We are extremely grateful to Jean-Antoine Lepesant and Nicolas Tissot who originally developed the protocol and shared some graphical elements of Figure 3 with us. We thank Fanny Roland-Gosselin who took the photos of Figure 4. We also thank other lab members for helpful discussions that contributed to the amelioration of this technique and Nathaniel Henneman for his comments that helped to improve this manuscript. We acknowledge the ImagoSeine core facility of the Institute Jacques Monod, member of France-BioImaging (ANR-10-INBS-04). Ma&#235;lys Loh is supported by a PhD fellowship from the French Ministry of Research (MESRI). Antoine Guichet and Fred Bernard were supported by the ARC (Grant PJA20181208148), the Association des Entreprises contre le Cancer (Grant Gefluc 2020 #221366) and by an Emergence grant from IdEx Universit&#233; de Paris (ANR-18-IDEX-0001).

1. Imaging medium preparation
<ol>
	<li>Prepare fresh media on the day of use. Pipette 200 &#181;L of Schneider medium (containing L-Glutamine and 0.40 g/L of NaHCO3 complemented with 10% heat-inactivated fetal calf serum, 100 U/mL of penicillin, and 100 mg/mL of streptomycin).</li>
	<li>Supplement with 30 &#181;L of insulin 10 mg/mL.</li>
	<li>Add 4 &#181;L of heat-inactivated fetal calf serum.</li>
</ol>
2. Observation-chamber preparation
<ol>
	<li>With a pipette tip, apply a small amount of silicone grease all around the hole on the underside of the punctured slide (Figure 4D).</li>
	<li>Position a 24 x 50 mm coverslip of 0.13-0.16 mm thickness.</li>
	<li>With the wide end of a pipette tip, apply pressure on the coverslip to flatten the silicone in order to seal the coverslip and create a silicone ring interior to the slide (Figure 4F,G).</li>
</ol>
3. Ovary dissection
<ol>
	<li>Anesthetize a female fly of the desired phenotype on a CO2 pad.</li>
	<li>Transfer the female in 150 &#181;L of the imaging medium in a dissecting well (Figure 4H).</li>
	<li>Open one female by grabbing its thorax with forceps and pinching the dorsal abdomen cuticle with a second pair of forceps.</li>
	<li>Isolate and detach the pair of ovaries, which should be readily visible upon cuticle opening.</li>
	<li>Carefully remove the uterus, oviduct, and muscle sheath (Figure 4I).</li>
	<li>Place a drop of 10-15 &#181;L of the imaging medium and transfer one ovary in the imaging chamber (Figure 4J,K).</li>
</ol>
4. Egg chamber isolation
<ol>
	<li>To separate the ovarioles, hold the posterior end of the ovary (toward the older stages) with the needle. Tease apart the ovarioles by carefully pulling on the germarium with another needle.</li>
	<li>Remove the remaining muscle sheath on the egg chambers; one needle holding the sheath and the other pulling on the ovariole through the larger chambers (stage 9 or older).</li>
	<li>Allow the unsheathed ovariole to sink and contact the coverslip.</li>
	<li>Remove late stages and the rest of the ovaries from the micro-chamber with the help of forceps. Carefully distance the ovarioles from the others with needles to facilitate the acquisition (Figure 4L).</li>
</ol>
5. Observation chamber closing
<ol>
	<li>Cut a small square (10 x 10 mm) of permeable membrane (Figure 4M,N).</li>
	<li>Carefully apply the membrane on top of the imaging medium to expel any air bubbles (Figure 4O).</li>
	<li>Hermetically seal the chamber with a thin layer of halocarbon oil around the well on the contour of the membrane (Figure 4P,Q).</li>
</ol>
6. Imaging
<ol>
	<li>Place the imaging set-up on the slide holder of the inverted microscope using a 40x objective (HCX PL Apo, 1.25NA, oil immersion).</li>
	<li>Locate and save positions of different stage 6 oocytes in which the nucleus is ready to migrate.</li>
	<li>Set-up the 488 and 561 nm lasers. With a measured output laser power of 150 mW, use 30% of the laser power and 300 ms and 500 ms exposure, respectively.</li>
	<li>Set-up the experiment. Take a time-lapse of 12 h with the interval of 15 min-41 sections with an interval of 1 &#181;m centered on the nucleus. 
	&#8203;NOTE: According to the exposure time described above, this setting allows the acquisition of one position in around 45 s. Since there is a delay due to position changing, it is recommended to set a maximum of 12 positions in these conditions.</li>
</ol>
7. Image analysis
<ol>
	<li>Process the movies on the software Fiji, using the plug-in Orthogonal view and manually track the nuclei.</li>
</ol>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/62688/62688fig04.jpg" /> 
Figure 4:&#160;Step by step micro-chamber mounting pictures. (A,B,C) Preparation of the needed tools: dissecting well plate, forceps, needles, imaging media, silicon grease, permeable membrane, and the aluminum slide. (D) Application of the silicon grease at the back of the aluminum slide with a pipette tip. (E) A glass coverslip is glued on the silicon grease to create the bottom of the chamber. (F,G) Pressure application on the coverslip with the wider extremity of a pipette tip to create a joint inside the chamber. (H,I)&#160;Dissection of the fly ovaries in the imaging media. (J) Pipetting of a drop of imaging media in the micro-chamber. (K,L) Separation of the ovarioles in the micro-chamber using needles. (M,N,O) Permeable membrane cut into a 10 x 10 mm square and placing over the drop of the medium in the micro-chamber. (P,Q) Sealing of the micro-chamber with halocarbon oil. The samples are ready to be imaged. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62688/62688fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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A., Guichet, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nucleus+positioning+within+Drosophila+egg+chamber.">Nucleus positioning within Drosophila egg chamber.</a> Seminars in Cell and Developmental Biology. 82, 25-33 (2017).</li><li>Koch, E. A., Spitzer, R. 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C. C., Starz-Gaiano, M., Melani, M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+culturing+drosophila+melanogaster+stage+9+egg+chambers+for+live+imaging.">A protocol for culturing drosophila melanogaster stage 9 egg chambers for live imaging.</a> Nature Protocols. 2 (10), 2467-2473 (2007).</li><li>Weil, T. T., Parton, R. M., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+individual+Drosophila+egg+chambers+for+live+imaging.">Preparing individual Drosophila egg chambers for live imaging.</a> Journal of Visualized Experiments: JoVE. (60), (2012).</li><li>Chanet, S., Huynh, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collective+cell+sorting+requires+contractile+cortical+waves+in+germline+cells.">Collective cell sorting requires contractile cortical waves in germline cells.</a> Current Biology. 30 (21), 4213-4226 (2020).</li><li>Zhao, T., Graham, O. 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The authors declare no competing interests.

