We thank Cara F&#228;rber for proofreading; Katharina Wollersheim, Martin Mahlberg, Lea Korte, and Jasmin Rebholz for their technical assistance.

The study was approved by the ethics committee at University Hospital Bonn (007/09). Written informed consent was obtained from each patient. The study group included human ovarian tissue from 50 patients with an average age of 27.4 years prior to cryopreservation, as indicated in Figure 1. The reagents and equipment used in this study are listed in the Table of Materials.
1. Preparation of loading devices
<ol>
	<li>Prepare surgical scissors and metal meshes for customization. Cut the meshes into strips measuring 25 mm x 8 mm, as indicated in Figure 2A.</li>
	<li>Place the customized metal meshes in a sterilization container and autoclave for 2 h (Figure 2B). After autoclaving, place the sterilization container and 1.8 mL vials under the laminar flow bench.</li>
	<li>Turn on the ultraviolet (UV) irradiation of the laminar flow bench for 30 min for additional sterilization at 254 nm wavelength.</li>
	<li>Open the sterilization container, remove the metal grids, and fit them into the caps of the 1.8 mL vials, as indicated in Figure 2C. Close the 1.8 mL vials. 
	NOTE: All tissue culture work described above should be performed in a Class II laminar flow hood. Clean the laminar flow bench with a surface disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
</ol>
2. Preparation of vitrification media
<ol>
	<li>Prepare one serological 10 mL pipette, one electronic pipette aid, and three 50 mL tubes for Vitrification Solution 1 (VS1), Vitrification Solution 2 (VS2), and Vitrification Solution 3 (VS3), as well as for recording the date of preparation. Turn on the ultraviolet (UV) radiation of the laminar flow bench for additional sterilization.</li>
	<li>For VS1 (15 mL), pipette 12 mL of oocyte handling medium (supplemented with human serum albumin by the manufacturer), 1.5 mL of SSS (Serum Substitute Supplement), and 1.5 mL of ethylene glycol into one 50 mL tube.</li>
	<li>For VS2 (15 mL), pipette 10.5 mL of oocyte handling medium (supplemented with human serum albumin by the manufacturer), 3 mL of ethylene glycol, and 1.5 mL of SSS into one 50 mL tube.</li>
	<li>For VS3, pipette 8.5 mL of oocyte handling medium (supplemented with human serum albumin by the manufacturer), 5.25 mL of ethylene glycol, and add 2.57 g of sucrose and 0.75 g of Polyvinylpyrrolidone (PVP) into one 50 mL tube. See Figure 3 for details. Mix all solutions by vortexing at 3,000 rpm. 
	NOTE: All tissue culture work described above should be performed in a Class II laminar flow hood. Clean the laminar flow bench with a disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
</ol>
3. Vitrification preparation
<ol>
	<li>Clean the laminar flow bench with a surface disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
	<li>Prepare one 6-well plate, a cell strainer, a scalpel, tweezers, one 2 mm biopsy punch, and two 90 mm round-bottom dishes. Also, prepare media: preservation solution for tissue transplantation and VS1-3. Pipette 5 mL of each chilled solution (VS1-3) into separate wells and wait 30 min for the solutions to reach room temperature.</li>
	<li>Sterilize liquid nitrogen with an automated UV irradiation system; alternatively, use the UV irradiation of the laminar flow hood for 120 min.</li>
</ol>
4. Vitrification of tissue
<ol>
	<li>Prepare ovarian cortex tissue by removing the medulla. Since the cortex is usually harder than the medulla, they can be easily distinguished and separated.
	<ol>
		<li>Cut the ovarian cortex tissue into the desired shapes (10 mm x 5 mm for transplantation; 2 mm x 2 mm punches for viable follicle counting prior to and after vitrification), as indicated in Figure 4A. 
		NOTE: Avoid damaging the tissue by pressing too hard, and handle it carefully. Tissue samples may vary between patients.</li>
	</ol>
	</li>
	<li>Place a cell strainer into the first well of the 6-well plate with VS1. Equilibrate the ovarian cortex tissue for 5 min in 5 mL of VS1 in well 1 of the 6-well plate using the cell strainer.
	<ol>
		<li>Then, move the cell strainer with the cortex tissue to well 2 containing 5 mL of VS2, and equilibrate for 5 min. Finally, equilibrate for 6 min in 5 mL of VS3, as indicated in Figure 3 and Table 1.</li>
	</ol>
	</li>
	<li>Open the prepared cryovials. Fill the vials with sterilized liquid nitrogen and place them into the cryo dewar vessel filled with liquid nitrogen. Load the tissue samples onto the metal grid of the vitrification device within 1 min (Figure 4B,C).
