The authors thank David Larson for developing the original protocol for imaging the live Drosophila pupal eye. We thank our colleagues Richard Carthew, Shoichiro Tsukita and Matthew Freeman, who developed the transgenic flies that we combined to generate our live-imaging fly lines. Brittany Baldwin-Hunter gave helpful comments on the manuscript. This work was supported by start-up funds awarded to Ruth Johnson by Wesleyan University.

Figure 3 shows a summary of the experimental procedure.
1. Tissue Preparation
<ol>
	<li>To determine the consequences of modified expression of a gene of interest, set up the following cross in duplicate and maintain at 25 &#176;C: UAS-transgene (males)X GMR-GAL4; UAS-&#945;-cat:GFP, ubi-DE-Cad:GFP ; + / SM5-TM6b (virgin females) NOTE: To obtain control tissue cross UAS-lacZ males to GMR-GAL4; UAS-&#945;-cat:GFP, ubi-DE-Cad:GFP females. &#946;-Galactosidase generates no defects in cell behavior in the retina.</li>
	<li>To enhance efficacy of RNAi transgenes, cross UAS-RNAi males to GMR-GAL4, UAS-Dcr-2; UAS-&#945;-cat:GFP, ubi-DE-Cad:GFP; + / SM5-TM6b virgin females. 
	NOTE: Occasional defects in cell behavior in retinae expressing ectopic Dcr-2 have been observed (data not shown). Vigilance is recommended if using this approach.</li>
	<li>Select white pre-pupae of appropriate genotypes and place in 1.5 ml microcentrifuge tubes. If the pupae express Dcr-2, select either male or female pupae: expression of UAS-Dcr-2, an insert on the X chromosome, is stronger in males. Pupae should be sexed at 0 hr APF before pigmentation of the pupal case &#8211; the gonads of males are visible as large translucent globes on the lateral sides of the pupa (approximately one-third from the posterior).</li>
	<li>Incubate microcentrifuge tubes containing pupae, in a clean plastic tip-box at 25 &#176;C. To prevent desiccation of pupae place a 10 cm2 piece of tissue, soaked in distilled water, into the tip-box. 
	NOTE: if the humidity in the tip-box becomes too high the pupal case may become soggy and difficult to dissect in subsequent steps.</li>
	<li>Incubate for appropriate time. To capture cell behaviors associated with patterning the eye, prepare pupae for imaging at around 17 hr APF, 20 hr APF or 24 hr APF. See Representative Results for further discussion on when specific cell behaviors are best observed.</li>
	<li>Place a piece of fresh double-sided tape onto a black sylgard dissection dish. Position a pupa, dorsal side up, with the head of the pupa adhered to the double-sided tape (Figure 1A).Try not to adhere the thoracic and abdominal regions of the pupa to the double-sided tape. With forceps carefully lift and remove the operculum to expose the pupal head. Tear along the side of the pupal case to expose the area around one eye.</li>
	<li>Gently remove the pupa from the adhesive tape. If the pupa remains adherent, apply a small volume of distilled water to the junction between the pupal case and tape to weaken adhesion.</li>
</ol>
2. Mounting
<ol>
	<li>Construct a small 15 mm2 frame from blotting paper and punch a hole in the middlethat is 5.5 mm in diameter (Figure 1B). Immerse in distilled water and place in the center of a microscope slide. Using a 30 cc syringe, squeeze out a uniform ring of petroleum jelly to surround the blotting paper frame. Ensure that the petroleum jelly ring is marginally higher than the width of the pupa and that the diameter of the ring is marginally less than the width of a cover slip.</li>
	<li>Add a small bead of petroleum jelly directly onto the slide, in the hole punched into the blotting paper(Figure 1C).Carefully arrange the pupa, on its side, supported by the petroleum jelly bead (Figure 1D).</li>
	<li>Place a coverslip on top of the petroleum jelly ring so that it contacts the epithelium above the eye neuroepithelium (Figure 1D). Gently compress the preparation to seal the coverslip against the petroleum jelly and generate a small flat contact surface between the pupal eye region and the coverslip.</li>
</ol>
3. Fluorescence Imaging
<ol>
	<li>Image the pupal preparation using a fluorescent microscope. Every 7 min capture serial sections through the apical domain of the eye neuroepithelium, in the region of the adherens junctions. 
	NOTE: a) Appropriate serial sections are usually 4-5 per&#160;z-stack&#160;comprised of 0.2 &#956;m z-steps. b) Irregularities in the topology of the epithelium, including mild bumps and folds, may make it challenging to acquire in-focus images of large regions of tissue. While imaging, one may find part of an area of interest to be in-focus, and a neighboring region to be out-of-focus. Including a small droplet of distilled water between the pupal eye and coverslip can eliminate some acute tissue folds but not the mildly uneven topology characteristic of the early stages of pupal eye morphogenesis. In these instances oversample the tissue by extending the z-stack until in-focus slices are acquired for the entire field. c) Capturing serial sections every 7 min should enable live-imaging for 3 to 4 hr without a significant reduction in the intensity of GFP fluorescence that may compromise image quality. Shorter time-intervals may reduce the total time of imaging.</li>
	<li>Continue imaging for 3 to 4 hr. Do not use an automatic focus and time function that is available on many fluorescent microscopes since pupal growth and pumping of the hemolymph moves the position of the retina and vigilant re-focusing is required every 14-21 min. NOTE that imaging beyond 4 hours may slow or stall morphogenesis of the eye.</li>