Other protocols describe how to prepare and culture Drosophila egg chambers ex vivo for live-imaging assay12,13. The novelty of this protocol is the use of an imaging chamber constructed using a hollowed aluminum slide, a coverslip, and an O2/CO2 permeable membrane. The main advantage of this set-up is to limit the movement in Z without exerting pressure on the sample. Thus, the oocyte can still move freely, and this is why first, t...

For several years, the Drosophila oocyte has emerged as a model system to study nuclear migration. The Drosophila oocyte develops in a multicellular structure called the egg chamber. Egg chambers encompass 16 germ cells (15 nurse cells and the oocyte) surrounded by an epithelial layer of follicular somatic cells. Egg chamber development has been subdivided into 14 stages (Figure 1A), during which the oocyte will grow and accumulate reserves necessary for the early development of the embryo. During the development, upon microtubule reorganization and asymmetric transport of maternal determinants, the oocyte polarizes along the antero-dorsal and dorso-ventral axes. These axes determine the subsequent polarity axes of the embryo and the adult arising from the fertilization of this oocyte1. During oogenesis, the nucleus adopts an asymmetric position in the oocyte. In stage 6, the nucleus is centered in the cell. Upon a yet to be identified signal emitted by the posterior follicular cells which is received by the oocyte, the nucleus migrates toward the intersection between the anterior and lateral plasma membranes in stage 7 (Figure 1B)2,3. This asymmetric position is required to induce the determination of the dorso-ventral axis.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62688/62688fig1v2.jpg" /> 
Figure 1: Drosophila melanogaster egg chambers. (A) Fixed ovariole from transgenic flies expressing Fs(2)Ket-GFP that labels the nuclear envelopes and ubi-PH-RFP that labels the plasma membranes. The ovariole is composed of developing egg chambers at different stages. Maturation increases along the antero-posterior axis with the germarium at the anterior tip (left) where the germ stem cell resides and the older stage at the posterior tip (right). (B) Z-projection of living egg chamber by spinning disk confocal microscopy at stage 6 of oogenesis (left), in which the nucleus is centered in the oocyte. The nucleus will migrate to adopt an asymmetrical position at stage 7 (right) in contact with the anterior plasma membrane (between the oocyte and the nurse cell) and the lateral plasma membrane (between the oocyte and the follicular cells). This position will induce the determination of the dorsal side and, thus, the dorso-ventral axis of the egg chamber. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62688/62688fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
For many decades, this nuclear migration has been studied on fixed tissues by immunostaining. This approach has notably made it possible to demonstrate that this process depends on a dense network of microtubules4,5. More recently, we developed a protocol offering conditions compatible with live imaging of the oocyte during several hours making it possible to study this process dynamically6.
Hence, for the first time, we have been able to describe that the nucleus has preferential and characteristic trajectories during its migration, one along the anterior plasma membrane (APM) and another along the lateral plasma membrane (LPM) of the oocyte (Figure 2). These latest results underline the importance of live-imaging protocols when studying dynamic processes such as nuclear migration.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/62688/62688fig02.jpg" /> 
Figure 2: Schematic representation of the different migration paths of the nucleus. At stage 6 of the oogenesis, the oocyte is a large cell with a central nucleus. At this stage, the antero-posterior polarity axis is set with a posterior/lateral plasma membrane of the oocyte in contact with the follicular cells and the anterior plasma membrane (in yellow) is in contact with the nurse cells2. We have previously reported that the nucleus could migrate either along the anterior plasma membrane (APM), along the lateral plasma membrane (LPM), or through the cytoplasm (STAD, straight to the antero-dorsal cortex)6. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62688/62688fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The oocyte nucleus migration is a phenomenon of about 3 h6, and so far, the event triggering the start of the actual migration is unknown. The start of the migration can also be delayed by protein mutants used to study this mechanism. These unknown variables motivated us to acquire images over long time periods (10-12 h). It is, therefore, important to ensure that the oocytes remain alive. As the egg chamber develops, it elongates along the antero-posterior axis from a spherical to an elliptical shape. This elongation is driven by the rotation of follicular cells, which occurs from stage 1 to stage 8, perpendicular to the antero-posterior axis7. In addition, a tubular sheath of muscle with pulsatile property surrounds the egg chambers. Its physiological function is to push the developing follicles toward the oviduct continuously8. In order to limit the movements that induce oscillations of the egg chambers after their dissection, we designed an observation micro-chamber measuring 150 &#181;m in height (Figure 3A). This height is marginally higher than the size of a follicle at stages 10 and 11. It considerably limits the vertical movements of the sample while preserving the rotation of the egg chamber, thereby resulting in limited defects in follicle development. We then perform live imaging for 12 h on dissected egg chambers by multi-position time-lapse acquisitions using a spinning-disk confocal microscope. Here we describe our protocol for studying the oocyte nuclear migration between stages 6 and 7.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62688/62688fig3v3.jpg" /> 
Figure 3: Schematic representation of the observation chamber. (A) (Top view) Precise dimensions of the aluminum slide with the heights (A&#39;) and circumferences (A&#39;&#39;) of the well drilled in the middle of the slide. (B) (Bottom view) A coverslip blocking the well is sealed to the slide with silicon grease. (C) (Top view) Dissected ovarioles develop in an imaging medium that is covered by a gas permeable membrane. Halocarbon oil is used to stabilize the membrane. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62688/62688fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
In order to follow the nuclear migration and precisely assess trajectories in the oocyte, markers for both the nuclear envelope and plasma membrane are needed. With this aim, two transgenes that have a high signal/noise ratio and do not fade over the course of live imaging have been selected. To label the plasma membrane, the use of a P[ubi-PH-RFP] that encodes the Pleckstrin Homology (PH) domain of the Human Phospholipase C &#8706;1 (PLC&#8706;1) fused to RFP is recommended. This PH domain binds to the phosphoinositide PI(4,5)P2 distributed along the plasma membrane of the oocyte9. For the nuclear envelope, the P[PPT-un1]Fs(2)Ket-GFP protein-trap strain where GFP is inserted within the gene encoding the Drosophila &#223;-importin displays a homogeneous and an intense signal10. Young flies (1-2 days old) are placed in fresh vials containing dry yeast 24-48 h prior to ovary dissection.
For this live-imaging assay, a 1 mm thick piece of aluminum, which is nonreactive for the sample, has been cut into the dimensions of a microscopy slide. It has a 16 mm diameter hole in the center of the slide that has been counterbored to 0.85 mm. This counterbore has an additional 6 mm diameter hole with a depth of 150 &#181;m (Figure 3A). A coverslip is glued with silicone grease (inert for the sample) at the bottom of the aluminum chamber (Figure 3B). After placing the samples in the medium-filled well, a membrane permeable to O2/CO2 exchange is placed over the medium and surrounded by halocarbon oil (Figure 3C).
For the dissection, it is recommended to use stainless steel forceps with a tip dimension of 0.05 x 0.02 mm, and 0.20 mm diameter needles for the separation of the ovarioles (Figure 4B,C). The migrating nuclei are imaged on a spinning-disk confocal inverted microscope CSU-X1 equipped with a camera. Multi-position images were acquired by time-lapse every 15 min at 24 &#176;C. A 15 min interval allows performing multi-position acquisitions with limited photobleaching of the fluorescent proteins and phototoxicity for the samples. Furthermore, a shorter interval would not provide much more informative data to follow the nuclear trajectories. The movies are processed and analyzed via Fiji software11.

