We are grateful to Laura Bampton, Alex Davidson, Richard Parton, Arita Acharya for assistance during the preparation of this manuscript; Mariana Wolfner for discussions on egg activation; Matt Wayland for imaging support; and Nan Hu for general support in the laboratory.
This work was supported by the University of Cambridge, ISSF to T.T.W. [grant number 097814].

Note: Carry out all steps at room temperature unless stated otherwise.
1. Preparing for Dissection
Note: The steps described here are in accordance with E. Gavis, Princeton University &#38; Weil et. al. 18. Perform the following steps two days before imaging.
<ol>
	<li>Take a fly vial containing approximately 10 ml of solid food and add a small amount of dried yeast to one corner, less than 1 g is sufficient. 
	Note: For information about fly vials or food see &#39;Drosophila Maintenance&#39; 19.</li>
	<li>Add 1 - 3 drops of water to hydrate the yeast.</li>
	<li>Set the vial aside at a 45&#176; angle and allow 3 - 5 min for the yeast to take up the water.</li>
	<li>While the yeast is absorbing the water, select a bottle of flies expressing UASt-myrGCaMP5 20 driven by tub-VP16GAL4 (referred to as GCaMP5) that have recently emerged (a myristoylated form of the GCaMP5 was chosen to provide a high signal to noise ratio through tethering the GECI to the membrane in the egg).</li>
	<li>Anaesthetize the flies in the bottle with CO2 gas. Be sure to keep the bottle inverted so as not to lose flies in the wet food.</li>
	<li>Tip the anesthetized flies onto a CO2 pad and sort 10 - 15 females and 5 males using a paintbrush under a dissecting microscope. Note: It is standard to include males to provide healthy females for dissection.</li>
	<li>When the vial from step 1.3 is ready, add the selected flies to the vial. Take care to keep the vial horizontal until the flies become active.</li>
	<li>Place the vial at 25 &#176;C for approximately 36 hr.</li>
	<li>Return the remaining flies from the CO2 pad to the bottle or discard.</li>
</ol>
2. Dissecting Drosophila Ovaries
Note: The steps described here are in accordance with Weil et. al. 18.
<ol>
	<li>After 36 hr, anaesthetize the flies from the yeasted vial with CO2.</li>
	<li>Once the flies inside the vial are anesthetized tip them onto the CO2 pad to be sorted with a paint brush under a dissecting microscope.</li>
	<li>Select a female.</li>
	<li>Isolate the two ovaries from the female. For full detailed protocol see Weil et. al. 18. In summary:
	<ol>
		<li>Place a small drop of oxygenated halocarbon oil (series 95) on a No. 1.5, 22 x 40 mm coverslip.</li>
		<li>Grasp the female with forceps at the anterior of the abdomen.</li>
		<li>Holding the female under the dissecting microscope, use the second set of forceps to gently remove the posterior of the female.</li>
		<li>Wipe off the posterior tissue from the second forceps.</li>
		<li>Squeeze on the abdomen cavity with the second forceps from the anterior to the posterior of the abdomen. The two opaque ovaries should stick to the forceps when outside of the female.</li>
		<li>Touch the ovaries into the oil.</li>
		<li>Remove any tissue from the forceps by wiping off the forceps.</li>
		<li>Repeat these steps (2.4.1 - 2.4.7) to obtain three coverslips, each containing ovaries from one fly.</li>
	</ol>
	</li>
</ol>
3. Isolating Individual Mature Eggs
<ol>
	<li>Under a standard dissecting microscope with the magnification set to 4X, on the first coverslip from step 2.4, separate the two ovaries by tearing the oviduct.</li>
	<li>Introduce a pair of closed forceps into the center of a single ovary, taking care not to puncture any egg chambers.</li>
	<li>Insert a standard dissecting probe into the center of the ovary next to the closed forceps.</li>
	<li>Gently drag the dissecting probe through the ovary towards the posterior pole, where the mature eggs (stage 14 egg chambers) are located.</li>
	<li>Repeat steps 3.1 - 3.4 2 - 5 times depending on the size of the ovary and number of eggs. 
	Note: Addition of CO2 typically causes the deposition of a single egg which will appear larger with raised dorsal appendages than pre-deposited eggs. This egg should be discarded for imaging since it has already been activated inside the female. 
	Note: Mature eggs are likely to separate easily from ovarian connective tissue.</li>
	<li>If no mature eggs are visible outside of the ovary, continue gently teasing with the dissecting probe.</li>
	<li>Once the mature eggs are visible on the coverslip outside of the ovary, maneuver 3 - 5 in a line in the center of the coverslip. 
	Note: For best results, run the probe gently along the side of the eggs. 
	Note: Avoid pulling on the dorsal appendages as this can damage the egg. 
	Note: Depending on the future imaging set-up, align the eggs approximately 200 &#181;m apart perpendicular to the coverslip for maximum imaging area.</li>
	<li>Once aligned, remove the rest of the ovarian tissue by dragging it away from the aligned eggs using the forceps.</li>
	<li>Remove excess oil by dabbing with the corner of a medical wipe. Be careful not to touch the mature eggs with the medical wipe as mature eggs that are contacted will be lost.</li>
	<li>Draw a crosshair on the coverslip with a marker to simplify locating the sample under the microscope.</li>
