This work was supported by Department of Defense start-up funds to J.T.S.

1. Prepare Animals, Tools, and Media for Dissection
<ol>
	<li>Cross male and female flies to obtain progeny of the desired genotype. Useful transgenic markers for identification and staging of meiotic spermatocytes include GFP-tubulin to label the meiotic spindle and RFP-histone 2A to label chromsomes8. Use five to ten females per cross and maintain crosses on standard fly food in vials in a 25 &#176;C incubator until third instar larvae begin crawling up the sides of the vial (4&#8211;5 days). Early white pupae will begin forming one day later.</li>
	<li>Obtain two pairs of straight forceps with fine tips. Ensure that the tips are sharp, even in length, and straight (Figure 1A). Use these forceps only for dissections, and cap with 10 &#956;L pipette tips when not in use.</li>
	<li>Prepare a scalpel tool.
	<ol>
		<li>Insert the blunt end of a black anodized steel insect pin into a pin holder, and twist the screw of the holder to secure the pin in place. Using a pair of forceps and under a dissecting microscope, grasp the insect pin in the middle and bend it to form an internal angle of 135&#176; (Figure 1B). The sharp end is for cutting tissue, while the edge is used for transferring testes between drops of media.</li>
		<li>Cap with a 1,000 &#956;L pipette tip when not in use.</li>
	</ol>
	</li>
	<li>Working in a tissue culture hood, prepare 5 mL aliquots of Schneider&#8217;s media and store at 4 &#176;C until the time of use. When ready to begin dissections, warm one 5 mL media aliquot to room temperature. Then transfer the warmed dissection media to a 10 mL syringe and attach a 0.22 &#181;m syringe filter.</li>
</ol>
2. Dissection of Testes from Larvae and Early Pupae
<ol>
	<li>Use the media filled filter syringe to expel three drops of media (50 &#181;L each) at the top, middle and bottom of a glass slide.</li>
	<li>Use a dissecting probe to transfer five to ten third instar larvae or early pupae (pupae that are still white or yellow) to the top drop. Gently agitate the larvae and pupae in the media with the dissecting probe to remove any food or debris.</li>
	<li>Under a dissecting microscope, turn a single larva or pupa on its side with a dissecting probe to identify the two bilateral testes if present, which appear as oval shaped translucent structures in the posterior third of the body (Figure 2A, B). Animals that lack testes are female and should be discarded.</li>
	<li>When a male larva or pupa is found and is clean, move it to the second drop of media. Repeat until 3 or 4 male larvae and/or pupae have been transferred to the second drop of media.</li>
	<li>Working in the second drop of media, grasp a single male larva or pupa with a pair of forceps at its mid-region, just anterior to the testes. Use the second pair of forceps to gently tear the animal in half.</li>
	<li>Hold the posterior end of the larva or pupa down on the glass slide with a pair of forceps. Starting just next to the forceps, push down on the cuticle using the edge of the scalpel tool and move the scalpel toward the cut end of the animal. This will expel the animal&#8217;s internal organs, which include the guts, fat bodies, and testes.</li>
	<li>Gently tease apart the testes from the rest of the organs. The testes are clear, oval shaped organs embedded in ribbons of fat body (Figure 2C).</li>
	<li>Transfer the testes one at a time to the third drop of media. To accomplish this, insert the edge of the scalpel under the fat body, lift the tissue out of the media, and quickly move it to the third drop.</li>
	<li>Alternatively, if there is not enough fat body still attached to the testes to successfully lift the tissue, gently transfer the tissue using a glass Pasteur pipette that has been pre-wetted with dissection media.</li>
	<li>Use the sharp end of the scalpel tool to gently slice away excess fat body from the testes, leaving just a small rim of fat body around the edges (Figure 2C). It is not necessary to remove all of the fat body, and attempting to do so will likely result in damaging or rupturing of the testes.</li>
	<li>Repeat the above steps for all of the male larvae on the glass slide.</li>
	<li>Proceed either to step 3 or step 5.</li>
</ol>
3. Mounting Testes for Live Microscopic Imaging
NOTE: This mounting procedure was adapted from a recently published protocol for imaging of Drosophila larval brain neuroblasts20. Additional details can be found in this reference.
<ol>
	<li>Use the filter syringe to deposit a single drop of Schneider&#8217;s media (about 30 &#181;L) onto the center of a 50 mm gas-permeable culture dish.</li>
	<li>Use a pre-wetted Pasteur pipette to transfer the prepared testes to the drop on the gas-permeable dish. The testes will generally float to the surface of the drop.</li>
	<li>Use the scalpel tool to gently push the testes down onto the gas-permeable membrane. Take care not to rupture the gas-permeable membrane with the sharp point of the scalpel. Push all of the testes to the center of the drop.</li>
	<li>Use a 1 mL syringe filled with halocarbon oil to make four drops of oil, about 30 &#956;L each, on the gas permeable membrane. Space the drops of oil around the drop of media containing the testes to correspond to the four corners of a 22 mm glass coverslip.</li>
	<li>Line up the corners of a 22 mm glass coverslip with the four drops of halocarbon oil and gently lower the coverslip onto the media and oil. Allow the coverslip to settle and for the media containing the testes to spread between the drops of oil.</li>