<table><tbody><tr><td>Anesthetize CO2 pad</td><td>Dutscher</td><td>789060</td><td>Anesthetize flies</td></tr><tr><td>Coverslip (24x50 mm)</td><td>Knittel Glass</td><td>VD12450Y100A</td><td>Observation-chamber preparation</td></tr><tr><td>Forceps Dumont #5</td><td>Carl Roth</td><td>K342.1</td><td>Dissection</td></tr><tr><td>Stainless steel needles</td><td>Entosphinx</td><td>20</td><td>Dissection</td></tr><tr><td>Heat-inactivated fetal calf serum</td><td>SIGMA-ALDRICH</td><td>F7524</td><td>Imaging medium</td></tr><tr><td>Insulin solution bovine pancreas</td><td>SIGMA-ALDRICH</td><td>10516 - 5ml</td><td>Imaging medium</td></tr><tr><td>Penicilin/Streptomycin solution</td><td>SIGMA-ALDRICH</td><td>P0781</td><td>Imaging medium</td></tr><tr><td>Permeable membrane</td><td>Leica</td><td>11521746</td><td>Observation-chamber preparation</td></tr><tr><td>Schneider Medium</td><td>Pan Biotech</td><td>P04-91500</td><td>Imaging medium</td></tr><tr><td>Silicon grease</td><td>BECKMAN COULTER</td><td>335148</td><td>Observation-chamber preparation</td></tr><tr><td>Spinning disk confocal</td><td>Zeiss</td><td>CSU-X1</td><td>Nuclear migration observation</td></tr><tr><td>Voltalef oil 10S</td><td>VWR</td><td>24627 - 188</td><td>Observation-chamber preparation</td></tr></tbody></table>

nuclear migration in the drosophila oocyte

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton rearrangements, or polarity patterning within the cells. When studying oocyte nucleus positioning, live-imaging techniques are essential to capture the dynamic events of this process. The Drosophila egg chamber is a multicellular structure and an excellent model system to study this phenomenon because of its large size and availability of numerous genetic tools. During Drosophila mid-oogenesis, the nucleus migrates from a central position within the oocyte to adopt an asymmetric position mediated by microtubule-generated forces. This migration and positioning of the nucleus are necessary to determine the polarity axes of the embryo and the subsequent adult fly. One characteristic of this migration is that it occurs in three dimensions (3D), creating a necessity for live imaging. Thus, to study the mechanisms that regulate nuclear migration, we have developed a protocol to culture the dissected egg chambers and perform live imaging for 12 h by time-lapse acquisitions using spinning-disk confocal microscopy. Overall, our conditions allow us to preserve Drosophila egg chambers alive for a long period of time, thereby enabling the completion of nuclear migration to be visualized in a large number of samples in 3D.

