We thank Rebecca Williams and Johanna Dela Cruz from the Cornell BRC Imaging and Andrew Recknagel for helpful suggestions and technical assistance. This work was supported to National Institutes of Health grant S10-OD018516 (to Cornell&#39;s Imaging Facility), T32HD057854 to J.C.B. and R01-GM45415 to J.C.S.

Cornell University&#39;s Institutional Animal Care and Use Committee (IACUC) has approved all the methods described here, under protocol 2004-0038 to JCS.
1. Preparation of Instruments and Culture Media
<ol>
	<li>Wipe the working area clean with 10% bleach and allow the bleach to remain on the work area surface for at least 5 min. After 5 min, remove the excess bleach with clean paper towels and 70% ethanol (EtOH). Clean the dissecting microscope thoroughly with 70% EtOH. 
	NOTE: It is important that the work area be at least semi-sterile to avoid contamination of organ cultures.</li>
	<li>Wrap clean forceps and scissors with a paper towel dampened with 70% EtOH until the time of use. 
	NOTE: The ideal dissection tools may vary according to the age/size of the ovary. Best results are obtained using one sharp micro dissecting scissor, two fine tip forceps (either number 5 or 55) and one micro dissecting iris scissor.</li>
	<li>In a conical tube, make 50 mL of ovary culture media and warm in a 37 &#176;C water bath.
	<ol>
		<li>To make ovary culture medium4, supplement 1x Minimal Essential Media (MEM) with 10% Fetal Bovine Serum (FBS), 25 mM (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), 100 units/mL penicillin, 100 &#181;g/mL streptomycin, 0.25 &#181;g/mL Amphotericin B (e.g., 45 mL of MEM, 5 mL of heat inactivated FBS, 1.25 mL of HEPES, and 0.5 mL of 100x stock of penicillin, streptomycin, and Amphotericin B).</li>
	</ol>
	</li>
	<li>Prepare plates and inserts prior to the ovary culture.
	<ol>
		<li>In a tissue culture hood, add 1 mL of ovary culture medium to a 35 mm tissue culture plate.</li>
		<li>Pre-soak insert membrane with the culture medium. Add sufficient media, usually about 0.55 mL per well, to the 24-well carrier tissue culture plate, and place a cell culture insert in the well. Make sure the insert&#39;s membrane touches the media.</li>
		<li>Prepare the number of 35 mm plates and wells with cell culture inserts according to the expected number of ovaries in the experiment. Place the 35 mm plates containing 1 mL of culture media and the 24-well carrier plate containing culture media and inserts in a 37 &#176;C, 5% CO2, and atmospheric O2 incubator.</li>
	</ol>
	</li>
	<li>Arrange a hot plate next to the dissection scope and set the temperature to 37 &#176;C. Keep a 37 &#176;C, 5% CO2 and the atmospheric O2 incubator is close to the dissection room.</li>
</ol>
2. Ovary Dissection and Organ Culture
NOTE: Ovaries can be dissected at room temperature as long as the exposure to the non-ideal temperature is minimal.
<ol>
	<li>Euthanize the female mice at post-natal day 5, by decapitation, one at a time. 
	NOTE: Euthanasia must be conducted according to IACUC guidelines present at the facility where the research is being performed.</li>
	<li>Spray the animal with 70% EtOH. Place the animal on the top of the dry paper towel under the dissection microscope. Place a 35 mm tissue culture dish containing ovary culture media close to the dissection microscope.</li>
	<li>With forceps or a micro dissection iris scissor, make an incision through the skin of the animal and into the abdominal cavity, using caution not to cut into the intestines. Push the intestines out of the abdominal cavity and locate the ovaries. Extract the ovaries and place them into the 35-mm tissue culture dish containing culture media.</li>
	<li>Once the ovaries have been dissected from the animal, increase the magnification of the dissection microscope in order to better visualize ovaries and any attached tissue. With clean fine tip forceps, remove all non-ovarian tissue, such as bursa and fatty tissue. Be careful to keep the ovaries intact when removing the attached non-ovarian tissue.</li>
	<li>Once the ovaries are isolated, place the pair into the pre-soaked cell culture inserts of the carrier 24-well plate (refer to steps 1.4.2 and 1.4.3) and adjust the volume of the medium to ensure that it is sufficient to keep the organ moist (without completely submerging it). 
	Caution: Be careful not to poke a hole in the membrane of the cell culture insert when transferring the ovaries.</li>
	<li>Place explanted organs in a 37 &#176;C, 5% CO2, and atmospheric O2 incubator.</li>