	<li>Remove excess media to allow the coverslip to settle onto the surface of the testes. While observing the testes under a dissecting microscope, insert the corner of a delicate task wipe into the media under the coverslip to wick away a small amount of liquid. Wick away enough media until the glass coverslip just makes contact with the surface of the largest testis. It is critical not to remove too much media and lower the coverslip too far, as this will exert pressure on the testes and cause them to rupture (see Figure 5).</li>
</ol>
4. Live Imaging of Meiotic Spermatocytes
NOTE: Spermatocytes can be imaged using a laser scanning or spinning disk confocal microscope. The system must have adequate speed and sensitivity to avoid photobleaching of fluorescent proteins or photodamage of the tissue.
<ol>
	<li>Place a drop of immersion oil on the glass coverslip just above the testes. Flip over the dish and place it on the microscope stage and move the microscope objective (40x or 60x) into the oil until it is just below the testes. Use transmitted light to find a single testis and bring it into focus.</li>
	<li>While capturing fast images with the confocal, move the testis around to examine the organization of the cells. One end of the testis will have many small, densely packed cells. This end contains the germline stem cells and spermatogonia. Moving toward the other end of the organ, identify progressively larger cells organized into discrete clusters or cysts. These large cells are the spermatocytes (Figure 3A).</li>
	<li>If imaging GFP-tubulin, identify spermatocytes that will commence meiosis within 30&#8211;60 min by the presence of two bright microtubule asters (i.e., centrosomes) located at the cortex of the cell (Figure 3B, C).</li>
	<li>Once a cyst of meiotic spermatocytes has been identified, set up acquisition parameters to capture z-stacks of images over time. Typically, acquisition of 15&#8211;20 images spaced 1.5 &#956;m apart in the z-dimension is sufficient to capture the full volume of several spermatocytes within the same cyst.</li>
	<li>Acquire full z-stacks every 2 min if imaging the entire first meiotic division, which takes about 1.5 hours. It is possible to use faster acquisition rates, particularly if only a specific event of meiosis is to be analyzed. However, take care not to cause photobleaching or photodamage due to repeated exposure of the sample to laser excitation.</li>
	<li>Use a continuous, automatic focus control system to prevent focal drift over the long time-course of acquisition. Temperature control of the sample on the microscope stage generally is not necessary, assuming room temperature of approximately 22&#8211;23 &#176;C. If temperature control is available, maintain the sample at 25 &#176;C on the microscope stage.</li>
	<li>If meiotic spermatocytes are not found, store the sample for several hours or overnight in a 25 &#176;C incubator and image again.</li>
</ol>
5. Fixation and Immunostaining
<ol>
	<li>From step 2.10 above, use a glass Pasteur pipette to transfer testes to a well in a 9-well glass dissecting dish containing 0.5 mL of 8% paraformaldehyde in PBSTx (phosphate buffered saline with 0.3% Triton X-100). Ensure that the testes are laying on the bottom of the dish. 
	NOTE: Paraformaldehyde is toxic and should be handled with gloves in a fume hood.</li>
	<li>Fix the testes for 20 min at room temperature.</li>
	<li>Transfer the testes to a well containing 0.5 mL PBSTx. Wash for 5 min with gentle agitation. Repeat the wash step in new wells with fresh PBSTx two more times.</li>
	<li>Dilute primary antibody to the desired concentration in 0.5 mL PBSTx containing 5% bovine serum albumin (BSA) in a 1.5 mL microcentrifuge tube. Transfer testes from the 9-well glass dissecting dish to the tube of diluted primary antibody and incubate overnight at 4 &#176;C with gentle rocking or nutation.</li>
	<li>Wash the testes in 0.5 mL of PBSTx for 5 min in a well of a 9-well glass dissecting dish. Repeat the wash step with fresh PBSTx two more times.</li>
	<li>Dilute secondary antibody in 250 &#956;L of PBSTx containing 5% BSA and dispense into a clean well of a 9-well glass dissecting dish. If desired, add DAPI to stain DNA. Transfer testes to the well containing secondary antibody, cover with plastic wrap, and incubate in the dark with gentle agitation for 4 hours at room temperature.</li>
	<li>Transfer the testes to a clean well filled with 0.5 mL of PBSTx. Wash with fresh PBSTx twice for 5 min and a third time for 30 min.</li>
</ol>
6. Mounting Fixed Testes
<ol>
	<li>Transfer the testes from the last wash onto a glass microscopy slide using a Pasteur pipette. If necessary, use the edge of the scalpel tool to push the testes down onto the surface of the slide.</li>
	<li>Use the corner of a delicate task wipe to wick away excess liquid from the testes. Remove as much liquid as possible.</li>
	<li>Apply a 30&#8211;50 &#181;L drop of microscopy mounting medium to the testes.</li>
	<li>Place a 22 mm coverslip onto the drop of mounting medium and allow the coverslip to settle and the mounting medium to spread under the coverslip. Apply gentle pressure to the coverslip if necessary to squeeze out excess mounting medium and allow the coverslip to rest on the surface of the testes.</li>
	<li>Use nail polish to seal the edges of the coverslip.</li>
</ol>