	<ol>
		<li>Insert the tissue samples loaded on the metal grid into the liquid nitrogen in the prepared grid-based cryovials (Figure 4D). 
		NOTE: Rapid freezing of the tissue loaded on the metal grid is essential for the success of this method. All tissue culture work described above should be performed in a Class II laminar flow hood. Clean the laminar flow bench with a surface disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
	</ol>
	</li>
</ol>
5. Preparation of rapid warming media
<ol>
	<li>Prepare the solutions as noted in Figure 5 and Table 2. Weigh the sucrose for the rapid warming solution (RWS) and the equilibriation solution (ES) and transfer it to the tube. Add the oocyte handling medium (supplemented with human serum albumin by the manufacturer) using a serological pipette and a pipetting aid. Add SSS and sucrose as listed in Table 2.</li>
	<li>Close the tubes and let them agitate on a rolling shaker until fully dissolved. 
	NOTE: All tissue culture work described above should be performed in a Class II laminar flow hood. Clean the laminar flow bench with a surface disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
</ol>
6. Rapid warming preparation
<ol>
	<li>One day before rapid warming, prepare the consumables and decontaminate them using UV light in the laminar flow bench: Prepare the sample beaker for RWS, the 6-well plate for ES, RS1, and RS2, and the sample beaker for tissue transportation after rapid warming. Place sterile, disposable tweezers, pipetting equipment, and other consumables in the laminar flow bench.</li>
	<li>Preheat a heating plate to 37.2 &#176;C. Incubate RWS at 37 &#176;C for at least 1 h.</li>
	<li>Prepare a sterile 6-well plate with 6 mL of ES, 6 mL of RS for &#34;RS1,&#34; and another 6 mL of RS for &#34;RS2&#34; in the laminar flow bench. Refer to Figure 5 for a schematic illustration of the 6-well plate. Let these incubate at room temperature for 1 h before warming the tissue.</li>
	<li>Transfer RWS to a sterile sample beaker under sterile conditions and place it on the heating plate. The RWS should be maintained at 37 &#176;C. 
	NOTE: Rapid warming requires a fast warming rate. It is crucial to hold the RWS at 37 &#176;C to ensure the success of the first step.</li>
</ol>
7. Rapid warming of ovarian tissue
<ol>
	<li>Transport the cryopreserved vials containing vitrified ovarian cortical tissue in liquid nitrogen to the laminar flow bench. Open the vials while they are partially submerged under liquid nitrogen. Rapidly submerge the mesh with the vitrified tissue into the RWS and let the tissue sit there for 1 min at 37 &#176;C. 
	NOTE: This is the most critical step during rapid warming and must be conducted without any delay.</li>
	<li>Using sterile forceps, transfer the tissue to the ES and incubate it for 3 min while gently shaking it on a rocking shaker.</li>
	<li>Rinse the tissue at room temperature for 10 min each with RS1 and RS2 on a rocking shaker, as indicated in Figure 5.</li>
	<li>Transfer the warmed tissue to a sterile sample beaker containing 5 mL of preservation solution for tissue transplantation, which should be held at 4 &#176;C for transportation to the transplantation site. 
	NOTE: All tissue culture work described above should be performed in a Class II laminar flow hood. Clean the laminar flow bench with a surface disinfectant while wearing disposable laboratory gloves; non-powdered gloves are recommended.</li>
</ol>
8. Determination of follicular viability
<ol>
	<li>Bring dimethyl sulfoxide (DMSO) to room temperature overnight to obtain a liquid state. Pipette 100 &#181;L of DMSO into one vial of calcein and resuspend to dissolve the calcein. Pipette 3 &#181;L of the dissolved calcein into the bottom of one 1.5 mL tube and store the vials at -20 &#176;C.</li>
	<li>Place 0.007 g of collagenase in one 1.5 mL tube and store it at -20 &#176;C.</li>
	<li>Add 997 &#181;L of DPBS to the prealiquoted and frozen 3 &#181;L of calcein in one 1.5 mL tube, and resuspend to dissolve the calcein. Add 0.007 g of collagenase to obtain a 1000 &#181;L of working solution.</li>
	<li>Use 500 &#181;L of the working solution to digest two pieces of 2 mm ovarian cortex fragments for 90 min at 37 &#176;C, shielded from light, in a 4-well dish. After 60 min and 70 min, resuspend the solution repeatedly. Finally, determine the follicular viability via fluorescence microscopy, as shown in Figure 6.</li>
</ol>