	<li>Repeat steps 2.1 - 2.6 until the ovaries of each animal are dissected and placed on the cell culture insert of the 24-well plate containing the ovary culture medium.</li>
	<li>Change the ovary culture medium every 2 days, using warmed ovary media aliquots from a 37 &#176;C water bath and proceed to Part 3 of the protocol at the desired experimental time point.</li>
</ol>
3. Tissue Fixation
<ol>
	<li>In a 15 mL conical tube, prepare 4% paraformaldehyde (PFA) in 1x phosphate-buffered saline (PBS) (e.g., add 2 mL of 16% PFA electron microscopy grade to 6 mL of 1x PBS). 
	Caution: PFA is a hazardous substance and should be handled under a chemical hood.</li>
	<li>Fix the cultured ovaries, without removing the tissue from the culture insert, by covering the tissue with the freshly prepared 4% PFA solution. Seal the edges of the plate with plastic adhesive (to avoid evaporation) and place the samples at 4 &#176;C overnight. 
	NOTE: In order to facilitate handling and tissue integrity, avoid displacing ovaries from the cell culture insert throughout the entire procedure. Cultured ovaries attached to the surface of the insert are advantageous for handling without damaging or losing the organ.</li>
	<li>Rinse the tissue 3x with 70% EtOH and either proceed to step 4.1 or store it in scintillation vials filled with 70% EtOH at 4 &#176;C until further processing. 
	NOTE: If using tissue endogenously expressing fluorescent proteins, storing the samples with 70% EtOH may greatly reduce the signal. Alternatively, tissue can be rinsed and stored in the PBS with 0.2% sodium azide (NaN3). NaN3 is, however, highly toxic and poses a serious inhalation hazard. Make the stock solution in the fume hood and handle wearing appropriate personal protective equipment such as a laboratory coat and nitrile gloves.</li>
</ol>
4. Whole Mount Immunofluorescence
<ol>
	<li>Place the fixed tissue in 1x PBS at RT for a minimum of 4 h prior to the permeabilization step 4.3.</li>
	<li>Prepare permeabilization solution and blocking solution.
	<ol>
		<li>Prepare permeabilization solution as described. To 1x PBS, add 0.2% polyvinyl alcohol (PVA), 0.1% sodium borohydride (NaBH4) solution (from 12 wt.% in 14 M sodium hydroxide (NaOH) solution), and 1.5% polyethylene glycol tert-octylphenyl ether (non-ionic surfactant detergent). In a 50 mL conical tube, add 5 mL of 10x PBS, 0.1 g of PVA, 50 &#181;L of NaBH4, and 750 &#181;L of non-ionic surfactant detergent, then add ultrapure water up to 50 mL and agitate well at room temperature until the mixture is entirely in solution.</li>
		<li>Prepare blocking solution as described. To 1x&#160;PBS, add 0.1% non-ionic surfactant detergent, 0.15% of 2.5M glycine pH 7.4, 10% normal goat serum, 3% bovine serum albumin (BSA), 0.2% NaN3, 100 units/mL penicillin, 100 &#181;g/mL streptomycin, and 0.25 &#181;g/mL Amphotericin B. Prepare a stock solution of 2.5 M glycine by dissolving 93.8 g of glycine in ultrapure water to a 500 mL final volume (pH 7.4) and a stock solution of 10% NaN3 by dissolving 5 g in 50 mL of ultrapure water; then, in a 50 mL conical tube prepare the blocking solution by adding 5 mL of 10x PBS, 50 &#181;L of non-ionic surfactant detergent, 750 &#181;L of glycine stock solution, 5 mL of goat serum, 1.5 g of BSA, 1 mL of NaN3 stock solution, and 0.5 mL of 100x stock of penicillin, streptomycin, and Amphotericin B . Add ultrapure water up to a final volume of 50 mL and syringe filter the final solution.</li>
	</ol>
	</li>
	<li>Remove and discard the 1x PBS from the vial, and then add enough permeabilization solution to completely submerge the ovaries. Place the sample in permeabilization solution on an orbital shaker (e.g., nutator) at room temperature for 4 h. 
	NOTE: Incubation time may vary according to the organ thickness.</li>
	<li>Replace the permeabilization solution with enough blocking solution to completely submerge the ovaries. Seal the vial with the plastic adhesive and leave the tissue incubating for a minimum of 12 h at room temperature on an orbital shaker.</li>
	<li>Prepare primary antibody dilution in blocking solution as per the manufacturer&#39;s datasheet. 
	NOTE: Note that not all antibodies are compatible with the clearing agent. The average volume needed per sample is approximately 750 &#181;L.
	<ol>
		<li>For oocyte quantification, use TAp63 (nuclear oocyte marker) and MVH (cytoplasmic germ cell marker). 
		NOTE: The antibodies used in the immunofluorescence images are mouse anti-p63 and rabbit anti-MVH). The primary antibody solution can be reused 2x or more if stored at 4 &#176;C between staining protocols and properly handled to avoid bacterial growth.</li>