	<li>Use appropriate deconvolution software to reduce background and enhance contrast of the serial section images. For each z-stack file, perform the following in LAS AF software (see Table of Materials): Navigate to the Tools panel, select 3D Deconvolution and click Apply.</li>
	<li>Generate a maximum projection (MP) image for each deconvoluted stack file: Navigate to the Tools panel, select 3D Projection and click Apply. 
	NOTE: The Maximum Projection algorithm may fail to generate a uniformly in-focus image for tissue that has been oversampled. A technique for sharpening out-of-focus regions is outlined in step 5.2.</li>
</ol>
4. Post-imaging Pupal Rescue and Phenotype Verification
<ol>
	<li>To address whether artifacts were introduced or the pupa harmed during imaging, carefully remove the pupa from the imaging rig: using forceps remove the coverslip and transfer the pupa to a clean vial of food. Store at room temperature or 25 &#176;C until the adult fly emerges.</li>
	<li>Carefully examine the phenotype of the adult eye and compare to the eyes of adult flies emerging from the original fly cross.</li>
</ol>
5. Image Processing
<ol>
	<li>Automatic Image Alignment: 
	NOTE: The live Drosophila tissue will shift and grow throughout the imaging process. Consequently, the centers of images gathered at successive time points may not correspond to the same point in the Drosophila tissue. In addition the entire retinal disc rotates about 30&#176; between ~21 and 23 hr APF. To highlight individual cell behaviors and reduce distractions caused by organismal growth,align each MP image.While this can be performed manually,the image editing software used here (see Table of Materials) has built-in algorithms that can expedite the process.
	<ol>
		<li>From the File tab of the main menu, open Scripts and select Load Files into Stack.</li>
		<li>Import the MP image files into the Load Layers panel and select OK. Make sure that the Attempt to Automatically Align Source Images option is not selected. 
		NOTE: If this option is selected, the software may use an alignment algorithm that distorts the image data.</li>
		<li>Once all MP files have loaded into the Layers pane, make sure that all images are in chronological order such that the earliest time point is at the top of the layer stack. If the layers are not in the correct order, drag and drop them until they are ordered correctly.</li>
		<li>Select Auto-Align Layers from the Edit tab of the main menu. Choose Reposition and click OK.</li>
		<li>If the Auto-Align algorithm fails to orient the frames to a relevant focal point, make minor adjustments using the Move tool.</li>
	</ol>
	</li>
	<li>Composite Sharpening: 
	NOTE: Out-of-focus regions of an initial MP image can be sharpened by &#8216;cropping&#8217; the corresponding deconvoluted stack in LAS AF software. Cropping a stack file allows one to manually restrict the interval of a z-stack that is subjected to the Maximum Projection algorithm.
	<ol>
		<li>Locate the Crop function in the Tools panel (under the Process tab). Manually restrict the initial and final slices such that they span only the adhesion belt within the initially out-of-focus region. Click Apply to generate a new stack file that is optimized for this region.</li>
		<li>Generate a new Maximum Projection for the locally optimized stack and export as a TIFF file.</li>
		<li>In the image editing software (see Table of Materials), open Scripts (located in the File tab of the main menu) and select Load Files into Stack.</li>
		<li>Import the initial MP image file and locally optimized MP file for a particular time point into the Load Layers panel and select OK. Make sure that the Attempt to Automatically Align Source Images option is not selected.</li>
		<li>Select both layers in the Layers pane and then choose Auto-Blend Layers (located in the Edit tab of the main menu) to automatically mask the appropriate regions of the two images, yielding a uniformly in-focus retinal field. Save this composite image as a TIFF file.</li>
		<li>Repeat steps 5.2.1 through 5.2.5 for all suboptimal MP images.</li>
	</ol>
	</li>
	<li>Further Adjustments: Rotate, crop, and Adjust the Levels of the Movie Layers:
	<ol>
		<li>With all of the layers selected, use the Transform tool (Edit &#62; Free Transform) to rotate the images so that the dorsal-ventral axis of the retina is aligned to the y-axis of the movie frame.</li>
		<li>If necessary, use the Crop tool to reduce the field of view to a desired size and shape.</li>
		<li>Use level adjustment (of individual frames) to help enhance contrast between the cell membrane and cell body.</li>
		<li>For stylistic purposes and to emphasize specific cell behaviors, introduce false color to highlight points of interest in each frame.</li>
	</ol>
	</li>
	<li>Animation: Convert the Images into a Movie:
	<ol>
		<li>Open the Animation pane from the Windows tab of the main menu. Select Make Frames from Layers from the menu located in the upper right hand corner of the Animation pane.</li>
		<li>Note that the layers are added to the animation pane in sequential order beginning from the bottom of the stack. To reverse the order of the frames, first select all of the frames and then select Reverse Frames from the Animation pane menu.</li>
		<li>Select a frame delay time for each frame using the dropdown menu below the frame thumbnail. 
		NOTE: The frame delay for Movies 1 to 4 presented in this paper is 0.8 sec.</li>
		<li>From the File tab of the main menu, open Export and select Render Video. Choose QuickTime Export under File Options and select a desired video format. Under Render Options set Frame Rate to Custom, enter a frame rate of 15, and click Render.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests.