Follicular Viability Assessment in Vitrified Human Ovarian Cortex Tissues

Here, a protocol for high-throughput vitrification of human ovarian cortex tissue, suitable for clinical routine, is presented. Similar to the vitrification of oocytes or embryos, the successful application of the procedure requires detailed adherence to the protocol concerning the temperature of the vitrification and warming solutions, as well as the equilibration period. Compliance with EU tissue directives37 regarding air quality and sterility is also essential.
The vitrification procedure results in a non-crystalline, amorphous, or glassy state. Overall, vitrification is a versatile process with significant implications in various scientific and technological domains. The primary benefit of vitrification is its ability to convert tissue into a glassy state, thereby preventing ice crystal formation38,39,40, which can negatively affect tissue integrity and its components.
Supplementation of cryoprotective agents (CPAs) with polyvinylpyrrolidone (PVP) allows for a reduction in CPA concentration without compromising the quality of the vitrification solutions41,42. Furthermore, the use of metal grids provides high thermal conductivity compared to plastic-based carrier systems. The grid structure also facilitates surface adhesion, ensuring the safe and secure cryopreservation of tissue samples and small cortex punches for quality measures. If cryovessels from other manufacturers are used, it is important to test the size of the metal grids beforehand to ensure stability and proper grip within the cryovessel lid, as well as a good fit into the vessel.
Critical steps to ensure successful vitrification include rapid vitrification by immersing the tissue in sterilized liquid nitrogen and performing rapid warming without delay to avoid adverse results. In terms of cost-effectiveness, tissue vitrification is less demanding compared to the slow freezing procedure, which may influence personnel deployment planning. Additionally, vitrification eliminates the need to purchase and service equipment required for slow freezing.
Biological measures and meta-analyses have demonstrated the comparability or even advantages of vitrification compared to slow freezing43. However, differences in results after vitrification may be attributed to the lack of standardization in both the vitrification device and the protocol, including the solutions used, which vary across studies. Future research should explore the potential for follicle culture from vitrified/rapid-warmed tissue to monitor growth in vitro, as successfully demonstrated in mouse ovarian tissue by several groups44,45,46,47,48,49,50.
In summary, vitrification of ovarian tissue is a significant alternative to the widely used slow freezing protocol, supported by five successful deliveries reported by Suzuki51 (Japan), Silber52 (USA), and S&#228;nger53 (Germany). In contrast to commercially available vitrification media and kits for cells, there are few FDA/CE-approved systems for ovarian tissue, which may limit their application in clinical settings. Therefore, the development of FDA/CE-approved kits and media for the vitrification and rapid warming of ovarian tissue is recommended30.

