This work was supported by NIH Grant GM59354 awarded to Sharon E. Bickel. We thank Huy Q. Nguyen for assistance in developing the protocol for generating fluorescent arm probes, Ann Lavanway for help with confocal microscopy, and J. Emiliano Reed for technical assistance. We also thank numerous colleagues in the Drosophila community for helpful discussions and advice.

<ol>
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B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disjunction+of+homologous+chromosomes+in+meiosis+I+depends+on+proteolytic+cleavage+of+the+meiotic+cohesin+Rec8+by+separin.">Disjunction of homologous chromosomes in meiosis I depends on proteolytic cleavage of the meiotic cohesin Rec8 by separin.</a> Cell. 103 (3), 387-398 (2000).</li><li>Bickel, S. E., Orr-Weaver, T., Balicky, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+sister-chromatid+cohesion+protein+ORD+is+required+for+chiasma+maintenance+in+Drosophila+oocytes.">The sister-chromatid cohesion protein ORD is required for chiasma maintenance in Drosophila oocytes.</a> Curr. Biol. 12 (11), 925-929 (2002).</li><li>Hodges, C. A., Revenkova, E., Jessberger, R., Hassold, T. J., Hunt, P. 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The authors declare no competing financial interests.

The use of FISH probes to assess the state of arm cohesion in prometaphase I and metaphase I Drosophila oocytes is a significant advancement in the field of Drosophila meiosis. Historically, Drosophila researchers have been limited to genetic tests to infer premature loss of arm cohesion in mature oocytes11,18,19. Now, with the methods presented here, the state of arm cohesion can be assayed directly u...

Proper segregation of chromosomes during mitosis and meiosis requires that sister chromatid cohesion be established, maintained, and released in a coordinated fashion1,2. Cohesion is established during S phase and is mediated by the cohesin complex, which forms physical linkages that hold the sister chromatids together. In meiosis, cohesion distal to a crossover also functions to hold recombinant homologs together and this physical association helps ensure proper orientation of the bivalent on the metaphase I spindle (Figure 1)3,4,5. Release of arm cohesion at anaphase I allows the homologs to segregate to opposite spindle poles. However, if arm cohesion is lost prematurely, recombinant homologs will lose their physical connection and segregate randomly, which can result in aneuploid gametes (Figure 1).
In human oocytes, errors in chromosome segregation are the leading cause of miscarriages and birth defects, such as Down Syndrome6, and their incidence increases exponentially with maternal age7. Sister chromatid cohesion is established in fetal oocytes and meiotic recombination is completed before birth. Oocytes then arrest in mid-prophase I until ovulation and during this arrest, the continued physical association of recombinant homologs relies on sister chromatid cohesion. Therefore, accurate segregation during meiosis and normal pregnancy outcomes require that cohesion remain intact for up to five decades.
Premature loss of cohesion during the prolonged meiotic arrest of human oocytes has been proposed to contribute to the maternal age effect and multiple lines of evidence support this hypothesis8,9. However, given the challenges of studying meiotic cohesion in human oocytes, much of our understanding of this phenomenon relies on the use of model organisms5,10,11,12,13,14,15.
Drosophila melanogaster oocytes offer numerous advantages for the study of meiotic cohesion and chromosome segregation. A simple genetic assay allows one to recover progeny from aneuploid gametes and measure the fidelity of X-chromosome segregation on a large scale11,16,17. Moreover, one may also determine whether chromosome segregation errors arise because recombinant homologs missegregate during meiosis I, a phenotype that is consistent with premature loss of arm cohesion11,18,19. Direct observation of the state of meiotic cohesion in Drosophila oocytes is also possible using fluorescence in situ hybridization (FISH). Although fluorescent oligonucleotides that hybridize to repetitive satellite sequences have been used for over a decade to monitor pericentromeric cohesion in mature Drosophila oocytes4,20, analysis of arm cohesion has been much more challenging. Visualization of the state of arm cohesion requires a probe that spans a large region of single copy sequences and is bright enough to result in visible signals for individual sister chromatids when arm cohesion is absent. In addition, the oocyte fixation conditions and size of the labeled DNAs must facilitate penetration21 into the large mature Drosophila oocyte (200 &#181;m wide by 500 &#181;m long). Recently, an arm probe was successfully utilized to visualize Drosophila oocyte chromatids during anaphase I, but the authors stated that they could not detect a signal in prometaphase or metaphase I arrested oocytes22. Here we provide a detailed protocol for the generation of X-chromosome arm FISH probes and oocyte preparation conditions that have allowed us to assay for premature loss of sister chromatid cohesion in prometaphase I and metaphase I oocytes. These techniques, which have enabled us to identify gene products that are required for the maintenance of meiotic cohesion, will allow others to assay for sister chromatid cohesion defects in mature Drosophila oocytes of different genotypes.