	<li>Set the coverslip in a safe place and leave the prepared egg chambers on the coverslip to rest for 5 - 10 min. This allows for the egg to settle in the oil. Samples can be used up to 1 hr post dissection.</li>
	<li>Repeat steps 3.1 - 3.11 for the rest of the coverslips with ovaries on them.</li>
</ol>
4. Preparing for Imaging
<ol>
	<li>Bring prepared activation buffer 15 to room temperature. 
	Note: Activation buffer is a hypotonic buffer: 3.3 mM NaH2PO4, 16.6 mM KH2PO4, 10 mM NaCl, 50 mM KCl, 5% polyethylene glycol 8,000, 2 mM CaCl2, brought to pH 6.4 with a 1:5 ratio of NaOH:KOH. For more information see Mahowald, 1983 15. Aliquots of activation buffer can be stored at -20 &#176;C for months and at 4 &#176;C for up to 2 weeks.</li>
	<li>Carefully transport prepared coverslips, activation buffer and glass pipettes to the imaging facility. 
	Note: A wide-field, confocal, spinning disc or light sheet microscope can be used. We recommend using an inverted microscope. Our representative results were obtained using a confocal microscope.</li>
	<li>An objective suitable for imaging the entire mature egg should be chosen (here: 20X 0.7 NA).</li>
	<li>Mount the first prepared coverslip on to the microscope stage. Take care not to break the coverslips on the stage.</li>
	<li>Qualitatively select a field of view with a mature egg chamber for activation. Do not image punctured egg chambers or those with blebbing in the membrane. 
	Note: For steps 4.6 - 4.7, software commands will vary depending on microscope and manufacturer.</li>
	<li>Set imaging parameters for standard GFP excitation (488 nm) and emission (500 - 580 nm). Also set up a bright-field view in order to visualize and orientate the mature egg and displaced oil.</li>
	<li>Select the lowest Z-plane where mature eggs are visible and set a full Z-stack to be taken for 40 &#181;m at approximately 2 &#181;m per step. Set the parameters so there is no delay between Z-stack acquisition. Set the time series to capture for approximately 30 min.</li>
</ol>
5. Imaging Ex Vivo Egg Activation
<ol>
	<li>Start image capture.</li>
	<li>Wait for 1 - 2 full Z-stacks to be acquired. This shows the mature egg prior to activation.</li>
	<li>Withdraw approximately 300 &#181;l of activation buffer into a glass pipette.</li>
	<li>Positioning the tip of the glass pipette containing activation buffer at a 45&#176; angle above the coverslip, slowly move the pipette into the beam path until it is directly above the mature egg being imaged. The glass pipette should be 1 - 2 cm above the oil.</li>
	<li>Add 1 - 2 drops of activation buffer in less than 5 sec from the glass pipette. This should displace the oil. Avoid adding excess activation buffer or touching the egg with the glass pipette as it can cause displacement of the mature egg. Take care not to touch the coverslip with the pipette as this can cause a change in the focal plane or air bubbles to form in the immersion oil between the objective and coverslip.</li>
	<li>Whilst adding activation buffer during image acquisition, check the bright-field channel to ensure that the oil has been displaced by the activation buffer. 
	Note: This will first appear as a shadow due to the pipette breaking the beam path but will also result in the appearance of small oil droplets. Some oil droplets can remain associated with the mature egg which will affect experimental outcomes and image quality.</li>
	<li>If the oil is not displaced from the initial addition of activation buffer repeat steps 5.4 - 5.6.</li>
	<li>Once the oil is successfully displaced, allow the time series to run until completion. 
	Note: Due to swelling of oocytes at activation, some egg chambers will move dramatically out of the focal plane or field of view upon addition of activation buffer. In this situation, stop the acquisition and mount the next coverslip.</li>
	<li>After the image sequence has completed acquisition, save data.</li>
</ol>
6. Post-acquisition Image Processing
<ol>
	<li>Open the data set using Fiji (http://fiji.sc/Downloads#Fiji), or equivalent image processing software, and select the relevant files for analysis.</li>
	<li>To build a projection of the acquired data select: Image &#62; Stacks &#62; Z-project.</li>
	<li>Select a start Z-slice and a stop Z-slice, typically the whole section. Select a projection type, typically set to &#39;Max Intensity&#39;. Check tick box &#39;All Time Frames&#39; to apply the selected settings to the whole series.</li>
	<li>If more than one fluorescent channel has been imaged, use the Channels tool to enable movement between colors, make a composite, or make the image grayscale. Select Image &#62; Colors &#62; Channels tool.</li>
	<li>To adjust the image brightness and contrast select Image &#62; Adjust &#62; Brightness/Contrast.</li>
	<li>To export a single image, move timescale slider to relevant frame and select File &#62; Save As &#62; Jpeg, or any preferred format.</li>
	<li>Choose a location and title for the exported file, then save. 
	Note The frame rate from the imaging software will be required to give a time stamp on the image.</li>
	<li>To export the time series as a movie select: File &#62; Save As &#62; AVI, then select frame rate (~ 7 frames per second).</li>
	<li>Choose a location and title for the exported file, then save.</li>
	<li>To save the adjusted file in Fiji format, select File &#62; Save.</li>
</ol>

