The authors would like to thank all members of the Balboula laboratory for their valuable help and discussions. The authors thank Melina Schuh for kindly sharing&#160;the mCherry-Cep192 construct. This study was supported by R35GM142537 (NIGMS, NIH) to AZB.

All the methods described here were approved by the University of Missouri (Animal Care Quality Assurance Ref. Number 9695). Cep192-eGFP reporter female mice aged 6-8 weeks old were used in the present study. To generate Cep192-eGFP reporter mice, CRISPR/Cas9-mediated homology-directed repair was used to integrate the EGFP reporter gene into the CF-1 mouse genome. The EGFP reporter was fused at the C-terminus of Cep192 (an integral component of MTOCs)15. To maintain the mouse colony, homozygous Cep192-eGfp reporter mice were used. All the animals were maintained in cages (up to four animals/cage) at 21 &#176;C and 55% humidity, with a 12 h light/dark cycle and ad libitum access to food and water.
1. Mouse oocyte collection
<ol>
	<li>Prepare the culture medium (Chatot, Ziomek, and Bavister, CZB19, see the Table of Materials), and incubate it at 37 &#176;C and 5% CO2 overnight. 
	NOTE: The CZB medium can be stored at 4 &#176;C for 1 month (see Supplementary File 1 for the media composition).</li>
	<li>Supplement the CZB medium with glutamine (1 mM) and milrinone (2.5 mM) (CZB + M) (see Table of Materials), and place it in the incubator. 
	&#8203;NOTE: Milrinone is a phosphodiesterase inhibitor that maintains the oocytes arrested at prophase I and prevents meiotic resumption20.</li>
	<li>Prepare the collection medium (bicarbonate-free minimal essential medium, MEM) containing 3 mg/mL polyvinylpyrolidone (PVP), 25 mM HEPES (pH 7.3) (Supplementary File 1), and milrinone (2.5 mM) (MEM/PVP + M, see the Table of Materials).</li>
	<li>Prepare the collection and culture dishes, make four microdrops (100 &#181;L) of collection medium (MEM/PVP + M) and two microdrops (100 &#181;L) of culture (CZB + M) medium in 60 mm and 35 mm Petri dishes, respectively, and cover them with mineral oil (see Table of Materials). Keep the collection dish on the slide warmer, and put the culture dish into the incubator at 37 &#176;C and 5% CO2.</li>
	<li>Inject intraperitoneally 5 IU of pregnant mare&#39;s serum gonadotropin (PMSG, see the Table of Materials) into sexually mature (6-8 week-old) Cep192-eGFP reporter female mice 44-48 h before oocyte collection.</li>
	<li>Sacrifice the mice by cervical dislocation, identify and remove the ovaries21, and transfer them into a watch glass containing prewarmed collection medium (MEM/PVP + M) at 37 &#176;C.</li>
	<li>Fix the ovary by touching a 1 mL syringe to the bottom of the watch glass and puncturing it several times (~40 times per ovary) using sewing needles bundled together to release the oocytes into the medium.</li>
	<li>Using a plastic transfer pipet, transfer all the medium containing the cumulus-oocyte complexes (COCs) from step 1.7 into an empty 100 mm plastic Petri dish.</li>
	<li>Under&#160;a stereomicroscope, collect the COCs using a Pasteur glass pipet, and transfer them to the collection dish that contains MEM/PVP + M.</li>
	<li>Using a narrow Pasteur glass pipet (approximately 100 &#181;m in diameter), denude the oocytes mechanically by gentle repetitive pipetting, followed by transferring and washing them in four microdrops of (100 &#181;L) MEM/PVP + M (the collection dish) prior to their transfer into the CZB + M culture dish.</li>
	<li>Incubate the denuded oocytes for 1 h at 37 &#176;C with 5% of CO2 in the air.</li>
</ol>
2. Oocyte microinjection
<ol>
	<li>Place a 250 &#181;L drop of MEM/PVP + M into a 100 mm plastic Petri dish lid, and cover it with mineral oil. 
	NOTE: The Petri dish lid has a lower rim that provides more space for adjusting the micromanipulator.</li>
	<li>Turn on the microinjection system (see the Table of Materials).</li>
	<li>Place the microinjection dish on the warming stage (37 &#176;C) of the microscope.</li>
	<li>Load the injection needle with 0.5 &#181;L of mCh-Cep192cRNA using microloader pipette tips (see Table of Materials), and secure the injection needle to the micromanipulator.</li>
	<li>Transfer the denuded oocytes (step 1.11) to the 250 &#181;L microdrop (step 2.1).</li>
	<li>Under a 20x or 40x objective lens, adjust the position and the focus of the injection and holding needles according to the oocyte position (Figure 1).</li>
	<li>Set up the microinjection unit, and set the injection pressure (pi), compensation pressure (pc), and injection time (ti) to be able to inject 5-10 pl of mCh-Cep192 cRNA (to exogenously label the MTOCs).</li>
	<li>Carefully inject the oocytes without touching the nucleus.</li>
	<li>Once all the oocytes are injected, wash them in three microdrops of CZB + M, transfer them to the culture dish (CZB + M), and incubate them for 3 h at 37 &#176;C to allow mCh-Cep192 expression.</li>
</ol>