	</ol>
	</li>
	<li>Place the insert containing the tissue in a 24-well plate and add about 750 &#181;L of primary antibody solution. Ensure that ovaries are completely submerged. Seal the plate with plastic adhesive to minimize evaporation and leave the tissue incubating for 4 days at room temperature on an orbital shaker.</li>
	<li>Prepare washing solution.
	<ol>
		<li>Prepare washing solution as described. Make 1x PBS, add 0.2% PVA and 0.15% non-ionic surfactant detergent. In a 50 mL conical tube, add 5 mL of 10x PBS, 0.1 g of PVA, 75 &#181;L of non-ionic surfactant detergent and ultrapure water up to a final volume of 50 mL.</li>
	</ol>
	</li>
	<li>Remove primary antibody dilution and add a generous amount of washing solution. Always make sure to submerge the ovaries. Allow the ovaries to soak in the solution overnight at room temperature on the orbital shaker.</li>
	<li>Replace the washing solution and incubate on the orbital shaker for 2 h or more.</li>
	<li>Repeat the step 4.9 one additional time.</li>
	<li>Prepare the secondary antibody dilution in blocking solution. 
	NOTE: The secondary antibodies used are anti-mouse 488 fluorescent dye raised in goat and anti-rabbit 594 fluorescent dye raised in goat. See step 4.5. for the average volume needed per sample.</li>
	<li>Remove washing solution from the ovaries and add secondary antibody solution. Protect the samples from light (wrap plate with aluminum foil) and seal the plate with plastic adhesive to minimize evaporation. Incubate samples on an orbital shaker at room temperature for 2 days. 
	NOTE: Continue protecting the samples from light using aluminum foil during all subsequent incubation steps.</li>
	<li>Remove secondary antibody solution from the samples and add washing solution. Incubate on an orbital shaker for 8 h. 
	NOTE: If 4,6-Diamidino-2-phenylindole (DAPI) staining is desired, add 50 ng/mL of DAPI to the first wash step (~8 h incubation).</li>
	<li>Replace the washing solution for an overnight wash.</li>
	<li>While preparing the clearing solution (see section 5), replace the overnight washing solution with fresh solution and keep the samples on the orbital shaker at room temperature.</li>
</ol>
5. Ovary Clearing and Imaging
<ol>
	<li>Prepare ScaleS(0) clearing solution.
	<ol>
		<li>Prepare ScaleS(0) clearing solution10 by adding reagents in the given proportion: 40% D-(-) sorbitol (w/v), 10% glycerol, 4.3 M urea, and 20% dimethyl sulfoxide (DMSO), pH 8.1 (e.g. in a 50 mL conical tube, add 2.5 mL of glycerol, 10 g of sorbitol, 5 mL of DMSO, and 12 mL of 9 M urea for a total volume of 25 mL. Mix solution by inversion at 50 &#176;C for 30 min and degas prior to use.)</li>
	</ol>
	</li>
	<li>Remove the washing solution and submerge the samples in clearing solution. For better results maintain samples on the orbital shaker.</li>
	<li>Replace the clearing solution 2x daily until the tissue becomes transparent (approximately 2 days).</li>
	<li>Once the tissue is cleared, prepare the sample for imaging by carefully removing the insert membrane containing the cultured ovaries with a fine tip scalpel and placing the sample on a glass slide.</li>
	<li>Proceed to imaging the samples on a microscope capable of optical sectioning.</li>
</ol>
6. Oocyte Quantification
NOTE: There are many different computer programs that are able to quantify cells. Described below is a protocol for oocyte quantification using the non-commercial image processing package, FIJI-ImageJ14.
<ol>
	<li>Using a flattened maximum intensity projection of the Z-stack image series for the specific cellular markers of interest (without the DAPI channel), convert the image into a black and white 8-bit image under Type in the Image drop-down menu.</li>
	<li>Under the Image drop-down menu, highlight the Adjust tab and then select Threshold.</li>
	<li>Manually adjust the threshold to a level that best identifies each individual oocyte. Once the level is determined, click Apply to generate a black and white image. Once completed, quantify the sample by delineating using the Oval or Freehand icon. 
	NOTE: FIJI-ImageJ has different options to fine tune the image that will be used for the particle quantification. For instance, in Process | Binary tab, there are different options that can be used to improve the detection of what will be counted. In Figure 4, the options used to better individualize follicles was Erode, followed by Fill holes, and lastly Watershed.</li>
	<li>Under the Analyze drop-down menu, select Analyze particles. Set the parameters according to the desired resolution.</li>
</ol>