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The authors have nothing to disclose.

The first critical step in this protocol is isolating the mature eggs without damage. This can be achieved by gentile maneuvering of the eggs with the dissecting probe. Practice will enable this manipulation to be executed without damage to the eggs. A second crucial step is avoiding the loss of the sample when activation buffer is applied to the mature egg in oil. Application of the activation buffer should be done slowly and without contact with the coverslip. This step can be challenging if the microscope set-up does not allow for an easy access to the sample. Moving axillary parts of the microscope, such as a temperature incubator, is advisable. 
Modifications to this protocol can be made in order to visualize other fluorescently tagged factors, instead of Ca2+ at egg activation. More generally, different stages of Drosophila development or the results of adding a different buffer could be analyzed using this protocol. 
Troubleshooting may be required if the settings on the microscope are not optimal for visualizing the signal from the sample. This can be achieved by increasing or decreasing the laser power, altering capture range and adjusting the Z-stack parameters. Another issue might be eggs consistently moving out of the field of view upon the addition of activation buffer. If this happens regularly, allow the mature eggs to settle on the coverslips for a longer period of time, 15 to 20 minutes. Beware that using a higher viscosity oil will alter the displacement of the oil by activation buffer. 
This technique presented here is limited by the working distance of the objective, dehydration of the mature eggs and the ability to dissect the mature eggs without damage. However, when compared to in vivo imaging methods where the mature oocyte passes through the adult female and is deposited 13, our method enables more spatial resolution, the option to physically manipulate the mature egg before activation and the ability to test the role of egg activation without fertilization. 
There are many potential future applications of this technique, including testing reporter constructs for cellular components at egg activation and visualizing the Ca2+ wave in mutant backgrounds 14,24. Together with experimental and genetic tools, the ex vivo egg activation assay presented here enables the study of the trigger, propagation and downstream effects of the universally conserved Ca2+ wave.

