This study was supported by the National Research, Development and Innovation Office of Hungary (grant 116912 to &#193;H), the COST office (Food and Agriculture COST Action FA1205: AQUAGAMETE), the Stipendium Hungaricum Scholarship Programme (grant to ZM) and the New Hungarian National Excellence Predoctoral Fellowship (grant to EK).

All methods described here have been approved by the Hungarian Animal Welfare Law.
1. Reagent Preparation
<ol>
	<li>Stock solutions
	<ol>
		<li>Prepare 1 M trehalose by adding 0.378 g of trehalose dihydrate into 1 mL of dH2O. Mix well until it dissolves completely.</li>
		<li>Prepare 1 M sucrose by adding 0.342 g of sucrose into 1 mL of dH2O. Mix well until it dissolves completely.</li>
		<li>Prepare 1 M HEPES by&#160;adding 0.238 g of HEPES to 1 mL of dH2O. Mix well until it dissolves completely.</li>
		<li>Prepare 20 mg/mL collagenase (290 U/mg) by&#160;adding 100 mg to 5 mL of PBS. Mix thoroughly until it dissolves. Sterile filter through 0.2 &#181;m filters. Aliquot 50 &#181;L into 0.2 mL PCR tubes and freeze at - 20 &#176;C.</li>
		<li>Prepare 15 mg/mL trypsin (~ 10000 U/mg) by&#160;adding 75 mg to 5 mL of PBS. Mix thoroughly until it dissolves. Sterile filter through 0.2 &#181;m filters. Aliquot 50 &#181;L into 0.2 mL PCR tubes and freeze at - 20 &#176;C.</li>
		<li>Prepare 1 mg/mL DNase I (~ 3000 U/mg) by&#160;adding 5 mg to 5 mL of PBS. Mix thoroughly until it dissolves. Sterile filter through 0.2 &#181;m filters. Aliquot 50 &#181;L into 0.2 mL PCR tubes and freeze at - 20 &#176;C.</li>
		<li>Prepare 0.4% trypan blue by&#160;adding 40 mg of trypan blue to 10 mL of PBS. Sterile filter through 0.2 &#181;m filters. Aliquot 1 mL into 1.5 mL tubes.</li>
		<li>Prepare 200 mg/L MS-222 (Tricaine methane sulfonate) by&#160;adding 200 mg of MS-222 into 1 L of dH2O. Mix well until it dissolves completely. Additionally, buffer the MS-222 solution with sodium bicarbonate to a neutral pH of 7.0.</li>
	</ol>
	</li>
	<li>Solutions for vitrification
	<ol>
		<li>Prepare Equilibration solution (ES): Prepare 2 mL of ES by mixing 121.5 &#181;L of methanol (MeOH; 1.5 M), 220 &#181;L of propylene glycol (PG; 1.5 M), 200 &#181;L of FBS (10%), 200 &#181;L of trehalose stock solution (0.1 M), 50 &#181;L of HEPES stock solution (25 mM) and 1208.5 &#181;L of L-15 into a 2 mL tube.</li>
		<li>Prepare Vitrification solution (VS): Prepare 2 mL of VS by mixing 439 &#181;L of PG (3 M), 426 &#181;L of dimethyl sulfoxide (Me2SO; 3 M), 200 &#181;L of FBS (10%), 200 &#181;L of trehalose stock solution (0.1 M), 50 &#181;L of HEPES stock solution (25 mM) and 685 &#181;L of L-15 into a 2 mL tube.</li>
	</ol>
	</li>
	<li>Solutions for warming
	<ol>
		<li>Prepare warming solution 1 (WS1): Prepare 1.5 mL of WS1 in a 2 mL tube by mixing 150 &#181;L of FBS (10%), 450 &#181;L of sucrose stock solution (3 M) and 900 &#181;L of L-15.</li>
		<li>Prepare warming solution 2 (WS2): Prepare 1.5 mL of WS2 in a 2 mL tube by mixing 150 &#181;L of FBS (10%), 150 &#181;L of sucrose stock solution (1 M) and 1200 &#181;L of L-15.</li>
		<li>Prepare warming solution 3 (WS3): Prepare 1.5 mL of WS2 in a 2 mL tube by mixing 150 &#181;L FBS (10%) and 1350 &#181;L of L-15.</li>
	</ol>
	</li>
	<li>Prepare digestion solution: for each sample prepare 500 &#181;L of the digestion solution by mixing 50 &#181;L of collagenase stock solution (2 mg/mL), 50 &#181;L of trypsin stock solution (1.5 mg/mL), 10 &#181;L of DNase I (20 &#181;g/mL) and 390 &#181;L of L-15 in a 2 mL tube.</li>
	<li>Obtain dissection tools: scissors, microscissors, tweezers, curved tweezers, and acupuncture needles.</li>
</ol>
2. Testes Collection