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The authors have nothing to disclose.

We have described a protocol for the preparation of larval or early pupal Drosophila testes, optimized for long-term, live imaging of spermatocyte cell division. This is a powerful method for analysis of cell division in the physiological context of intact tissue. The power of this method is further expanded when combined with Drosophila genetic tools, such as specific gene mutations, tissue-specific RNAi-mediated suppression, and fluorescently labeled protein and organelle markers. In addition, the sma...

Cell division is often studied using cell lines grown in culture1. While we have gained a wealth of invaluable insight and understanding of fundamental mechanisms from these studies2, cells grown in culture cannot fully recapitulate the physiology of cell division as it occurs in intact, living tissue. For example, in intact tissues and organs, cells must divide at the right place and at the right time so that progeny cells are properly situated within the tissue, so that they can undergo appropriate differentiation or functional programs, and so that cell proliferation is properly coordinated with tissue growth or homeostasis3. For cells grown in culture on the other hand, cell division is generally regulated by growth factors in the culture medium4, and thus we cannot learn from these cells how in vivo environmental factors such as tissue architecture or developmental signaling influence the division process. It is also important to note that many of the cell lines used to study cell division, such as HeLa and U2OS cells, were derived from metastatic tumors5. Therefore, many aspects of the basic physiology of these cancer cells, such as cell cycle regulatory mechanisms and chromosomal stability, have likely been altered compared to healthy cells. Complete understanding of cell division physiology, therefore, depends on our ability to study dividing cells in their native, in vivo environments that preserve physiological regulatory mechanisms and tissue architecture.
Advances in understanding how cell division operates within intact tissues and organs are hampered by difficulties inherent to in vivo or ex vivo analysis. First, it can be difficult or impossible to access dividing cells for microscopic analysis within large organs or thick tissues. Second, it is often difficult to predict when individual cells will divide in vivo. Third, tissue physiology may rapidly deteriorate during ex vivo culture. In this protocol, we describe readily accessible methods for live and fixed analysis of Drosophila melanogaster spermatocytes as they undergo meiotic cell divisions within fully intact testes. These cells are ideal for live, ex vivo analysis because they are readily accessible with standard optical methods such as confocal microscopy, they divide at predictable times and locations, and intact testes can be maintained in ex vivo culture for up to about 24 h. In addition, Drosophila spermatocytes are large round cells (approximately 20&#8211;30 &#181;m in diameter) that do not change shape when they divide, making them ideal for high resolution, time-lapse imaging of cellular components such as the spindle apparatus and cytoplasmic organelles. Although these cells undergo meiosis as opposed to mitosis, many of the essential cell division processes are very similar between these two cell division mechanisms6. These advantages, combined with powerful Drosophila genetic tools, have made Drosophila spermatocytes an extensively used model for ex vivo analysis of essential cell division processes including spindle formation and regulation, cytokinesis, and organelle remodeling and partitioning7,8,9,10,11.
Spermatocytes are cells that are at the meiotic stage of spermatogenesis. In Drosophila, spermatogenesis begins with a group of germline stem cells that are located in a small region or &#8220;hub&#8221; at one pole of the testis12,13. These cells divide by an asymmetric mitosis, giving rise to a single differentiated spermatogonium. Spermatogonia then undergo four synchronous mitoses to produce a group of 16 cells that remain closely associated within a single cyst. At this stage, the cells transition from a mitotic to a meiotic cell cycle and are referred to as spermatocytes. Spermatocytes spend about two to three days in an extended G2 stage of the cell cycle, during which they grow dramatically and undergo cytological changes in preparation for the two meiotic divisions and subsequent spermiogenesis13,14. The entire group of 16 spermatocytes within a single cyst then enter the first meiotic division at the same time. Thus, several meiotic spermatocytes can be imaged simultaneously as they proceed through cell division. The first meiotic division proceeds for approximately 1.5 h and is followed almost immediately by the second meiotic division, yielding 64 total spermatids that go on to differentiate into mature spermatozoa.
A unique advantage of using spermatocytes to study cell division in live, intact tissue is that groups or cysts of cells constantly progress through the different stages of spermatogenesis, and cells at all stages of spermatogenesis can usually be identified in any given testis (see Figure 3A). Therefore, it is relatively easy to find meiotic cells in whole testes. We generally focus our analyses on the first as opposed to second meiosis because these cells are much larger and more amenable to high resolution imaging, but the entire process encompassing both meiotic divisions can be successfully imaged. It should also be noted that the general protocol for testis preparation and culture can be used to analyze other processes of spermatogenesis as well, such as the earlier mitotic stem cell or spermatogonial divisions or the cytological changes that occur as spermatids mature into spermatozoa15. Many of these aspects of spermatogenesis are highly conserved between Drosophila and humans16.
Drosophila spermatocytes begin to reach the meiotic phase of spermatogenesis during the third instar larval stage of Drosophila development13. Therefore, testes isolated from lifecycle stages beginning with third instar larvae and including pupae and adults can be used for analysis of dividing spermatocytes. Several excellent protocols are available for extraction of testes from adult male flies for live and fixed analysis of spermatogenesis17,18,19. These protocols may be preferable for studying late stages of spermatogenesis or if genetic markers that are only visible in adults must be used. The protocol focuses instead on testis preparation from larvae and early pupae, because testes at these stages have several advantages that are specifically pertinent to cell division analysis by high resolution, time-lapse microscopy. First, testes from larvae and pupae are smaller than those from adults, and the cells within the organ are less crowded. Because of this, dividing spermatocytes can often be imaged near the outer surface of larval testes, without having to penetrate through multiple layers of light scattering tissue. Second, adult testes move rhythmically due to contractions of the attached accessory organs, and these movements make time-lapse imaging of individual cells challenging. And third, larval testes are advantageous when studying gene mutations that cause pupal or adult lethality. Our methods are optimized for long-term culture of testes on the microscope stage, allowing for imaging of multiple rounds of cell division or the progression of individual cells through multiple stages of spermatogenesis in the same preparation. We also describe a protocol for fixation and immunostaining of whole testes. Overall, our protocols are particularly useful for those interested in studying cell division in intact tissue, and the ability to combine spermatocyte analysis with highly tractable Drosophila genetic tools makes this an especially powerful approach.