Cryopreservation of ovarian tissue is an important option for fertility preservation. Explanted tissue containing ovarian follicles, in which oocytes are embedded, is cryopreserved. After storage, the ovarian tissue can be thawed, warmed, and reimplanted in the patient. For viable cells or tissue, two cryopreservation methods are available: slow freezing and vitrification1.
Vitrification is used to preserve biological materials, such as embryos and oocytes, with superior survival rates compared to the slow freezing protocol1,2,3,4. Slow freezing has limitations, such as ice crystal formation, which can potentially damage cell and tissue structures. However, slow freezing is an important cryopreservation approach that facilitates the long-term storage of biological samples, and the functionality of this method has been widely proven5. Vitrification induces a glassy state of aggregation, preventing ice crystal formation6,7. On a technical level, vitrification significantly simplifies the cryopreservation procedure by reducing equipment maintenance, decreasing the likelihood of technical errors, and shortening the duration of the cryopreservation process8,9. In female fertility preservation, ovarian tissue cryopreservation is a decisive approach prior to cancer treatment10. Different groups have successfully demonstrated the concept of cryopreservation, thawing, and transplantation of tissue based on the slow freezing protocol11,12,13,14, which is currently regarded as the standard approach15.
Vitrification of ovarian tissue is regarded as a promising alternative method16,17,18,19,20,21, in terms of resource-saving22, follicular survival rates, DNA fragmentation levels, and balanced angiogenic potential23,24,25,26,27. This is substantiated by successful deliveries in Japan28, the USA29, and Germany30.
Comparing the two options for ovarian tissue cryopreservation (OTC)-vitrification versus the standard procedure of slow freezing-results are partially conflicting in current meta-analyses16. Several factors may have contributed to this, as current vitrification protocols vary greatly. These differences include the choice of cryoprotectant or combination of protectants, their concentration, the composition of the OTC media, the size of tissue fragments, and the device used as a tissue carrier. Accordingly, there is no standardized warming protocol.
As the authors found a method that yields convincing results in terms of handling, viability, apoptosis onset, release of angiogenic factors, and even a report of a birth after reimplantation9,27, a very detailed description of the protocol is provided. The described method offers a valid and effective protocol that may contribute to the standardization of vitrification of ovarian tissue.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1.8 mL vials</td>
			<td>VWR International GmbH</td>
			<td>479-6837</td>
			<td></td>
		</tr>
		<tr>
			<td>10 mL serological pipette</td>
			<td>Sarstedt</td>
			<td>86.1254.001</td>
			<td></td>
		</tr>
		<tr>
			<td>4 well plate</td>
			<td>Gynemed</td>
			<td>GYOOPW-FW04</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL Tube</td>
			<td>Sarstedt</td>
			<td>62.559.001</td>
			<td></td>
		</tr>
		<tr>
			<td>6 well plates</td>
			<td>Sarstedt</td>
			<td>83.3920</td>
			<td></td>
		</tr>
		<tr>
			<td>Bacillol AF</td>
			<td>Hartmann</td>
			<td>973385</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcein AM</td>
			<td>Merck</td>
			<td>17783</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase type 1A</td>
			<td>Merck</td>
			<td>C2674</td>
			<td></td>
		</tr>
		<tr>
			<td>Cryosure DMSO</td>
			<td>WAK Chemie</td>
			<td>WAK-DMSO-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Custodiol</td>
			<td>Dr. Franz K&#246;hler Chemie</td>
			<td>00867288</td>
			<td></td>
		</tr>
		<tr>
			<td>DPBS CTS</td>
			<td>Gibco Life technologies</td>
			<td>A12856-01</td>
			<td></td>
		</tr>
		<tr>
			<td>ErgoOne pipette aid</td>
			<td>Starlab</td>
			<td>S7166-0010</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylene glycol</td>
			<td>Sigma Aldrich</td>
			<td>102466</td>
			<td></td>
		</tr>
		<tr>
			<td>Euronda sterilization container</td>
			<td>euronda</td>
			<td>282021</td>
			<td></td>
		</tr>
		<tr>
			<td>G-MOPS+</td>
			<td>Vitrolife</td>
			<td>10130</td>
			<td></td>
		</tr>
		<tr>
			<td>Metal meshes</td>
			<td>Sigma Aldrich</td>
			<td>S0770</td>
			<td></td>
		</tr>
		<tr>
			<td>Metzenbaum scissors</td>
			<td>world precision instruments</td>
			<td>501262102</td>
			<td></td>
		</tr>
		<tr>
			<td>N-Bath System</td>
			<td>Nterilizer</td>
			<td>N-Bath 3.0</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyvinylpyrrolidone (PVP)</td>
			<td>SAGE</td>
			<td>ART-4005</td>
			<td></td>
		</tr>
		<tr>
			<td>Serum substitute supplement (SSS)</td>
			<td>Fujifilm Irvine scientific</td>
			<td>99193</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile cup</td>
			<td>Sarstedt</td>
			<td>75.562.105</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile forceps</td>
			<td>Carl Roth</td>
			<td>KL05.1</td>
			<td></td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Merck</td>
			<td>S0389</td>
			<td></td>
		</tr>
	</tbody>
</table>

vitrification of ovarian cortex tissue to achieve a glassy state of aggregation

Ovarian tissue cryopreservation (OTC) is an important option for fertility preservation. For patients whose gonadotoxic treatments cannot be postponed or for pre-pubertal girls, it is often the only option for fertility protection. Cryopreservation can be performed either by vitrification or by slow freezing. Slow freezing is currently the standard approach. An increasing number of studies indicate that vitrification can replace slow freezing in the state-of-the-art in vitro fertilization (IVF) laboratories, significantly improving thawing survival rates and simplifying the technical aspects of cryopreservation. A metal grid-based, high-throughput protocol for rapid vitrification of ovarian cortex tissue, suitable for clinical routine, is described. The sterilization of metal grids and liquid nitrogen ensures high quality, meeting good manufacturing practice (GMP) standards. Vitrification was conducted to ensure ultra-rapid cooling rates. Instead of slowly thawing, samples were rapidly warmed. To assess follicular viability, calcein staining was performed both prior to cryopreservation and after rapid warming. The successful application of vitrification and rapid warming using metal grids is reported. No significant differences in follicular viability were observed prior to vitrification and after rapid warming. These results substantiate the high capacity of tissue vitrification for clinical routine applications as a potential substitute for the widely used slow-freezing method.

