how to microinject and image the embryos and how to analyze the bulk flow of LDs using particle image velocimetry (PIV, adapted from6). It provides advice on troubleshooting to ensure embryo survival and create the best system for imaging. Also discussed is how the protocol can be modified to simultaneously image LDs and microtubules or to apply it to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the endoplasmic reticulum (ER).

<table><tbody><tr><td>AUTODO</td><td>abcepta</td><td>SM1000a</td><td>Stains lipid droplets in violet/blue</td></tr><tr><td>BODIPY 493/503</td><td>ThermoFisher</td><td>D3922</td><td>Stains lipid droplets in green</td></tr><tr><td>Desiccant Beads &amp;ndash;Desiccant-Anhydrous Indicating Drierite</td><td>W.A. Hammond Drierite Company</td><td>21001</td><td></td></tr><tr><td>Dissecting microscope SteREO DiscoveryV20 with transillumination base</td><td>Zeiss</td><td>4350030000000000</td><td></td></tr><tr><td>Double sided tape-Permanent Double-Sided Tape</td><td>Scotch (3M)</td><td>-</td><td>Sold by many vendors</td></tr><tr><td>ER-Tracker Green (BODIPY FL Glibenclamide)</td><td>ThermoFisher</td><td>E34251</td><td>Stains the ER/nuclear envelope</td></tr><tr><td>Femptotip II</td><td>Eppendorf</td><td>H129354N</td><td>Ready to use</td></tr><tr><td>Glass capillaries -Glass Thinw w/fil 1.0mm 4in</td><td>World Precision Instruments</td><td>TW100F-4</td><td>Must be pulled</td></tr><tr><td>Glass coverslip</td><td>-</td><td>-</td><td>Buy the appropriate refractive index for your objective lens</td></tr><tr><td>Glass slides -Double Frosted Microscope Slides precleaned</td><td>FisherBrand</td><td>12-550-343</td><td></td></tr><tr><td>Halocarbon oil 27</td><td>Sigma-Aldrich</td><td>H8773-100ML</td><td></td></tr><tr><td>Halocarbon oil 700</td><td>Sigma-Aldrich</td><td>H8898-50ML</td><td></td></tr><tr><td>Heptane&lt;br/&gt; greener alternative&lt;br/&gt; anhydrous, 99%</td><td>Sigma-Aldrich</td><td>246654-1L</td><td>Heptane is considered toxic by the USA&#039;s OSHA</td></tr><tr><td>Confocal microscope</td><td>Leica</td><td>Sp5</td><td>Many different types of confocal microscopes will work.</td></tr><tr><td>LipidSpot 610</td><td>Biotium</td><td>#70069</td><td>Stains lipid droplets in far red</td></tr><tr><td>LysoTracker Red</td><td>ThermoFisher</td><td>L7528</td><td>Stains lysosomes in red</td></tr><tr><td>MitoView 633</td><td>Biotium</td><td>#70055</td><td>Stains mitochondria</td></tr><tr><td>Needle loading pipette tips - 20uL microloader tips</td><td>Eppendorf</td><td># 5242956.003</td><td></td></tr><tr><td>P-1000, Next Generation Micropipette Puller</td><td>Sutter Instruments</td><td>P-1000</td><td>The settings used are heat-470, pull-70, velocity-60, delay-90, pressure-200, ramp-479 at 1x1. They refer to the intensity and length of the heating, the timing of pulling force and whether the force is applied linearly. Using these conditions, one capillary generates two usable needles with fine openings at the tip.</td></tr><tr><td>SiR-Tubulin</td><td>Cytosketon Inc</td><td>CY-SC014</td><td>Made by Spirochrome; Cytoskeleton Inc. is the North American distributor. Stains microtubules in far red</td></tr><tr><td>TransferMan</td><td>Eppendorf</td><td>&amp;nbsp;5178 NK</td><td></td></tr><tr><td>ViaFluor 405</td><td>Biotium</td><td>#70064</td><td>Stains microtubules in violet/blue</td></tr></tbody></table>

