Name: evaluation of the spindle assembly checkpoint integrity in mouse oocytes
Uploaded: 2022-09-13T14:50:00.000+00:00
Duration: 10 min 9 s
Description: Watch this Scientific Journal Video about Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes at JoVE.com

Funding for this project was provided by the National Institutes of Health (R35GM136340 to KS).

All mice used in these protocols were housed and raised according to the Rutgers University Institutional Animal Use and Care Committee (Protocol 201702497) and National Institutes of Health guidelines. These regulatory bodies approved all experimental procedures involving animal studies. All mice used in the present study were 6-8-week-old CF-1 females.
1. Experimental preparation
<ol>
	<li>Before starting the SAC evaluations, collect mouse oocytes following the previously published report12. Divide collected oocytes into three evenly sized groups and keep them in Chatot, Ziomek, and Bavister (CZB) culture media containing 2.5 &#181;M milrinone (see Table of Materials) to avoid meiotic resumption13. 
	NOTE: CZB composition: 81.6 mM NaCL; 4.8 mM KCl; 1.2 mM KH2PO4; 1.2 mM MgSO4-7H2O; 0.27 mM Pyruvate; 1.7 mM CaCl2; 30.8 mM DL-Lactic Acid; 7 mM Taurine; 0.1 mM EDTA; 25 mM NaHCO3; Gentamicin; and 0.3% BSA.</li>
	<li>Prepare the cRNA for microinjection as described previously12,14. Amplify the mouse Securin gene from the cDNA cloned from mouse germinal vesicle oocytes, followed by subcloning into the pMDL2 vector containing Gfp sequence15,16.
	<ol>
		<li>To prepare Securin-Gfp cRNA, linearize the plasmid with Nde I digestion. Perform in vitro transcription by using T3 RNA polymerase and purifying the cRNA16. 
		&#8203;NOTE: Store cRNA in aliquots of 2-3 &#181;L at -80 &#176;C until use.</li>
	</ol>
	</li>
</ol>
2. Nocodazole treatment and live imaging
<ol>
	<li>Prepare 1 mL of culture media (CZB) containing 5 &#181;M nocodazole (NOC), 1 mL of culture media with 5 &#181;M of NOC plus 0.5 &#181;M reversine (NOC + REV), and 1 mL of culture media with Dimethyl sulfoxide (DMSO) (1:2000) as a control (see Table of Materials).</li>
	<li>For oocyte maturation and live imaging, use a 96-well plate pre-warmed in the incubator at 37 &#176;C, 5% CO2,&#160;and 80% humidity. In the first well, load 150 &#181;L of the control DMSO treatment; in the second well, load 150 &#181;L of NOC treatment; and in the third well, load 150 &#181;L of NOC + REV treatment. Keep the plate in the incubator under the same conditions as above until needed. 
	NOTE: In the 96-well plate, avoid using the first row and the first column because the shadow of the border interferes with the quality of the images.</li>
	<li>To start meiotic maturation, remove milrinone. While viewing the oocytes under a stereomicroscope using magnification between 30x-64x, wash milrinone out of the media by sequentially transferring the oocytes through six drops of 100 &#181;L of milrinone-free culture media containing DMSO and place them into the corresponding well of the 96-well plate using a hand- or mouth-operated pipette.
	<ol>
		<li>Count oocytes by picking them up using a hand- or mouth-operated pipette and delivering them to the next drop to avoid losing any. The oocytes are single, large (~80 &#181;m in diameter), and round cells. The nucleus will appear like a button in the center of the cell, and the membrane and zona pellucida surround the oocyte. 
		NOTE: Transfer the oocytes among drops while moving the least amount of liquid possible. This allows for optimal removal of the milrinone from the culture media. If oocytes fail to resume meiosis, it is likely that the milrinone was not effectively removed.</li>
	</ol>
	</li>
	<li>Repeat the same process (step 2.3) for NOC and NOC + REV treatment.</li>
	<li>Image the oocytes using a brightfield microscope equipped with an incubator chamber with a controlled environment under these conditions: 37 &#176;C, 5% CO2, and 80% humidity. Capture images at the middle plane of the oocytes at intervals of 20 min for 24 h.</li>
	<li>Quantify the number of oocytes that extrude a polar body (PB) by identifying the cells that go through asymmetrical cytokinesis. The result will be a small cell (PB) next to the egg and within the shared zona pellucida. View the images using imaging software (ImageJ, see Table of Materials).</li>
