Name: planarian ovary dissection for ultrastructural analysis and antibody staining
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Description: Watch this Scientific Journal Video about Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining at JoVE.com

The work was supported by the Howard Hughes Medical Institute (LK and ASA) and the Helen Hay Whitney Foundation (LHG). 
 
Original data underlying this manuscript can be accessed from the Stowers Original Data Repository at http://www.stowers.org/research/publications/libpb-1628

1. Preparation
<ol>
	<li>Prepare worms: feed sexual planarians twice a week with organic liver paste to achieve sexual maturity. 
	NOTE: Generally, such worms are bigger than 1 cm in length and have a gonopore posterior to the pharynx opening.</li>
	<li>Prepare solutions.
	<ol>
		<li>Prepare the following reagents: 16% paraformaldehyde (PFA); 50% glutaraldehyde (GA) aqueous solution; N-acetyl-L-cysteine (NAC); 1x phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 8 mM Na2HPO4, and 2 mM KH2PO4.</li>
		<li>Dilute 50% GA and 16% PFA with PBS to a final mixture of 2.5% GA and 2% PFA. Dilute 16% PFA with PBS to a final concentration of 4% PFA.</li>
		<li>Dissolve 5 g of NAC in 100 mL of PBS to make a 5% working solution.</li>
	</ol>
	</li>
</ol>
2. Collecting ovaries
<ol>
	<li>Treat sexually mature worms with 5% NAC in a dish at room temperature for 5 min. 
	NOTE: Large worms may curl up. Brushes or pipette tips can be used to flatten the worms. The amount of 5% NAC used is minimal, enough to cover the surface of the Petri dish.</li>
	<li>Replace NAC with 10 mL of freshly prepared 4% paraformaldehyde and fix the worms for 1 h at room temperature with occasional shaking. Start the dissection 10 min after the addition of 4% PFA, and perform the dissection in 4% PFA as the worms are being fixed.</li>
	<li>Observe the epithelial pigmentation to locate the ventral nerve cords, which appear as two lines with lighter pigmentation running from the anterior cephalic ganglia toward the tail.</li>
	<li>Locate the ovaries. 
	NOTE: The ovary is located right next to the nerve cord on the medial side (Figure 1A). Rely on epithelia pigmentation to locate the position of the ovary in the anterior-posterior direction. Ideally, the pigmentation of the ventral epithelia in the ovary region is lighter than the surrounding regions. In some worms, the pigmentation of the ventral epithelia does not provide enough confidence. These worms can be discarded as some may not have well-developed ovaries.</li>
	<li>Validate the ovaries&#39; positions once the putative ovaries are located by pigmentation. 
	NOTE: First, the ovaries are posterior to the cephalic ganglia. Second, dark-colored testes are lateral to the dorsal surface above the ovary. Ovaries locate in front of the most anterior testis (Figure 1B). Sometimes, testis lobes are positioned at the level of or slightly anterior to the ovaries.</li>
	<li>Use two surgical knives to cut posterior and anterior to the two ovaries (Figure 1C) to remove the anterior and posterior worm fragments completely. Use one knife to anchor the worm and clean the other knife used to make the cuts.</li>
	<li>Cut in the middle line of the ovary fragment to separate the two ovaries. Flip the fragment to have the ventral side down.</li>
	<li>Peel off the dorsal tissue to expose the gut (Figure 1D) with two pairs of pointed tweezers (type 5-SA). 
	NOTE: The dome-shaped ovary is located beneath the gut branches.</li>
	<li>Gently remove the gut branches sitting above the ventral tissues with the tip of the tweezers or a soft brush to expose the ovary (Figure 1E).</li>
	<li>Remove the surrounding tissues to take out the ovary (Figure 1F,G).</li>
	<li>Take the ovaries out and transfer them to a 1.5 mL tube to wash with 1 mL of PBS.</li>
</ol>
3. Fixation for TEM
<ol>
	<li>Wash the ovaries in PBS for 10 min with gentle shaking. Keep the tubes upright, and let the ovaries sink to the bottom by gravity. Repeat the wash once. 
	NOTE: If the ovaries do not sink, apply a gentle spin for 15 s with a mini-benchtop centrifuge (maximum speed 2000 &#215; g).</li>
	<li>Replace the PBS with 2% PFA/2.5% GA/PBS.</li>
	<li>Fix the samples at room temperature for 1 h on a shaker at 40 rpm.</li>
	<li>Keep the samples in fixative at 4 &#176;C overnight on a shaker at 40 rpm.</li>
</ol>
4. Sample processing for TEM
<ol>
	<li>Wash the ovaries in PBS 3 times for 10 min each.</li>
