UD, WFM and BR are supported by the Center for Cellular Construction, a National Science Foundation (NSF) Science and Technology Center, under grant agreement DBI-1548297. BR is also supported through an NSF CAREER award, 1553695.

<ol>
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The authors have nothing to disclose.

The protocol described here is highly versatile and easily adaptable for studying biological events in a variety of stages in Drosophila embryonic development. This protocol is well suited for studying biological processes that occur over long periods of time. For example, studies of cellularization29,30 or the formation of heterochromatin domains in Drosophila31,32 could benefit from utilizing the methods described in this protocol since these processes occur over an extended period of time. In addition, sample size can be increased by decreasing magnification to study questions that require less optical resolution, such as the timing between syncytial divisions33 or duration of gastrulation34. &#160;Likewise, a 20x objective allows for two embryos to be imaged within one imaging plane. The methods in this protocol provide a dynamic platform to ask biological questions using embryonic development as a model system for quantitative analysis.
There are 3 critical steps in this protocol. Post dechorionation in 50% bleach it is imperative that embryos are rinsed and dried vigorously 5 times (2.3.4). This step removes any leftover small pieces of chorion. Leftover chorion will obstruct the field of view when imaging the embryo. When mounting the embryos onto the glass coverslip (3.5), it is important to press the embryos onto the glue with uniform pressure. This will place the embryos on the same z plane. When cutting open the microinjection needle, it is important to retract the plunger immediately to adjust the flow rate of the needle or you will lose all your injectant.
Our study was limited by two factors. The first factor was not having an automated segmentation process. We designed a manual segmentation script using MATLAB, however; with a genetically encoded spindle marker this process could be automated. In the future we would like to produce CNN-RFP21, Rtln1-GFP and CNN-RFP; ReepB-GFP transgenic lines to enable automated spindle segmentation. For more information regarding our MATLAB script and analysis pipeline, refer to supplemental materials of Diaz et al.15. Secondly, we were also limited by our acquisition interval of 35 s, which only allowed us to image 6 embryos simultaneously. A longer acquisition interval would allow us to image more embryos simultaneously. Although drug delivery throughput can be increase by replacing microinjection with a permeation step, we found this to be unnecessary since our sample size was already limited by acquisition interval.
While this protocol was used to image Drosophila embryonic development, the overall approach is adaptable for studying embryonic development in other multicellular model systems, such as C. elegans, Sea Urchins, and Xenopus. Embryos are extremely useful for quantitative analysis since they are easily synchronized via collection or fertilization. In addition, embryos don&#8217;t move or swim away from a field of view, enabling the use of a simple multipoint acquisition capable software to produce multiple time lapse videos for quantitative analysis. Although we used a high content analyzer in our study, any inverted microscope system capable of multipoint acquisition can be paired with the methods referred to herein.
Similar results can be obtained using a multipoint acquisition capable inverted confocal microscope; however, the advantages of a high content analyzer emerge as one increases sample size to study questions that require less resolution in time. An obvious advantage to using a high content analyzer is the ability to image 4 separate slides simultaneously compared to 1 slide on traditional systems. With a full set of 4 slides and 40 embryos on each slide, 160 embryos can be imaged over several hours, so long as the acquisition interval is at least 15 min between frames. Another important advantage to using a high content analyzer is that the samples are completely sealed off from any external light sources, which is necessary for fluorescence intensity-based measurements. Lastly, the high content analyzer used in our study has a 5.5-megapixel CMOS camera which provides a generous 221.87 &#956;m field of view at 60x magnification. This larger field of view enabled us to image half an embryo per acquisition point.&#160;&#160;