3. Oocyte maturation
<ol>
	<li>Prepare the maturation medium by adding glutamine (1 mM) to CZB medium pre-equilibrated in an incubator with 5% CO2 in air at 37 &#176;C for at least 3 h.</li>
	<li>Make two microdrops (100 &#181;L) of the maturation medium in a 35 mm Petri dish, and cover them with mineral oil (maturation dish).</li>
	<li>Wash prophase I-arrested oocytes at least three times in 100 &#181;L milrinone-free CZB microdrops to completely remove the milrinone and allow meiotic resumption.</li>
	<li>Transfer the oocytes to the maturation dish, and incubate them for 5 h (prometaphase I stage) in a humidified incubator with 5% CO2 in air at 37 &#176;C.</li>
</ol>
4. Oocyte preparation for ablation and imaging
<ol>
	<li>Transfer the prometaphase I oocytes (step 3.4) to a glass-bottom culture dish containing 100 &#181;L of maturation medium (step 3.1) covered with mineral oil.</li>
</ol>
5. Microscope preparation for ablation
<ol>
	<li>At least 30 min before ablation, turn on the temperature controller for the stage incubator (see Table of Materials) and set it at 37 &#176;C.</li>
	<li>Turn on the CO2 controller and set it at 5% CO2.</li>
	<li>Select a 40x oil immersion apochromatic objective, and apply a small drop of the immersion oil.</li>
	<li>Mount the glass-bottom culture dish with the oocytes on a stage-top incubator, and cover the incubator with a gas lid.</li>
	<li>In the image acquisition software (see the Table of Materials), select an option enabling the saving of multiple stage positions (e.g., &#34;Define mark and find experiment&#34;). Using transmitted light brightfield illumination, center on the individual oocytes and save their positions.</li>
	<li>In the image acquisition software, select the XYZ scanning mode, set the image format to 256 x 256 pixels, set the zoom factor to 2.5x - 3.0x, and select a scanning frequency of 600 Hz. This corresponds to a pixel dwell time of approximately 1.6 &#181;s.</li>
	<li>Set a 488 nm excitation laser line at approximately 10% of the laser power (which corresponds to 4.0 &#181;W at the sample level) (see Table of Materials), and use a spectral bandwidth of 500-550 nm to observe the GFP signal from the MTOCs. For the simultaneous observation of fluorescence from the mCherry-tagged Cep192, set a 585 nm excitation laser line at approximately 8% of the laser power (which corresponds to 10.7 &#181;W at the sample level), and use another detector set to a spectral bandwidth of 595-645 nm. 
	NOTE: It is helpful to use the confocal microscope&#39;s transmitted light detector (TLD) for concurrently detecting oocyte boundaries.</li>
	<li>Using a live scanning mode and the manual control of the z-drive, screen the oocyte for MTOCs.</li>
	<li>Once an mcMTOC is detected, draw a square region of interest (ROI) around it.</li>
</ol>
6. Ablation of mcMTOCs
<ol>
	<li>Set a femtosecond laser to a 740 nm wavelength. 
	NOTE: The laser wavelength can be changed according to the microscope and the laser conditions.</li>
	<li>Use the electro-optic modulator of the multi-photon microscope (see Table of Materials) to set the laser power to 70%-80%, corresponding to 60-70 mW power at the sample plane. 
	NOTE: If the laser is equipped with a femtosecond pulse compensator, use it to correct for group delay dispersion (GDD). GDD correction reduces the amount of laser power required for efficient mcMTOC ablation and minimizes photodamage to the oocyte. The GDD control is usually integrated with the image acquisition software of commercial multi-photon microscopes.</li>
	<li>Use the same image format, zoom factor, and scanning frequency as in step 5.6. Set the line and frame averaging parameters to 1.</li>
	<li>Click on the Scan button in the software to perform the ablation of the mcMTOC by conducting a single laser scan of the selected ROI.</li>
	<li>Use the channel settings from step 5.7 to review the results of the ablation by comparing the images of the GFP-labeled structures taken before and after the multi-photon laser exposure. If the ablation is successful, the intensity of the GFP fluorescence in the targeted mcMTOC decreases to the levels observed in the background.</li>
	<li>If some fragments of a GFP-labeling structure remain, repeat step 6.4 one or more times by focusing on various z planes of the mcMTOC.</li>
	<li>Use the channel settings from step 5.7 to verify the ablation efficiency by the complete loss of mcMTOC-associated mCherry signals (mCh-Cep192).</li>
	<li>Repeat step 5.8 to step 6.7 until all the MTOCs of an oocyte (at various focal planes) are ablated. 
	NOTE: This protocol uses mCh-Cep192 microinjection as a strategy to confirm mcMTOC depletion following multi-photon laser exposure and exclude the possibility that the loss of GFP fluorescence is caused by GFP photobleaching. However, the microinjection of this probe is optional and not required for the ablation procedure.</li>
</ol>

