This work was made possible through Victorian State Government Operational Infrastructure Support and Australian Government NHMRC IRIISS and supported by funding from National Health and Medical Research Council (ALW #1120300) and Australian Research Council (KJH #FT190100265). The authors would like to acknowledge the technical support of the Monash Animal Research Platform, Monash Histology Platform and Monash Micro Imaging facility.

Ovaries were collected from female C57BL6J mice. All animal procedures and experiments were performed in accordance with the NHMRC Australian Code of Practice for the Care and Use of Animals and approved by the Monash Animal Research Platform Animal Ethics Committee.
NOTE: A chemotherapy agent shown to deplete primordial follicle oocytes, as determined using stereology11 and direct counts12,13 was used in this report to compare the two counting methods in the same animal. Female, 8-week-old (young adult) mice were weighed prior to a single intraperitoneal injection of 75 mg/kg/bodyweight of cyclophosphamide, or saline vehicle control (n=5/group). This dose has been shown to cause an approximate 50% depletion of primordial follicles, but not reported to cause morbidity or mortality in mice14. Ovaries were harvested 48 hours after treatment. One ovary from each animal was fixed in 10% (v/v) neutral buffered formalin solution for 24 hours, and the other fixed in Bouin&#8217;s solution for 24 hours. Tissue was then embedded in either glycolmethacrylate resin and serially sectioned at 20 &#181;m, or in paraffin and serially sectioned at 5 &#181;m. All tissues were stained with periodic acid Schiff and haematoxylin.
1. Histological preparation: fixation, processing, embedding and sectioning mouse ovaries
<ol>
	<li>Dissect mouse ovaries by trimming the oviduct and all surrounding fat, without damaging or cutting the ovary itself. If necessary, use a dissecting microscope and fine blade for this step (Figure 1A).</li>
	<li>Fix tissues immediately by placing into a small labelled tube containing either Bouin&#8217;s fixative (stereology), or formalin fixative (direct counts) for 24 hours (Figure 1B), before transferring tissues into 70% ethanol. 
	NOTE: Follicle morphology is conserved best within Bouin&#8217;s fixed ovarian tissue, embedded into glycolmethacrylate resin (Figure 2).</li>
	<li>Process whole fixed ovaries and then embed in glycolmethacrylate resin for stereology (Supplementary File 1), or paraffin wax for direct counts using a standard histological protocol. 
	CAUTION: Resin is toxic, so ensure all tissue processing steps are performed in a fume hood and gloves are worn at all times.</li>
	<li>Use a specialized resin methacrylate microtome (Figure 1C) fitted with a glass knife (Figure 1D) to exhaustively cut thick glycolmethacrylate resin sections (e.g., 20 &#181;m). Collect the sections at a regular interval (e.g., every 3rd section) onto glass microscope slides for stereology.</li>
	<li>Use a standard microtome to cut thin paraffin sections (e.g., 4-6 &#181;m). Collect tissue sections at a regular interval (e.g., every 9th section) onto a glass microscope slide for direct follicle counts.</li>
	<li>Stain the slides with periodic acid Schiff and haematoxylin (Supplementary File 2).</li>
	<li>Coverslip with standard DPX for paraffin sections, or thick DPX for glycolmethacrylate resin sections (Figure 1E). 
	CAUTION: Glycolmethacrylate resin DPX is hazardous, so perform this step in to fume hood. 
	NOTE: Glycolmethacrylate resin DPX is extremely viscous. The glass coverslip must be adhered firmly by pressing it down with a spatula to ensure the DPX is evenly and thinly dispersed, and there are no air bubbles present under the coverslip (Figure 1F).</li>
</ol>
2. Stereological estimation of primordial follicle number using the optical fractionator
<ol>
	<li>Turn on the computer, the multi-control unit, the camera and the light source within the stereology setup and set the microscope objective to a low magnification (e.g., 10x).</li>
	<li>Open the stereology software.</li>
	<li>Put the first slide securely on the microscope stage.</li>