visualizing cytoskeleton-dependent trafficking of lipid-containing organelles in drosophila embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of embryogenesis, these organelles undergo dramatic movements powered by actin-based cytoplasmic streaming and motor-driven trafficking along microtubules. The development of a multitude of small, organelle-specific fluorescent probes (FPs) makes it possible to visualize a wide range of different lipid-containing structures in any genotype, allowing live imaging without requiring a genetically encoded fluorophore. This protocol shows how to inject vital dyes and molecular probes into Drosophila embryos to monitor the trafficking of specific organelles by live imaging. This approach is demonstrated by labeling lipid droplets (LDs) and following their bulk movement by particle image velocimetry (PIV). This protocol provides a strategy amenable to the study of other organelles, including lysosomes, mitochondria, yolk vesicles, and the ER, and for tracking the motion of individual LDs along microtubules. Using commercially available dyes brings the benefits of separation into the violet/blue and far-red regions of the spectrum. By multiplex co-labeling of organelles and/or cytoskeletal elements via microinjection, all the genetic resources in Drosophila are available for trafficking studies without the need to introduce fluorescently tagged proteins. Unlike genetically encoded fluorophores, which have low quantum yields and bleach easily, many of the available dyes allow for rapid and simultaneous capture of several channels with high photon yields.