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The authors declare no competing interests.

Different methods exist to disturb the cytoskeleton-related structures within cells22,23,24,25. However, finding efficient techniques to selectively perturb the targeted structure without compromising the cell viability is challenging. The multi-photon laser ablation method presented here is an efficient strategy to induce a selective mechanical perturbation to mcMTOCs within the oocyte without altering the oocyte viability.
Laser ablation has been extensively used to understand the molecular mechanisms controlling chromosome segregation during mitosis and meiosis22,23,26,27. Due to the relatively large size (&#62;80 mm in diameter) of mammalian oocytes compared to somatic cells28, the ablation of their intracellular structures represents a challenge. Moreover, the average mcMTOC volume in metaphase I oocytes is ~20 &#181;m15, representing an additional challenge. An efficient method that offers deeper tissue penetration must be adopted to overcome these challenges. The main advantage of using the multi-photon laser for ablation is its ability to reach deeper into the cell while minimizing off-target effects29.
To verify the efficacy and efficiency of the laser ablation method to deplete the targeted structure, it is recommended to use a fluorescently labeled protein to identify the targeted structure over time (before and after ablation)23. It is important to notice that laser ablation depletes the whole mcMTOC as a structure, and although smaller mcMTOCs require only a single laser exposure to be depleted, larger mcMTOCs may require more than one laser exposure at different focal planes. It is also recommended to fix and immunolabel a subset of control and mcMTOC-ablated oocytes with an MTOC marker (such as &#947;-tubulin, pericentrin, or Cep192) to confirm the efficiency of the mcMTOC ablation further. In control oocytes, areas of the cytoplasm just adjacent to but not overlapping with mcMTOCs will be exposed to the laser.
This experiment requires several mcMTOC ablations while moving among different focal planes within the oocytes. Therefore, it is highly recommended to practice this technique several times before executing the experiment to minimize the experiment time, thereby increasing oocyte viability. Moreover, it is important to use the minimum laser power that is sufficient to deplete the mcMTOCs without affecting oocyte viability.
This technique has some limitations. First, multi-photon confocal microscopes are relatively more expensive than regular confocal microscopes. Second, perturbing all mcMTOCs at different focal planes is more time-consuming than chemical or genetic perturbations. Third, this protocol requires technical skills to ablate all mcMTOCs in the shortest possible time. However, once mastered, the use of the multi-photon laser provides an excellent strategy to perturb several intracellular structures in mouse oocytes, including mcMTOCs, contributing to the understanding of molecular mechanisms regulating spindle positioning and its timely migration in mammalian oocytes.