	<li>Adjust the light exposure by checking Automatic under Exposure in the Camera Settings panel (Supplementary Figure 1A). Alternatively, manually adjust the light exposure.</li>
	<li>Use the joystick to locate the first tissue sample and bring the sample into focus.</li>
	<li>Adjust the white balance, either by clicking on More Settings (located bottom right of the Camera Settings panel), and then click White Balance and click on Automatic (Supplementary Figure 1B). Alternatively, click the White Balance button adjacent to the More Settings button (or Select Area in More Settings), to set the white balance manually by selecting a white area on the section.</li>
	<li>Go to the Probes drop down menu and click on Optical Fractionator Workflow. Then click Start New Project and click OK.
	<ol>
		<li>If an existing sampling configuration has been saved, under Sampling Parameters click Yes | &#8230; and select the desired sampling configuration.</li>
		<li>If not, click No and manually enter the serial section information (Supplementary Figure 1C) and define the probe configuration at step 2.13.</li>
	</ol>
	</li>
	<li>Click on Next, set the microscope to Low Magnification and choose 10x magnification from the dropdown menu.</li>
	<li>Click on Next, and then trace around the entire ovarian section &#8211; start by left clicking around the section and at the end, right click and choose Close Contour.</li>
	<li>Click on Next, set the microscope to High Magnification and choose 100x Oil magnification from the dropdown menu.</li>
	<li>Place a drop of oil on the tissue section on the slide and move the microscope objective to 100x magnification.</li>
	<li>Adjust the light exposure (as in step 2.4) and click Next.</li>
	<li>Set up the Sampling Parameters to define the probe configuration. Here, the counting frame was set to 47.5 &#181;m x 47.5 &#181;m (2,256.25 &#181;m2) and the step length was set to 100 &#181;m x 100 &#181;m (10,000 &#181;m2) (Supplementary Figure 2). Once the sampling parameters are established, save the template and re-open during subsequent analysis sessions at step 2.7.</li>
	<li>Click on Start Counting (Supplementary Figure 1D). Focus to the top of the sample, click OK and begin counting.</li>
	<li>Use the focusing knob to move through the 10 &#181;m sampling depth and count any primordial follicles whose oocyte nucleus comes into focus. Click Next to move to the next area.
	<ol>
		<li>Classify follicles as a primordial if the oocyte is surrounded by squamous (flattened) granulosa cells, but no cuboidal granulosa cells (Figure 2A). 
		NOTE: Primordial follicles are distinct from intermediate/transitional follicles, which comprise a combination of cuboidal and squamous granulosa cells (Figure 2B), and primary follicles, which are surrounded predominantly by cuboidal granulosa cells (Figure 2C). These follicle classes should be quantified separately.</li>
		<li>Only count follicles in which the oocyte nucleus is visible. The oocyte nucleus must appear within the counting frame or be touching the green inclusion lines of the counting frame (Supplementary Figure 1E,F).</li>
		<li>If the oocyte nucleus falls outside the counting frame (Supplementary Figure 1G) or touches the red exclusion lines of the counting frame, do not count this follicle.</li>
		<li>When assessing primordial follicle depletion in response to an exogenous chemical (e.g., chemotherapy), ensure all follicles counted are healthy and thus have normal morphology (Figure 2). Count any abnormal or atretic follicles separately. Often, follicle death is induced by insults such as chemotherapy, and quantification of the atretic follicles should be obtained separately in order to distinguish between healthy and atretic follicles, as only healthy follicles comprise the ovarian reserve15.</li>
	</ol>
	</li>
	<li>Once counting is complete on that section, do one of the following:
	<ol>
		<li>Click Add New Section, and then return to step 2.3 to set up the next section for counting.</li>
		<li>Click I&#8217;ve Finished Counting to end the session. Return the objective to 10x, exit the stereology software and turn off the light source, camera, multi-control unit and computer.</li>