<ol>
	<li>Euthanize the fish by placing it into a dish containing 200 mg/L MS-222 (pH = 7). Before continuing with the dissection, make sure that the gills have stopped moving and that at least 10 min have passed since that moment to ensure death by hypoxia.</li>
	<li>Dry the fish by patting it on a paper towel and place it on its dorsal side on a dissecting mat.
	<ol>
		<li>Firstly cut off the pelvic and pectoral fins for easier dissection.</li>
		<li>Horizontally snip the skin on the belly of the fish between the pectoral fins and cut the skin and underlying muscle along the belly until the anal fin.</li>
		<li>Open the body cavity of the fish and pin the left and right body wall to the dissection mat with needles. 
		NOTE: Testes are located above the gastrointestinal organs on both sides of the swimming bladder (Figure 1).</li>
	</ol>
	</li>
	<li>To prevent contamination, do not take out the gastrointestinal organs, but gently push them to one side and remove the testis from the opposing side. Testes are connected to the body wall so remove them carefully and cut the connecting peritoneum by microscissors.</li>
	<li>Additionally, in order to prevent contamination, firstly sterilize the testes by putting them in 70% ethanol for 2 s, and then place them in L-15 on a 96-well plate on ice. 
	NOTE: If it is necessary, clean testes from fatty tissue and large blood vessels precisely under the stereomicroscope</li>
</ol>
3. Ultra-Rapid Cooling of the Testicular Tissue
<ol>
	<li>Mark needles with appropriate labels depending on the sample type and prepare the equilibration (ES) and vitrification solutions (VS).</li>
	<li>Transfer the testes to a larger well plate or dish and pin two to three testes (or more depending on the size of the needle) on a sterile acupuncture needle.
	<ol>
		<li>Firstly position the testis on top of curved tweezers. Place the point of the needle in the middle of the testis and carefully pull the testis upward with the tweezers.</li>
		<li>After puncturing the testis, gently pull it upwards toward the top of the needle. Make sure the testes are separated from each other and that they are not falling off or sliding down.</li>
		<li>Place the needles with pinned testes in a 2 mL tube filled with L-15 until vitrification (at 25 &#730;C; storage time should not exceed 1 h). 
		NOTE: Pinning the testes on the needle should be done fast to prevent the testes from drying out.</li>
	</ol>
	</li>
	<li>Transfer the needle with testes into the equilibration solution and incubate for 5 min at 25&#730;C.</li>
	<li>Transfer the needle into the vitrification solution and incubate for 30 s at 25&#730;C.</li>
	<li>Remove the needle from the vitrification solution, quickly and gently absorb the remaining solution from the tissue by a sterile paper towel. Use caution to avoid the testes sticking to the paper towel or falling off.</li>
	<li>Quickly place the needle into liquid nitrogen placed in a Styrofoam box. 
	NOTE: Do this step quickly to avoid exposure of tissue to the liquid nitrogen vapor. Use low volumes of liquid nitrogen to prevent cross-contamination of samples.</li>
	<li>Keep the needle in liquid nitrogen in a closed Styrofoam box for 5 - 10 min.</li>
	<li>Precool an opened 4.5 mL cryotube and its cap in the same box.</li>
	<li>After 5 - 10 min, transfer each needle into a separate cryotube under the surface of liquid nitrogen and close the cryotube. 
	Caution: Always use protective clothing (such as insulated gloves) when working with liquid nitrogen as exposure can lead to severe frostbite.</li>