Following injection, the dye will be localized only at the site where the needle tip was inserted. The dye will then diffuse away from the injection site depending on its diffusive characteristics. Figure 1 shows injection of BODIPY 493/503, soon after injection (panel A) and 24 min later (panel B). After 24 min, the dye has made it to roughly the midpoint of the embryo&#39;s long axis.
Analyzing organelle motility can be achieved through dye injections and time-lapse imaging. In Figure 2, an embryo was co-injected with BODIPY 493/503 (Figure 2A) and LysoTracker Red (Figure 2B) and imaged using laser excitation at 488 and 596 nm, respectively. This embryo was then time-lapse imaged (one frame every 30 s for 30 min, 5 min analyzed). The time series was then run through PIV analysis, the output of which is shown via streamlines in Figure 2A,B. Note that the streamlines do not represent the trajectory of individual particles, but the cytoplasmic flow is inferred from analyzing all the particles in that region of the cytoplasm. Through labeling of two independent cellular structures (LDs and acidic organelles), the PIV analysis finds similar flows, with both labels converging on the central region of the embryo where the cytoplasm is flowing into the embryo&#39;s interior6.
Currently available FPs allow the labeling of many other organelles and cellular structures. Figure 3 shows the labeling of the ER via ER tracker Green. ER tracker provides a nice resolution of the nuclear envelope, allowing visualization of major cell cycle stages. ER tracker Green is imaged using 488 nm excitation.
Labeling of the mitochondria is tricky, as most dyes tested seem to be trapped in the first mitochondrion they enter. On the other hand, no sign of dye toxicity was detected, making it possible to follow labeled mitochondria through cellularization and the ectodermal nuclear cycle 15. Figure 4 shows an ectodermal cell several hours post-injection with Mitoview 633 (excitation wavelength 633 nm).
BODIPY, Lysotracker, and LipidSpot are robust and can be used to acquire ~500+ images at 512 x 512, line average 3, frame rates from 1/s to 0.1/s. ER tracker Green, SiR-tubulin, and the mitochondrial dyes mentioned are less robust and yield ~50-200 images under the same conditions.
Starting at blastoderm stages, LDs move bidirectionally along microtubules, powered by the opposite-polarity motors kinesin-1 and cytoplasmic dynein5. This motion can be visualized by co-labeling LDs and microtubules via injecting both BODIPY 493/503 and SiR-Tubulin (Figure 5). As LDs frequently reverse their direction of movement (as they switch between kinesin-1 and cytoplasmic dynein), higher frame rates during acquisition better capture critical details of LD motility.
Live imaging of autofluorescent yolk vesicles is possible without any form of dye injection (Figure 6). However, the autofluorescence is dim, and the excitation laser is phototoxic. Thus, live imaging of autofluorescent yolk has a poor signal-to-noise ratio relative to dye injection.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63291/63291fig01.jpg" /> 
Figure 1: Diffusion of BODIPY 493/503 through the embryo. The dye was injected along the lateral edge toward the anterior end (top right) and diffuses from this injection site into the embryo, labeling LDs. (A) The dye has diffused through portions of the embryo adjacent to the injection site. (B) Roughly 24 min/2 nuclear cycles later, the dye has diffused past the midpoint of the embryo. Scale bar: 100 &#181;m. A 1024 x 1024 frame (line average 4) was acquired every 30 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63291/63291fig02.jpg" /> 
Figure 2: Particle image velocimetry (PIV) for LDs and acidic organelles. An embryo was injected with both BODIPY 493/503 and LysoTracker Red. (A) BODIPY channel. (B) LysoTracker Red channel. (A&#39;,B&#39;) Streamline diagrams generated by PIV analysis of the two channels generated from 10 sequential frames, including those shown in A and B. A&#39; corresponds to the flow of LDs, and B&#39; corresponds to the flow of acidic organelles. Note that both A&#39; and B&#39; show a left-of-center confluence where embryonic contents are flowing out of the plane of view, into the center of the embryo. Also, note that BODIPY has diffused more than LysoTracker as different dyes have different diffusive properties. Scale bar: 100 &#181;m. A 1024 x 1024 frame (line average 4) was acquired every 30 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63291/63291fig03.jpg" /> 
Figure 3: ER tracker labels syncytial nuclear divisions. A syncytial blastoderm embryo was microinjected with ER tracker and a portion of its surface was imaged over time. (A) Spindle assembly during a nuclear division. (B) Abscission of the nuclear envelope during the same division. (C)&#160;Subsequent interphase. (D) The onset of the next division is indicated by centrosome appearance (occurrence of circular ER-free regions, marked by arrowheads). Note the gradual dye bleaching. Excitation wavelength: 488 nm. Scale bar: 5 &#181;m. A: initial frame, B: 3 min elapsed, C: 10 min elapsed, D: 13 min elapsed. A 1024 x 1024 frame (line average 4) was acquired every 30 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63291/63291fig04.jpg" /> 
Figure 4: Mitoview 633 labeling of mitochondria. An embryo was injected with Mitoview 633 during the syncytial blastoderm stage and imaged 4 h later, after cellularization. The image shows a neuroectodermal cell of an embryo in germ-band extension. Scale bar: 5 &#181;m. A 1024 x 1024 frame (line average 4) was acquired every 30 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63291/63291fig05.jpg" /> 
Figure 5: Co-labeling of LDs and microtubules. A cellularizing embryo was injected with a mixture of BODIPY 493/503 (yellow A,B,C) and SiR Tubulin (magenta A&#39;,B&#39;,C&#39;). A&#39;&#39;, B&#39;&#39;, C&#39;&#39; show the merged channels. Panels A, A&#39;, and A&#39;&#39; show the initial frame, panels B, B&#39; and B&#39;&#39; show the frame after 5 s, and panels C, C&#39; and C&#39;&#39; show the frame after 10 s. Scale bar: 5 &#181;m. A 512 x 512 frame (line average 3) was acquired every 2.5 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63291/63291fig06.jpg" /> 
Figure 6: Imaging yolk vesicle autofluorescence during syncytial cleavage stages. (A) At the start of the acquisition. (B) After 8 min. (C) After 16 min. Excitation wavelength: 405 nm. Low excitation intensity was used to keep the embryo alive. Scale bar: 100 &#181;m. A 1024 x 1024 frame (line average 4) was acquired every 30 s. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/63291fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental data. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/63291/PIV_analysis_script_Kilwein_Welte_JoVE_2021.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos at JoVE.com

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

Early Drosophila embryos are large cells containing a vast array of conventional and embryo-specific organelles. During the first three hours of ...

visualizing-cytoskeleton-dependent-trafficking-lipid-containing

Research

JoVE Journal

Developmental Biology

2.1K Views.  University of Rochester. In the early Drosophila embryo, many organelles are motile. In principle, they can be imaged live via specific fluorescent probes, but the eggshell prevents direct application to the embryo. This protocol describes how to introduce such probes via microinjection, and then analyze bulk organelle motion via particle image velocimetry.