	</ol>
	</li>
	<li>Obtain the sum raw follicle number (Q) from each tissue section sampled from the entire ovary, then using a spreadsheet, use the equation below to obtain the final value from each replicate animal (N)4. 
	<img alt="Equation 1" src="/files/ftp_upload/61782/61782equ01.jpg" />, where: 
	N = Total estimated number of follicles within the ovary. 
	Q = Raw primordial follicle count. 
	f1 = Sampling interval. Every 3rd section was sampled thus <img alt="Equation 2" src="/files/ftp_upload/61782/61782equ02.jpg" />. 
	f2 = Relationship between the counting frame (sample area) and stepper, calculated as <img alt="Equation 3" src="/files/ftp_upload/61782/61782equ03.jpg" />. Since the sample area was 2256 &#181;m2 (47.5 &#181;m x 47.5 &#181;m) and the stepper area was 10000 &#181;m2 (100 &#181;m x 100 &#181;m), <img alt="Equation 4" src="/files/ftp_upload/61782/61782equ04.jpg" />. 
	f3 = Fraction of ovarian section sampled. Since 10 &#181;m of the 20 &#181;m section was sampled, <img alt="Equation 5" src="/files/ftp_upload/61782/61782equ05.jpg" />. 
	Therefore, <img alt="Equation 6" src="/files/ftp_upload/61782/61782equ06.jpg" />. 
	NOTE: This protocol describes how to apply these principles of stereological analyses using a widely cited stereology software (Table of Materials); however, other stereological software is available. The principles applied during stereological analyses of ovarian follicles are the same, regardless of the software used to set up the parameters. Stereology is most accurate when 100 or more objects are counted in an adult mouse ovary4, as this gives a coefficient of error for the estimate below 10%16. The sampling parameters outlined here have been optimized to ensure a minimum of approximately 100 objects (i.e., primordial, transitional and primary follicles) can be counted in control adult wild-type C57BL6J ovaries. A pilot study can be conducted including a small sample size to establish the optimal sampling parameters, such as the interval and number of sections to be analyzed and the number of optical dissectors within the sampled sections17.</li>
</ol>
3. Estimation of primordial follicle number by direct ovarian follicle counts
<ol>
	<li>Place the microscope slide securely under a standard light microscope and perform direct counts to obtain raw primordial follicle number.
	<ol>
		<li>Classify follicles as a primordial if the oocyte is clearly visible and is surrounded by squamous (flattened) granulosa cells, but no cuboidal granulosa cells (Figure 2D). 
		NOTE: Primordial follicles are distinct from intermediate/transitional follicles, which comprise a combination of cuboidal and squamous granulosa cells (Figure 2E), and primary follicles, which are surrounded predominantly by cuboidal granulosa cells (Figure 2F). These follicle classes should be quantified separately.</li>
		<li>Ensure all follicles counted are healthy and thus have normal morphology (Figure 2). Count any abnormal or atretic follicles separately, as only healthy follicles comprise the ovarian reserve.</li>
	</ol>
	</li>
	<li>Alternatively, take multiple, or stitched high-power (e.g., 20x) photomicrographs of the entire ovarian tissue section to perform counts by opening the image file(s). This can be done manually, or using an automated slide scanner.</li>
	<li>Obtain the sum raw follicle number (Q) from each tissue section sampled from the entire ovary at the predetermined interval. Multiply this number by the inverse of the sampling fraction to obtain the final value for each replicate animal (N), using the equation below. 
	<img alt="Equation 7" src="/files/ftp_upload/61782/61782equ07.jpg" />, where: 
	N = Total estimated number of follicles within the ovary. 
	Q = Raw follicle count (of each individual type, calculated separately). 
	f1 = Sampling interval. Every 9th section was sampled thus <img alt="Equation 8" src="/files/ftp_upload/61782/61782equ08.jpg" />. 
	Therefore, <img alt="Equation 9" src="/files/ftp_upload/61782/61782equ09.jpg" />.</li>
</ol>