<table><tbody><tr><td>&lt;strong&gt;Kits&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>Midi Prep kit</td><td>Qiagen</td><td>12143</td><td>Prep BAC clone DNA</td></tr><tr><td>GenomePlex Complete Whole Genome Amplification (WGA) Kit</td><td>Sigma</td><td>WGA2</td><td>Amplify BAC clone DNA</td></tr><tr><td>ARES Alexa Fluor 647 DNA labeling kit</td><td>Invitrogen</td><td>A21676</td><td>Label BAC clone DNA</td></tr><tr><td>PCR purification kit</td><td>Qiagen</td><td>28104</td><td>Remove non-conjugated dye following labeling of BAC clone DNA</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Chemicals &amp;amp; Solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Note: &lt;/strong&gt;All solutions are prepared using sterile ultrapure water and should be sterilized either by autoclave or filter sterilization.</td><td></td><td></td><td></td></tr><tr><td>Bovine serum albumin (BSA)</td><td>Fisher Scientific</td><td>BP1600-100</td><td>Prepare 10% stock&lt;br/&gt; Freeze aliquots</td></tr><tr><td>Calcium chloride</td><td>Fisher Scientific</td><td>C75-500</td><td></td></tr><tr><td>DAPI (4&amp;rsquo;, 6-Diamidino-2-Phenylindole, Dihydrochloride)</td><td>Invitrogen</td><td>D1306</td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection. Prepare 100&amp;micro;g/ml stock in 100% ethanol, store in aliquots at -20 &amp;deg;C. Prepare 1 &amp;micro;g/ml solution in 2X SSCT before use.</td></tr><tr><td>Dextran sulfate</td><td>Sigma</td><td>D-8906</td><td></td></tr><tr><td>Drierite</td><td>Drierite Company</td><td>23001</td><td></td></tr><tr><td>Dithiothreitol (DTT)</td><td>Invitrogen</td><td>15508-013</td><td>Prepare 10 mM stock</td></tr><tr><td>dTTP (10 &amp;micro;mol, 100 &amp;micro;l)</td><td>Boehringer Mannheim</td><td>1277049</td><td>Prepare 1 mM stock</td></tr><tr><td>EDTA (Disodium ethylenediamine tetraacetic acid)</td><td>Fisher Scientific</td><td>S311-500</td><td>Prepare 250 mM stock</td></tr><tr><td>100% ethanol (molecular grade, 200 proof)</td><td>Decon Laboratories</td><td>2716</td><td></td></tr><tr><td>Tris (Ultra Pure)</td><td>Invitrogen</td><td>15504-020</td><td></td></tr><tr><td>EGTA (Ethylenebis(oxyethylenenitrilo)tetraacetic acid)</td><td>Sigma</td><td>E-3889</td><td></td></tr><tr><td>16% formaldehyde</td><td>Ted Pella, Inc.</td><td>18505</td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection</td></tr><tr><td>Formamide</td><td>Invitrogen</td><td>AM9342</td><td>&lt;strong&gt;Toxic: &lt;/strong&gt;wear appropriate protection</td></tr><tr><td>Glucose</td><td>Fisher Scientific</td><td>D16-1</td><td></td></tr><tr><td>Glycogen</td><td>Roche</td><td>901393</td><td></td></tr><tr><td>HEPES</td><td>Boehringer Mannheim</td><td>737-151</td><td></td></tr><tr><td>Heptane</td><td>Fisher Scientific</td><td>H350-4</td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection.</td></tr><tr><td>Hydrochloric acid</td><td>Millipore</td><td>HX0603-4</td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection.</td></tr><tr><td>Hydroxylamine</td><td>Sigma</td><td>438227</td><td>Prepare 3 M stock</td></tr><tr><td>4.9 M Magnesium chloride</td><td>Sigma</td><td>104-20</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4 &lt;/sub&gt;&amp;Yuml; 7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Fisher Scientific</td><td>S373-500</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4 &lt;/sub&gt;&amp;Yuml; 2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Fisher Scientific</td><td>S369-500</td><td></td></tr><tr><td>Poly-L-lysine (0.1mg/ml)</td><td>Sigma</td><td>P8920-100</td><td></td></tr><tr><td>Potassium acetate</td><td>Fisher Scientific</td><td>BP364-500</td><td></td></tr><tr><td>Sodium acetate</td><td>Fisher Scientific</td><td>S209-500</td><td>Prepare 3M stock</td></tr><tr><td>Sodium cacodylate</td><td>Polysciences, Inc.