	<li>Wash the ovaries in double-distilled water (ddH2O) 3 times for 10 min each.</li>
	<li>Post-fix the ovaries in 2% aqueous OsO4 for 1-2 h at room temperature or overnight at 4 &#176;C. 
	NOTE: The sample containers need to be sealed with parafilm during this step. Prepare 2% OsO4 with reverse-osmosis-treated water (see the Table of Materials).</li>
	<li>Rinse the samples with ddH2O 3 times, 10 min each.</li>
	<li>Pre-stain the ovaries in 2% aqueous uranyl acetate (UA) overnight at 4 &#176;C. 
	NOTE: The UA solution needs to be filtered before use; avoid light during the en bloc staining. Prepare 2% OsO4 with reverse-osmosis-treated water (see the Table of Materials).</li>
	<li>Rinse the samples in ddH2O 4 times with gentle agitation, 10 min each.</li>
	<li>Dehydrate the ovaries in a graded ethanol series (30%, 50%, 70%, 95%, and two times 100%, 10 min each solution) and then equilibrate them in two incubations (10 min) in propylene oxide.</li>
	<li>For infiltration with resin, incubate the samples in 50% propylene oxide/50% liquid epoxy resin mixture overnight at 4 &#176;C. Infiltrate the samples with 100% epoxy resin with 3 changes (1 h each change) with gentle agitation.</li>
	<li>Embed the ovaries in 100% epoxy resin and polymerize the resin at 60 &#176;C for 48 h.</li>
</ol>
5. Ultramicrotomy
<ol>
	<li>Block trimming and sample check with a light microscope
	<ol>
		<li>Trim the sample blocks into a pyramid shape with a razor blade.</li>
		<li>Cut sections of 1-2 &#956;m thickness with a glass or diamond knife.</li>
		<li>Transfer the sections onto a slide with a drop of water, then heat the slide on a hot plate until the sections flatten and adhere to the slide surface during water evaporation.</li>
		<li>Cover the sections with a drop of 1% toluidine blue O and heat them on a hot plate for ~10 s. Rinse the slide with running water, then let it dry. Check the sections under a regular light microscope.</li>
	</ol>
	</li>
	<li>Ultrathin sections cutting, collection, and post-staining.
	<ol>
		<li>Cut ultrathin sections of 50-70 nm thickness with a diamond knife. Transfer the sections from the knife water boat onto mesh or single-slot copper grids.</li>
		<li>Stain the sections in 2% UA for 8 min. Wash the grids in running ddH2O for 30 s.</li>
		<li>Stain the sections with 1% lead solution for 6 min. Wash the sections in running ddH2O for 1 min and air-dry.</li>
	</ol>
	</li>
</ol>
6. Data collection and analysis
<ol>
	<li>Collect TEM data using appropriate software, e.g., Digital Micrograph (see the Table of Materials for more details).</li>
	<li>Operate the scope at 80 kV. Insert the sample grid into the scope when the vacuum is ready.</li>
	<li>Turn on the beam by clicking Light on the menu. Turn the Intensity knob on the left control pad until the Auto Exposure time is ~1 s. Click on Start Acquire to get a final image.</li>
	<li>Construct three-dimensional EM models using the open-source image processing, modeling, and display (IMOD) package.</li>
</ol>
7. Antibody staining
<ol>
	<li>Wash the dissected ovaries (step 2.16) in a 1.5 mL microfuge tube with 1 mL of 1x PBS supplemented with 0.5% Triton X-100 (PBST). Place the tubes on a shaker for agitation at 40 rpm for 10 min. Let the tubes stand in a rack and the ovaries sink by gravity. Replace the wash with fresh PBST, and repeat twice.</li>
	<li>Digest the tissues with 2 &#181;g/mL of Proteinase K and 0.1% sodium dodecylsulfate for 10 min at room temperature in PBST.</li>
	<li>Wash the digested ovaries in PBST three times for 10 min each wash.</li>
	<li>Incubate the ovaries with 10% horse serum in PBST for 1 h at room temperature.</li>
	<li>Dilute primary antibodies 1 to 100 in the blocking solution (10% horse serum in PBS with 0.5% Triton X-100). Incubate the ovaries with primary antibodies overnight at 4 &#176;C with gentle shaking.</li>
	<li>Wash the ovaries in PBST three times for 10 min each wash.</li>
	<li>Incubate the ovaries with secondary antibodies overnight at 4 &#176;C with gentle shaking. Dilute all secondary antibodies 1:300 in the blocking solution.</li>
	<li>Wash the ovaries in PBST three times, ensuring that the first and third washes last for 10 min. Stain the ovarian nuclei with Hoechst 33342 at 1:300 dilution in PBST for 30 min at room temperature in the second wash.</li>
	<li>Mount the ovaries onto slides. 
	NOTE: Anti-fading mountant can be used to prolong the fluorescence signals.</li>
</ol>