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The authors have nothing to disclose.

This article provides a step-by-step protocol for the gold standard technique for enumerating mouse primordial follicles, stereology, and the more commonly employed method of direct follicle counting. Chemotherapy treatment was used to compare and contrast the results obtained from these two different methods within the left and right ovary from the same animal. Both methods revealed high inter-animal variability in primordial follicle numbers. A significant depletion of the ovarian reserve was recorded using stereology,...

The immature, meiotically-arrested oocytes stored within primordial follicles in the ovary are the storage unit for the female germline and comprise an individual&#8217;s lifetime ovarian reserve. Primordial follicle numbers decline naturally with age1, or alternatively, can be prematurely depleted following exposure to exogenous chemicals, including some pharmaceuticals and environmental toxicants in air, food and water2. Given that the primordial follicle number is finite, the quantity and quality of follicles present within the ovary largely determine female fertility and offspring health. Thus, accurate quantification of primordial follicle number in women is essential for evaluating the off-target impacts of exogenous insults on the ovarian reserve.
In women, analysis of the whole ovary is generally not possible, thus non-invasive surrogate measures of the ovarian reserve must be utilized in a clinical setting. Anti-M&#971;llerian hormone (AMH) is the most widely used surrogate biomarker clinically3. Serum AMH levels are often measured in women of advanced maternal age, or before and after cancer treatment, such as chemotherapy. However, AMH is produced by growing follicles and not by primordial follicles, and thus, serum levels do not inform on absolute primordial follicle number.
With the absence of methods to accurately determine primordial follicle number in women in situ, counting ovarian follicles in small animal models, such as rodents, remains an essential research tool to assess the degree by which exogenous insults impact on primordial follicles and thus, fertility. Unfortunately though, reports throughout the literature of primordial follicle numbers in rodent models are highly variable4. A major reason for this is widely reported technical differences in the counting method employed. Predominately, there are two different techniques described in the literature for enumerating primordial follicles in mice. These include stereology, which employs the fractionator optical dissector method, and direct follicle counts.
Stereology is widely regarded the gold standard as it uses systematic uniform random sampling5, making it the most accurate method of quantifying primordial follicle number in whole mouse, or rat ovaries4,6,7. Stereology is unbiased, as it accounts for the three-dimensional structure of the object of interest8. Using an optical dissector/fractionator method, three levels of sampling are applied to quantify primordial follicles using thick tissue sections (e.g., 20 &#181;m) within a known fraction of the total mouse ovary. Firstly, the sampling interval is chosen (e.g., every 3rd section) at a random start (sampling fraction 1, f1)4. Sections are then sampled in a systematic, uniform manner from this point through the whole ovary. Then, an unbiased counting frame is superimposed over the ovarian section and progressively moved along a defined, randomized counting grid (sampling fraction 2, f2)8. Lastly, a known fraction of the section thickness is optically sampled (e.g., 10 &#181;m) and follicles within this area are counted (sampling fraction 3, f3)4. The raw follicle number is multiplied by the inverse of these sampling fractions to obtain the final value. This method requires expert training and equipment, including a microscope with a motorized stage driven by stereological software. Tissues should be preserved in a specialized Bouin&#8217;s fixative, and embedded in glycolmethacrylate resin to allow for thick tissue sections to be cut using a glycolmethacrylate resin microtome with a glass knife. This method is designed to account for tissue shrinkage and deformation, to best preserve the three-dimensional morphological structure of the ovary and follicles9.
Direct follicle counting is the most frequently used method for counting follicles10. More common fixatives (i.e., formalin) can be used, followed by paraffin wax embedding and exhaustive serial sectioning using a standard microtome at a thickness of between 4-6 &#181;m. Follicles are systematically counted in the entire tissue section at a defined interval, and then multiplied by the inverse of the sampling interval to obtain the total follicle estimate. This method is quick, easy, can be performed using archived tissues, and prepared using standard histological techniques. It requires only a light microscope with standard imaging capabilities. However, despite these advantages, direct follicle counting lacks the accuracy and strict counting parameters of stereology, making it more prone to investigator bias. Additionally, tissues may undergo shrinkage and deformation during processing, disrupting the integrity and morphology of the ovary and thus making follicle classification and quantification difficult.
The aim of this article is to describe two commonly-used methods to quantitatively assess primordial follicle number in mouse ovaries: stereology and direct follicle counting. We will provide detailed protocols for these two methods and highlight some of their strengths and weaknesses, in order to enhance reproducibility in our field and allow researchers to make an informed decision of the most appropriate counting method for their studies.

