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.

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	<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>KH2PO4</td>
			<td>Sigma</td>
			<td>P5655</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica SP8 equipped with a 63&#215;, 1.40 NA oil immersion objective</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4&#183;7H20</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>Na2HPO4</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>NaN3</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...

Watch this Scientific Journal Video about Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes at JoVE.com

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 ...

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1.9K Views.  Rutgers, The State University of New Jersey. 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.

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 ...

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 ...

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 ...

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

Live Imaging of Mouse Secondary Palate Fusion

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

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

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

A Neural Network-Based Identification of Developmentally Competent or Incompetent Mouse Fully-Grown Oocytes

Infertility clinics would benefit from the ability to select developmentally competent vs. incompetent oocytes using non-invasive procedures, thus improving the overall pregnancy outcome. We recently developed a classification method based on microscopic live observations of mouse oocytes during their in vitro maturation from the germinal vesicle (GV) to the metaphase II stage, followed by the analysis of the cytoplasmic movements occurring during this time-lapse period. Here, we present detailed protocols of this procedure. Oocytes are isolated from fully-grown antral follicles and cultured for 15 h inside a microscope equipped for time-lapse analysis at 37 &#176;C and 5% CO2. Pictures are taken at 8 min intervals. The images are analyzed using the Particle Image Velocimetry (PIV) method that calculates, for each oocyte, the profile of Cytoplasmic Movement Velocities (CMVs) occurring throughout the culture period. Finally, the CMVs of each single oocyte are fed through a mathematical classification tool (Feed-forward Artificial Neural Network, FANN), which predicts the probability of a gamete to be developmentally competent or incompetent with an accuracy of 91.03%. This protocol, set up for the mouse, could now be tested on oocytes of other species, including humans.

Here, we present a protocol for non-invasive assessment of oocyte developmental competence performed during their in vitro maturation from the germinal vesicle to the metaphase II stage. This method combines time-lapse imaging with particle image velocimetry (PIV) and neural network analyses.

Infertility clinics would benefit from the ability to select developmentally competent vs. incompetent oocytes using non-invasive procedures, thus ...

Live Cell Imaging of Chromosome Segregation During Mitosis

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

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

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

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

Determining Bile Duct Density in the Mouse Liver

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques are used, including serum chemistry, histological analysis, and immunostaining for specific markers. Although these techniques can provide important information about the biliary system, they often do not present a full picture of bile duct (BD) developmental defects across the whole liver. This is in part due to the robust ability of the mouse liver to drain the bile even in animals with significant impairment in biliary development. Here we present a simple method to calculate the average number of BDs associated with each portal vein (PV) in sections covering all lobes of mutant/transgenic mice. In this method, livers are mounted and sectioned in a stereotypic manner to facilitate comparison among various genotypes and experimental conditions. BDs are identified via light microscopy of cytokeratin-stained cholangiocytes, and then counted and divided by the total number of PVs present in liver section. As an example, we show how this method can clearly distinguish between wild-type mice and a mouse model of Alagille syndrome. The method presented here cannot substitute for techniques that visualize the three-dimensional structure of the biliary tree. However, it offers an easy and direct way to quantitatively assess BD development and the degree of ductular reaction formation in mice.

We present a rather simple and sensitive method for accurate quantification of bile duct density in the mouse liver. This method can aid in determining the effects of genetic and environmental modifiers and the effectiveness of potential therapies in mouse models of biliary diseases.

Mouse is broadly used as a model organism to study biliary diseases. To evaluate the development and function of the biliary system, various techniques ...

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous system and protects neural tissue from toxins and pathogens. Defects in the formation of this barrier have been correlated with neuropathies, and the breakdown of this barrier has been observed in many neurodegenerative diseases. Therefore, it is critical to identify the genes that regulate the formation and maintenance of the blood-brain barrier to identify potential therapeutic targets. In order to understand the exact roles these genes play in neural development, it is necessary to assay the effects of altered gene expression on the integrity of the blood-brain barrier. Many of the molecules that function in the establishment of the blood-brain barrier have been found to be conserved across eukaryotic species, including the fruit fly, Drosophila melanogaster. Fruit flies have proven to be an excellent model system for examining the molecular mechanisms regulating nervous system development and function. This protocol describes a step-by-step procedure to assay for blood-brain barrier integrity during the embryonic and larval stages of D. melanogaster development.

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster

Blood-brain barrier integrity is critical for nervous system function. In Drosophila melanogaster, the blood-brain barrier is formed by glial cells during late embryogenesis. This protocol describes methods to assay for blood-brain barrier formation and maintenance in D. melanogaster embryos and third instar larvae.

Proper nervous system development includes the formation of the blood-brain barrier, the diffusion barrier that tightly regulates access to the nervous ...

Evaluation of the Spindle Assembly Checkpoint Integrity in Mouse Oocytes

Summary

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Chapters in this video

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

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

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

Generation of Genetically Modified Mice through the Microinjection of Oocytes

Live Imaging of Mouse Secondary Palate Fusion

A Neural Network-Based Identification of Developmentally Competent or Incompetent Mouse Fully-Grown Oocytes

Live Cell Imaging of Chromosome Segregation During Mitosis

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Determining Bile Duct Density in the Mouse Liver

Assay for Blood-brain Barrier Integrity in Drosophila melanogaster