	<li>Import image sequence into the image analysis software and advance the frames until one or more cells go through asymmetrical cytokinesis. Calculate the percentage of oocytes that extrude a PB of the total of oocytes17.</li>
</ol>
3. Monitoring the pattern of Securin-gfp degradation during meiotic maturation
<ol>
	<li>Centrifuge 3 &#181;L of Securin-Gfp cRNA(step 1.2) at 19,283 x g for 30 min at 4 &#176;C18. 
	NOTE: Use the supernatant to avoid loading impurities that could cause needle blockage during microinjection.</li>
	<li>Follow step-by-step oocyte microinjection, extensively described in reference12. Microinject prophase I-arrested oocytes with 100 ng/&#181;L of Securin-gfp cRNA prepared in step 1.2.</li>
	<li>After microinjection, allow the oocytes to recover and translate the RNA in the CO2 incubator for at least 3 h. Check the expression by observing the oocytes in an automated multichannel fluorescence imaging system (see Table of Materials) to view the GFP signal.</li>
	<li>As described in step 2.2, load 150 &#181;L of culture media with or without 5 &#181;M of nocodazole and 150 &#181;L of culture media with 0.5 &#181;M of reversine in three different wells of a 96-well plate.</li>
	<li>Wash the microinjected oocytes through six drops of milrinone-free culture media and transfer 1/3 of the oocytes into each treatment.</li>
	<li>Keep the plate in the incubator at 37 &#176;C, with 5% CO2 and 80% humidity until the oocytes resume meiosis, around 3 h post milrinone washing. 
	NOTE: Use a stereomicroscope with magnification between 30x-64x to evaluate the nuclear envelope breakdown as a hallmark of meiotic resumption.</li>
	<li>Record images of oocyte maturation using a fluorescence microscope equipped with an incubator chamber as described in step 2.6. Use brightfield settings to find the oocytes in the well. Use a 488 filter to detect the Securin-gfp signal and adjust the fluorescence intensity to avoid over-exposing the cells.
	<ol>
		<li>If the imaging system is automated, save the positions of the oocytes in each well. Because Securin-gfp is evenly distributed into the cytoplasm, capture images in the middle plane of the oocytes at intervals of 20 min for 24 h. To capture groups of oocytes in one picture, use a lower magnification objective lens such as 10x.</li>
	</ol>
	</li>
	<li>Use image analysis software to quantify Securin-gfp destruction. Open the images of the GFP channel. Start at time point 1. In the preferred analysis software, use a selection or shape tool to mark each oocyte and generate a region of interest (ROI) for each cell.
	<ol>
		<li>Use the same ROI to select a background area to be subtracted later. Measure GFP pixel intensity in each ROI.</li>
	</ol>
	</li>
	<li>Using the ROIs for each oocyte generated in step 3.8, measure the GFP intensity in all the time points. 
	NOTE: In some software programs, an &#34;Analyze&#34; tab allows the selection of the Measure function.
	<ol>
		<li>Then, advance to the next time frame and repeat this process to measure the intensity of GFP in each time frame. Lastly, to each GFP value, subtract the measurement of the ROI in the background selected in step 3.8. This analysis will give a value for Securin-gfp intensity for each oocyte at each time point.</li>
	</ol>
	</li>
	<li>Extract different parameters from the pattern of Securin destruction: (a) the time that Securin-gfp degradation began. This tells when SAC silencing started; (b) the time of Securin-gfp minimum signal. This tells when SAC silencing has dropped below a threshold level to allow high APC/C activation and full Securin-GFP degradation; (c) the rate of Securin-gfp destruction19. This tells how rapidly SAC activity drops below a threshold level for APC/C activation and Securin-GFP degradation.</li>
</ol>
4. Recruitment of MAD2 at kinetochores by immunofluorescence during meiotic maturation
NOTE: For oocyte collection and maturation, refer to the previously published report12.
<ol>