<table><tbody><tr><td>1-Butanol (HPLC)</td><td>Fisher Chemical</td><td>#A383-1</td><td></td></tr><tr><td>Acid alcohol</td><td>Amber Scientific</td><td>#ACDL</td><td></td></tr><tr><td>Bouin&amp;rsquo;s fixative</td><td>Sigma-Aldrich</td><td>#HT10132</td><td>Picric acid 0.9% (w/v), formaldehyde 9% (v/v), acetic acid 5% (w/v)</td></tr><tr><td>Cyclophosphamide</td><td>Sigma-Aldrich</td><td>#C0768-5G</td><td></td></tr><tr><td>Dibutylphthalate Polystyrene Xylene (DPX)</td><td>Sigma-Aldrich</td><td>#06522</td><td></td></tr><tr><td>Ethanol</td><td>Amber Scientific</td><td>#ETH</td><td>Ethanol 100%</td></tr><tr><td>Micro Feather opthalmic scalpel with aluminium handle</td><td>Designs for Vision</td><td>#FEA-745-SR</td><td>Feather blade for dissections (seen in Figure 1)</td></tr><tr><td>Formalin fixative</td><td>Australian Biostain</td><td>#ANBFC</td><td></td></tr><tr><td>Glass coverslip</td><td>Thermo Scientific</td><td>#MENCS22501GP</td><td>22 mm x 50 mm</td></tr><tr><td>Glycomethacrylate resin RM2165 microtome</td><td>Leica Microsystems</td><td>#RM2165</td><td></td></tr><tr><td>Glycolmethacrylate DPX</td><td>*made in house</td><td></td><td>*Mix 1.5 L Xylene; 800 g polystyrene pellets; 100mL Dibutyl phthalate for 3 weeks</td></tr><tr><td>Histolene</td><td>Trajan</td><td>#11031</td><td></td></tr><tr><td>Mayer&amp;rsquo;s haematoxylin</td><td>Amber Scientific</td><td>#MH</td><td></td></tr><tr><td>Olympus BX50 microscope</td><td>Olympus</td><td>#BX50</td><td>Brightfield microscope fitted with 10x dry &amp;amp; 100x oil immersion objective (numerical aperture 1.3)</td></tr><tr><td>Olympus immersion oil type-F</td><td>Olympus</td><td>#IMMOIL-F30CC</td><td></td></tr><tr><td>Olympus TH4-200 light source</td><td>Olympus</td><td>#TH4-200</td><td></td></tr><tr><td>Paraffin wax</td><td>Sigma-Aldrich</td><td>#03987</td><td></td></tr><tr><td>Periodic acid</td><td>Trajan</td><td>#PERI1%</td><td>Periodic acid 1%</td></tr><tr><td>Rotary Microtome CUT 4060</td><td>MicroTec</td><td>#4060R/F</td><td>Used to cut paraffin sections</td></tr><tr><td>Schiff&amp;rsquo;s reagent</td><td>Trajan</td><td>#SCHF</td><td></td></tr><tr><td>Scott&#039;s tap water</td><td>Amber Scientific</td><td>#SCOT</td><td>Potassium carbonate, magnesium sulphate, water</td></tr><tr><td>StereoInvestigator Stereological System</td><td>MBF Bioscience</td><td></td><td>Includes StereoInvestigator software, multi-control unit, automatic stage and joystick</td></tr><tr><td>Superfrost microscope slides</td><td>Thermo Scientific</td><td>#MENSF41296SP</td><td>1 mm, 72 pcs</td></tr><tr><td>Technovit 7100 Plastic embedding system</td><td>Emgrid Australia</td><td>#64709003</td><td>500 mL/5 x 1 g/40 mL</td></tr><tr><td>Technovit 3040 yellow</td><td>Emgrid Australia</td><td>#64708805</td><td>100 g/80 mL</td></tr></tbody></table>