	<li>Place the cryotube onto a metal cane and place it in the cryobank canister as fast as possible. Store the samples in a canister storage dewar until further use.</li>
</ol>
4. Warming Procedure
<ol>
	<li>Warm each needle separately. All warming solutions should be at 25 &#176;C.</li>
	<li>Detach the cryotube from the metal cane and plunge it into liquid nitrogen stored in a Styrofoam box.</li>
	<li>Open the cryotube in the liquid nitrogen vapor (using forceps) and release the needle into the liquid nitrogen.</li>
	<li>Transfer the needle very fast into the first warming solution and incubate for 1 min (at 25 &#176;C). Make sure to transfer the needle quickly to prevent its warming in the air during transfer.</li>
	<li>Transfer the needle into the second warming solution and incubate for 3 min (at 25 &#176;C).</li>
	<li>Transfer the needle into the third warming solution and incubate for 5 min (at 25 &#176;C).</li>
	<li>Release the testes from the needle by sliding them downwards with curved tweezers. Place each warmed testis individually into a separate well (96-well plate) filled with 250 &#181;L of L-15 supplemented with 10% FBS. Make sure that the wells are labeled according to the sample. Keep the well plate on ice until all samples are warmed and until further work.</li>
</ol>
5. Tissue Digestion
<ol>
	<li>Prepare 500 &#181;L of the dissociation solution for each sample in separate 2 mL tubes. Label the tubes according to the code of the sample. Incubate the prepared dissociation solution for 5 min at room temperature.</li>
	<li>Add the warmed tissue into the digestion solution and cut it into small pieces by at least 30 movements of small scissors.</li>
	<li>Incubate the cut tissue on a shaking plate for 90 min at 25 &#730;C.</li>
	<li>Stop the digestion process by adding 400 &#181;L of L-15 and 100 &#181;L of FBS (10% FBS v/v). Briefly shake the solution and incubate for 1 min at room temperature.</li>
	<li>Filter the obtained solution through 50 &#181;m filters into a 1.5 mL tube. Label the tubes accordingly.</li>
	<li>Centrifuge the filtered solution at 200 &#215;g for 10 min at 25 &#730;C.</li>
	<li>Carefully remove the supernatant and resuspend the pellet in 20 &#181;L of L15 supplemented with 10% FBS. Manually shake the tube until the pellet is completely dissolved. Keep the cell suspension on ice or at 4 &#730;C until further use.</li>
</ol>
6. Viability Evaluation and Cell Counting
<ol>
	<li>Mix 5 &#181;L of cell suspension and 5 &#181;L of 0.4% trypan blue solution in an appropriately labeled 0.2 mL PCR tube. If different volumes are used, make sure that the dilution ratio of 1:1 remains.</li>
	<li>Incubate this suspension for 1 to 3 min at room temperature (25 &#176;C).</li>
	<li>Check the viability of each sample in a hemocytometer under the microscope.</li>
	<li>Count the number of live cells (unstained by trypan blue) in 15 fields. Calculate the total number of cells in 20 &#181;m of the cell suspension. The suspension is now ready for any downstream application. 
	NOTE: When optimizing the procedure for other species with larger body size, use testis fragments of equal weight. Use testes from one individual for all experimental groups in order to avoid individual variability in the results. In the case of zebrafish, we took advantage of the fact that left and right testis should contain an equal (or very similar) number of germ cells13 (see the Representative Results). The left testis was kept as a control while the right testis was vitrified/warmed. Viability percentage was calculated the number of cells isolated from the vitrified/warmed testis compared to the number of cells isolated from the fresh testis.</li>
</ol>