Haploid gametes (sperm and oocyte) are produced through meiosis, which entails one round of DNA replication followed by two consecutive divisions that are necessary for chromosome number reduction prior to fertilization. In mammals, during early fetal life, the oocyte undergoes a lengthy arrest (until puberty) at the diplotene stage of prophase I of the first meiotic division, a stage called the germinal vesicle (GV) stage. Following meiotic resumption, the GV oocyte undergoes nuclear envelope breakdown (NEBD), and the spindle is assembled mainly at the oocyte center1,2,3. Later, driven by F-actin, the spindle migrates in a timely manner from the oocyte center to the cortex to ensure highly asymmetrical division, resulting in an egg with a tiny polar body (PB)4,5,6.
In mitotic cells, the centrosomes consist of a pair of centrioles surrounded by peri-centriolar material components (PMC), such as pericentrin, &#947;-tubulin, Cep152, and Cep1927. These centriole-containing centrosomes contribute to the fidelity of bipolar spindle formation8. However, centrioles are lost during early oogenesis in various species, including rodents9. Therefore, mouse oocytes adopt a centriole-independent spindle assembly pathway using numerous acentriolar microtubule (MT) organizing centers (MTOCs)9,10. Upon meiotic resumption, the perinuclear MTOCs undergo three distinct steps of recondensation, stretching, and fragmentation into a large number of smaller MTOCs11,12. The fragmented MTOCs are then clustered and sorted to organize a bipolar spindle10,13,14. Another pool of MTOCs is located in the cytoplasm during NEBD. Some of these cytoplasmic MTOCs migrate and form spindle poles (polar MTOCs, pMTOCs)10,11. Recently, another subset of cytoplasmic MTOCs was discovered, termed metaphase cytoplasmic MTOCs (mcMTOCs), that do not contribute to spindle pole formation but remain in the oocyte cytoplasm during metaphase I (Met I)15. Depleting mcMTOCs by multi-photon laser ablation or abnormally increasing their numbers by autophagy inhibition perturbs spindle positioning and migration and increases the incidence of aneuploidy in metaphase II oocytes15.
Interestingly, mcMTOCs differ from pMTOCs in many aspects15. For example, in contrast to pMTOCs, which mainly originate from perinuclear MTOCs, mcMTOCs originate from the oocyte cortex. When the spindle is still at the oocyte center, the mcMTOCs are localized asymmetrically opposite to the side to which the spindle migrates for PB extrusion15. Astral-like MTs cannot reach the cortex in the relatively large oocyte cell. Therefore, these mcMTOCs nucleate MTs to anchor the spindle (via astral-like MTOCs) to the cortex. These findings suggest a model in which the mcMTOC-nucleated MT force counteracts the F-actin-mediated force that drives spindle migration toward the cortex. The balance between these two opposing forces is essential to regulate the central spindle positioning and timely spindle migration15.
To date, all the examined PMC proteins (pericentrin, g-tubulin, Cep192, and Aurora kinase A) localize to both MTOC pools: mcMTOCs and pMTOCs15. Therefore, there is no chemical or genetic approach to selectively perturb mcMTOCs without perturbing pMTOCs. These limitations can be circumvented by selectively targeting the mcMTOCs with laser ablation. Among the laser-based technologies developed for microablation, pulsed multi-photon femtosecond lasers show great potential due to their precision impact limited to the focal plane, the high penetration depth of near-infrared light, and the reduced phototoxicity and thermal damage to the cell16,17,18. This work describes a selective approach to ablate mcMTOCs in mouse oocytes using a multi-photon laser coupled to an inverted microscope.