accurate follicle enumeration in adult mouse ovaries

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial follicles, the storage unit of the female germline. They are non-renewable, thus their number at birth and subsequent rate of loss largely dictates the female fertile lifespan. Accurate quantification of primordial follicle numbers in women and animals is essential for determining the impact of medicines and toxicants on the ovarian reserve. It is also necessary for evaluating the need for, and success of, existing and emerging fertility preservation techniques. Currently, no methods exist to accurately measure the number of primordial follicles comprising the ovarian reserve in women. Furthermore, obtaining ovarian tissue from large animals or endangered species for experimentation is often not feasible. Thus, mice have become an essential model for such studies, and the ability to evaluate primordial follicle numbers in whole mouse ovaries is a critical tool. However, reports of absolute follicle numbers in mouse ovaries in the literature are highly variable, making it difficult to compare and/or replicate data. This is due to a number of factors including strain, age, treatment groups, as well as technical differences in the methods of counting employed. In this article, we provide a step-by-step instructional guide for preparing histological sections and counting primordial follicles in mouse ovaries using two different methods: [1] stereology, which relies on the fractionator/optical dissector technique; and [2] the direct count technique. Some of the key advantages and drawbacks of each method will be highlighted so that reproducibility can be improved in the field and to enable researchers to select the most appropriate method for their studies.

A well-characterized model of follicle depletion was used, whereby young adult female mice were administered a single dose of cyclophosphamide chemotherapy, or saline vehicle control (n=5/group) and both ovaries were harvested from each animal after 48 hours. One ovary per animal was prepared as described in Step 1 for each of the two methods: stereology or direct counts. The left and right ovary from each animal was randomly assigned to each group. These data show that when using stereology, a significant depletion of m...

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Accurate Follicle Enumeration in Adult Mouse Ovaries

Here, we describe, compare, and contrast two different techniques for accurate follicle counting of fixed mouse ovarian tissues.

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial ...

accurate-follicle-enumeration-in-adult-mouse-ovaries

Universidad de Cartagena UDEC

Research

JoVE Journal

Developmental Biology

8.1K Views.  Monash University. Here, we describe, compare, and contrast two different techniques for accurate follicle counting of fixed mouse ovarian tissues.

Video: Accurate Follicle Enumeration in Adult Mouse Ovaries

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 ...

Retrieval of Mouse Oocytes

To date, only a few studies have reported successful manipulations of Peromyscus embryogenesis or reproductive biology.  Together with the Peromyscus Genetic Stock Center (http://stkctr.biol.sc.edu), we are characterizing the salient differences needed to develop this system.  A primary goal has been to optimize oocyte/early embryo retrieval.

This protocol illustrates the technique for extracting oocytes or early-stage fertilized embryos from the oviduct of mice. The ability to identify the infindibulum and insert a blunt end needle into it is essential to correctly performing the procedure.

To date, only a few studies have reported successful manipulations of Peromyscus embryogenesis or reproductive biology.  Together with the Peromyscus ...