<table>
	<tbody>
		<tr>
			<td>5&#34; Dissecting Probe</td>
			<td>Fisher</td>
			<td>08-965-A</td>
			<td></td>
		</tr>
		<tr>
			<td>5.75&#34; Glass Pasteur Pipet</td>
			<td>Fisher</td>
			<td>13-678-20A</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA)</td>
			<td>Fisher</td>
			<td>BP9703-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fine Science Tools</td>
			<td>11252-20</td>
			<td>Straight forcepts with fine tips</td>
		</tr>
		<tr>
			<td>Frosted Microscope Slides</td>
			<td>Fisher</td>
			<td>12-544-2</td>
			<td>Slides for mounting fixed tissue, with frosted writing surface</td>
		</tr>
		<tr>
			<td>Halocarbon oil 700</td>
			<td>Sigma</td>
			<td>H8898-100 mL</td>
			<td></td>
		</tr>
		<tr>
			<td>Lumox Dish 50</td>
			<td>Sarstedt</td>
			<td>SAR946077410</td>
			<td>Gas-permeable tissue culture dish</td>
		</tr>
		<tr>
			<td>Microscope Cover Glass</td>
			<td>Fisher</td>
			<td>12-541-B</td>
			<td>22x22 mm, #1.5 glass coverslip</td>
		</tr>
		<tr>
			<td>Minutien Pins</td>
			<td>Fine Science Tools</td>
			<td>26002-15</td>
			<td>Insect pins used to make scalpel tool</td>
		</tr>
		<tr>
			<td>Nickel Plated Pin Holder</td>
			<td>Fine Science Tools</td>
			<td>26018-17</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde 32% Solution, EM Grade</td>
			<td>Electron Microsocopy Sciences</td>
			<td>15714</td>
			<td></td>
		</tr>
		<tr>
			<td>Plan Beveled Edge Microscope Slides</td>
			<td>Fisher</td>
			<td>12-549-5</td>
			<td>Slides used for dissections</td>
		</tr>
		<tr>
			<td>PYREX Spot Plates</td>
			<td>Fisher</td>
			<td>13-748B</td>
			<td>9-well glass dissecting dish</td>
		</tr>
		<tr>
			<td>Schneider&#39;s Drosophila medium</td>
			<td>Fisher</td>
			<td>21720-024</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Filter, 0.22 &#181;m</td>
			<td>EMD Millipore</td>
			<td>SLGS033SB</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Vectashield</td>
			<td>Vector Labs</td>
			<td>H-1000</td>
			<td>Microscopy mounting medium</td>
		</tr>
	</tbody>
</table>