Over the past 30 years, advances in imaging technologies and molecular probes for live cell imaging have advanced our understanding of the complexity and inner workings of the cell. Accompanying this headway in technology is a greater reliance on the use of ex vivo cell culture models for the acquisition of high throughput imaging data. Cell culture models provide several advantages, including control of the microenvironment through microfluidic systems, and the ability to image multiple cells simultaneously, enabling the use of quantitative approaches necessary for high-throughput screening1,2. However, there are also significant disadvantages associated with cell culture models in the context of morphological studies, such as two-dimensional glass or plastic surfaces producing confounding effects on cell morphology and its regulation3,4,5. These effects can provide false or misleading data with regards to the study of biological processes regulating cellular morphology.
Although the alternative has been to study cellular morphology in the context of an intact organism6, few suitable intact organisms facilitate high throughput quantitative live cell imaging due to difficulties in keeping whole organisms alive through out imaging, and challenges associated with imaging inside a large multicellular organism. As a result, cell biological studies in intact organisms have generally focused on observing a single organism over time, which greatly limits sample size. In addition, this one-animal-at-a-time limitation makes live imaging difficult and costly to employ in the form of a screen and restricts the ability to apply quantitative analytical tools since the number of samples is generally too small. Thus, arises the need for a multicellular model organism that can be used for high content based quantitative approaches.
This protocol adapts the Drosophila embryo for plate-based high content imaging to generate experimental sample sizes large enough for quantitative analysis. Drosophila melanogaster is commonly known as the first model organism. Research using Drosophila has led to breakthroughs in cell and molecular biology, including the transmission of genetic information, identification of cell signaling pathways, and genetic requirements of embryonic development7,8,9. In addition, the Drosophila embryo has been used to explore in vivo microtubule dynamics during cellular division10,11,12. The use of microinjection to deliver small molecules and molecular probes provides temporal control that makes Drosophila embryonic development particularly powerful for probing cellular processes in vivo13,14. However, generating a significant number of experimental repeats with a traditional imaging approach is extremely labor intensive and has limited use in high-throughput imaging screens and quantitative analysis. Our approach makes use of a high content analyzer typically reserved for single cell analysis and adapts in vivo imaging analysis in the syncytium of the early Drosophila embryo.
This protocol was used to collect, stage, and microinject Drosophila embryos to explore the mechanisms that drive morphological changes in the Endoplasmic Reticulum (ER) during mitosis15. Although many studies have outlined the dynamic nature of the ER during the cell cycle16,17,18,19, it has been difficult to compare the effects of multiple perturbations across time on ER morphology. To identify the components that drive changes in ER morphology across cell division, the methods listed in this protocol were used to probe the role of microtubules and other cytoskeletal factors on mitotic ER reorganization using the cortical syncytial division in the early Drosophila embryo. Taken together, the availability of genetic perturbation within the Drosophila community, the ability to microinject drugs and molecular probes at precise times during development, and the inexpensive cost of working with Drosophila makes this approach highly amenable to high-throughput image-based screening. The methods described here are applicable for studying a wide variety of cellular and developmental processes in vivo using the Drosophila embryo20. Specifically, this protocol is well adapted to study biological events that occur over an extended period and within 100 microns of the Drosophila embryo cortex.