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The authors have nothing to disclose.

These fixation-based procedures for dissecting planarian ovaries will facilitate the understanding of oocyte meiosis as well as ovary development and regeneration. The sizes of the oocytes and their somatic supportive cells can range from 20 &#181;m to 50 &#181;m. Dissection-based methods will provide accessibility to intact single-ovary cells that sectioning or whole-mount-based methods cannot achieve. This protocol will facilitate the study of intact planarian ovary anatomy and oocyte cell biology at cellular and subce...

Planarian anatomy has been examined by using TEM in many tissues1,2,3,4,5,6. However, little attention has been given to ovaries or oocytes. The paucity of oocyte literature is partly due to the difficulty accessing these cells, leaving the biology of planarian oocytes largely unexplored. Molecular tools have uncovered many regulatory mechanisms of ovary development in the planarians&#160;using light or fluorescence microscopy7,8,9,10,11,12,13,14,15,16,17,18,19,20. All these experiments were performed on whole worms or histological sections of whole worms. The antibody staining and in situ hybridization protocols on whole worms involve extensive bleaching, washing, and tissue clearing steps, which are time-consuming and will take several days.
The overall goal of the method described here is to provide accessibility to intact, dissected planarian ovaries and oocytes, which will remove the necessity of bleaching or histological sectioning and shorten the time for washing and tissue clearing in antibody staining and in situ hybridization. The dissected ovaries will also improve probe or antibody penetration and increase imaging depth and quality for light and electron microscopes. Accessibility to the dissected ovaries and oocytes allows cell biology research at cellular and subcellular resolution with whole intact oocytes. A recent study on dissected planarian ovaries characterized planarian oocyte meiosis for the first time with TEM and confocal microscopy21. The work provided a comprehensive description of a new phenomenon during meiosis called nuclear envelope vesiculation.
Here, we present the detailed procedures in the dissection of planarian ovaries. A fixation step was sufficient to preserve the ovary cell structure for dissection and downstream manipulation (i.e., processing for TEM and light microscope analysis). Given their similarity in body plans and tissue architecture, this protocol should also be broadly informative for studying oocytes and their nuclei in several other Platyhelminthes species (e.g., the genus of Dugesia or Polycelis). This protocol is likely irrelevant for Macrostomum lignano for their small sizes and almost transparent body architecture, which will allow for direct observation of the ovary and oocytes22,23,24. The body area containing the ovaries is more optically distinguishable (e.g., darker pigmented or lighter pigmented) in some species (e.g., Dugesia ryukyuensis9,25) than S. mediterranea. Studies in these species can rely less on the guidelines for locating the ovaries in S. mediterranea presented here but take advantage of the fixation and dissection conditions.