This protocol presents the procedures for the preparation of vitrification media, loading devices, vitrification, preparation of rapid warming media, rapid warming, and determination of follicular viability. A direct comparison of follicular viability and angiogenic factors between slow freezing and vitrification has been validated and published31,32.
The overall success of the described vitrification protocol was assessed by comparing the vital follicle count before and after vitrification/rapid warming. The experimental setup is shown in Figure 1, and the results are presented in Figure 6. In 50 patients, a mean count of 77.98 vital follicles was observed before vitrification and 62.99 after vitrification/rapid warming, reflecting a survival rate of 80.8%. This was not significantly different according to the Wilcoxon test33.
Metal grids are individually customized with sharp scissors to a size of 25 mm x 8 mm, which fits into the caps of 1.8 mL cryovials, as shown in Figure 2A,C. After sterilization using steam autoclaving, the grids and caps of 1.8 mL vials are assembled under a laminar flow bench, as indicated in Figure 2B. This setup provides a secure hold in the cap without the need for additional actions and offers sufficient area for tissue of various sizes. Typically, tissue pieces of 5 mm x 10 mm for transplantation and 2 mm diameter pieces for assessing vital follicle count after rapid thawing are vitrified. Both sizes fit perfectly onto the metal grids.
Equilibration with vitrification solutions (VS1, VS2, and VS3) is conducted in a 6-well plate on a rocking shaker at room temperature under a laminar flow bench, as shown in Figure 3. The time frames in the scheme ensure the effective uptake of the cryoprotectant ethylene glycol. Using a 6-well plate over single vessels is recommended as it facilitates quick movement of the tissue between solutions and helps prevent mix-ups.
For rapid vertical vitrification in liquid nitrogen, ovarian cortex tissue is cut into suitable pieces (Figure 4A). Ovarian cortex samples are placed on loading devices (Figure 4B,C) and submerged vertically into liquid nitrogen (Figure 4D) to achieve a glassy state of aggregation through vitrification. The chosen metal grids allow for vertical handling of the tissue, as shown in Figure 4D. Additionally, the metal grids are highly thermally conductive, ensuring rapid cooling rates from 22 &#176;C to -196 &#176;C, which is a critically important step in the vitrification process.
For rapid warming, RWS is prepared in a sterile cup at 37 &#176;C. ES, RS1, and RS2 are prepared in a 6-well plate on a rocking shaker at room temperature, as shown in Figure 5. The high volume of pre-warmed RWS prevents the solution from cooling down excessively upon the addition of the vitrified tissue and ensures a consistently warm environment for the tissue throughout the rapid warming process.
To assess and ensure high quality standards, 2 mm x 2 mm biopsy punches are stained with calcein prior to vitrification and after rapid warming to determine follicular viability using fluorescence microscopy34,35 (Figure 6). Viable follicles emit green fluorescence at 495 nm after the intracellular uptake of calcein. Alternatively, follicular viability (Table 3) can be assessed using neutral red dyes36.
By following the described steps in this protocol, ovarian tissue is transformed into a glassy state of aggregation, which facilitates high survival rates after rapid warming, as confirmed by fluorescence microscopy.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66801/66801fig01.jpg" /> 
Figure 1: Study design. Ovarian cortical samples from 50 patients were examined before (fresh) and after vitrification and rapid warming for the number of viable follicles. For each group, two tissue pieces of 2 mm diameter were cultured for 24 h before follicle count assessment. The tissue was digested with collagenase and stained with calcein to assess viability. The number of viable follicles was determined using a microscope. <a href="https://www.jove.com/files/ftp_upload/66801/66801fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66801/66801fig02.jpg" /> 
Figure 2: Preparation of loading devices. Metal grids were cut to a size of 8 mm x 25 mm (A). The customized metal grids were sterilized by autoclaving (B). The sterilized metal grids were then inserted into caps of 1.8 mL vials, ready for use (C). <a href="https://www.jove.com/files/ftp_upload/66801/66801fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66801/66801fig03.jpg" /> 
Figure 3: Preparation of vitrification solutions. Vitrification solutions (VS) were prepared and transferred to a 6-well plate. The wells show the volume of each solution and the individual incubation times used in the vitrification protocol. <a href="https://www.jove.com/files/ftp_upload/66801/66801fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66801/66801fig04.jpg" /> 
Figure 4: Ovarian cortex processing and vitrification. Ovarian cortex tissue was processed for cryopreservation by removing the medulla and cutting the tissue into 5 mm x 10 mm pieces (A). After incubation in the vitrification solutions shown in Figure 3, the tissue was loaded onto the vitrification loading device (B,C). For rapid vertical vitrification of ovarian cortex samples, the caps with the loaded tissue were quickly inserted into sterilized liquid nitrogen (D). <a href="https://www.jove.com/files/ftp_upload/66801/66801fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66801/66801fig05.jpg" /> 
Figure 5: Preparation of rapid warming solutions. Rapid warming solution (RWS), equilibration solution (ES), and rinsing solutions (RS) 1 and 2 were prepared and transferred to a 6-well plate. The vessels and wells show the volume of each solution and the corresponding incubation times. Note that RWS is maintained at 37.2 &#176;C on a heating plate. <a href="https://www.jove.com/files/ftp_upload/66801/66801fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66801/66801fig06.jpg" /> 
Figure 6: Viability count. To assess the number of viable follicles, tissue pieces of 2 mm diameter were digested with collagenase and stained with calcein. The representative images show (A) calcein staining of 3 recovered follicles prior to vitrification. (B) A recovered viable follicle after vitrification and rapid warming. (C) A recovered viable follicle prior to slow freezing. (D) Recovered viable follicles after slow freezing and thawing. Follicular viability is indicated by calcein, a green fluorescent dye that emits green fluorescence when enzymatically converted by viable cells at 495 nm. Scale bar = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/66801/66801fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>VS 1 (15 mL)</td>
			<td>Ethylene glycol</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>SSS</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>12 mL</td>
		</tr>
		<tr>
			<td>VS 2 (15 mL)</td>
			<td>Ethylene glycol</td>
			<td>20%</td>
			<td>3 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>SSS</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>10.5 mL</td>
		</tr>
		<tr>
			<td>VS 3 (15 mL)</td>
			<td>Ethylene glycol</td>
			<td>35%</td>
			<td>5.25 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>SSS</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>Sucrose</td>
			<td>0.5 mol/L</td>
			<td>2.57 g</td>
		</tr>
		<tr>
			<td></td>
			<td>PVP</td>
			<td>5 % (w/v)</td>
			<td>0.75 g</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>ad 15 mL</td>
		</tr>
	</tbody>
</table>
Table 1: Composition of vitrification solutions (VS).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>RWS (30 mL)</td>
			<td>Sucrose</td>
			<td>0.8 mol/L</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>8.22 g</td>
		</tr>
		<tr>
			<td></td>
			<td>SSS</td>
			<td>10%</td>
			<td>3 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>ad 30 mL</td>
		</tr>
		<tr>
			<td>ES (15 mL)</td>
			<td>Sucrose</td>
			<td>0.4 mol/L</td>
			<td>2.05 g</td>
		</tr>
		<tr>
			<td></td>
			<td>SSS</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>ad 15 mL</td>
		</tr>
		<tr>
			<td>RS 1&#38;2 (15 mL)</td>
			<td>SSS</td>
			<td>10%</td>
			<td>1.5 mL</td>
		</tr>
		<tr>
			<td></td>
			<td>G-MOPS+</td>
			<td></td>
			<td>ad 15 mL</td>
		</tr>
	</tbody>
</table>
Table 2: Composition of rapid warming solution (RWS), equilibration solution (ES), and rinsing solutions (RS). This table provides the components and concentrations for the rapid warming solution (RWS), equilibration solution (ES), and rinsing solutions (RS) used in the post-vitrification processing.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Parameter</td>
			<td>Fresh</td>
			<td>Interval</td>
			<td>Rapid warmed after vitrification</td>
			<td>Interval</td>
			<td>n</td>
			<td>*P-value</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>SD</td>
			<td></td>
			<td>SD</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Follicular viability count [n]</td>
			<td>77.98</td>
			<td>0-386</td>
			<td>62.99</td>
			<td>0.5-349</td>
			<td>50</td>
			<td>0.130</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td>77.95</td>
			<td></td>
			<td>80.02</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>*Wilcoxon test</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 3: Representative results of follicular viability. This table presents the results of follicular viability assessments before cryopreservation and after rapid warming. Two tissue pieces of 2 mm diameter per patient were used to count the number of viable follicles before and after vitrification/rapid warming. Paired tissue samples from 50 patients were analyzed using the Wilcoxon test.