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.

When this protocol is successfully executed, testes will remain fully intact for imaging by confocal microscopy or other fluorescence microscopy methods. As seen in Figure 3A, the cellular organization of the testes is preserved, and the progression of cell differentiation from one end of the testis to the other including spermatogonia, spermatocytes, and haploid spermatids is visible. GFP-tubulin is a useful marker for identifying dividing spermatocytes and for live imaging of the progressi...

Watch this Scientific Journal Video about Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue at JoVE.com

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.

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 ...

preparation-drosophila-larval-pupal-testes-for-analysis-cell-division

Research

JoVE Journal

Developmental Biology

8.0K Views.  Uniformed Services University of the Health Sciences. 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.

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

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 ...

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often compromise the lives of those who suffer from these complex diseases. Detecting mitochondrial metabolic changes is vital to the understanding of mitochondrial disorders and mitochondrial responses to pharmacological agents. Although mitochondrial metabolism is at the core of metabolic regulation, the detection of subtle changes in mitochondrial metabolism may be hindered by the overrepresentation of other cytosolic metabolites obtained using whole organism or whole tissue extractions. 
Here we describe an isolation method that detected pronounced mitochondrial metabolic changes in Drosophila that were distinct between whole-fly and mitochondrial enriched preparations. To illustrate the sensitivity of this method, we used a set of Drosophila harboring genetically diverse mitochondrial DNAs (mtDNA) and exposed them to the drug rapamycin. Using this method we showed that rapamycin modifies mitochondrial metabolism in a mitochondrial-genotype-dependent manner. However, these changes are much more distinct in metabolomics studies when metabolites were extracted from mitochondrial enriched fractions. In contrast, whole tissue extracts only detected metabolic changes mediated by the drug rapamycin independently of mtDNAs.

Preparation of Mitochondrial Enriched Fractions for Metabolic Analysis in Drosophila

Mitochondria play central roles in the regulation of metabolism and homeostasis. Subtle changes in mitochondrial metabolism that affect organismal physiology could be difficult to detect in whole organism metabolomics studies. Here we describe an isolation method that enhances the detection of subtle metabolic shifts in Drosophila melanogaster.

Since mitochondria play roles in amino acid metabolism, carbohydrate metabolism and fatty acid oxidation, defects in mitochondrial function often ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

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 ...

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

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 ...

Live Cell Imaging of Chromosome Segregation During Mitosis

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled by multiple mechanisms that include the Spindle Assembly Checkpoint (SAC). The SAC is part of a complex feedback system that is responsible for prevention of a cell progress through mitosis unless all chromosomal kinetochores have attached to spindle microtubules. Chromosomal lagging and abnormal chromosome segregation is an indicator of dysfunctional cell cycle control checkpoints and can be used to measure the genomic stability of dividing cells. Deregulation of the SAC can result in the transformation of a normal cell into a malignant cell through the accumulation of errors during chromosomal segregation. Implementation of the SAC and the formation of the kinetochore complex are tightly regulated by interactions between kinases and phosphatase such as Protein Phosphatase 2A (PP2A). This protocol describes live cell imaging of lagging chromosomes in mouse embryonic fibroblasts isolated from mice that had a knockout of the PP2A-B56&#947; regulatory subunit. This method overcomes the shortcomings of other cell cycle control imaging techniques such as flow cytometry or immunocytochemistry that only provide a snapshot of a cell cytokinesis status, instead of a dynamic spatiotemporal visualization of chromosomes during mitosis.

This protocol describes an easy and convenient method to label and visualize live chromosomes in mitotic cells using Histone2B-GFP BacMam 2.0 label and a spinning disc confocal microscopy system.

Chromosomes must be reliably and uniformly segregated into daughter cells during mitotic cell division. Fidelity of chromosomal segregation is controlled ...

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 ...

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 ...