Video: Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Neuroscience

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Biology

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an embryonic fillet preparation by a microinjector. Each motor neuron whose membrane is contacted by the droplet can then be rapidly labeled. Individual motor neurons are continuously labeled, enabling fine structural details to be clearly visualized. Given that lipophilic dyes come in various colors, the technique also provides a means to get adjacent neurons labeled in multicolor. This tracing technique is therefore useful for studying neuronal morphogenesis and synaptic connectivity in the motor neuron system of Drosophila.

Retrograde Tracing of Drosophila Embryonic Motor Neurons Using Lipophilic Fluorescent Dyes

We describe a method for retrograde tracing of the Drosophila embryonic motor neurons using lipophilic fluorescent dyes.

We describe a technique for retrograde labeling of motor neurons in Drosophila. We use an oil-dissolved lipophilic dye and deliver a small droplet to an ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

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

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

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

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

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

Enhancing Protein Visualization with Antibody-Mediated Live Imaging

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

Visualization of proteins in living cells using GFP (Green Fluorescent Protein) and other fluorescent tags has greatly improved understanding of protein localization, dynamics, and function. Compared to immunofluorescence, live imaging more accurately reflects protein localization without potential artifacts arising from tissue fixation. Importantly, live imaging enables quantitative and temporal characterization of protein levels and localization, crucial for understanding dynamic biological processes such as cell movement or division. However, a major limitation of fluorescent tagging approaches is the need for sufficiently high protein expression levels to achieve successful visualization. Consequently, many endogenously tagged fluorescent proteins with relatively low expression levels cannot be detected. On the other hand, ectopic expression using viral promoters can sometimes lead to protein mislocalization or functional alterations in physiological contexts. To address these limitations, an approach is presented that utilizes highly sensitive antibody-mediated protein detection in living embryos, essentially performing immunofluorescence without the need for tissue fixation. As proof of principle, endogenously GFP-tagged Notch receptor that is barely detectable&#160;in living embryos can be successfully visualized after antibody injection. Furthermore, this approach was adapted to visualize post-translational modifications (PTMs) in living embryos, allowing the detection of temporal changes in tyrosine phosphorylation patterns during early embryogenesis and revealing a novel subpopulation of phosphotyrosine (p-Tyr) underneath apical membranes. This approach can be modified to accommodate other protein-specific, tag-specific, or PTM-specific antibodies and should be compatible with other injection-amenable model organisms or cell lines. This protocol opens new possibilities for live imaging of low-abundance proteins or PTMs that were previously challenging to detect using traditional fluorescent tagging methods.

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

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

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

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally important for cell function and cell integrity of neurons. Drosophila photoreceptor cells have become a model for studying membrane protein trafficking. Besides rhodopsin, which upon illumination becomes internalized from the photoreceptor membrane and is degraded, the transient receptor potential-like (TRPL) ion channel in Drosophila exhibits a light-dependent translocation between the rhabdomeral photoreceptor membrane (where it is located in the dark) and the photoreceptor cell body (to which it is transported upon illumination). This intracellular transport of TRPL can be studied in a simple and non-invasive way by expressing eGFP-tagged TRPL in photoreceptor cells. The eGFP fluorescence can then be observed either in the deep pseudopupil or by water immersion microscopy. These methods allow detection of fluorescence in the intact eye and are therefore useful for high-throughput assays and genetic screens for Drosophila mutants defective in TRPL translocation. Here, the preparation of flies, the microscopic techniques, as well as quantification methods used to study this light-triggered translocation of TRPL are explained in detail. These methods can be applied also for trafficking studies on other Drosophila photoreceptor proteins, for example, rhodopsin. In addition, by using eGFP-tagged rhabdomeral proteins, these methods can be used to assess the degeneration of photoreceptor cells.

Studying Membrane Protein Trafficking in Drosophila Photoreceptor Cells Using eGFP-Tagged Proteins

Here, non-invasive methods are described for localization of photoreceptor membrane proteins and assessment of retinal degeneration in the Drosophila compound eye using eGFP fluorescence.

Membrane protein trafficking regulates the incorporation and removal of receptors and ion channels into the plasma membrane. This process is fundamentally ...

Visualizing Cytoskeleton-Dependent Trafficking of Lipid-Containing Organelles in Drosophila Embryos

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