</td><td>1131</td><td>&lt;strong&gt;Toxic: &lt;/strong&gt;wear appropriate protection. Prepare 400mM stock</td></tr><tr><td>Sodium citrate</td><td>Fisher Scientific</td><td>BP327-1</td><td></td></tr><tr><td>Sodium chloride</td><td>Fisher Scientific</td><td>S271-3</td><td>Sodium chloride</td></tr><tr><td>Sucrose</td><td>Fisher Scientific</td><td>S5-500</td><td></td></tr><tr><td>10% Tween 20</td><td>Thermo Scientific</td><td>28320</td><td>Surfact-Amps</td></tr><tr><td>10% Triton X-100</td><td>Thermo Scientific</td><td>28314</td><td>Surfact-Amps</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Note: &lt;/strong&gt;All solutions are prepared using sterile ultrapure water and should be sterilized either by autoclave or filter sterilization.</td><td></td><td></td><td></td></tr><tr><td>TE buffer</td><td></td><td></td><td>10 mM Tris, 1 mM EDTA, pH = 8.0</td></tr><tr><td>20X SSC (Saline Sodium Citrate)</td><td></td><td></td><td>3 M NaCl, 300 mM sodium citrate</td></tr><tr><td>2X cacodylate fix solution</td><td></td><td></td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection. 200 mM sodium cacodylate, 200 mM sucrose, 80 mM sodium acetate, 20 mM EGTA</td></tr><tr><td>1.1X Hybridization buffer</td><td></td><td></td><td>3.3X SSC, 55% formamide, 11% dextran sulfate</td></tr><tr><td>Fix solution</td><td></td><td></td><td>&lt;strong&gt;Toxic: &lt;/strong&gt;wear appropriate protection. 4% formaldehyde, 1X cacodylate fix solution</td></tr><tr><td>PBSBTx</td><td></td><td></td><td>1X PBS, 0.5% BSA, 0.1% Trition X-100</td></tr><tr><td>PBSTx</td><td></td><td></td><td>1X PBS, 1% Trition X-100</td></tr><tr><td>Extraction buffer (PBSTx + Rnase)</td><td></td><td></td><td>1X PBS, 1% Trition X-100, 100 &amp;micro;g/mL RNase</td></tr><tr><td>2X SSCT</td><td></td><td></td><td>2X SSC, 0.1% Tween 20</td></tr><tr><td>2X SSCT + 20% formamide</td><td></td><td></td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection. 2X SSC, 0.1% Tween 20, 20% formamide</td></tr><tr><td>2X SSCT + 40% formamide</td><td></td><td></td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection. 2X SSC, 0.1% Tween 20, 40% formamide</td></tr><tr><td>2X SSCT + 50% formamide</td><td></td><td></td><td>&lt;strong&gt;Toxic:&lt;/strong&gt; wear appropriate protection. 2X SSC, 0.1% Tween 20, 50% formamide</td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Catalog Number&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>&lt;strong&gt;Enzymes&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>AluI</td><td>New England Biolabs</td><td>R0137S</td><td></td></tr><tr><td>HaeIII</td><td>New England Biolabs</td><td>R0108S</td><td></td></tr><tr><td>MseI</td><td>New England Biolabs</td><td>R0525S</td><td></td></tr><tr><td>MspI</td><td>New England Biolabs</td><td>R0106S</td><td></td></tr><tr><td>RsaI</td><td>New England Biolabs</td><td>R0167S</td><td></td></tr><tr><td>BfuCI</td><td>New England Biolabs</td><td>R0636S</td><td></td></tr><tr><td>100X BSA</td><td>New England Biolabs</td><td></td><td>Comes with NEB enzymes</td></tr><tr><td>10X NEB buffer #2</td><td>New England Biolabs</td><td></td><td>Restriction enzyme digestion buffer. 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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.