	<li>Split oocytes into three groups. Transfer each group of oocytes into 100 &#181;L drops of milrinone-free culture media. Cover the drops with mineral oil and place them in the incubator (37 &#176;C, 5% CO2, and 80% humidity) for 3 h, 5 h, and 7 h to reach early prometaphase I, late prometaphase I, and metaphase I stage, respectively. 
	NOTE: Place each time point in a different dish to avoid taking out the same dish from the incubator several times, which affects the regular timing of meiotic maturation.</li>
	<li>At the assigned time points, fix the oocytes by transferring them into 500 &#181;L drops of 2% PFA in 1x PHEM buffer for 20 min at room temperature, using a 9-well glass dish as previously described20. Then, transfer the cells to a clean well containing a 500 &#181;L drop of blocking solution (PBS + 0.3% BSA + 0.01% Tween-20 + 0.02% NaN3). 
	NOTE: PHEM composition: 60 mM PIPES; 25 mM HEPES; 10 mM EGTA; and 2 mM MgCl2. 
	NOTE: One can stop at this point and store the oocytes in blocking solution in the 9-well plate at 4 &#176;C until the convenient time to complete immunofluorescence.</li>
	<li>To continue MAD2 detection, transfer the oocytes to a clean well containing a 500 &#181;L drop of permeabilization solution (PBS + 0.3% BSA + 0.1% TritonX-100 + 0.02% NaN3). Incubate for 20 min at room temperature, and then transfer the cells to a new well of blocking solution and incubate for 10 min.</li>
	<li>For the rest of the immunofluorescence steps, use a 96-well dish lid with indentations as previously described20. Use a humidified dark chamber to avoid light exposure and evaporation. Transfer the oocytes into a 30 &#181;L drop of blocking solution with anti-MAD2 antibody (1:1000, rabbit) and anti-centromeric antibody (ACA) (1:30, human) (see Table of Materials) and incubate for 2 h at room temperature. 
	NOTE: The 96-well dish lid allows room for the processing of several proteins and groups at the same time.</li>
	<li>To wash excess primary antibodies, transfer the cells to a 30 &#181;L drop of 1x PHEM buffer supplemented with 0.5% Triton and incubate for 10 min in the humidified chamber. Repeat this step two more times.</li>
	<li>Perform the fourth wash, transferring the cells to a 30 &#181;L drop of 1x PHEM buffer without 0.5% 1x Triton, and incubate for 10 min.</li>
	<li>Transfer the cells to a 30 &#181;L drop of blocking solution containing secondary antibodies such as anti-human-633 (1:200) and anti-rabbit-568 (1:200) (see Table of Materials) and incubate for 1 h at room temperature. 
	NOTE: Choose the combination of fluorophores based on your microscope lasers or filters.</li>
	<li>To wash excess secondary antibodies, repeat steps 4.5-4.6.</li>
	<li>To mount the cells on the microscope slide, transfer the cells to a 10 &#181;L drop of mounting media containing DAPI (0.1 mg/mL) (see Table of Materials). Add small dots of petroleum jelly to each corner of the coverslip, carefully place them on top of the mounting media drop, and press slowly to distribute. Use clear nail polish to seal the coverslip to the slide. For a more detailed description of the mounting process, see reference20.</li>
	<li>Image kinetochores using a confocal microscope (see Table of Materials) equipped with a 40x or 63x objective. Using the ACA signal, determine the z-range that allows imaging of the entire chromosome area. 
	NOTE: An optical zoom of 4.0 and z-step size of 0.5 &#181;m can be used on some imaging systems. These parameters may vary from system to system, and optimization will be necessary.</li>
	<li>Analyze MAD2 intensity at kinetochores using image analysis software. Make a z-stack maximum projection and then split the channels.</li>
	<li>First, create a mask using the ACA channel by selecting the ACA channel to establish a threshold that identifies all the kinetochore signals. Go to the edit tab and create a selection. Then, create an ROI with this selection.</li>
	<li>Select the MAD2 channel and bring in the selection created in step 4.12. Select a threshold method that better adapts to the MAD2 signal. Measure the intensity. 
	NOTE: Select the threshold method in the control treatment and keep it constant when analyzing different treatments.</li>
	<li>To make relative pixel intensity calculations, divide the intensity of each cell by the average intensity of the WT oocytes in the experiment.</li>
</ol>