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The authors have nothing to disclose.

The study of mammalian reproduction requires using and quantifying specialized cells that are not routinely amenable to cell culture. However, ex vivo culture systems are effective at maintaining ovary and follicle viability1,15. During ovary culture, the tissue requires a larger surface area for exchange of nutrients through diffusion. Therefore, 5-day old mouse ovaries are ideal in size and shape for organ culture. This protocol was optimized for ovari...

In order to study the cellular composition and morphological features of mammalian ovaries, scientists often rely on in vivo experiments followed by immunohistological staining of paraffin embedded ovaries. More recently though, whole ovary organ culture has proven to be an effective alternative to study ovarian function1,2,3,4 because the technique can be coupled with better visualization and quantification tools. Traditionally, analysis of ovarian morphology depends on reconstructing three-dimensional ovarian architecture from paraffin embedded serial sections, but, in addition to being laborious and time consuming, serially sectioning paraffin embedded tissue does not guarantee proper reconstruction of the organ, and sections are often lost or mis-ordered in the process.
In addition to the technical challenges associated with serial sectioning, there are also variations in the methods routinely used to quantify follicle numbers per ovary5,6. The methodological variability currently used impairs meta-analysis of ovarian reserve across studies5,7. For example, follicle numbers from different research articles can vary by 10-fold or more between similar developmental ages within a specific strain6. These large differences in reported follicle quantification can lead to confusion and have hindered cross-study comparisons. Experimentally, traditional approaches to follicle quantification from serial sections are performed by counting follicles of a pre-defined number of sections (e.g., every fifth, tenth, or other section). Variability in follicle counts using this approach arises not only from the periodicity in which sections are counted but also from variations in section thickness, and technical experience in generating serial sections5,6. In addition to its variability, another disadvantage of traditional tissue sectioning is that the sectioning of small ovaries from young animals is especially challenging and highly dependent on tissue orientation8.
The protocol below describes a routinely used ovary culture technique1 but greatly improves upon traditional follicle quantification by substituting physical sectioning with tissue clearing and optical sectioning using confocal microscopy8,9. Clearing using tissue immersion (without the need for transcranial perfusion or electrophoresis) in a urea- and sorbitol-based solution (e.g., ScaleS(0)10) proved compatible with the immunostaining and allowed for the reduction of clearing time without compromising depth of imaging. Other reported methods (e.g., ScaleA28,10, SeeDB11, ClearT12, and ClearT212) are either more time consuming or do not allow in-depth optical resolution of the sample. Optical sectioning is advantageous because it is less labor intensive and maintains the organ&#39;s three-dimensional architecture7,8. Another benefit of this approach is that preparation of the samples does not require costly reagents to clear the tissue and can be conducted with relative ease.
Specifically, the protocol described has been optimized for cultured mouse ovaries at postnatal day five but has been conducted on ovaries ranging from postnatal day 0 - 10. The method makes use of an ovary culture system in which the tissue naturally attaches to the membrane on which it is cultured, facilitating organ handling and manipulation. The culture system described can be used to maintain explanted ovaries for up to 10 days and to assess how different experimental conditions may interfere with oocyte survival13. The quantification procedure described is performed using the non-commercial image processing package FIJI-ImageJ14 and can be conducted on most personal computers. Furthermore, images used for quantification can be made widely available for the scientific community, thus allowing for future meta-analysis.