A change in intracellular Ca2+ concentration at egg (oocyte) activation is a conserved component of all studied organisms 1,2. This event initiates a wide range of Ca2+-dependent processes, including resumption of the cell cycle and translation of stored mRNAs. Due to this requirement for Ca2+, visualizing the transient changes in intracellular Ca2+ concentrations has been of key interest.
Historically, model organisms were selected for studies on egg activation based on the size and availability of their eggs. Various visualization approaches have been utilized to follow and quantify changes in intracellular Ca2+ concentrations in these systems, including: the photoprotein aequorin in medaka fish 3; Ca2+ sensitive fluorescent dyes such as Fura-2 in sea urchin and hamster 4,5; and calcium green-1-dextran in Xenopus 6.
The generation and improvement of genetically encoded calcium indicators (GECIs) has transformed the ability to visualize Ca2+ dynamics in vivo 7. These genetic constructs are expressed in specific tissues and limit the need for invasive tissue preparations 8.
GCaMPs are a green fluorescent protein (GFP)-based class of GECIs that have been very effective due to their high Ca2+ affinity, signal-to-noise ratio and capacity to be customized 9-11. In the presence of Ca2+, the GCaMP complex undergoes a series of conformational changes, starting with the binding of Ca2+ to calmodulin, that results in an increased fluorescent intensity of the GFP component 9.
GCaMPs have been extensively used in research on Drosophila neurons to visualize changes in intracellular Calcium12. The recent application of GCaMP technology to visualize Ca2+ in mature Drosophila eggs has revealed a single transient Ca2+ wave at egg activation 13,14. The Ca2+ wave can be visualized at low magnification during ovulation in vivo 13 or at higher magnification using an ex vivo activation assay 13,14. In the ex vivo assay, individual mature oocytes are isolated from the ovaries and activated using a hypotonic solution, referred to as activation buffer, which has been shown to recapitulate the events of in vivo activation 15-17.
This ex vivo assay enables easy high resolution visualization of the Ca2+ wave under different experimental conditions including pharmacological disruption, physical manipulations and genetic mutants. This article demonstrates the preparation of mature Drosophila eggs for ex vivo activation and the subsequent microscopy used to visualize the Ca2+ wave using GCaMP. This approach can be used to test the initiation and control of the Ca2+ wave and to probe downstream outcomes.

<table border="0" cellpadding="0" cellspacing="0" width="999">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Dry active yeast</td>
			<td style="width:373px;">Fleischman&#8217;s</td>
			<td style="width:243px;">#2192</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Dissecting microscope</td>
			<td>Leica</td>
			<td>MZ6</td>
			<td>Various will work</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Lamp</td>
			<td>Mircotec Fibre optic</td>
			<td>MF0-90</td>
			<td>Various will work</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Medical wipes</td>
			<td>Kimberly-Clark Corporation</td>
			<td>KLEW3020&#160;</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Marker pen</td>
			<td>Stabilo</td>
			<td>841/46</td>
			<td>OHPen universal, black - available in most stationary shops. Superfine Width 0.4mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">CO2 pad</td>
			<td>&#160;Genesee Scientific</td>
			<td>59-114 (789060 Dutscher)</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cover slip No. 1.5, 22 x 50mm</td>
			<td>International- No1-VWR Cat=631-0137-22-50mm</td>
			<td>Menzel-Glaser-22-50mm-MNJ 400-070T&#160;</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="40" rowspan="2" style="height:40px;">Halocarbon oil, series 95</td>
			<td rowspan="2">Halocarbon Products Corp.</td>
			<td>Distributor in Europe:</td>
			<td rowspan="2"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Solvadis (GMBH)</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Oil 10 S</td>
			<td>VWR Chemicals</td>
			<td align="right">24627.188</td>
			<td>Can be used instead of Halocarbon oil, series 95</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Forceps</td>
			<td>Fine Science Tools</td>
			<td>11252-00</td>
			<td>Whilst these tool should be clean, use of an autoclave is not necessary, ethanol would be sufficient.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Probe</td>
			<td>Fine Science Tools</td>
			<td>10140-01</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Glass pipette</td>
			<td>Fisherbrand,</td>
			<td>FB50251</td>
			<td>Pasteur Pipettes, glass, unplugged, 150mm length&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Vial</td>
			<td style="width:373px;">Regina Industries</td>
			<td style="width:243px;">P1014</td>
			<td>GPPS&#160; 80mm x 25mm</td>
		</tr>
	</tbody>
</table>