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The authors have no conflicts to disclose.

The spindle assembly checkpoint is a critical control mechanism during cell division designed to prevent chromosome segregation errors. It allows the cell to have enough time to correct improper KT-MT attachments. Meiosis in oocytes is an error-prone process, where most of the chromosome mis-segregation occurs during meiosis I, leading to the generation of aneuploid eggs that are the main cause of early miscarriages and infertility in humans1,24. Several hypothes...

Aneuploidy, which arises from errors in chromosome segregation, is the leading cause of early miscarriages and is highly linked to mistakes in meiosis1. Meiosis is distinct from mitosis because it consists of two rounds of cell division without an intervening DNA replication step. In meiosis I, homologous chromosomes separate while sister chromatids remain together. In oocytes, this step is error-prone, leading to aneuploid egg production2.
To prevent chromosome segregations errors, most cell types activate a surveillance mechanism that pauses the cell cycle, called the spindle assembly checkpoint (SAC). This mechanism senses kinetochore (KT)-microtubule (MT) attachments and the tension is generated when chromosomes are oriented in a bipolar manner3. Unattached kinetochores trigger a SAC response which starts with the recruitment of MPS1, the master regulator of the SAC, to kinetochores3,4. MPS1 initiates the recruitment of other SAC components, acting as a platform to form the mitotic checkpoint complex (MCC). The MCC, composed of MAD1, MAD2, BUB3, and BUBR1, diffuses into the cytoplasm and inhibits APC/C activation by sequestrating its activator CDC20. Once all kinetochores are stably attached to MTs and chromosomes are aligned at the metaphase plate, the SAC is silenced, and the MCC disassembles and releases CDC20, thereby allowing APC/C activation. Active APC/C degrades Securin and Cyclin B, two key steps in triggering anaphase onset5,6. In somatic cells, the SAC is stringent because it is activated by a single unattached kinetochore and is sufficient to induce cell-cycle arrest6. However, during oocyte meiosis, the SAC is more permissive, and oocytes can enter anaphase I with one or more unattached kinetochores6,7,8,9,10. Understanding why the SAC is more permissive in oocytes is an ongoing area of focus in the field. Mechanisms that cause defects in SAC activation or SAC silencing could lead to errors in chromosome segregation or prolonged cell cycle arrest and cell death. Therefore, evaluating the mechanisms that maintain SAC integrity in oocytes is important to understanding the process of forming healthy, euploid eggs.
This protocol describes techniques to comprehensively evaluate the SAC integrity in mouse oocyte meiosis by examining different critical steps of the checkpoint. First, the evaluation of the SAC response after inducing SAC activation is described. This activation is achieved by generating unattached kinetochores using nocodazole, a drug that depolymerizes MTs11. Second, a method to monitor SAC silencing is described by tracking the dynamics of Securin degradation during oocyte maturation. Finally, an immunofluorescence-based assay is employed to measure the recruitment of MAD2, one of the MCC components, to kinetochores. Together, these assays comprehensively assess SAC integrity during oocyte meiotic maturation.

<table><tbody><tr><td>Bovine serum albumin (BSA)</td><td>Sigma</td><td>A4503</td><td></td></tr><tr><td>DAPI</td><td>Life Technologies</td><td>D1306</td><td></td></tr><tr><td>Dimethyl-sulfoxide (DMSO)</td><td>Sigma</td><td>D5879</td><td></td></tr><tr><td>Donkey-anti-rabbit-Alexa-568</td><td>Life Technologies</td><td>A10042</td><td></td></tr><tr><td>EVOS FL Auto Imaging System</td><td>Life Technologies</td><td></td><td>Fluorescence microscope</td></tr><tr><td>EVOS Onstage Incubator</td><td>Life Technologies</td><td></td><td>Incubator chamber</td></tr><tr><td>Glass Bottom 96- well plates N 1.5 uncoated</td><td>MatTek Corporation</td><td>P96G-1.5-5-F</td><td></td></tr><tr><td>goat-anti-human-Alexa-633</td><td>Life Technologies</td><td>A21091</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H3537</td><td></td></tr><tr><td>Human anti-ACA</td><td>Antibodies Incorporated</td><td>15-234</td><td>Dilution 1/30</td></tr><tr><td>ImageJ</td><td>NIH</td><td></td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P5405</td><td></td></tr><tr><td>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>P5655</td><td></td></tr><tr><td>Leica SP8 equipped with a 63&amp;times;, 1.40 NA oil immersion objective</td><td>Leica</td><td></td><td></td></tr><tr><td>MgSO4&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;0</td><td>Sigma</td><td>M7774</td><td></td></tr><tr><td>Milrinone</td><td>Sigma</td><td>M4659</td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>S2429</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S5886</td><td></td></tr><tr><td>NaN&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S2002</td><td></td></tr><tr><td>Nocodazole</td><td>Sigma</td><td>M1404</td><td></td></tr><tr><td>Paraformaldhyde (PFA)</td><td>Sigma</td><td>P6148</td><td></td></tr><tr><td>PIPES</td><td>Sigma</td><td>P6757</td><td></td></tr><tr><td>Rabbit anti- MAD2</td><td>Biolegend</td><td>924601</td><td>Dilution 1/1000; previously Covance #PRB-452C</td></tr><tr><td>Reversine</td><td>Cayman Chemical</td><td>10004412</td><td></td></tr><tr><td>Triton-X</td><td>Sigma</td><td>274348</td><td></td></tr><tr><td>Tween-20</td><td>Sigma</td><td>X100</td><td></td></tr><tr><td>Vectashield</td><td>Vector laboratories</td><td>H-1000</td><td></td></tr></tbody></table>