Author Spotlight: Advancing Fertility Preservation in Young Female Cancer Patients Through Ovarian Tissue Vitrification and In Vitro Follicular Growth

A protocol for the vitrification of ovarian tissue, as an alternative cryopreservation method to the widely used slow freezing protocol, is presented.

Vitrification of Ovarian Cortex Tissue to Achieve a Glassy State of Aggregation

Ovarian tissue cryopreservation (OTC) is an important option for fertility preservation. For patients whose gonadotoxic treatments cannot be postponed or ...

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Research

JoVE Journal

Medicine

1.1K Views.  University Hospital of Bonn. A protocol for the vitrification of ovarian tissue, as an alternative cryopreservation method to the widely used slow freezing protocol, is presented.

Video: Author Spotlight: Advancing Fertility Preservation in Young Female Cancer Patients Through Ovarian Tissue Vitrification and In Vitro Follicular Growth

In vitro Mesothelial Clearance Assay that Models the Early Steps of Ovarian Cancer Metastasis

Ovarian cancer is the fifth leading cause of cancer related deaths in the United States1. Despite a positive initial response to therapies, 70 to 90 percent of women with ovarian cancer develop new metastases, and the recurrence is often fatal2. It is, therefore, necessary to understand how secondary metastases arise in order to develop better treatments for intermediate and late stage ovarian cancer. Ovarian cancer metastasis occurs when malignant cells detach from the primary tumor site and disseminate throughout the peritoneal cavity. The disseminated cells can form multicellular clusters, or spheroids, that will either remain unattached, or implant onto organs within the peritoneal cavity3 (Figure 1, Movie 1). 
All of the organs within the peritoneal cavity are lined with a single, continuous, layer of mesothelial cells4-6 (Figure 2). However, mesothelial cells are absent from underneath peritoneal tumor masses, as revealed by electron micrograph studies of excised human tumor tissue sections3,5-7 (Figure 2). This suggests that mesothelial cells are excluded from underneath the tumor mass by an unknown process. 
Previous in vitro experiments demonstrated that primary ovarian cancer cells attach more efficiently to extracellular matrix than to mesothelial cells8, and more recent studies showed that primary peritoneal mesothelial cells actually provide a barrier to ovarian cancer cell adhesion and invasion (as compared to adhesion and invasion on substrates that were not covered with mesothelial cells)9,10. This would suggest that mesothelial cells act as a barrier against ovarian cancer metastasis. The cellular and molecular mechanisms by which ovarian cancer cells breach this barrier, and exclude the mesothelium have, until recently, remained unknown. 
Here we describe the methodology for an in vitro assay that models the interaction between ovarian cancer cell spheroids and mesothelial cells in vivo (Figure 3, Movie 2). Our protocol was adapted from previously described methods for analyzing ovarian tumor cell interactions with mesothelial monolayers8-16, and was first described in a report showing that ovarian tumor cells utilize an integrin &#x2013;dependent activation of myosin and traction force to promote the exclusion of the mesothelial cells from under a tumor spheroid17. This model takes advantage of time-lapse fluorescence microscopy to monitor the two cell populations in real time, providing spatial and temporal information on the interaction. The ovarian cancer cells express red fluorescent protein (RFP) while the mesothelial cells express green fluorescent protein (GFP). RFP-expressing ovarian cancer cell spheroids attach to the GFP-expressing mesothelial monolayer. The spheroids spread, invade, and force the mesothelial cells aside creating a hole in the monolayer. This hole is visualized as the negative space (black) in the GFP image. The area of the hole can then be measured to quantitatively analyze differences in clearance activity between control and experimental populations of ovarian cancer and/ or mesothelial cells. This assay requires only a small number of ovarian cancer cells (100 cells per spheroid X 20-30 spheroids per condition), so it is feasible to perform this assay using precious primary tumor cell samples. Furthermore, this assay can be easily adapted for high throughput screening.

In vitro Mesothelial Clearance Assay that Models the Early Steps of Ovarian Cancer Metastasis

The mesothelial clearance assay described here takes advantage of fluorescently labeled cells and time-lapse video microscopy to visualize and quantitatively measure the interactions of ovarian cancer multicellular spheroids and mesothelial cell monolayers. This assay models the early steps of ovarian cancer metastasis.

Ovarian cancer is the fifth leading cause of cancer related deaths in the United States1. Despite a positive initial response to therapies, 70 to 90 ...

Heterotypic Three-dimensional In Vitro Modeling of Stromal-Epithelial Interactions During Ovarian Cancer Initiation and Progression

Epithelial ovarian cancers (EOCs) are the leading cause of death from gynecological malignancy in Western societies. Despite advances in surgical treatments and improved platinum-based chemotherapies, there has been little improvement in EOC survival rates for more than four decades 1,2. Whilst stage I tumors have 5-year survival rates &gt;85%, survival rates for stage III/IV disease are &lt;40%. Thus, the high rates of mortality for EOC could be significantly decreased if tumors were detected at earlier, more treatable, stages 3-5. At present, the molecular genetic and biological basis of early stage disease development is poorly understood. More specifically, little is known about the role of the microenvironment during tumor initiation; but known risk factors for EOCs (e.g. age and parity) suggest that the microenvironment plays a key role in the early genesis of EOCs. We therefore developed three-dimensional heterotypic models of both the normal ovary and of early stage ovarian cancers. For the normal ovary, we co-cultured normal ovarian surface epithelial (IOSE) and normal stromal fibroblast (INOF)&nbsp;cells, immortalized by retrovrial transduction of the catalytic subunit of human telomerase holoenzyme (hTERT) to extend the lifespan of these cells in culture. To model the earliest stages of ovarian epithelial cell transformation, overexpression of the CMYC oncogene in IOSE cells, again co-cultured with INOF cells. These heterotypic models were used to investigate the effects of aging and senescence on the transformation and invasion of epithelial cells. Here we describe the methodological steps in development of these three-dimensional model; these methodologies aren't specific to the development of normal ovary and ovarian cancer tissues, and could be used to study other tissue types where stromal and epithelial cell interactions are a fundamental aspect of the tissue maintenance and disease development.

Heterotypic Three-dimensional In Vitro Modeling of Stromal-Epithelial Interactions During Ovarian Cancer Initiation and Progression

We describe methodologies for establishing in vitro heterotypic three-dimensional models comprising ovarian fibroblasts and normal ovarian surface or ovarian cancer epithelial cells. We discuss the use of these models to study stromal-epithelial interactions that occur during ovarian cancer development.

Epithelial ovarian cancers (EOCs) are the leading cause of death from gynecological malignancy in Western societies. Despite advances in surgical ...

A Method to Study the Impact of Chemically-induced Ovarian Failure on Exercise Capacity and Cardiac Adaptation in Mice

The risk of cardiovascular disease (CVD) increases in post-menopausal women, yet, the role of exercise, as a preventative measure for CVD risk in post-menopausal women has not been adequately studied. Accordingly, we investigated the impact of voluntary cage-wheel exercise and forced treadmill exercise on cardiac adaptation in menopausal mice. The most commonly used inducible model for mimicking menopause in women is the ovariectomized (OVX) rodent. However, the OVX model has a few dissimilarities from menopause in humans. In this study, we administered 4-vinylcyclohexene diepoxide (VCD) to female mice, which accelerates ovarian failure as an alternative menopause model to study the impact of exercise in menopausal mice. VCD selectively accelerates the loss of primary and primordial follicles resulting in an endocrine state that closely mimics the natural progression from pre- to peri- to post-menopause in humans. To determine the impact of exercise on exercise capacity and cardiac adaptation in VCD-treated female mice, two methods were used. First, we exposed a group of VCD-treated and untreated mice to a voluntary cage wheel. Second, we used forced treadmill exercise to determine exercise capacity in a separate group VCD-treated and untreated mice measured as a tolerance to exercise intensity and endurance.