Before migration, the nucleus is dynamic and oscillates around a central position during a period defined as pre-migration. These small movements reflect a balance of pushing and pulling forces that maintain equilibrium in the middle of the oocyte. By quantifying the trajectories of the nuclei, we have shown that the APM and LPM trajectories had similar proportions. We define the nature of the trajectory by the first contact between the nucleus and the plasma membrane6. Thus, the nucleus reaches e...

Watch this Scientific Journal Video about Nuclear Migration in the Drosophila Oocyte at JoVE.com

Nuclear Migration in the Drosophila Oocyte

In Drosophila, the oocyte nucleus undergoes microtubule-dependent migration during oogenesis. Here, we describe a protocol that was developed to follow the migration by performing live imaging on egg chambers ex-vivo. Our procedure maintains egg chambers alive for 12 h to acquire multi-position 3D time-lapse movies using spinning-disk confocal microscopy.

Nuclear Migration in the Drosophila Oocyte

Live cell imaging is particularly necessary to understand the cellular and molecular mechanisms that regulate organelle movements, cytoskeleton ...

nuclear-migration-in-the-drosophila-oocyte

Research

JoVE Journal

Developmental Biology

3.7K Views.  Université de Paris, CNRS, Institut Jacques Monod. In Drosophila, the oocyte nucleus undergoes microtubule-dependent migration during oogenesis. Here, we describe a protocol that was developed to follow the migration by performing live imaging on egg chambers ex-vivo. Our procedure maintains egg chambers alive for 12 h to acquire multi-position 3D time-lapse movies using spinning-disk confocal microscopy.

Video: Nuclear Migration in the Drosophila Oocyte

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and transmitted from one generation to the next through the female oocyte is of fundamental importance. Because of the genetic tractability, and the elegant, ordered simplicity by which oocyte development proceeds, Drosophila oogenesis has become an invaluable system for mitochondrial study. An EdU (5-ethynyl-2&#180;-deoxyuridine) labeling method was utilized to detect mitochondrial DNA (mtDNA) replication in Drosophila ovaries. This method is superior to the BrdU (5-bromo-2'-deoxyuridine) labeling method in that it allows for good structural preservation and efficient fluorescent dye penetration of whole-mount tissues.
Here we describe a detailed protocol for labeling replicating mitochondrial DNA in Drosophila adult ovaries with EdU. Some technical solutions are offered to improve the viability of the ovaries, maintain their health during preparation, and ensure high-quality imaging. Visualization of newly synthesized mtDNA in the ovaries not only reveals the striking temporal and spatial pattern of mtDNA replication through oogenesis, but also allows for simple quantification of mtDNA replication under various genetic and pharmacological perturbations.

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

Drosophila oogenesis continues to be exceptionally useful in the study of mitochondrial proliferation and inheritance. This manuscript describes a detailed protocol used to label the replicating mitochondrial DNA (mtDNA) in Drosophila adult ovaries with 5-ethynyl-2&#180;-deoxyuridine (EdU), which facilitates uncovering mechanisms associated with mitochondrial inheritance that were previously debatable.

The mitochondrial genome is inherited exclusively through the maternal line. Understanding of how the mitochondrion and its genome are proliferated and ...

Genetics

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 ...