imaging calcium in drosophila at egg activation

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

Here we have demonstrated how to prepare mature Drosophila eggs for ex vivo activation. Eggs expressing a GECI enable imaging of Ca2+ dynamics at egg activation and the beginning of embryonic development (Figure 1). It should be noted that depending on the GCaMP used, specifically the presence of a myristoyl group, results may have slight qualitative differences13 14. We have also demonstrated a role for functional actin during egg activation by the addition of an inhibitor of actin polymerization, cytochalasin D, to the activation buffer (Figure 2) 14. 
&#160; 
A requirement for the cytoskeleton at egg activation is conserved. In C. elegans, following fertilization, cytoskeletal components are reorganized to prepare the one-cell embryo for the first asymmetrical division 21,22. In sea urchin, disruption of cytoskeletal re-organization following activation has been shown to affect the cell cycle by preventing contractile ring formation 23. The role of actin in mediating a Ca2+ wave and its downstream function at egg activation in Drosophila remains to be fully elucidated. 
&#160; 
In Drosophila, co-visualization of the Ca2+ wave and actin cytoskeleton can be achieved using this ex vivo activation assay. Time series of multiple channels can be acquired and dynamics of the intracellular Ca2+ can be matched with changes in actin organization in the oocyte (Figure 3).
<img alt="Figure 1" src="/files/ftp_upload/54311/54311fig1.jpg" /> 
Figure 1: A Single Ca2+ Wave at Drosophila Egg Activation. Time series of ex vivo mature Drosophila egg expressing UASt-myrGCaMP5 following the addition of activation buffer (A-H). The rise in cytoplasmic Ca2+ originates at the posterior pole (B) and propagates (B-F) with an average velocity of approximately 1.5 &#181;m/sec. Initiation of the wave is typically observed within 3 min of the addition of activation buffer. Following activation, the intracellular calcium levels of the mature egg returns to pre-activation levels (G,H). See supplemental&#160;movie 1 (total time 33 min). Scale bars = 50 &#181;m. Max projection = 40 &#181;m. <a href="https://www.jove.com/files/ftp_upload/54311/54311fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<img alt="Figure 2" src="/files/ftp_upload/54311/54311fig2.jpg" /> 
Figure 2: Addition of Cytochalasin D to Activation Buffer Perturbs Ca2+ Wave Propagation. Time series of ex vivo mature Drosophila egg expressing UASt-myrGCaMP5 following addition of activation buffer containing 10 &#181;g/ml cytochalasin D final concentration (A-H). Whilst the Ca2+ wave is initiated at the posterior pole (A), it is compromised and does not reach the anterior of the egg (F) (white arrowheads denote the front of the wave). No further Ca2+ changes were observed over 30 min). See supplemental movie 2 (total time 30 min). Scale bars = 50 &#181;m. Max projection = 40 &#181;m. <a href="https://www.jove.com/files/ftp_upload/54311/54311fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/54311/54311fig3.jpg" /> 
Figure 3: Co-visualization of Actin and Ca2+ at Egg Activation in Drosophila. Time series of ex vivo mature Drosophila egg expressing UASt-myrGCaMP5 (Cyan) and UASp-F-Tractin.tdTomato (Magenta) following addition of activation buffer (A-H). The Ca2+ wave initiates from the posterior pole and recovers as in Figure 1. Actin appears to be changing over time, white arrowheads (A vs H) . The lack of Ca2+ signal detection in the center of the mature egg is due to movement of the sample during image capture. Scale bars = 50 &#181;m. Max projection = 40 &#181;m. <a href="https://www.jove.com/files/ftp_upload/54311/54311fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Imaging Calcium in Drosophila at Egg Activation at JoVE.com

Imaging Calcium in Drosophila at Egg Activation

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

Imaging Calcium in Drosophila at Egg Activation

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

imaging-calcium-in-drosophila-at-egg-activation

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

7.8K Views.  University of Cambridge. This article describes an adaptable ex vivo protocol for visualizing Ca2+ during egg activation in Drosophila.