The description of wild-type development (above) forms the basis for comparisons of patterning events in RNAi or overexpression genotypes. Comparisons of live tissue development are invaluable when determining precisely which cell behaviors are regulated by a protein of interest. Further, and not described here, live cell imaging enables one to make qualitative and/or quantitative descriptions of the role of a gene of interest in the events that pattern the eye. By 30-32 hr APF most pigment cells have acquired stable pos...

The compound Drosophila eye is characterized by the stereotyped arrangement of its ~750 ommatidia separated by a honeycomb-lattice of accessory pigment cells 4,5. These pigment cells are patterned by a coordinated combination of events: local cell movements, cell growth, changes in cell shape, and apoptosis. Live visualization of this epithelium allows one to study the molecular mechanisms underpinning these events in a physiologically relevant and unperturbed three-dimensional context.
In contrast to previous protocols 6,7, the technique outlined here incorporates an efficient method for stabilizing extraneous tissue movement that cannot be uncoupled from the imaging process. This method enhances studies of cell behavior in the developing Drosophila pupal eye epithelium &#8211; a tissue that grows, rotates, and shifts over the course of imaging. In addition the motion-stabilization technique described here will be useful for studying cells in other contexts where extraneous movement occurs.
To visualize cell boundaries in the Drosophila retinal field, transgenic fly lines were generated that express ubi-DE-Cadherin:GFP as well as UAS-&#945;-catenin:GFP under control of the eye-specific driver GMR-GAL4 1-3. The use of two GFP-tagged membrane markers allows for the visualization of cell boundaries at lower intensity light. This minimizes tissue damage and photobleaching on repeated exposure to high-energy wavelengths, enabling increased frame rate and movie duration. To enhance efficacy of RNAi transgenes, UAS-Dcr-2 was also incorporated into a second fly line 8.
In a third update from previous protocols, a simpler imaging rig that is easily assembled in most laboratories is described. This apparatus obviates the requirement to have a specialized imaging rig generated by a university&#8217;s &#8216;machine shop&#8217; or similar service. This imaging rig is similar to that used to image other pupal tissues 9,10.
Presented here is a simple live-cell imaging protocol that can be used to directly assess the morphogenetic events that contribute to eye patterning from ~17 to 42 hr after puparium formation (hr APF). Specifically, this protocol enables one to determine the consequences of modifying gene expression during pupal development.