Figure 5 presents images obtained with an arm probe that hybridizes to cytological region 6E-7B on the X chromosome. This probe results in a signal that co-localizes with that of DAPI, is easily distinguishable from the background, and has been used successfully to quantify arm cohesion defects in different genotypes19. Quantification of cohesion defects was restricted to prometaphase I and metaphase I stages; oocytes prior to...

Watch this Scientific Journal Video about Using Fluorescence In Situ Hybridization FISH to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes at JoVE.com

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

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.

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 ...

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11.3K Views.  Dartmouth College. 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.

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

Using Mouse Oocytes to Assess Human Gene Function During Meiosis I

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively correlated with maternal age, age alone is not always predictive of the risk of generating an aneuploid embryo. Therefore, gene variants might account for incorrect chromosome segregation during oogenesis. Given that access to human oocytes is limited for research purposes, a series of assays were developed to study human gene function during meiosis I using mouse oocytes. First, messenger RNA (mRNA) of the gene and gene variant of interest are microinjected into prophase I-arrested mouse oocytes. After allowing time for expression, oocytes are synchronously released into meiotic maturation to complete meiosis I. By tagging the mRNA with a sequence of a fluorescent reporter, such as green fluorescent protein (Gfp), the localization of the human protein can be assessed in addition to the phenotypic alterations. For example, gain or loss of function can be investigated by establishing experimental conditions that challenge the gene product to fix meiotic errors. Although this system is advantageous in investigating human protein function during oogenesis, adequate interpretation of results should be undertaken given that protein expression is not at endogenous levels and, unless controlled for (i.e. knocked out or down), murine homologs are also present in the system.

As the genetic variants associated with human disease begin to become uncovered, it is becoming increasingly important to develop systems with which to rapidly evaluate the biological significance of those identified variants. This protocol describes methods for evaluating human gene function during female meiosis I using mouse oocytes.

Embryonic aneuploidy is the major genetic cause of infertility in humans. Most of these events originate during female meiosis, and albeit positively ...

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), quantitative, real-time RT-PCR (RT-qPCR) and RNA sequencing. When these techniques are performed using a single oocyte or embryo, low-copy mRNAs are not reliably detected. To overcome this problem, oocytes or embryos can be pooled together for analysis; however, this often leads to high variability amongst samples. In this protocol, we describe the use of fluorescence in situ hybridization (FISH) using branched DNA chemistry. This technique identifies the spatial pattern of mRNAs in individual cells. When the technique is coupled with Spot Finding and Tracking&#160;computer software, the abundance of mRNAs in the cell can also be quantified. Using this technique, there is reduced variability within an experimental group and fewer oocytes and embryos are required to detect significant differences between experimental groups. Commercially available branched-DNA SM-FISH kits have been optimized to detect mRNAs in sectioned tissues or adherent cells on slides. However, oocytes do not effectively adhere to slides and some reagents in the kit were too harsh resulting in oocyte lysis. To prevent this lysis, several modifications were made to the FISH kit. Specifically, oocyte permeabilization and wash&#160;buffers designed for the immunofluorescence of oocytes and embryos replaced the proprietary buffers. The permeabilization, washes, and incubations with probes and amplifier were performed in 6-well plates and oocytes were placed on slides at the end of the protocol using mounting media. These modifications were able to overcome the limitations of the commercially available kit, in particular, the oocyte lysis. To accurately and reproducibly count the number of mRNAs in individual oocytes, computer software was used. Together, this protocol represents an alternative to PCR and sequencing to compare the expression of specific transcripts in single cells.

Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes

To reproducibly count the numbers of mRNAs in individual oocytes, single molecule RNA fluorescence in situ hybridization (RNA-FISH) was optimized for non-adherent cells. Oocytes were collected, hybridized with the transcript specific probes, and quantified using an image quantification software.

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

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

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

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

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

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.

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.

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

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

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

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

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

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Live Imaging of 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 ...

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Assessing the expression pattern of a gene, as well as the subcellular localization properties of its transcribed RNA, are key features for understanding its biological function during development. RNA in situ hybridization (RNA-ISH) is a powerful method used for visualizing RNA distribution properties, be it at the organismal, cellular or subcellular levels 1. RNA-ISH is based on the hybridization of a labeled nucleic acid probe (e.g. antisense RNA, oligonucleotides) complementary to the sequence of an mRNA or a non-coding RNA target of interest 2. As the procedure requires primary sequence information alone to generate sequence-specific probes, it can be universally applied to a broad range of organisms and tissue specimens 3. Indeed, a number of large-scale ISH studies have been implemented to document gene expression and RNA localization dynamics in various model organisms, which has led to the establishment of important community resources 4-11. While a variety of probe labeling and detection strategies have been developed over the years, the combined usage of fluorescently-labeled detection reagents and enzymatic signal amplification steps offer significant enhancements in the sensitivity and resolution of the procedure 12. Here, we describe an optimized fluorescent in situ hybridization method (FISH) employing tyramide signal amplification (TSA) to visualize RNA expression and localization dynamics in staged Drosophila embryos. The procedure is carried out in 96-well PCR plate format, which greatly facilitates the simultaneous processing of large numbers of samples.

Whole Mount RNA Fluorescent in situ Hybridization of Drosophila Embryos

Here we describe a whole-mount fluorescent in situ hybridization (FISH) protocol for determining the expression and localization properties of RNAs expressed during embryogenesis in the fruit fly, Drosophila melanogaster.

Assessing  the expression pattern of a gene, as well as the subcellular localization  properties of its transcribed RNA, are key features for ...

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 ...

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

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