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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.

Multi-photon laser ablation provides an efficient method to selectively ablate intracellular structures. The current study employed multi-photon laser ablation to deplete mcMTOCs in mouse oocytes selectively. Laser ablation depleted the mcMTOCs efficiently, as shown by the reduction in the endogenous GFP fluorescence in the targeted mcMTOCs to a level comparable to the background. Exogenous mCh-Cep192 in the targeted mcMTOCs was also abolished following laser ablation. The laser ablation of mcMTOCs should be performed at different focal planes within the oocyte (where mcMTOCs are located, Figure 2) to ensure that all the mcMTOCs within the oocyte are depleted (Figure 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64439/64439fig01.jpg" /> 
Figure 1: Oocyte microinjection. Microinjection of a prophase I-arrested mouse oocyte with mCh-Cep192 cRNA. Scale bar: 10 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64439/64439fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64439/64439fig02.jpg" /> 
Figure 2: Depletion of mcMTOCs in mouse oocytes. Representative images of single focal planes. The white squares indicate an mcMTOC before and after multi-photon laser ablation. Scale bar: 40 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64439/64439fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64439/64439fig03.jpg" /> 
Figure 3: Depletion of mcMTOCs at different focal planes in a mouse oocyte. Representative maximum projection images of an mcMTOC-ablated mouse oocyte. Scale bar: 20 &#181;m. <a href="https://www.jove.com/files/ftp_upload/64439/64439fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Compositions of Chatot, Ziomek, and Bavister (CZB) and bicarbonate-free minimal essential medium (MEM/PVP). <a href="https://www.jove.com/files/ftp_upload/64439/64439_Supplementary File 1.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Multi-Photon Laser Ablation of Cytoplasmic Microtubule Organizing Centers in Mouse Oocytes at JoVE.com

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

This protocol offers an excellent strategy to selectively ablate cytoplasmic MTOCs, which are critical regulators for spindle positioning in mouse oocytes.Therefore contributing to our understanding of molecular mechanisms that regulates oocyte meiosis.This technique allows to perform targeted ablations within live oocytes with high spatial accuracy and with minimal phototoxic effects on the cell.This protocol can be used to induce a selective mechanical perturbation of different instructors without altering the cell viability.This protocol requires the technical skills to ablate all cytoplasmic MTOCs in the shortest possible time to avoid compromising the oocytes'viability.So practicing this technique several times before carrying out the actual experiment is essential.To begin supplement the CZB medium with glutamine and milrinone and place it in the incubator.Prepare the collection and culture dishes.Make four micro drops of collection medium and two micro drops of culture medium in 60 and 35 mm Petri dishes, respectively.And cover them with mineral oil.Keep the collection dish on the slide warmer, and incubate at 37

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1.6K Views.  University of Missouri. This protocol offers an excellent strategy to selectively ablate cytoplasmic MTOCs, which are critical regulators for spindle positioning in mouse oocytes.Therefore contributing to our understanding of molecular mechanisms that regulates oocyte meiosis.This technique allows to perform targeted ablations within live oocytes with high spatial accuracy and with minimal phototoxic effects on the cell.This protocol can be used to induce a selective mechanical perturbation of different instructors ...