evaluation of the spindle assembly checkpoint integrity in mouse oocytes

Aneuploidy is the leading genetic abnormality causing early miscarriage and pregnancy failure in humans. Most errors in chromosome segregation that give rise to aneuploidy occur during meiosis in oocytes, but why oocyte meiosis is error-prone is still not fully understood. During cell division, cells prevent errors in chromosome segregation by activating the spindle assembly checkpoint (SAC). This control mechanism relies on detecting kinetochore (KT)-microtubule (MT) attachments and sensing tension generated by spindle fibers. When KTs are unattached, the SAC is activated and prevents cell-cycle progression. The SAC is activated first by MPS1 kinase, which triggers the recruitment and formation of the mitotic checkpoint complex (MCC), composed of MAD1, MAD2, BUB3, and BUBR1. Then, the MCC diffuses into the cytoplasm and sequesters CDC20, an anaphase-promoting complex/cyclosome (APC/C) activator. Once KTs become attached to microtubules and chromosomes are aligned at the metaphase plate, the SAC is silenced, CDC20 is released, and the APC/C is activated, triggering the degradation of Cyclin B and Securin, thereby allowing anaphase onset. Compared to somatic cells, the SAC in oocytes is not as effective because cells can undergo anaphase despite having unattached KTs. Understanding why the SAC is more permissive and if this permissiveness is one of the causes of chromosome segregation errors in oocytes still needs further investigation. The present protocol describes the three techniques to comprehensively evaluate SAC integrity in mouse oocytes. These techniques include using nocodazole to depolymerize MTs to evaluate the SAC response, tracking SAC silencing by following the kinetics of Securin destruction, and evaluating the recruitment of MAD2 to KTs by immunofluorescence. Together these techniques probe mechanisms needed to produce healthy eggs by providing a complete evaluation of SAC integrity.

Evaluation of SAC responsiveness by nocodazole treatment 
The purpose of this experiment is to evaluate SAC activation and strength. By using nocodazole to depolymerize spindle microtubules, all kinetochores will be unattached, which will cause a SAC-mediated cell-cycle arrest. In the present imaging system, DMSO-treated control oocytes extruded a polar body around 14 h after release from milrinone (Figure 1, top panels). Consistent with SAC activation, nocodazole-treat...

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Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes

Error in chromosome segregation is a common feature in oocytes. Therefore, studying the spindle assembly checkpoint gives important clues about the mechanisms needed to produce healthy eggs. The present protocol describes three complementary assays to evaluate spindle assembly checkpoint integrity in mouse oocytes.

Aneuploidy is the leading genetic abnormality causing early miscarriage and pregnancy failure in humans. Most errors in chromosome segregation that give ...

evaluation-spindle-assembly-checkpoint-integrity-mouse

Research

JoVE Journal

Developmental Biology

1.9K Views.  Rutgers, The State University of New Jersey. Aneuploïdie is de belangrijkste genetische oorzaak van een vroege miskraam bij de mens. De meeste fouten in chromosoomsegregatie treden op tijdens meiose in eicellen. Daarom is het evalueren van het controlepunt voor de spindelassemblage in eicellen van cruciaal belang om te begrijpen waarom eicellen foutgevoelig zijn.Hier beschrijven we drie technieken om de SAC-integriteit in muizenoöcyten uitgebreid te evalueren door verschillende kritieke stappen op het checkpoint te onderzoeken ...

Video: Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Chromatin Spread Preparations for the Analysis of Mouse Oocyte Progression from Prophase to Metaphase II

Chromatin spread techniques have been widely used to assess the dynamic localization of various proteins during gametogenesis, particularly for spermatogenesis. These techniques allow for visualization of protein and DNA localization patterns during meiotic events such as homologous chromosome pairing, synapsis and DNA repair. While a few protocols have been described in the literature, general chromatin spread techniques using mammalian prophase oocytes are limited and difficult due to the timing of meiosis initiation in fetal ovaries. In comparison, prophase spermatocytes can be collected from juvenile male mice with higher yields without the need for microdissection. However, it is difficult to obtain a pure synchronized population of cells at specific stages due to the heterogeneity of meiotic and post-meiotic germ cell populations in the juvenile and adult testis. For later stages of meiosis, it is advantageous to assess oocytes undergoing meiosis I (MI) or meiosis II (MII), because groups of mature oocytes can be collected from adult female mice and stimulated to resume meiosis in culture. Here, methods for meiotic chromatin spread preparations using oocytes dissected from fetal, neonatal and adult ovaries are described with accompanying video demonstrations. Chromosome missegregation events in mammalian oocytes are frequent, particularly during MI. These techniques can be used to assess and characterize the effects of different mutations or environmental exposures during various stages of oogenesis. As there are distinct differences between oogenesis and spermatogenesis, the techniques described within are invaluable to increase our understanding of mammalian oogenesis and the sexually dimorphic features of chromosome and protein dynamics during meiosis.