Two exercise paradigms were tested on a newly developed chemically induced menopausal mouse model to examine the impact of menopause on exercise capacity and cardiac adaption to exercise.

The risk of cardiovascular disease (CVD) increases in post-menopausal women, yet, the role of exercise, as a preventative measure for CVD risk in ...

Method for Obtaining Primary Ovarian Cancer Cells From Solid Specimens

Reliable tools for investigating ovarian cancer initiation and progression are urgently needed. While the use of ovarian cancer cell lines remains a valuable tool for understanding ovarian cancer, their use has many limitations. These include the lack of heterogeneity and the plethora of genetic alterations associated with extended in vitro passaging.&#160;Here we describe a method that allows for rapid establishment of primary ovarian cancer cells form solid clinical specimens collected at the time of surgery. The method consists of subjecting clinical specimens to enzymatic digestion for 30 min. The isolated cell suspension is allowed to grow and can be used for downstream application including drug screening. The advantage of primary ovarian cancer cell lines over established ovarian cancer cell lines is that they are representative of the original specific clinical specimens they are derived from and can be derived from different sites whether primary or metastatic ovarian cancer.

This study describes a detailed method for isolation and characterization of primary ovarian cancer cells from solid clinical specimens. Ovarian cancer clinical specimens are subjected to enzymatic digestion to obtain viable, fibroblast-free epithelial ovarian cancer (EOC) cells highly suitable for downstream applications.

Reliable tools for investigating ovarian cancer initiation and progression are urgently needed. While the use of ovarian cancer cell lines remains a ...

State of the Art Cranial Ultrasound Imaging in Neonates

Cranial ultrasound (CUS) is a reputable tool for brain imaging in critically ill neonates. It is safe, relatively cheap and easy to use, even when a patient is unstable. In addition it is radiation-free and allows serial imaging. CUS possibilities have steadily expanded. However, in many neonatal intensive care units, these possibilities are not optimally used. We present a comprehensive approach for neonatal CUS, focusing on optimal settings, different probes, multiple acoustic windows and Doppler techniques. This approach is suited for both routine clinical practice and research purposes. In a live demonstration, we show how this technique is performed in the neonatal intensive care unit. Using optimal settings and probes allows for better imaging quality and improves the diagnostic value of CUS in experienced hands. Traditionally, images are obtained through the anterior fontanel. Use of supplemental acoustic windows (lambdoid, mastoid, and lateral fontanels) improves detection of brain injury. Adding Doppler studies allows screening of patency of large intracranial arteries and veins. Flow velocities and indices can be obtained. Doppler CUS offers the possibility of detecting cerebral sinovenous thrombosis at an early stage, creating a window for therapeutic intervention prior to thrombosis-induced tissue damage. Equipment, data storage and safety aspects are also addressed.

Cranial ultrasound (CUS) is a valuable tool for brain imaging in critically ill neonates. This video shows a comprehensive approach for neonatal (Doppler) CUS for both clinical and research purposes, including a bedside demonstration of the technique.

Cranial ultrasound (CUS) is a reputable tool for brain imaging in critically ill neonates. It is safe, relatively cheap and easy to use, even when a ...

Quantitation of Intra-peritoneal Ovarian Cancer Metastasis

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States. Mortality is due to diagnosis of 75% of women with late stage disease, when metastasis is already present. EOC is characterized by diffuse and widely disseminated intra-peritoneal metastasis. Cells shed from the primary tumor anchor in the mesothelium that lines the peritoneal cavity as well as in the omentum, resulting in multi-focal metastasis, often in the presence of peritoneal ascites. Efforts in our laboratory are directed at a more detailed understanding of factors that regulate EOC metastatic success. However, quantifying metastatic tumor burden represents a significant technical challenge due to the large number, small size and broad distribution of lesions throughout the peritoneum. Herein we describe a method for analysis of EOC metastasis using cells labeled with red fluorescent protein (RFP) coupled with in vivo multispectral imaging. Following intra-peritoneal injection of RFP-labelled tumor cells, mice are imaged weekly until time of sacrifice. At this time, the peritoneal cavity is surgically exposed and organs are imaged in situ. Dissected organs are then placed on a labeled transparent template and imaged ex vivo. Removal of tissue auto-fluorescence during image processing using multispectral unmixing enables accurate quantitation of relative tumor burden. This method has utility in a variety of applications including therapeutic studies to evaluate compounds that may inhibit metastasis and thereby improve overall survival.

Ovarian cancer metastasis is characterized by numerous diffuse intra-peritoneal lesions, such that accurate visual quantitation of tumor burden is challenging. Herein we describe a method for in situ and ex vivo quantitation of metastatic tumor burden using red fluorescent protein (RFP)-labeled tumor cells and optical imaging.

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States.  Mortality is due to diagnosis of 75% of ...

Transplantation Into the Mouse Ovarian Fat Pad

Orthotopic transplantation assays in mice are invaluable for studies of cell regeneration and neoplastic transformation. Common approaches for orthotopic transplantation of ovarian surface and tubal epithelia include intraperitoneal and intrabursal administration of cells. The respective limitations of these methods include poorly defined location of injected cells and limited space volume. Furthermore, they are poorly suited for long-term structural preservation of transplanted organs. To address these challenges, we have developed an alternative approach, which is based on the introduction of cells and tissue fragments into the mouse fat pad. The mouse ovarian fat pad is located in the immediate vicinity of the ovary and uterine tube (aka oviduct, fallopian tube), and provides a familiar microenvironment for cells and tissues of these organs. In our approach fluorescence-labeled mouse and human cells, and fragments of the uterine tube are engrafted by using minimally traumatic dorsal incision surgery. Transplanted cells and their outgrowths are easily located in the ovarian fat pad for over 40 days. Long-term transplantation of the entire uterine tube allows correct preservation of all principle tissue components, and does not result in adverse side effects, such as fibrosis and inflammation. Our approach should be uniquely applicable for answering important biological questions such as differentiation, regenerative and neoplastic potential of specific cell populations. Furthermore, it should be suitable for studies of microenvironmental factors in normal development and cancer.