Video: Imaging Calcium in Drosophila at Egg Activation

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Upright Imaging of Drosophila Egg Chambers

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

Upright Imaging of Drosophila Egg Chambers

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

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

Imaging Cleared Embryonic and Postnatal Hearts at Single-cell Resolution

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, and allow for the study of the cellular and molecular basis of normal and abnormal cardiac morphogenesis. Recent emerging technologies of retrospective single clonal analyses make the study of cardiac morphogenesis at single cell resolution feasible. However, tissue opacity and light scattering of the heart as imaging depth is increased hinder whole-heart imaging at single cell resolution. To overcome these obstacles, a whole-embryo clearing system that can render the heart highly transparent for both illumination and detection must be developed. Fortunately, in the last several years, many methodologies for whole-organism clearing systems such as CLARITY, Scale, SeeDB, ClearT, 3DISCO, CUBIC, DBE, BABB and PACT have been reported. This lab is interested in the cellular and molecular mechanisms of cardiac morphogenesis. Recently, we established single cell lineage tracing via the ROSA26-CreERT2; ROSA26-Confetti system to sparsely label cells during cardiac development. We adapted several whole embryo-clearing methodologies including Scale and CUBIC (clear, unobstructed brain imaging cocktails and computational analysis) to clear the embryo in combination with whole mount staining to image single clones inside the heart. The heart was successfully imaged at single cell resolution. We found that Scale can clear the embryonic heart, but cannot effectively clear the postnatal heart, while CUBIC can clear the postnatal heart, but damages the embryonic heart by dissolving the tissue. The methods described here will permit the study of gene function at a single clone resolution during cardiac morphogenesis, which, in turn, can reveal the cellular and molecular basis of congenital heart defects.

We describe a protocol to volumetrically image fluorescent protein labeled cells deep inside intact embryonic and postnatal hearts. Utilizing tissue-clearing methods in combination with whole mount staining, single fluorescent protein-labeled cells inside an embryonic or postnatal heart can be imaged clearly and accurately.

Single clonal tracing and analysis at the whole-heart level can determine cardiac progenitor cell behavior and differentiation during cardiac development, ...

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

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

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

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

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

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the number of novel cell lines is expected to increase. The DGRC aims to familiarize researchers with using Drosophila cell lines as an experimental tool to complement and drive their research agenda. Procedures for working with a variety of Drosophila cell lines with distinct characteristics are provided, including protocols for thawing, culturing, and cryopreserving cell lines. Importantly, this publication demonstrates the best practices required to work with Drosophila cell lines to minimize the risk of contaminations from adventitious microorganisms or from other cell lines. Researchers who become familiar with these procedures will be able to delve into the many applications that use Drosophila cultured cells including biochemistry, cell biology and functional genomics.

Thawing, Culturing, and Cryopreserving Drosophila Cell Lines

Drosophila cell lines are important reagents for both fundamental and biomedical research. This article provides protocols for thawing, subculturing, and the cryopreservation of commonly used Drosophila cell lines to assist researchers in incorporating the use of these reagents in their research.

There are currently over 160 distinct Drosophila cell lines distributed by the Drosophila Genomics Resource Center (DGRC). With genome engineering, the ...

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological processes. In particular, appropriate regulation of calcium activity patterns during embryogenesis is necessary for many aspects of vertebrate neural development, including proper neural tube closure, synaptogenesis, and neurotransmitter phenotype specification. While the observation that calcium activity patterns can differ in both frequency and amplitude suggests a compelling mechanism by which these fluxes might transmit encoded signals to downstream effectors and regulate gene expression, existing population-level approaches have lacked the precision necessary to further explore this possibility. Furthermore, these approaches limit studies of the role of cell-cell interactions by precluding the ability to assay the state of neuronal determination in the absence of cell-cell contact. Therefore, we have established an experimental workflow that pairs time-lapse calcium imaging of dissociated neuronal explants with a fluorescence in situ hybridization assay, allowing the unambiguous correlation of calcium activity pattern with molecular phenotype on a single-cell level. We were successfully able to use this approach to distinguish and characterize specific calcium activity patterns associated with differentiating neural cells and neural progenitor cells, respectively; beyond this, however, the experimental framework described in this article could be readily adapted to investigate correlations between any time-series activity profile and expression of a gene or genes of interest.

Fluorescent Calcium Imaging and Subsequent In Situ Hybridization for Neuronal Precursor Characterization in Xenopus laevis

We present a two-part protocol that combines fluorescent calcium imaging with in situ hybridization, allowing the experimenter to correlate patterns of calcium activity with gene expression profiles on a single-cell level.

Spontaneous intracellular calcium activity can be observed in a variety of cell types and is proposed to play critical roles in a variety of physiological ...