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

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the cephalochordate amphioxus, Branchiostoma lanceolatum. We use constructs for membrane and nuclear targeted mCherry and eGFP that have been modified to accommodate amphioxus codon usage and Kozak consensus sequences. We describe the type of injection needles to be used, the immobilization protocol for the unfertilized oocytes, and the overall injection set-up. This technique generates fluorescently labeled embryos, in which the dynamics of cell behaviors during early development can be analyzed using the latest in vivo imaging strategies. The development of a microinjection technique in this amphioxus species will allow live imaging analyses of cell behaviors in the embryo as well as gene-specific manipulations, including gene overexpression and knockdown. Altogether, this protocol will further consolidate the basal chordate amphioxus as an animal model for addressing questions related to the mechanisms of embryonic development and, more importantly, to their evolution.

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the ...

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 ...

Surgical Ablation Assay for Studying Eye Regeneration in Planarians

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their abundant pluripotent stem cell population and ability to regenerate all cell and tissue types after injuries that would be catastrophic for most animals. Recently, planarians have gained popularity as a model for eye regeneration. Their ability to regenerate the entire eye (comprised of two tissue types: pigment cells and photoreceptors) allows for the dissection of the mechanisms regulating visual system regeneration. Eye ablation has several advantages over other techniques (such as decapitation or hole punch) for examining eye-specific pathways and mechanisms, the most important of which is that regeneration is largely restricted to eye tissues alone. The purpose of this video article is to demonstrate how to reliably remove the planarian optic cup without disturbing the brain or surrounding tissues. The handling of worms and maintenance of an established colony is also described. This technique uses a 31 G, 5/16-inch insulin needle to surgically scoop out the optic cup of planarians immobilized on a cold plate. This method encompasses both single and double eye ablation, with eyes regenerating within 1-2 weeks, allowing for a wide range of applications. In particular, this ablation technique can be easily combined with pharmacological and genetic (RNA interference) screens for a better understanding of regenerative mechanisms and their evolution, eye stem cells and their maintenance, and phototaxic behavioral responses and their neurological basis.

This protocol shows how to consistently excise planarian eyes (optic cups) without disturbing surrounding tissues. Using an insulin needle and syringe, either one or both eyes can be ablated to facilitate investigations into the mechanisms regulating eye regeneration, the evolution of visual regeneration, and the neural basis of light-induced behavior.

In the study of adult stem cells and regenerative mechanisms, planarian flatworms are a staple in vivo model system. This is due in large part to their ...

Multi-Photon Time Lapse Imaging to Visualize Development in Real-time: Visualization of Migrating Neural Crest Cells in Zebrafish Embryos

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise to numerous cell types throughout the body. Understanding the biology of the neural crest has been limited, reflecting a lack of genetically tractable models that can be studied in vivo and in real-time. Zebrafish is a particularly important developmental model for studying migratory cell populations, such as the neural crest. To examine neural crest migration into the developing eye, a combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time videos of the developing eye in transgenic zebrafish embryos, namely Tg(sox10:EGFP) and Tg(foxd3:GFP), as sox10 and foxd3 have been shown in numerous animal models to regulate early neural crest differentiation and likely represent markers for neural crest cells. Multi-photon time-lapse imaging was used to discern the behavior and migratory patterns of two neural crest cell populations contributing to early eye development. This protocol provides information for generating time-lapse videos during zebrafish neural crest migration, as an example, and can be further applied to visualize the early development of many structures in the zebrafish and other model organisms.

A combination of the advanced optical techniques of laser scanning microscopy with long wavelength multi-photon fluorescence excitation was implemented to capture high-resolution, three-dimensional, real-time imaging of neural crest migration in Tg(sox10:EGFP) and Tg(foxd3:GFP) zebrafish embryos.

Congenital eye and craniofacial anomalies reflect disruptions in the neural crest, a transient population of migratory stem cells that give rise 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 ...