Biology

In Vitro Growth of Mouse Preantral Follicles Under Simulated Microgravity

14 day-old mouse ovarian tissue and preantral follicles isolated from same-aged mice were incubated in a simulated microgravity culture system. We quantitatively assessed follicle survival, measured follicle and oocyte diameters, and examined ultrastructure of the oocytes produced from the system. We observed decreased follicle survival, downregulation of expressions of proliferating cell nuclear antigen and growth differentiation factor 9, as indicators for the development of granulosa cells and oocytes, respectively, and oocyte ultrastructural abnormalities under the simulated microgravity condition. The simulated microgravity experimental setup needs to be optimized to provide a model for investigation of the mechanisms involved in the oocyte/follicle in vitro development.

In Vitro Growth of Mouse Preantral Follicles Under Simulated Microgravity

A highly promising technique to generate tissue constructs without using matrix is to culture cells in a simulated microgravity condition. Using a rotary culture system, we examined ovarian follicle growth and oocyte maturation in terms of follicle survival, morphology, growth, and oocyte function under the simulated microgravity condition.

14 day-old mouse ovarian tissue and preantral follicles isolated from same-aged mice were incubated in a simulated microgravity culture system. We ...

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 ...

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ovarian fitness is often assessed by visualizing and quantifying follicles and oocytes. Because the ovary is an opaque three-dimensional tissue, traditional approaches require laboriously slicing the tissue into numerous serial sections in order to visualize cells throughout the entire organ. Furthermore, because quantification by this method typically entails scoring only a subset of the sections separated by the approximate diameter of an oocyte, it is prone to inaccuracy. Here, a protocol is described that instead utilizes whole organ tissue clearing and immunofluorescence staining of mouse ovaries to visualize follicles and oocytes. Compared to more traditional approaches, this protocol is advantageous for visualizing cells within the ovary for numerous reasons: 1) the ovary remains intact throughout sample preparation and processing; 2) small ovaries, which are difficult to section, can be examined with ease; 3) cellular quantification is more readily and accurately achieved; and 4) the whole organ imaged.

Here, we present a protocol to quantify follicles in cultured ovaries of young mice without serial sectioning. Using whole organ immunofluorescence and tissue clearing, physical sectioning is replaced with optical sectioning. This method of sample preparation and visualization maintains organ integrity and facilitates automated quantification of specific cells.

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ...

Ovarian Tissue Culture to Visualize Phenomena in Mouse Ovary

Mammalian females periodically ovulate an almost constant number of oocytes during each estrus cycle. To sustain such regularity and periodicity, regulation occurs at the hypothalamic-pituitary-gonadal axis level and on developing follicles in the ovary. Despite active studies, follicle development mechanisms are not clear because of the several steps involved from the dormant primordial follicle activation to ovulation, and because of the regulation complexity that differs at each follicular stage. To investigate the mechanisms of follicle development, and the dynamics of follicles throughout the estrus cycle, we developed a mouse ovarian tissue culture model that can be used to observe follicle development using a microscope. Systematic follicle development, periodical ovulation, and follicle atresia can all be reproduced in the cultured ovary model, and the culture conditions can be experimentally modulated. Here, we demonstrate the usefulness of this method in the study of the regulatory mechanisms of follicle development and other ovarian phenomena.

Ovarian tissue cultures can be used as models of follicle development, ovulation, and follicle atresia and indicate regulatory mechanisms of dynamic ovarian processes.

Mammalian females periodically ovulate an almost constant number of oocytes during each estrus cycle. To sustain such regularity and periodicity, ...

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 ...

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 ...