Microinjection of Xenopus Laevis Oocytes

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic transport. Because of its large nucleus and high density of nuclear pore complexes (NPCs), nuclear transport can be easily visualized in the Xenopus oocyte. Much insight into the mechanisms of nuclear import and export has been gained through use of this system (reviewed by Pant&eacute;, 2006). In addition, we have used microinjection of Xenopus oocytes to dissect the nuclear import pathways of several viruses that replicate in the host nucleus. 
Here we demonstrate the cytoplasmic microinjection of Xenopus oocytes with a nuclear import substrate. We also show preparation of the injected oocytes for visualization by thin-sectioning EM, including dissection, dehydration, and embedding of the oocytes into an epoxy embedding resin. Finally, we provide representative results for oocytes that have been microinjected with the capsid of the baculovirus Autographa californica nucleopolyhedrovirus (AcMNPV) or the parvovirus Minute Virus of Mice (MVM), and discuss potential applications of the technique.

Here we demonstrate cytoplasmic microinjection of Xenopus laevis oocytes with a nuclear import substrate, as well as preparation of the injected oocytes for visualization by thin-sectioning electron microscopy.

Microinjection of Xenopus laevis oocytes followed by thin-sectioning electron microscopy (EM) is an excellent system for studying nucleocytoplasmic ...

Biology

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk of conceiving an aneuploid fetus increases dramatically and this phenomenon is known as the maternal age effect. One requirement for accurate chromosome segregation during the meiotic divisions is maintenance of sister chromatid cohesion during the extended prophase period that oocytes experience. Cytological evidence in both humans and model organisms suggests that meiotic cohesion deteriorates during the aging process. In addition, segregation errors in human oocytes are most prevalent during meiosis I, consistent with premature loss of arm cohesion. The use of model organisms is critical for unraveling the mechanisms that underlie age-dependent loss of cohesion. Drosophila melanogaster offers several advantages for studying the regulation of meiotic cohesion in oocytes. However, until recently, only genetic tests were available to assay for loss of arm cohesion in oocytes of different genotypes or under different experimental conditions. Here, a detailed protocol is provided for using fluorescence in situ hybridization (FISH) to directly visualize defects in arm cohesion in prometaphase I and metaphase I arrested Drosophila oocytes. By generating a FISH probe that hybridizes to the distal arm of the X chromosome and collecting confocal Z stacks, a researcher can visualize the number of individual FISH signals in three dimensions and determine whether sister chromatid arms are separated. The procedure outlined makes it possible to quantify arm cohesion defects in hundreds of Drosophila oocytes. As such, this method provides an important tool for investigating the mechanisms that contribute to cohesion maintenance as well as the factors that lead to its demise during the aging process.

Using Fluorescence In Situ Hybridization FISH to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

This manuscript presents a detailed method for generating X-chromosome arm probes and performing fluorescence in situ hybridization (FISH) to examine the state of sister chromatid cohesion in prometaphase and metaphase I arrested Drosophila oocytes. This protocol is suitable for determining whether meiotic arm cohesion is intact or disrupted in different genotypes.

In humans, chromosome segregation errors in oocytes are responsible for the majority of miscarriages and birth defects. Moreover, as women age, their risk ...

Dissection and Staining of Drosophila Larval Ovaries

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells form, and how they interact with their environment is therefore crucial for understanding development, homeostasis and disease. The ovary of the fruit fly Drosophila melanogaster has served as an influential model for the interaction of germ line stem cells (GSCs) with their somatic support cells (niche) 1, 2. The known location of the niche and the GSCs, coupled to the ability to genetically manipulate them, has allowed researchers to elucidate a variety of interactions between stem cells and their niches 3-12.
Despite the wealth of information about mechanisms controlling GSC maintenance and differentiation, relatively little is known about how GSCs and their somatic niches form during development. About 18 somatic niches, whose cellular components include terminal filament and cap cells (Figure 1), form during the third larval instar 13-17. GSCs originate from primordial germ cells (PGCs). PGCs proliferate at early larval stages, but following the formation of the niche a subgroup of PGCs becomes GSCs 7, 16, 18, 19. Together, the somatic niche cells and the GSCs make a functional unit that produces eggs throughout the lifetime of the organism.
Many questions regarding the formation of the GSC unit remain unanswered. Processes such as coordination between precursor cells for niches and stem cell precursors, or the generation of asymmetry within PGCs as they become GSCs, can best be studied in the larva. However, a methodical study of larval ovary development is physically challenging. First, larval ovaries are small. Even at late larval stages they are only 100&mu;m across. In addition, the ovaries are transparent and are embedded in a white fat body. Here we describe a step-by-step protocol for isolating ovaries from late third instar (LL3) Drosophila larvae, followed by staining with fluorescent antibodies. We offer some technical solutions to problems such as locating the ovaries, staining and washing tissues that do not sink, and making sure that antibodies penetrate into the tissue. This protocol can be applied to earlier larval stages and to larval testes as well.