Confocal Microscope-Based Laser Ablation and Regeneration Assay in Zebrafish Interneuromast Cells

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally replenished in non-mammalian vertebrates such as zebrafish. The zebrafish lateral line system is a useful model for characterizing sensory hair cell regeneration. The lateral line is comprised of hair cell-containing organs called neuromasts, which are linked together by a string of interneuromast cells (INMCs). INMCs act as progenitor cells that give rise to new neuromasts during development. INMCs can repair gaps in the lateral line system created by&#160;cell death. A method is described here for selective INMC ablation using a conventional laser-scanning confocal microscope and transgenic fish that express green fluorescent protein in INMCs. Time-lapse microscopy is then used to monitor INMC regeneration and determine the rate of gap closure. This represents an accessible protocol for cell ablation that does not require specialized equipment, such as a high-powered pulsed ultraviolet laser. The ablation protocol may serve broader interests, as it could be useful for the ablation of additional cell types, employing a tool set that is already available to many users. This technique will further enable the characterization of INMC regeneration under different conditions and from different genetic backgrounds, which will advance the understanding of sensory progenitor cell regeneration.

Laser ablation is a widely applicable technology for studying regeneration in biological systems. The presented protocol describes use of a standard laser-scanning confocal microscope for laser ablation and subsequent time-lapse imaging of regenerating interneuromast cells in the zebrafish lateral line.

Hair cells are mechanosensory cells that mediate the sense of hearing. These cells do not regenerate after damage in humans, but they are naturally ...

Minimum Volume Vitrification of Immature Feline Oocytes

In wild animals&#8217; conservation programs, gamete banking is crucial to safeguard genetic resources of valuable individuals and rare species and to promote biodiversity preservation. In felids, most species are threatened with extinction, and domestic breeds are used as a model to increase the efficiency of protocols for germplasm banking. Among oocyte cryopreservation techniques, vitrification is more and more popular in human and veterinary assisted reproduction. Cryotop vitrification, which was at first developed for human oocytes and embryos, has demonstrated to be well-suited for cat oocytes. This method offers several advantages, such as the feasibility in field conditions and the speed of the procedure, which can be helpful when several samples need to be processed. However, the efficiency is strongly dependent on the operator&#8217;s skills, and intra- and inter-laboratory standardization are needed, as well as personnel training. This protocol describes minimum volume vitrification of immature feline oocytes on a commercial support in a step by step field-friendly protocol, from oocyte collection to warming. Following the protocol, preservation of oocyte integrity and viability at warming (as high as 90%) can be expected, although there is still room for improvement in post-warming maturation and embryonic development outcomes.

This manuscript describes a protocol for the minimum volume vitrification of immature cat oocytes with laboratory-made media on commercial supports. It covers every step from oocyte isolation from ex vivo gonads to vitrification and warming.

In wild animals&#8217; conservation programs, gamete banking is crucial to safeguard genetic resources of valuable individuals and rare species and to ...

Separation of Follicular Cells and Oocytes in Ovarian Follicles of Zebrafish

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of oocytes and surrounding follicular cells. It is vital to separate both follicular cells and oocytes for various research purposes such as for primary culture of follicular cells, analysis of gene expression, oocyte maturation and in vitro fertilization, etc. The conventional method uses forceps to separate both compartments, which is laborious, time consuming and has high damage to the oocyte. Here, we have established a simple method to separate both compartments using a pulled glass capillary. Under a stereomicroscope, oocytes and follicular cells can be easily separated by pipetting in a pulled fine glass capillary (the diameter depends on the follicle diameter). Compared with the conventional method, this new method has high efficiency in separating both oocytes and follicular cells and has low damage to the oocytes. More importantly, this method can be applied to early-stage follicles including at the pre-vitellogenesis stage. Thus, this simple method can be used to separate follicular cells and oocytes of zebrafish.

Here, we present a simple method for separating follicular cells and oocytes in zebrafish ovarian follicles, which will facilitate investigations of ovarian development in zebrafish.

Zebrafish has become an ideal model to study the ovarian development of vertebrates. The follicle is the basic unit of the ovary, which consists of ...