Oogenesis in mammals is known to be error-prone, particularly due to chromosome missegregation. This manuscript describes chromatin spread preparation methods for mouse prophase, metaphase I and II-staged oocytes. These fundamental techniques allow for the study of chromatin-bound proteins and chromosome morphology throughout mammalian oogenesis.

Chromatin spread techniques have been widely used to assess the dynamic localization of various proteins during gametogenesis, particularly for ...

Genetics

Isolation and Characterization of Mouse Antral Oocytes Based on Nucleolar Chromatin Organization

This protocol describes a simple and quick method to isolate and characterize mouse antral GV (Germinal Vesicle) oocytes as able (SN, Surrounded Nucleolus) or unable (NSN, Not Surrounded Nucleolus) to develop to the blastocyst stage after in vitro maturation (IVM) and in vitro fertilization (IVF). It makes use of Hoeschst33342 (or any other DNA intercalating dye) able to bind to the heterochromatin of the nucleolus showing a ring in the SN oocytes or not, like in the NSN oocytes. This represents the easiest and quickest way to sort both antral oocytes that can be eventually used for IVM or IVF procedures. 
Briefly, the protocol consists of the following steps: hormone injection to stimulate follicular growth; isolation of the oocytes at the GV stage from the antral compartment by puncturing the ovary with a sterile needle; preparation of thin glass pipettes for mouth pipetting of the oocytes; sorting of the oocytes with Hoechst33342 prepared at a supravital concentration; IVM, IVF or any other molecular/cellular analysis.
Unfortunately there are still few evidences to sort SN and NSN oocytes using less invasive techniques. If and once they will be identified, they could be potentially applied to human assisted reproductive technologies, although with several aspects that should be modified. To date, this technique has potential implications to dramatically increase IVM and IVF successful procedures in both endangered and species with economic interest.

Here we present a protocol for the characterization of mouse antral oocytes based on nucleolar organization.

This protocol describes a simple and quick method to isolate and characterize mouse antral GV (Germinal Vesicle) oocytes as able (SN, Surrounded ...

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 ...

Analysis of Chromosome Segregation, Histone Acetylation, and Spindle Morphology in Horse Oocytes

The field of assisted reproduction has been developed to treat infertility in women, companion animals, and endangered species. In the horse, assisted reproduction also allows for the production of embryos from high performers without interrupting their sports career and contributes to an increase in the number of foals from mares of high genetic value. The present manuscript describes the procedures used for collecting immature and mature oocytes from horse ovaries using ovum pick-up (OPU). These oocytes were then used to investigate the incidence of aneuploidy by adapting a protocol previously developed in mice. Specifically, the chromosomes and the centromeres of metaphase II (MII) oocytes were fluorescently labeled and counted on sequential focal plans after confocal laser microscope scanning. This analysis revealed a higher incidence in the aneuploidy rate when immature oocytes were collected from the follicles and matured in vitro compared to in vivo. Immunostaining for tubulin and the acetylated form of histone four at specific lysine residues also revealed differences in the morphology of the meiotic spindle and in the global pattern of histone acetylation. Finally, the expression of mRNAs coding for histone deacetylases (HDACs) and acetyl-transferases (HATs) was investigated by reverse transcription and quantitative-PCR (q-PCR). No differences in the relative expression of transcripts were observed between in vitro and in vivo matured oocytes. In agreement with a general silencing of the transcriptional activity during oocyte maturation, the analysis of the total transcript amount can only reveal mRNA stability or degradation. Therefore, these findings indicate that other translational and post-translational regulations might be affected.
Overall, the present study describes an experimental approach to morphologically and biochemically characterize the horse oocyte, a cell type that is extremely challenging to study due to low sample availability. However, it can expand our knowledge on the reproductive biology and infertility in monovulatory species.

This manuscript describes an experimental approach to morphologically and biochemically characterize horse oocytes. Specifically, the present work illustrates how to collect immature and mature horse oocytes by ultrasound-guided ovum pick-up (OPU) and how to investigate chromosome segregation, spindle morphology, global histone acetylation, and mRNA expression.

The field of assisted reproduction has been developed to treat infertility in women, companion animals, and endangered species. In the horse, assisted ...