<table><tbody><tr><td>Micro dissecting scissor 4.5&quot;, straight, sharp points</td><td>Roboz</td><td>RS-5912</td><td>Micro dissecting scissor</td></tr><tr><td>Jewelers style forceps 4-3/8&quot;, style 5</td><td>Integra Miltex</td><td>17-305X</td><td>Fine tip forceps</td></tr><tr><td>Micro Iris Scissors 4&quot;, straight, sharp points</td><td>Integra Miltex</td><td>18-1618</td><td>Micro dissecting iris scissor or micro dissecting spring scissor</td></tr><tr><td>1X Minimal Essential Media (MEM)</td><td>ThermoFisher Scientific</td><td>11090-081</td><td></td></tr><tr><td>Fetal Bovine Serium</td><td>Corning</td><td>35011CV</td><td></td></tr><tr><td>HEPES (1M)</td><td>Gibco</td><td>15630080</td><td></td></tr><tr><td>Antibiotic-Antimycotic (100X)</td><td>Gibco</td><td>15240062</td><td>Used in Step 1.3.1</td></tr><tr><td>35mm Tissue Culture Treated Dish</td><td>Corning</td><td>430165</td><td></td></tr><tr><td>Nunc&amp;trade; 24-Well Carrier Plate for Cell Culture Inserts</td><td>ThermoFisher Scientific</td><td>141006</td><td>Pore size: 8&amp;mu;m</td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Electron Microscopy Sciences</td><td>15710</td><td>16% solution</td></tr><tr><td>1X Phosphate-Buffered Saline (PBS)</td><td>Gibco</td><td>10010023</td><td></td></tr><tr><td>Nutator mixer GyroTwister&amp;trade;</td><td>Labnet</td><td>S1000-A-B</td><td>three dimensional shaker</td></tr><tr><td>Normal Goat Serum</td><td>VWR</td><td>103219-578</td><td></td></tr><tr><td>Bovine Serum Albumen (BSA)</td><td>VWR</td><td>97061-416</td><td></td></tr><tr><td>Sodium Azide (NaN3)</td><td>Sigma-Aldrich</td><td>S2002-25G</td><td></td></tr><tr><td>Sodium borohydride solution (NaBH&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>452904-25ML</td><td>Use the solution, rather than the tablet or powder form&amp;nbsp;</td></tr><tr><td>Polyvinyl Alcohol (PVA)</td><td>Sigma-Aldrich</td><td>P8136-250G</td><td>Cold water soluble</td></tr><tr><td>Triton&amp;trade; X-100</td><td>Sigma-Aldrich</td><td>93443-100ML</td><td>polyethylene glycol tert-octylphenyl ether</td></tr><tr><td>Syringe filters</td><td>ThermoFisher Scientific</td><td>725-2520</td><td>25mm</td></tr><tr><td>10mL Syringes</td><td>BD</td><td>309695</td><td></td></tr><tr><td>Mouse anti-p63 antibody (4A4)</td><td>Biocare Medical</td><td>CM 163A</td><td>Dilution 1:500</td></tr><tr><td>Rabbit anti-MVH antibody</td><td>Abcam</td><td>ab13840</td><td>Dilution 1:600</td></tr><tr><td>Alexa Fluor&amp;reg; goat anti-mouse 594</td><td>ThermoFisher Scientific</td><td>A-11032</td><td>Dilution 1:1000</td></tr><tr><td>Alexa Fluor&amp;reg; goat anti-rabbit 488</td><td>ThermoFisher Scientific</td><td>A-11034</td><td>Dilution 1:1000</td></tr><tr><td>4,6-Diamidino-2-phenylindole (DAPI)&amp;nbsp;</td><td>ThermoFisher Scientific</td><td>62248</td><td></td></tr><tr><td>D-(-)sorbitol&amp;nbsp;</td><td>Sigma-Aldrich</td><td>240850-100G</td><td></td></tr><tr><td>Glycerol</td><td>Sigma-Aldrich</td><td>G9012-100ML</td><td></td></tr><tr><td>Urea</td><td>Sigma-Aldrich</td><td>U5378-500G</td><td></td></tr><tr><td>Dimethyl sulfoxide (DMSO)</td><td>Sigma-Aldrich</td><td>D2650-5X10ML</td><td></td></tr><tr><td>FIJI-ImageJ</td><td></td><td></td><td>Image processing software</td></tr><tr><td>Disposable plastic transfer pipettes</td><td>VWR</td><td>414044-034</td><td></td></tr></tbody></table>