We describe an ovarian fad pad transplantation assay suitable for studies of normal and transformed epithelia of the female reproductive tract. The mouse fat pad allows transplantation of large tissue fragments, is easily accessible for surgery and imaging, and provides the most favorable native environment for tissues of M&#252;llerian origin.

Orthotopic transplantation assays in mice are invaluable for studies of cell regeneration and neoplastic transformation. Common approaches for orthotopic ...

Use of MRI-ultrasound Fusion to Achieve Targeted Prostate Biopsy

Here, we present a protocol to perform targeted prostate biopsy using a magnetic resonance imaging-ultrasound (MRI/US) fusion system. Prostate cancer has traditionally been diagnosed via transrectal ultrasound (TRUS) biopsy. Though considered the gold standard, TRUS is unable to visualize most prostate cancer lesions and therefore requires sampling of the entire prostate. This biopsy method often undergrades prostate cancer and fails to detect up to 35% of cancers on initial biopsy. Prostate MRI has been shown to have excellent sensitivity in the detection of cancerous lesions, and advancements in MRI technology during the last decade have led to the development of targeted biopsy. In targeted biopsy, a software platform overlays MRI data onto live TRUS images to create a fused MRI/US three-dimensional model of the prostate. Regions suspicious for malignancy on MRI are contoured by a radiologist, uploaded into the fusion system, and then displayed within the live MRI/US fused model. The urologist is then able to directly biopsy these targets. When compared to conventional TRUS biopsy, MRI/US fusion technology has been demonstrated to improve the detection of clinically significant cancer while reducing insignificant cancer detection. This technology, therefore, has the potential to diagnose prostate cancer primarily in men who would benefit from treatment.

Presented herein is a protocol to perform targeted biopsy of the prostate using an MRI-ultrasound fusion system.

Here, we present a protocol to perform targeted prostate biopsy using a magnetic resonance imaging-ultrasound (MRI/US) fusion system. Prostate cancer has ...

Electroretinogram Recording for Infants and Children under Anesthesia to Achieve Optimal Dark Adaptation and International Standards

Electroretinogram (ERG) is the only clinical objective test available to assess retinal function. Full-field ERG (ffERG) measures the panretinal rod and cone photoreceptor function as well as inner retinal function and is an important measure in the diagnosis and management of inherited retinal diseases as well as inflammatory, toxic, and nutritional retinopathies. Adhering to international standards and maintaining retinal dark adaptation are critical to acquire valid and reliable dark-adapted (scotopic) and light-adapted (photopic) ffERG responses. Performing ffERG in infants and children is challenging and often requires general anesthesia in the operating room. However, maintaining retinal dark adaptation in the operating room is becoming increasingly difficult given the numerous light sources from anesthesiology monitoring systems and other equipment. A practical and widely applicable method for ffERG testing is described in the operating room that optimizes retinal dark adaptation. The method reduces operating room time by dark-adapting the patient before general anesthesiology is instituted. The operating room is modified for dark adaptation and any remaining light source in the darkened operating room is minimized with the use of a modified portable foldable darkroom that encloses the patient&#8217;s head and the ERG examiner during ffERG scotopic recordings. The simple method adheres to ffERG international standards and provides valid reliable scotopic and photopic ffERG recordings that are critical to assess objective retinal function in this young age group where subjective assessment of visual function such as visual acuity and visual fields are not possible. Furthermore, the ffERG is the gold standard clinical test in detecting early onset inherited retinal diseases including Leber congenital amaurosis where approved gene therapy has become available. In sedated conditions, very low amplitude ffERG signals can be detected due to minimal orbicularis muscle activity interference, which is particularly relevant in patients after gene therapy to detect improved amplitude responses.

Adhering to international standards and maintaining retinal dark adaptation are critical to acquire valid full-field electroretinogram responses in the diagnosis and management of inherited retinal diseases. A practical protocol using a portable darkroom is provided to obtain full-field&#160;electroretinogram for infants and children under sedation or general anesthesia in the operating room setting.

Electroretinogram (ERG) is the only clinical objective test available to assess retinal function. Full-field ERG (ffERG) measures the panretinal rod and ...

Fertility Preservation Through Oocyte Vitrification: Clinical and Laboratory Perspectives

Preserving female fertility is crucial in a multifunctional healthcare system that takes care of patients' future quality of life. Oocyte cryopreservation is recognized by several international scientific societies as the gold standard for fertility preservation in postpubertal women, for both medical and non-medical indications. The main medical indications are oncologic diseases, gynecologic diseases such as severe endometriosis, systemic diseases compromising the ovarian reserve, and genetic conditions involving premature menopause. This paper describes the whole clinical and laboratory work-up of a fertility preservation treatment by outlining recommendations for objective and evidence-based counseling. Furthermore, it focuses on the effectiveness of the procedure and describes the most appropriate strategies to fully exploit the ovarian reserve and maximize the number of oocytes retrieved in the shortest possible time. The evaluation of the ovarian reserve, the definition of an ideal stimulation protocol, as well as oocyte retrieval, denudation, and vitrification procedures have been detailed along with approaches to maximize their efficacy, efficiency, and safety.

Oocyte cryopreservation is recognized by several international scientific societies as the gold standard for fertility preservation in postpubertal women. Appropriate clinical and laboratory strategies ensure maximum efficacy, efficiency, and safety of fertility preservation treatments.

Preserving female fertility is crucial in a multifunctional healthcare system that takes care of patients' future quality of life. Oocyte cryopreservation ...