Dissection and Staining of Drosophila Larval Ovaries

How niches and stem cells form during development is an important question with practical implications. In the Drosophila ovary, germ line stem cells and their somatic niches form during larval development. This video demonstrates how to dissect, stain and mount female gonads from late third instar (LL3) Drosophila larvae.

Many organs depend on stem cells for their development during embryogenesis and for maintenance or repair during adult life. Understanding how stem cells ...

Preparing Individual Drosophila Egg Chambers for Live Imaging

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

Preparing Individual Drosophila Egg Chambers for Live Imaging

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

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

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The study of chromosome behavior in oocytes from model organisms holds much promise to uncover the molecular basis of the susceptibility of human oocytes to aneuploidy. Drosophila melanogaster is amenable to genetic manipulation, with over 100 years of research, community, and technique development. Visualizing chromosome behavior and spindle assembly in Drosophila oocytes has particular challenges, however, due primarily to the presence of membranes surrounding the oocyte that are impenetrable to antibodies. We describe here protocols for the collection, preparation, and imaging of meiosis I spindle assembly and chromosome behavior in Drosophila oocytes, which allow the molecular dissection of chromosome segregation in this important model organism.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The ...

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell imaging studies. Methods for detecting RNAs have expanded greatly since seminal studies linked site-specific mRNA localization to gene expression regulation. Dynamic mRNA processes can now be visualized via approaches that detect mRNAs, coupled with microscopy set-ups that are fast enough to capture the dynamic range of molecular behavior. The molecular beacon technology is a hybridization-based approach capable of direct detection of endogenous transcripts in living cells. Molecular beacons are hairpin-shaped, internally quenched, single-nucleotide discriminating nucleic acid probes, which fluoresce only upon hybridization to a unique target sequence. When coupled with advanced fluorescence microscopy and high-resolution imaging, they enable one to perform spatial and temporal tracking of intracellular movement of mRNAs. Although this technology is the only method capable of detecting endogenous transcripts, cell biologists have not yet fully embraced this technology due to difficulties in designing such probes for live cell imaging. A new software application, PinMol, allows for enhanced and rapid design of probes best suited to efficiently hybridize to mRNA target regions within a living cell. In addition, high-resolution, real-time image acquisition and current, open source image analysis software allow for a refined data output, leading to a finer evaluation of the complexity underlying the dynamic processes involved in the mRNA&#39;s life cycle.
Here we present a comprehensive protocol for designing and delivering molecular beacons into Drosophila melanogaster egg chambers. Direct and highly specific detection and visualization of endogenous maternal mRNAs is performed via spinning disc confocal microscopy. Imaging data is processed and analyzed using object detection and tracking in Icy software to obtain details about the dynamic movement of mRNAs, which are transported and localized to specialized regions within the oocyte.

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers

Here, we present a protocol for the visualization, detection, analysis and tracking of endogenous mRNA trafficking in live Drosophila melanogaster egg chamber using molecular beacons, spinning disc confocal microscopy, and open-source analysis software.

Fluorescence-based imaging techniques, in combination with developments in light microscopy, have revolutionized how cell biologists conduct live cell ...

Nuclear Migration in the Drosophila Oocyte

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Chapters in this video

In Situ Labeling of Mitochondrial DNA Replication in Drosophila Adult Ovaries by EdU Staining

Upright Imaging of Drosophila Egg Chambers

Microinjection of Xenopus Laevis Oocytes

Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

Dissection and Staining of Drosophila Larval Ovaries

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Imaging Through the Pupal Case of Drosophila melanogaster

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Visualizing and Tracking Endogenous mRNAs in Live Drosophila melanogaster Egg Chambers