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planarian ovary dissection for ultrastructural analysis and antibody staining

Accessibility to germ cells allows the study of germ cell development, meiosis, and recombination. The sexual biotype of the freshwater planarian, Schmidtea mediterranea, is a powerful invertebrate model to study the epigenetic specification of germ cells. Unlike the large number of testis and male germ cells, planarian oocytes are relatively difficult to locate and examine, as there are only two ovaries, each with 5-20 oocytes. Deeper localization within the planarian body and lack of protective epithelial tissues also make it challenging to dissect planarian ovaries directly. 
This protocol uses a brief fixation step to facilitate the localization and dissection of planarian ovaries for downstream analysis to overcome these difficulties. The dissected ovary is compatible for ultrastructural examination by transmission electron microscopy (TEM) and antibody immunostaining. The dissection technique outlined in this protocol also allows for gene perturbation experiments, in which the ovaries are examined under different RNA interference (RNAi) conditions. Direct access to the intact germ cells in the ovary achieved by this protocol will greatly improve the imaging depth and quality and allow cellular and subcellular interrogation of oocyte biology.

The method presented here has been described by Guo et al.21. The key to successful dissection is to identify the ovary pigmentation and position guides correctly. The strategy of the method is to move from broad positions to a specific location. First, to achieve this, rely on dorsal and ventral pigmentation patterns (Figure 1A,B). Ventral pigmentation, where the ovaries reside, will turn white after 5% NAC treatment and 4% PFA fixation. If the worm ...

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Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

This protocol presents steps taken to dissect ovaries in the freshwater planarians, Schmidtea mediterranea. The dissected ovaries are compatible for antibody immunostaining and ultrastructural analysis with transmission electron microscopy to study the cell biology of the oocytes and somatic cells, providing an imaging depth and quality that were previously inaccessible.

Accessibility to germ cells allows the study of germ cell development, meiosis, and recombination. The sexual biotype of the freshwater planarian, ...

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Research

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Biology

1.5K Views.  Stowers Institute for Medical Research. This protocol presents steps taken to dissect ovaries in the freshwater planarians, Schmidtea mediterranea. The dissected ovaries are compatible for antibody immunostaining and ultrastructural analysis with transmission electron microscopy to study the cell biology of the oocytes and somatic cells, providing an imaging depth and quality that were previously inaccessible.

Video: Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

The mammalian ovary is composed of ovarian follicles, each follicle consisting of a single oocyte surrounded by somatic granulosa cells, enclosed together within a basement membrane. A finite pool of follicles is laid down during embryonic development, when oocytes in meiotic arrest form a close association with flattened granulosa cells, forming primordial follicles. By or shortly after birth, mammalian ovaries contain their lifetime&#8217;s supply of primordial follicles, from which point onwards there is a steady release of follicles into the growing follicular pool.
The ovary is particularly amenable to development in vitro, with follicles growing in a highly physiological manner in culture. This work describes the culture of whole neonatal ovaries containing primordial follicles, and the culture of individual ovarian follicles, a method which can support the development of follicles from an immature through to the preovulatory stage, after which their oocytes are able to undergo fertilization in vitro. The work outlined here uses culture systems to determine how the ovary is affected by exposure to external compounds. We also describe a co-culture system, which allows investigation of the interactions that occur between growing follicles and the non-growing pool of primordial follicles.

This protocol describes the primary culture/co-culture of mouse ovarian tissue, using ovaries from neonatal mice and individual ovarian follicles from prepubertal mice. The culture techniques support development in a highly physiological manner, allowing investigation of the effect of extrinsic agents on the ovary, and of interactions between ovarian follicles.

The mammalian ovary is composed of ovarian follicles, each follicle consisting of a single oocyte surrounded by somatic granulosa cells, enclosed together ...

Developmental Biology

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Dissection and Staining of Drosophila Pupal Ovaries

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a pupal case. The challenge of dissecting pupal ovaries also lies in their physical location within the pupa: the ovaries are surrounded by fat body cells inside the pupal abdomen, and these fat cells must be removed to allow for proper antibody staining. To overcome these challenges, this protocol utilizes customized Pasteur pipets to extract fat body cells from the pupal abdomen. Moreover, a chambered coverglass is used in place of a microcentrifuge tube during the staining process to improve visibility of the pupae. However, despite these and other advantages of the tools used in this protocol, successful execution of these techniques may still involve several days of practice due to the small size of pupal ovaries. The techniques outlined in this protocol could be applied to time course experiments in which ovaries are analyzed at various stages of pupal development.