Human Egg Maturity Assessment and Its Clinical Application

The optimal timing of intracytoplasmic sperm injection (ICSI) is of a serious concern for fertility programs because untimely sperm entry diminishes the egg&#39;s developmental competence. Presence of the first polar body (PB) together with the meiotic spindle indicates completion of the oocyte maturation and the egg&#39;s readiness for fertilization. In clinical practice, it is customary to assume that all oocytes displaying a PB are mature metaphase (MII) oocytes. However, PB extrusion precedes the formation of the bipolar MII spindle. This asynchrony makes the mere presence of PB an unreliable marker of oocyte maturity. Noninvasive spindle imaging using polarized light microscopy (PLM) allows quick and easy inspection of whether the PB-displaying oocyte actually reassembled a meiotic spindle prior to ICSI. Here, we present a standard protocol to perform human egg maturity assessment in the clinical laboratory. We also show how to optimize the time of ICSI with respect to the oocyte&#39;s developmental stage in order to prevent premature sperm injection of late-maturing oocytes. Using this approach, even immature oocytes extruding PB in vitro can be clinically utilized. Affirmation that MII spindle is present prior to sperm injection and individual adjustment of the time of ICSI is particularly important in poor prognosis in-vitro fertilization (IVF) cycles with a low number of oocytes available for fertilization.

We provide an outline of the clinical protocol for non-invasive assessment of human egg maturity using polarized light microscopy.

The optimal timing of intracytoplasmic sperm injection (ICSI) is of a serious concern for fertility programs because untimely sperm entry diminishes the ...

Multi-Photon Laser Ablation of Cytoplasmic Microtubule Organizing Centers in Mouse Oocytes

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at prophase I of the first meiotic division. After puberty and upon meiotic resumption, the nuclear membrane disassembles (nuclear envelope breakdown), and the spindle is assembled mainly at the oocyte center. Initial central spindle positioning is essential to protect against abnormal kinetochore-microtubule (MT) attachments and aneuploidy. The centrally positioned spindle migrates in a time-sensitive manner toward the cortex, and this is a necessary process to extrude a tiny polar body. In mitotic cells, spindle positioning relies on the interaction between centrosome-mediated astral MTs and the cell cortex. On the contrary, mouse oocytes lack classic centrosomes and, instead, contain numerous acentriolar MT organizing centers (MTOCs). At the metaphase I stage, mouse oocytes have two different sets of MTOCs: (1) MTOCs that are clustered and sorted to assemble spindle poles (polar MTOCs), and (2) metaphase cytoplasmic MTOCs (mcMTOCs) that remain in the cytoplasm and do not contribute directly to spindle formation but play a crucial role in regulating spindle positioning and timely spindle migration. Here, a multi-photon laser ablation method is described to selectively deplete endogenously labeled mcMTOCs in oocytes collected from Cep192-eGfp reporter mice. This method contributes to the understanding of the molecular mechanisms underlying spindle positioning and migration in mammalian oocytes.

An optimized protocol is presented that enables the depletion of cytoplasmic microtubule organizing centers in mouse oocytes during metaphase I using a near-infrared femtosecond laser.

The fidelity of oocyte meiosis is critical for generating developmentally competent euploid eggs. In mammals, the oocyte undergoes a lengthy arrest at ...

Identifying DNA Damage in Prophase-Arrested Oocytes of Mice

Detection of DNA Double-Stranded Breaks in Mouse Oocytes

Oocytes are amongst the biggest and most long-lived cells in the female body. They are formed in the ovaries during embryonic development and remain arrested at the prophase of meiosis I. The quiescent state may last for years until the oocytes receive a stimulus to grow and obtain the competency to resume meiosis. This protracted state of arrest makes them extremely susceptible to accumulating DNA-damaging insults, which affect the genetic integrity of the female gametes and, therefore, the genetic integrity of the future embryo. 
Consequently, the development of an accurate method to detect DNA damage, which is the first step for the establishment of DNA damage response mechanisms, is of vital importance. This paper describes a common protocol to test the presence and progress of DNA damage in prophase-arrested oocytes during a period of 20 h. Specifically, we dissect mouse ovaries, retrieve the cumulus-oocyte complexes (COCs), remove the cumulus cells from the COCs, and culture the oocytes in &#924;2 medium containing 3-isobutyl-1-methylxanthine to maintain the state of arrest. Thereafter, the oocytes are treated with the cytotoxic, antineoplasmic drug, etoposide, to engender double-strand breaks (DSBs). 
By using immunofluorescence and confocal microscopy, we detect and quantify the levels of the core protein &#947;H2AX, which is the phosphorylated form of the histone H2AX. H2AX becomes phosphorylated at the sites of DSBs after DNA damage. The inability to restore DNA integrity following DNA damage in oocytes can lead to infertility, birth defects, and increased rates of spontaneous abortions. Therefore, the understanding of DNA damage response mechanisms and, at the same time, the establishment of an intact method for studying these mechanisms are essential for reproductive biology research.