<table border="0" cellpadding="0" cellspacing="0" width="672">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Name of Material/ Equipment</td>
			<td style="width:124px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td colspan="3" height="21" style="height:21px;">GMR-GAL4; UAS-a-cat:GFP, ubi-DE-Cadherin:GFP ; + / SM5-TM6b</td>
			<td>Available on request from authors</td>
		</tr>
		<tr height="21">
			<td colspan="3" height="21" style="height:21px;">UAS-dcr-2; UAS-a-cat:GFP, ubi-DE-Cadherin:GFP; + / SM5-TM6b&#160;</td>
			<td>Available on request from authors</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Scotch double stick tape</td>
			<td style="width:124px;">3M</td>
			<td style="width:119px;">665</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Black Sylgard dissection dish</td>
			<td style="width:124px;"></td>
			<td style="width:119px;"></td>
			<td>Fill glass petri dish to rim with Sylgard (colored black with finely-ground charcoal powder) or Sylgard 170; leave at room temperature for 24-48 h to polymerize</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Sylgard</td>
			<td style="width:124px;">Dow Corning</td>
			<td style="width:119px;">SYLG184</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Sylgard (Black)</td>
			<td style="width:124px;">Dow Corning</td>
			<td style="width:119px;">SYLG170</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Glass petri dish</td>
			<td style="width:124px;">Corning</td>
			<td style="width:119px;">7220-85</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">25 x 75 x 1 mm glass microscope slide</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:119px;">12-550-413</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">22 x 40 mm glass coverslip</td>
			<td style="width:124px;">VWR</td>
			<td>48393-172</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:263px;">Forceps&#160;</td>
			<td style="width:124px;">Fine Science Tools</td>
			<td style="width:119px;">91150-20</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:263px;">Whatman 3mm Chromatography Paper</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:119px;">05-713-336</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Vaseline</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:119px;">19-086-291</td>
			<td>Or purchase at local pharmacy.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">30ml syringe</td>
			<td style="width:124px;">Fisher Scientific</td>
			<td style="width:119px;">S7510-30</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:263px;">Adobe Photoshop CS5</td>
			<td style="width:124px;">Adobe</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:263px;">Leica TCS SP5 DM microscope</td>
			<td style="width:124px;">Leica Microsystems</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:263px;">LAS AF Version 2.6.0.7266 microscope software</td>
			<td style="width:124px;">Leica Microsystems</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
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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 pupal eye is a successful strategy to observe cell behaviors that contribute to patterning of the neuroepithelium. The role of specific proteins can be readily assessed by expressing transgenes that modify protein levels during eye development. To do this GMR-GAL4 is used to drive transgene expression behind the morphogenetic furrow. This offers the advantage of not perturbing earlier events that establish the eye field during the first two larval instars. In addition many RNAi transgenes req...

Watch this Scientific Journal Video about Live-imaging of the Drosophila Pupal Eye at JoVE.com

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.

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 ...

Research

JoVE Journal

Developmental Biology

9.3K Views.  Wesleyan University. 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.

Video: Live-imaging of the Drosophila Pupal Eye

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 ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a reliable protocol for the dissection of the delicate Drosophila pupal retina. Our surgical approach utilizes readily-available microdissection tools to open pupae and precisely extract eye-brain complexes. These can be fixed, subjected to immunohistochemistry, and retinas then mounted onto microscope slides and imaged if the goal is to detect cellular or subcellular structures. Alternatively, unfixed retinas can be isolated from brain tissue, lysed in appropriate buffers and utilized for protein gel electrophoresis or mRNA extraction (to assess protein or gene expression, respectively). Significant practice and patience may be required to master the microdissection protocol described, but once mastered, the protocol enables relatively quick isolation of mainly undamaged retinas.

Dissection of the Drosophila Pupal Retina for Immunohistochemistry, Western Analysis, and RNA Isolation

This paper presents a surgical method for dissecting Drosophila pupal retinas along with protocols for the processing of tissue for immunohistochemistry, western analysis, and RNA-extraction.&#160;

The Drosophila pupal retina provides an excellent model system for the study of morphogenetic processes during development. In this paper, we present a ...

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 ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, germ cell development, piRNA biology, and niche formation. Here, we present a dissection technique to live-image the gonad ex vivo during a period when in vivo live-imaging is highly ineffective. This protocol outlines how to transfer embryos to an imaging dish, choose appropriately-staged male embryos, and dissect the gonad from its surrounding tissue while still maintaining its structural integrity. Following dissection, gonads can be imaged using a confocal microscope to visualize dynamic cellular processes. The dissection procedure requires precise timing and dexterity, but we provide insight on how to prevent common mistakes and how to overcome these challenges. To our knowledge this is the first dissection protocol for the Drosophila embryonic gonad, and will permit live-imaging during an otherwise inaccessible window of time. This technique can be combined with pharmacological or cell-type specific transgenic manipulations to study any dynamic processes occurring within or between the&#160;cells in their natural gonadal environment.

Dissection and Live-Imaging of the Late Embryonic Drosophila Gonad

Here, we provide a dissection protocol required to live-image the late embryonic Drosophila male gonad. This protocol will permit observation of dynamic cellular processes under normal conditions or after transgenic or pharmacological manipulation.

The Drosophila melanogaster male embryonic gonad is an advantageous model to study various aspects of developmental biology including, but not limited to, ...