<table><tbody><tr><td>10 x 35mm Micro Dish</td><td>VWR</td><td>Cat #: 10799-192</td><td>N/A</td></tr><tr><td>100% grape juice concentrate</td><td>Welch&#039;s, 100% Concentrate Grape Juice</td><td>N/A</td><td>Purchase at Grocery Store</td></tr><tr><td>250ml Erlenmeyer Flasks, Narrow Mouth</td><td>VWR</td><td>Cat #: 10536-914</td><td>N/A</td></tr><tr><td>3&quot; Non-Metallic Sieve No. 140</td><td>Gilson Company</td><td>SV-165 #140</td><td>N/A</td></tr><tr><td>4 Slide Imaging Tray</td><td>Grace Biolabs</td><td>SKU: 400104</td><td>N/A</td></tr><tr><td>ACROS Organics , n-Heptane, 99+%, for spectroscopy</td><td>Fisher Scientific</td><td>Cat #: AC411255000</td><td>N/A</td></tr><tr><td>Acros Organics, Ethanol, 99.5%, ACS reagent, absolute, 200 proof</td><td>Fisher Scientific</td><td>Cat# : AC615095000</td><td>N/A</td></tr><tr><td>Acros Organics, Methyl 4-hydroxybenzoate, 99% . (methyl paraben )</td><td>Fisher Scientific</td><td>Cat#: AC126961000</td><td>N/A</td></tr><tr><td>Cover Slip, Rectangular, 25 x 75mm, #1 Float Glass</td><td>Fisher Scientific</td><td>Cat #: 50-143-782</td><td>N/A</td></tr><tr><td>Culture Well Silicone Sheet Material - 1.6mm thick</td><td>Grace Biolabs</td><td>SKU: 664273</td><td>N/A</td></tr><tr><td>DWK Life Sciences Wheaton 20mL PET Liquid Scintillation Vials</td><td>Fisher Scientific</td><td>Cat #: 03-341-71A</td><td>N/A</td></tr><tr><td>Embryo Collection Cage-Mini</td><td>Genesee Scientific</td><td>Cat #: 59-105</td><td>N/A</td></tr><tr><td>Eppendorf Microloader Pipette Tips</td><td>Fisher Scientific</td><td>Product Code: 10289651</td><td>N/A</td></tr><tr><td>Falcon 15mL Conical Centrifuge</td><td>Fisher Scientific</td><td>Cat #: 05-527-90</td><td>N/A</td></tr><tr><td>Falcon 50mL Conical Centrifuge Tubes</td><td>Fisher Scientific</td><td>Cat #: 14-432-22</td><td>N/A</td></tr><tr><td>Fleischmann&#039;s, Active Dry Yeast</td><td>Fleischmann&#039;s</td><td>N/A</td><td>Purchase at Grocery Store</td></tr><tr><td>Grape Plates</td><td>REFFER TO RECIPE PREP</td><td>N/A</td><td>N/A</td></tr><tr><td>Halocarbon 27 Oil</td><td>Genesee Scientific</td><td>Cat #: 59-133</td><td>N/A</td></tr><tr><td>Halocarbon 700 Oil</td><td>Genesee Scientific</td><td>Cat #: 59-131</td><td>N/A</td></tr><tr><td>Heptane Glue</td><td>REFFER TO RECIPE PREP</td><td>N/A</td><td>N/A</td></tr><tr><td>Industrial Razor Blades (Surgical Carbon Steel) Single Edged No. 9</td><td>VWR</td><td>Cat #: 55411-050</td><td>N/A</td></tr><tr><td>Parafilm M Laboratory Sealing Film, 4 inch x 250 foot Roll</td><td>Fisher Scientific</td><td>Cat #: 50-998-944</td><td>N/A</td></tr><tr><td>Scotch Double Sided Tape with Dispenser, Narrow Width, Engineered for Bonding, 1/2&quot; x 7 yds., 3/Pack Caddy</td><td>Fisher Scientific</td><td>Cat #: NC0879005</td><td>N/A</td></tr><tr><td>Silica gel beads, desiccant ~ 2 - 5 mm</td><td>Millipore Sigma</td><td>SKU: 1077351000</td><td>N/A</td></tr><tr><td>Stainless Steel Spoon/Spatula</td><td>VWR</td><td>Cat #: 470149-044</td><td>N/A</td></tr><tr><td>W.A. Hammond Drierite Indicating Absorbents</td><td>Fisher Scientific</td><td>Cat #: 23-116582</td><td>N/A</td></tr><tr><td>Yeast Paste</td><td>REFFER TO RECIPE PREP</td><td>N/A</td><td>N/A</td></tr><tr><td>&lt;strong&gt;EQUIPMENT&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Allegra X-12 Benchtop, Centrifuge</td><td>Beckman Coulter</td><td>N/A</td><td>You will need access to 15 ml conical centrifuge inserts.</td></tr><tr><td>Autoclave</td><td>N/A</td><td>N/A</td><td>Any autoclave will do fine.</td></tr><tr><td>Drummond Nanoject II Microinjector.</td><td>Drummond Scientific</td><td>N/A</td><td>This is a mineral oil based injection system.</td></tr><tr><td>GE Healthcare IN Cell Analyzer 6000.</td><td>GE healthcare</td><td>N/A</td><td>A fast confocal microscope capable of multipoint acquisition may be substituted.</td></tr><tr><td>Refrigerator (4C&amp;deg;)</td><td>N/A</td><td>N/A</td><td>Fly media and yeast paste must be stored at (4C&amp;deg;).</td></tr><tr><td>Zies Stemi 508 Stereo Microscope with Transillumination base 300 and overhead gooseneck lighting.</td><td>Zeiss Microscopy</td><td>N/A</td><td>The transillumination base and gooseneck lighting are necessary.</td></tr></tbody></table>

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.