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.

This protocol includes 6 major steps following dissection of the ovaries, as outlined in Figure 1. Figure 2, Figure 3, Figure 4 highlight the most novel features of this protocol, which include optimization of tissue clearing for ovaries and whole tissue oocyte quantification using FIJI-ImageJ. Figur...

Watch this Scientific Journal Video about Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries at JoVE.com

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

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, ...

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Research

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Developmental Biology

10.6K Views.  Cornell University. 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.

Video: Whole Mount Immunofluorescence and Follicle Quantification of Cultured 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 ...

Reprogramming Mouse Embryonic Fibroblasts with Transcription Factors to Induce a Hemogenic Program

This protocol details the induction of a hemogenic program in mouse embryonic fibroblasts (MEFs) via overexpression of transcription factors (TFs). We first designed a reporter screen using MEFs from human CD34-tTA/TetO-H2BGFP (34/H2BGFP) double transgenic mice. CD34+ cells from these mice label H2B histones with GFP, and cease labeling upon addition of doxycycline (DOX). MEFS were transduced with candidate TFs and then observed for the emergence of GFP+ cells that would indicate the acquisition of a hematopoietic or endothelial cell fate. Starting with 18 candidate TFs, and through a process of combinatorial elimination, we obtained a minimal set of factors that would induce the highest percentage of GFP+ cells. We found that Gata2, Gfi1b, and cFos were necessary and the addition of Etv6 provided the optimal induction. A series of gene expression analyses done at different time points during the reprogramming process revealed that these cells appeared to go through a precursor cell that underwent an endothelial to hematopoietic transition (EHT). As such, this reprogramming process mimics developmental hematopoiesis "in a dish," allowing study of hematopoiesis in vitro and a platform to identify the mechanisms that underlie this specification. This methodology also provides a framework for translation of this work to the human system in the hopes of generating an alternative source of patient-specific hematopoietic stem cells (HSCs) for a number of applications in the treatment and study of hematologic diseases.

The protocol described here details the induction of a hemogenic program in mouse embryonic fibroblasts via overexpression of a minimal set of transcription factors. This technology may be translated to the human system to provide platforms for future study of hematopoiesis, hematologic disease, and hematopoietic stem cell transplant.

This protocol details the induction of a hemogenic program in mouse embryonic fibroblasts (MEFs) via overexpression of transcription factors (TFs). We ...

Organ Culture and Whole Mount Immunofluorescence Staining of Mouse Wolffian Ducts

Tubal morphogenesis is a fundamental requirement for the development of most mammalian organs, including the male reproductive system. The epididymis, an integral part of the male reproductive tract, is responsible for sperm storage, maturation, and transport. The adult epididymis is a highly coiled tube that develops from a simple and straight embryonic precursor known as Wolffian duct (WD). Proper coiling of the epididymis is essential for male fertility, as sperm in the testis are unable to fertilize an oocyte. However, the mechanism responsible for epididymal development and coiling remains unclear, partially due to the lack of whole organ culture and imaging methods. In this study, we describe an in vitro culture system and whole mount immunofluorescence protocol to better visualize the process of WD coiling and development, which may also be applied to study other tubular organs.