Dissection and Staining of Drosophila Pupal Ovaries

The Drosophila ovary is an excellent model system for studying stem cell niche development. Though methods for dissecting larval and adult ovaries have been published, pupal ovary dissections require different techniques that have not been published in detail. Here we outline a protocol for dissecting, staining, and mounting pupal ovaries.

Unlike adult Drosophila ovaries, pupal ovaries are relatively difficult to access and examine due to their small size, translucence, and encasing within a ...

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. Whole mount preparations preserve the structure of meiotic nuclei, and importantly, each gonad arm contains all stages of meiosis, organized in a temporal-spatial progression that makes it easy to identify nuclei at different stages. Adult hermaphrodites have two gonad arms, each organized as a closed tube with proliferating germline stem cells at the distal closed end and cellularized oocytes at the proximal open end, which join in the center at the uterus. Dissection releases one or both gonad arms from the body cavity, allowing the entirety of meiosis to be visualized. Here, a common protocol for immunofluorescence against a protein of interest is presented, followed by DAPI staining to mark all chromosomes. Young adults are immobilized in levamisole and quickly dissected using two syringe needles. After germline extrusion, the sample is fixed before undergoing a freeze crack in liquid nitrogen, which helps permeabilize the cuticle and other tissues. The sample can then be dehydrated in ethanol, rehydrated, and incubated with primary and secondary antibodies. DAPI is added to the sample in the mounting medium, which allows reliable visualization of DNA and makes it easy to find animals to image under a fluorescent microscope. This technique is readily adopted by those familiar with handling C. elegans after a few hours spent practicing the dissection method itself. This protocol has been taught to high-schoolers and undergraduates working in a research lab and incorporated into a course-based undergraduate research experience at a liberal arts college.

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

This is a commonly used method for C. elegans gonad dissection followed by freeze crack, which produces germline samples for immunofluorescence via antibody staining, or for simple DAPI staining to visualize DNA. This protocol has been successful for undergraduates in a research lab and in a course-based undergraduate research experience.

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. ...

Genetics

Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart disease. To study the formation of PAAs, we developed a protocol using whole-mount immunofluorescence coupled with benzyl alcohol/benzyl benzoate (BABB) tissue clearing, and confocal microscopy. This allows for the visualization of the pharyngeal arch endothelium at a fine cellular resolution as well as the 3D connectivity of the vasculature. Using software, we have established a protocol to quantify the number of endothelial cells (ECs) in PAAs, as well as the number of ECs within the vascular plexus surrounding the PAAs within pharyngeal arches 3, 4, and 6. When applied to the whole embryo, this methodology provides a comprehensive visualization and quantitative analysis of embryonic vasculature.

Here, we describe a protocol to visualize and analyze the pharyngeal arch arteries 3, 4, and 6 of mouse embryos using whole-mount immunofluorescence, tissue clearing, confocal microscopy, and 3D reconstruction.

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart ...

Dissection for Identifying Migratory Insects via Ovarian Reproductive Analysis

Dissection and Grading of Ovarian Development in Wild-Type Female Insects

Migratory insect pests pose serious challenges to food production and security all over the world. The migratory pests can be monitored and captured using searchlight traps. One of the most important techniques for migratory pest forecasting is to identify the migratory species. However, in most cases, it is difficult to get the information just by appearance. Therefore, using knowledge acquired by systematic analysis of the female reproductive system can help to understand the combined anatomic morphology of the ovarian mating sac and ovary developmental grading of wild-type migratory insects captured with searchlight traps. To demonstrate the applicability of this method, ovarian development status and egg grain development stages were directly assessed in Helicoverpa armigera, Mythimna separata, Spodoptera litura, and Spodoptera exigua for the ovarian anatomy, and the ovarian mating sacs were studied in Agrotis ipsilon, Spaelotis valida, Helicoverpa armigera, Athetis lepigone, Mythimna separata, Spodoptera litura, Mamestra brassicae, and Spodoptera exigua, to explore their relationships. This work shows the specific dissection method to predict wild-type migratory insects, comparing the unique reproductive system of different migratory insects. Then, both tissues, namely, the ovary and mating sacs, were further investigated. This method helps to predict the dynamics and the structural development of reproductive systems in wild-type female migratory insects.