The approach presented in this protocol was used to examine the role of microtubules in Endoplasmic Reticulum (ER) reorganization during mitosis in the Drosophila embryo15. During mitosis, the structure of the ER displays a dramatic reorganization, however, the forces that drive these changes are poorly understood. Recently, a family of ER shaping proteins were shown to promote the formation of ER tubule25,26,27,28, which are known for their close proximity with microtubules28. To study the roll of microtubules on ER morphology, the methods provided in the protocol were used to generate data to quantitatively compare the effects of several microinjected drug treatments on ER reorganization during mitosis. Colchicine, which prevents new microtubule polymerization, was found to drastically reduce the reorganization of the ER during mitosis as shown in Figure 5 and Figure 6. Furthermore, the approach described here enabled the production of time laps imaging data for 32 embryos. Mean and max intensity measurements were produced from 12,800 regions of interest using a custom MATLAB script, which also generated descriptive and inferential statistics vital for making comparisons between drug treatments. Due to the availability of molecular probes and the ease of microinjections, the methods described here are easily adaptable to study a variety of biological processes via fluorescence intensity quantification.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/61589/61589fig5.jpg" /> 
Figure 5: Representative Images of Rtnl1-GFP and ReepB-GFP enrichment at the spindle poles during mitosis. (A) 60x field of view of ReepB-GFP during mitosis in cell cycle 10. The blue square represents the field of view used for panels B-E. Panel A is processed with a threshold binary filter. This filter was not applied to Panels B-E since it excludes data from pixels below the set threshold. (B,D) ReepB-GFP in control, and in colchicine. (C,E) Rtnl1-GFP in control, and in colchicine. This figure has been modified from Diaz et al.15. <a href="https://www.jove.com/files/ftp_upload/61589/61589fig5large.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/61589/61589fig6.jpg" /> 
Figure 6: Quantitation of Rtnl1-GFP and ReepB-GFP enrichment at the spindle poles during mitosis. 20 spindles and 20 cytoplasm ROIs across 10 frames were analyzed for 32 embryos in the following conditions. (A) 10% DMSO control injections for Rtnl1-GFP. Control embryos showed a 0.1-fold increase in enrichment (p&#60;0.001 for all samples at T210) compared to non-injected samples (p &#60; 0.001 for all samples at T210). (B) ReepB-GFP embryos injected with 10% DMSO to serve as control. Control embryos showed a 0.1-fold increase in enrichment (p&#60;0.001 for all samples at T210) compared to non-injected samples (Figure 1E; p &#60; 0.001 for all samples at T210). (C) Rtnl1-GFP embryos injected with 5 &#956;M Colchicine. Here we measured a reduction in Rtnl1-GFP enrichment to spindles (p &#60; 0.001 for all samples at T210) compared to controls (A) (p &#60; 0.001 for all samples at T210). (D) ReepB-GFP embryos injected with 5 &#956;M Colchicine. We measured a reduction in ReepB-GFP enrichment to spindles (p&#60;0.001 for all samples at T210) compared to controls (B) (p&#60;0.001 for all samples at T210). This figure has been modified from Diaz et al.15. <a href="https://www.jove.com/files/ftp_upload/61589/61589fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>1. Yeast Paste21</td>
			<td>2. Grape Plates21</td>
			<td>3. Heptane Glue24</td>
		</tr>
		<tr>
			<td>1.1 Add 7 grams of active dry yeast to a 50 ml conical. Add 9 milliliters of DI water and mix to even constancy with a metal spatula.</td>
			<td>2.1 Make solution a: Add 6 grams of agar to 140ml of DI water in a 250ml flask and autoclave it.</td>
			<td>3.1 Fill a glass scintillation vial with 50 inches of double-sided tape.</td>
		</tr>
		<tr>
			<td>1.1 Add 7 grams of active dry yeast to a 50 ml conical. Add 9 milliliters of DI water and mix to even constancy with a metal spatula.</td>
			<td>2.2 Make solution b: Mix 0.1 grams of methyl paraben in 2ml of ethanol, then add this solution to 60ml of grapefruit juice concentrate.</td>
			<td>3.2 Add 20 ml of heptane and seal glass scintillation vial with parafilm. Rotate overnight at room temperature.</td>
		</tr>
		<tr>
			<td></td>
			<td>2.3 Immediately after autoclaving solution a, mix in solution b and pour mixture into 10 x 35mm perti dishes. (makes 35 grape plates)</td>
			<td>3.3 Remove plastic tape debris by transferring glue into 2 15 ml conical tubes and centrifuging at 3000 RPM for 15 minutes.</td>
		</tr>
		<tr>
			<td></td>
			<td>2.4 Allow grape plates to set at room temperature overnight with plate lids off. Cover plates with lids and store in 4C&#176; after they set.</td>
			<td>3.4 Transfer glue supernatant into 1.5 ml microcentrifuge tubes for storage.</td>
		</tr>
	</tbody>
</table>
Table 1: Recipes of media and solutions.