Developing an Ovariectomized Mouse Model with Elevated FSH Level

Exploring Independent Effects of Follicle-Stimulating Hormone In Vivo in a Mouse Model

During the transition from a reproductive to a nonreproductive phase (menopause), many women experience significant physiological and pathological changes, including decreased bone mass, increased blood lipids, and increased visceral adiposity. Levels of follicle-stimulating hormone (FSH) rise during the menopausal transition. Many studies have shown that FSH in various extragonadal tissues and organs is associated with the pathogenesis of multiple diseases. Thus, building an animal model that can help study the independent effects of FSH in vivo is particularly important. In this study, C57BL/6 female mice were ovariectomized and supplemented with estradiol valerate (OVX + E2) to eliminate the effect of the hypothalamic-pituitary-gonadal axis. The OVX + E2 mice received solvent (N.S.) or different doses of recombinant FSH via intraperitoneal injection to create a mouse model (OVF) characterized by relatively stable estrogen and rising FSH levels. Thus, we successfully generated an experimental mouse model to mimic the early stage of menopause transition, characterized by elevated serum FSH levels. The OVF model has the advantages of being stable, low cost, and easy to operate, which is suitable for studies to explore the extragonadal actions of FSH. Here, we describe detailed protocols for the mouse OVF model.

Author Spotlight: Investigating the Relationship Between FSH and Pathophysiological Changes in Perimenopausal Women - Insights from a Mouse Model 

Follicle-stimulating hormone (FSH) in various extragonadal tissues and organs is associated with the pathogenesis of multiple diseases. The ovariectomized and FSH-treated mouse model (OVF) can be used to explore the extragonadal actions of FSH.

During the transition from a reproductive to a nonreproductive phase (menopause), many women experience significant physiological and pathological ...

Intact Ovary Harvest from Mouse: A Surgical Procedure to Isolate Complete Ovary from Murine Model

Before beginning the procedure, confirm the appropriate level of sedation by toe pinch, and apply ointment to the animal&#39;s eyes. Place the mouse in the supine position on a stack of heated paper towels, and starting just below the ribs and moving distally, use number 40 blade hair clippers to remove a 2 to 3-centimeter square of hair a few millimeters lateral to the midline on each side of the animal, with a 1 to 2 2-centimeter strip of hair covering the midline.
After administering analgesia, transfer the animal onto a heated sterile surgical field in the supine position, and use a 2 by 2 centimeter piece of 70% ethanol soaked gauze followed by a betadine scrub to disinfect the surgical area. Next, make a 1 to 1.5-centimeter paralumbar incision in one of the surgical areas, and two scissors to blunt dissect the skin from the underlying fascia.
Then, make another incision through the fascia, and blunt dissect laterally under the fascia as superficially as possible until the abdominal cavity is reached. Place a 2 by 2 centimeter gauze surgical drape over the incision, and wet the gauze with sterile saline.
Using small forceps, locate the adipose tissue that surrounds the ovary, and gently extract the fat pad and ovary onto the drape. The ovarian fat pad will appear whiter than the surrounding adipose tissue. Place a 1.5 inch Bulldog clamp onto the fat pad, taking care not to obstruct the ovarian bursa, to hold the tract in place during the procedure.
Use a dissecting microscope to identify the ovarian bursa. Then, grasp the bursa with two watchmaker forceps and incise the bursa on the opposite side from the ovarian hilum to expose the ovary.
Before deciding where to tear open the bursa, try to visualize a pocket in the bursa where you will place the new ovary prior to suturing. Only open the bursa as far as necessary to remove the endogenous ovary.
Gently remove the ovary from the bursa, and clamp the ovarian hilum with watchmaker forceps to excise the ovary and to prevent bleeding. Then, place the donor ovary in a watch glass containing 4 degrees Celsius sterile saline, and place a loose cover over the glass.

Accurate Follicle Enumeration in Adult Mouse Ovaries

Summary

Explore More Videos

Chapters in this video

Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

Retrieval of Mouse Oocytes

In Vitro Growth of Mouse Preantral Follicles Under Simulated Microgravity

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

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Ovarian Tissue Culture to Visualize Phenomena in Mouse Ovary

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

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

Exploring Independent Effects of Follicle-Stimulating Hormone In Vivo in a Mouse Model

Intact Ovary Harvest from Mouse: A Surgical Procedure to Isolate Complete Ovary from Murine Model