Vitrification of In Vitro Matured Oocytes Collected from Adult and Prepubertal Ovaries in Sheep

In livestock, in vitro embryo production systems can be developed and sustained thanks to the large number of ovaries and oocytes that can be easily obtained from a slaughterhouse. Adult ovaries always bear several antral follicles, while in pre-pubertal donors the maximal numbers of oocytes are available at 4 weeks of age, when ovaries bear peak numbers of antral follicles. Thus, 4 weeks old lambs are considered good donors, even if the developmental competence of prepubertal oocytes is lower compared to their adult counterpart. 
Basic research and commercial applications would be boosted by the possibility of successfully cryopreserving vitrified oocytes obtained from both adult and prepubertal donors. The vitrification of oocyte collected from prepubertal donors would also allow shortening the generation interval and thus increasing the genetic gain in breeding programs. However, the loss of developmental potential after cryopreservation makes mammalian oocytes probably one of the most difficult cell types to cryopreserve. Among the available cryopreservation techniques, vitrification is widely applied to animal and human oocytes. Despite recent advancements in the technique, exposures to high concentrations of cryoprotective agents as well as chilling injury and osmotic stress still induce several structural and molecular alterations and reduce the developmental potential of mammalian oocytes. Here, we describe a protocol for the vitrification of sheep oocytes collected from juvenile and adult donors and matured in vitro prior to cryopreservation. The protocol includes all the procedures from oocyte in vitro maturation to vitrification, warming and post-warming incubation period. Oocytes vitrified at the MII stage can indeed be fertilized following warming, but they need extra time prior to fertilization to restore damage due to cryopreservation procedures and to increase their developmental potential. Thus, post-warming culture conditions and timing are crucial steps for the restoration of oocyte developmental potential, especially when oocyte are collected from juvenile donors.

The protocol aims at providing a standard method for the vitrification of adult and juvenile sheep oocytes. It includes all the steps from the preparation of the in vitro maturation media to the post-warming culture. Oocytes are vitrified at the MII stage using Cryotop to ensure the minimum essential volume.

In livestock, in vitro embryo production systems can be developed and sustained thanks to the large number of ovaries and oocytes that can be easily ...

Spatiotemporal Subcellular Manipulation of the Microtubule Cytoskeleton in the Living Preimplantation Mouse Embryo using Photostatins

The microtubule cytoskeleton forms the framework of a cell and is fundamental for intracellular transport, cell division, and signal transduction. Traditional pharmacological disruption of the ubiquitous microtubule network using, for instance, nocodazole can have devastating consequences for any cell. Reversibly photoswitchable microtubule inhibitors have the potential to overcome the limitations by enabling drug effects to be implemented in a spatiotemporally-controlled manner. One such family of drugs is the azobenzene-based photostatins (PSTs). These compounds are inactive in dark conditions, and upon illumination with UV light, they bind to the colchicine-binding site of &#946;-tubulin and block microtubule polymerization and dynamic turnover. Here, the application of PSTs in the 3-dimensional (3D) live preimplantation mouse embryo is set out to disrupt the microtubule network on a subcellular level. This protocol provides instructions for the experimental setup, as well as light activation and deactivation parameters for PSTs using live-cell confocal microscopy. This ensures reproducibility and enables others to apply this procedure to their research questions. Innovative photoswitches like PSTs may evolve as powerful tools to advance the understanding of the dynamic intracellular microtubule network and to non-invasively manipulate the cytoskeleton in real-time. Furthermore, PSTs may prove useful in other 3D structures such as organoids, blastoids, or embryos of other species.

Typical microtubule inhibitors, used widely in basic and applied research, have far-reaching effects on cells. Recently, photostatins emerged as a class of photoswitchable microtubule inhibitors, capable of instantaneous, reversible, spatiotemporally precise manipulation of microtubules. This step-by-step protocol details the application of photostatins in a 3D live preimplantation mouse embryo.