Watch this Scientific Journal Video about Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer at JoVE.com

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

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 ...

drosophila-embryo-preparation-microinjection-for-live-cell-microscopy

Research

JoVE Journal

Developmental Biology

8.5K Views.  San Francisco State University. Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

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

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Biology

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

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 ...

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.

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.

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

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 ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

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 ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Simultaneous Live Imaging of Multiple Insect Embryos in Sample Chamber-Based Light Sheet Fluorescence Microscopes

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample chamber-based setups, which are specifically designed to preserve the three-dimensional integrity of the specimen and usually feature sample rotation, are the best choice in developmental biology. For instance, they have been used to document the entire embryonic morphogenesis of the fruit fly Drosophila melanogaster and the red flour beetle Tribolium castaneum. However, many available live imaging protocols provide only experimental frameworks for single embryos. Especially for comparative studies, such approaches are inconvenient, since sequentially imaged specimens are affected by ambient variance. Further, this limits the number of specimens that can be assayed within a given time. We provide an experimental framework for simultaneous live imaging that increases the throughput in sample chamber-based setups and thus ensures&#160;similar ambient conditions for all specimens. Firstly, we provide a calibration guideline for light sheet fluorescence microscopes. Secondly, we propose a mounting method for multiple embryos that is compatible with sample rotation. Thirdly, we provide exemplary three-dimensional live imaging datasets of Drosophila, for which we juxtapose three transgenic lines with fluorescently labeled nuclei, as well as of Tribolium, for which we compare the performance of three transgenic sublines that carry the same transgene, but at different genomic locations. Our protocol is specifically designed for comparative studies as it pro-actively addresses ambient variance, which is always present in sequential live imaging. This is especially important for quantitative analyses and characterization of aberrational phenotypes, which result e.g., from knockout experiments. Further, it increases the overall throughput, which is highly convenient when access to light sheet&#160;fluorescence microscopes is limited. Finally, the proposed mounting method can be adapted for other insect species and further model organisms, e.g., zebrafish, with basically no optimization effort.

Light sheet-based fluorescence microscopy is the most valuable tool in developmental biology. A major issue in comparative studies is ambient variance. Our protocol describes an experimental framework for simultaneous live imaging of multiple specimens and, therefore, addresses this issue pro-actively.

Light sheet-based fluorescence microscopy offers efficient solutions to study complex processes on multiple biologically relevant scales. Sample ...

Drosophila melanogaster Embryo and Larva Harvesting and Preparation

Drosophila melanogaster embryos and larvae are easy to manipulate and develop rapidly by mechanisms that are analogous to other organisms, including mammals. For these reasons, many researchers utilize fly embryos and larvae to answer questions in diverse fields ranging from behavioral to developmental biology. Prior to experimentation, however, the embryos and larvae must first be collected.
This video will first demonstrate how "egg-laying cups" are used to collect Drosophila embryos on agar plates. The harvest and dechorionation of embryos will then be described. Next, the video will demonstrate how to identify and manipulate Drosophila in one of the three larval stages that follow the embryo stage. Finally, examples of some of the ways in which fly embryos and larvae are used in biological research are provided.

Drosophila melanogaster Embryo and Larva Harvesting and Preparation

Drosophila melanogaster embryos and larvae are easy to manipulate and develop rapidly by mechanisms that are analogous to other organisms, including ...

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

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The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

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

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

Preparing Individual Drosophila Egg Chambers for Live Imaging

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

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Simultaneous Live Imaging of Multiple Insect Embryos in Sample Chamber-Based Light Sheet Fluorescence Microscopes

Drosophila melanogaster Embryo and Larva Harvesting and Preparation