We present a method for isolation and culture of the mouse Wolffian duct (WD). We also demonstrate a detailed procedure for whole mount immunostaining of cultured/freshly isolated WDs with fluorescently tagged antibodies. Together, these techniques enable the study of WD development, coiling, and differentiation.

Tubal morphogenesis is a fundamental requirement for the development of most mammalian organs, including the male reproductive system. The epididymis, an ...

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), quantitative, real-time RT-PCR (RT-qPCR) and RNA sequencing. When these techniques are performed using a single oocyte or embryo, low-copy mRNAs are not reliably detected. To overcome this problem, oocytes or embryos can be pooled together for analysis; however, this often leads to high variability amongst samples. In this protocol, we describe the use of fluorescence in situ hybridization (FISH) using branched DNA chemistry. This technique identifies the spatial pattern of mRNAs in individual cells. When the technique is coupled with Spot Finding and Tracking&#160;computer software, the abundance of mRNAs in the cell can also be quantified. Using this technique, there is reduced variability within an experimental group and fewer oocytes and embryos are required to detect significant differences between experimental groups. Commercially available branched-DNA SM-FISH kits have been optimized to detect mRNAs in sectioned tissues or adherent cells on slides. However, oocytes do not effectively adhere to slides and some reagents in the kit were too harsh resulting in oocyte lysis. To prevent this lysis, several modifications were made to the FISH kit. Specifically, oocyte permeabilization and wash&#160;buffers designed for the immunofluorescence of oocytes and embryos replaced the proprietary buffers. The permeabilization, washes, and incubations with probes and amplifier were performed in 6-well plates and oocytes were placed on slides at the end of the protocol using mounting media. These modifications were able to overcome the limitations of the commercially available kit, in particular, the oocyte lysis. To accurately and reproducibly count the number of mRNAs in individual oocytes, computer software was used. Together, this protocol represents an alternative to PCR and sequencing to compare the expression of specific transcripts in single cells.

Use of Single Molecule Fluorescent In Situ Hybridization SM-FISH to Quantify and Localize mRNAs in Murine Oocytes

To reproducibly count the numbers of mRNAs in individual oocytes, single molecule RNA fluorescence in situ hybridization (RNA-FISH) was optimized for non-adherent cells. Oocytes were collected, hybridized with the transcript specific probes, and quantified using an image quantification software.

Current methods routinely used to quantify mRNA in oocytes and embryos include digital reverse-transcription polymerase chain reaction (dPCR), ...

Genetics

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 ...

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and optical transparency. In particular, the zebrafish embryo has become an important organism to study vertebrate kidney organogenesis as well as multiciliated cell (MCC) development. To visualize MCCs in the embryonic zebrafish kidney, we have developed a combined protocol of whole-mount fluorescent in situ hybridization (FISH) and whole mount immunofluorescence (IF) that enables high resolution imaging. This manuscript describes our technique for co-localizing RNA transcripts and protein as a tool to better understand the regulation of developmental programs through the expression of various lineage factors.

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Cilia development is vital to proper organogenesis. This protocol describes an optimized method to label and visualize ciliated cells of the zebrafish.

In recent years, the zebrafish embryo has emerged as a popular model to study developmental biology due to traits such as ex utero embryo development and ...

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, ...

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.

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 ...

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 ...

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Summary

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Chapters in this video

Culture and Co-Culture of Mouse Ovaries and Ovarian Follicles

Reprogramming Mouse Embryonic Fibroblasts with Transcription Factors to Induce a Hemogenic Program

Organ Culture and Whole Mount Immunofluorescence Staining of Mouse Wolffian Ducts

Use of Single Molecule Fluorescent In Situ Hybridization (SM-FISH) to Quantify and Localize mRNAs in Murine Oocytes

In Vitro Growth of Mouse Preantral Follicles Under Simulated Microgravity

Visualizing Multiciliated Cells in the Zebrafish Through a Combined Protocol of Whole Mount Fluorescent In Situ Hybridization and Immunofluorescence

Ovarian Tissue Culture to Visualize Phenomena in Mouse Ovary

Accurate Follicle Enumeration in Adult Mouse Ovaries

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