Author Spotlight: Understanding DNA Damage Response in Mammalian Oocytes and Preimplantation Embryos 

Maintaining oocyte genome integrity is necessary to ensure the genetic fidelity in the resulting embryo. Here, we present an accurate protocol for detecting DNA double-strand breaks in mammalian female germ cells.

Oocytes are amongst the biggest and most long-lived cells in the female body. They are formed in the ovaries during embryonic development and remain ...

Mouse Oocyte Microinjection, Maturation and Ploidy Assessment

Mistakes in chromosome segregation lead to aneuploid cells. In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to infertility, miscarriages or developmental disorders like Down syndrome. Haploid gametes form through species-specific developmental programs that are coupled to meiosis. The first meiotic division (MI) is unique to meiosis because sister chromatids remain attached while homologous chromosomes are segregated. For reasons not fully understood, this reductional division is prone to errors and is more commonly the source of aneuploidy than errors in meiosis II (MII) or than errors in male meiosis 1,2. 
In mammals, oocytes arrest at prophase of MI with a large, intact germinal vesicle (GV; nucleus) and only resume meiosis when they receive ovulatory cues. Once meiosis resumes, oocytes complete MI and undergo an asymmetric cell division, arresting again at metaphase of MII. Eggs will not complete MII until they are fertilized by sperm. Oocytes also can undergo meiotic maturation using established in vitro culture conditions 3. Because generation of transgenic and gene-targeted mouse mutants is costly and can take long periods of time, manipulation of female gametes in vitro is a more economical and time-saving strategy. 
Here, we describe methods to isolate prophase-arrested oocytes from mice and for microinjection. Any material of choice may be introduced into the oocyte, but because meiotically-competent oocytes are transcriptionally silent 4,5 cRNA, and not DNA, must be injected for ectopic expression studies. To assess ploidy, we describe our conditions for in vitro maturation of oocytes to MII eggs. Historically, chromosome-spreading techniques are used for counting chromosome number 6. This method is technically challenging and is limited to only identifying hyperploidies. Here, we describe a method to determine hypo-and hyperploidies using intact eggs 7-8. This method uses monastrol, a kinesin-5 inhibitor, that collapses the bipolar spindle into a monopolar spindle 9 thus separating chromosomes such that individual kinetochores can readily be detected and counted by using an anti-CREST autoimmune serum. Because this method is performed in intact eggs, chromosomes are not lost due to operator error.

Oocytes are prone to aneuploidy due to errors in chromosome segregation during meiotic maturation. Aneuploid eggs can cause infertility, miscarriages or developmental disorders like Down syndrome. Here, we describe methods to introduce materials of choice into oocytes and methods to study meiotic maturation and assess ploidy.

Mistakes in chromosome segregation lead to aneuploid cells.  In somatic cells, aneuploidy is associated with cancer but in gametes, aneuploidy leads to ...

Biology

Meiotic Spindle Assessment in Mouse Oocytes by siRNA-mediated Silencing

Errors in chromosome segregation during meiotic division in gametes can lead to aneuploidy that is subsequently transmitted to the embryo upon fertilization. The resulting aneuploidy in developing embryos is recognized as a major cause of pregnancy loss and congenital birth defects such as Down&#8217;s syndrome. Accurate chromosome segregation is critically dependent on the formation of the microtubule spindle apparatus, yet this process remains poorly understood in mammalian oocytes. Intriguingly, meiotic spindle assembly differs from mitosis and is regulated, at least in part, by unique microtubule organizing centers (MTOCs). Assessment of MTOC-associated proteins can provide valuable insight into the regulatory mechanisms that govern meiotic spindle formation and organization. Here, we describe methods to isolate mouse oocytes and deplete MTOC-associated proteins using a siRNA-mediated approach to test function. In addition, we describe oocyte fixation and immunofluorescence analysis conditions to evaluate meiotic spindle formation and organization.

Here, we present a protocol for specific siRNA-mediated mRNA depletion followed by immunofluorescence analysis to evaluate meiotic spindle assembly and organization in mouse oocytes. This protocol is suitable for in vitro depletion of transcripts and functional assessment of different spindle and/or MTOC-associated factors in oocytes.

Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes

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