Author Spotlight: Analysis of Ovarian Anatomy in Migratory Insects to Overcome Experimental Challenges 

The protocol demonstrates a simple and easy dissection method, suitable for wild-type migratory female insects captured with searchlight traps. This technique can significantly clarify the same species by comparing both reproductive tissues, namely the mating sac and ovarian development of wild-type female insects.

Migratory insect pests pose serious challenges to food production and security all over the world. The migratory pests can be monitored and captured using ...

Behavior

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Microdissection and Dissociation of the Murine Oviduct: Individual Segment Identification and Single Cell Isolation

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental treatments. However, because of their small body size, some structures, such as the oviduct with a diameter of 200-400 &#956;m, have proven to be relatively difficult to study except by immunohistochemistry. Recently, immunohistochemical studies have uncovered more complex differences in oviduct segments than were previously recognized; thus, the oviduct is divided into four functional segments with different ratios of seven distinct epithelial cell types. The different embryological origins and ratios of the epithelial cell types likely make the four functional regions differentially susceptible to disease. For example, precursor lesions to serous intraepithelial carcinomas arise from the infundibulum in mouse models and from the corresponding fimbrial region in the human fallopian tube. The protocol described here details a method for microdissection to subdivide the oviduct in such a way to yield a sufficient amount and purity of RNA necessary for downstream analysis such as reverse transcription-quantitative PCR (RT-qPCR) and RNA sequencing (RNAseq). Also described is a mostly non-enzymatic tissue dissociation method appropriate for flow cytometry or single cell RNAseq analysis of fully differentiated oviductal cells. The methods described will facilitate further research utilizing the murine oviduct in the field of reproduction, fertility, cancer, and immunology.

A method for microdissection of the mouse oviduct that allows collection of the individual segments while maintaining RNA integrity is presented. In addition, non-enzymatic oviductal cell dissociation procedure is described. The methods are appropriate for subsequent gene and protein analysis of the functionally different oviductal segments and dissociated oviductal cells.

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental ...

Dissection and Staining of Drosophila Larval Ovaries

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

Dissection and Staining of Drosophila Larval Ovaries

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

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

Three-dimensional Reconstruction of the Vascular Architecture of the Passive CLARITY-cleared Mouse Ovary

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine system, but the complex structural relationships and three-dimensional (3D) vasculature architectures of the ovary are not well described. In order to visualize the 3D connections and architecture of blood vessels in the intact ovary, the first important step is to make the ovary optically clear. In order to avoid tissue shrinkage, we used the hydrogel fixation-based passive CLARITY (Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging/ Immunostaining/In situ-hybridization-compatible Tissue Hydrogel) protocol method to clear an intact ovary. Immunostaining, advanced multiphoton confocal microscopy, and 3D image-reconstructions were then used for the visualization of ovarian vessels and follicular capillaries. Using this approach, we showed a significant positive correlation (P &#60;0.01) between the length of the follicular capillaries and volume of the follicular wall.

Here we present an adaptation of the passive CLARITY and 3D reconstruction method for visualization of the ovarian vasculature and follicular capillaries in intact mouse ovaries.

The ovary is the main organ of the female reproductive system and is essential for the production of female gametes and for controlling the endocrine ...

Planarian Ovary Dissection for Ultrastructural Analysis and Antibody Staining

Summary

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Chapters in this video

Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

Upright Imaging of Drosophila Egg Chambers

Dissection and Staining of Drosophila Pupal Ovaries

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Dissection and Grading of Ovarian Development in Wild-Type Female Insects

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Microdissection and Dissociation of the Murine Oviduct: Individual Segment Identification and Single Cell Isolation

Dissection and Staining of Drosophila Larval Ovaries

Three-dimensional Reconstruction of the Vascular Architecture of the Passive CLARITY-cleared Mouse Ovary