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The authors have nothing to disclose.

The main purpose of this study was to adapt the needle immersed ultra-rapid cooling procedure developed for avian and mammalian species10,11,12 to the cryopreservation of fish testis (zebrafish as a model organism). Most of the previous studies regarding cryopreservation of zebrafish genetic resources were primarily focused on cryopreservation of zebrafish sperm2,3,...

Novel trends in science and biotechnology have led to the creation of thousands of new mutant lines of mice, Drosophila, zebrafish and other species used as model organisms in biomedical and other sciences1. Furthermore, as new technologies are developed and become available, the numbers of mutant lines steadily increase2. This leads to the necessity for a safe storage of genetic resources beyond the common practices of keeping breeding colonies. As a method which enables safe storage of genetic resources for an indefinite period of time, cryopreservation offers many advantages such as extension of reproductive season, circumvents the need for continuous maintenance of broodstock, and it is more cost- and labor-efficient2.
Protocols for sperm cryopreservation developed during the past several years2,3,4 offer the opportunity for successful storage of zebrafish male genetic material. However, cryopreservation of eggs or embryos in fish is not yet possible due to their complex structure and large amounts of yolk material. Recently, the practice of transplantation of primordial germ cells (PGCs) or spermatogonial stem cells (SSCs) offers a bypass to this barrier by developing into functional sperm and eggs after transplantation5. Therefore, cryopreservation of SSCs offers a new frontier in conservation of rare and valuable genetic resources.
Even though cryopreservation offers many advantages, the slow-rate freezing process generates several conditions that may lead to cell damage2. These include intracellular and extracellular ice formation, dehydration, cryoprotectant toxicity and others. Intracellular ice damages the cells, extracellular ice may lead to mechanical crushing of cells, while water diffusion from the cells during slow-rate freezing may lead to dehydration6. Recently, vitrification as a technique which prevents the negative effects of ice formation has been applied in the cryopreservation of fish gametes7,8,9. It presents an ultra-rapid cooling technique through which the internal and external media turn into an amorphous/glassy state without crystalizing into ice7,10. Successful vitrification of testicular and ovarian tissue has been evidenced in avian and mammalian species10,11,12, thus opening possibilities for its application in fish, as well.
In this study, we present the needle immersed vitrification (NIV) procedure for the cryopreservation of whole zebrafish testes. We demonstrate a reliable method for the isolation of zebrafish early-stage germ cells without contamination and a cryopreservation process that yields relatively high amounts of early-stage germ cells with a low presence of other cells, especially spermatozoa. To the best of our knowledge, this is the first study to demonstrate a detailed visualized protocol for ultra-rapid cooling of fish gonadal tissue and zebrafish germline cells. Additionally, repeatability of the method is demonstrated in five different zebrafish strains: AB wild type, casper (roy-/-; nacre-/-), leopard (leot1/t1), vasa [Tg(vas::eGFP)] and Wilms tumor [Tg(wt1b::eGFP 1)] transgenic line.

<table border="0" cellpadding="0" cellspacing="0" width="948">
	<tbody>
		<tr height="24">
			<td height="24" style="height:24px;width:232px;">Leibovitz media (L-15)</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td style="width:200px;">L1518</td>
			<td style="width:296px;">Supplemented with L-glutamine</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:232px;">Fetal bovine serum (FBS)</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td style="width:200px;">F9665</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:232px;">Tricaine methanesulfonate (MS-222)</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td>E10521</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:232px;">HEPES</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td style="width:200px;">H3375</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:232px;">Sucrose</td>
			<td style="width:220px;">Acros Organics</td>
			<td>57-50-1</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Trehalose</td>
			<td style="width:220px;">Acros Organics</td>
			<td>99-20-7</td>
			<td style="width:296px;">Dihydrate</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Methanol</td>
			<td style="width:220px;">Reanal</td>
			<td style="width:200px;">20740-0-08-65</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Propylene glycol</td>
			<td style="width:220px;">Reanal</td>
			<td style="width:200px;">08860-1-08-65</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Dimethyl sulfoxide</td>
			<td style="width:220px;">Reanal</td>
			<td style="width:200px;">00190-1-01-65</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Collagenase</td>
			<td style="width:220px;">Gibco</td>
			<td style="width:200px;">9001-12-1</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Trypsin</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td style="width:200px;">T8003</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="24">
			<td height="24" style="height:24px;width:232px;">DNase I</td>
			<td style="width:220px;">Panreac AppliChem</td>
			<td style="width:200px;">A3778</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Trypan blue</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td style="width:200px;">T6146</td>
			<td style="width:296px;"></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:232px;">Phospate buffered saline (PBS)</td>
			<td style="width:220px;">Sigma-Aldrich</td>
			<td>P4417</td>
			<td>Tablets for preparation of 200 ml PBS solution</td>
		</tr>
	</tbody>
</table>

cryopreservation of zebrafish spermatogonia by whole testes needle immersed ultra-rapid cooling

Current trends in science and biotechnology lead to creation of thousands of new lines in model organisms thereby leading to the necessity for new methods for safe storage of genetic resources beyond the common practices of keeping breeding colonies. The main purpose of this study was to adapt the needle immersed vitrification (NIV) procedure to cryopreserve whole zebrafish testes. Cryopreservation of early-stage germ cells by whole testes NIV offers possibilities for the storage of zebrafish genetic resources, especially since after transplantation they can mature into both male and female gametes. Testes were excised, pinned on an acupuncture needle, equilibrated in two cryoprotective media (equilibration solution containing 1.5 M methanol and 1.5 M propylene glycol; and vitrification solution containing 3 M dimethyl sulfoxide and 3 M propylene glycol) and plunged into liquid nitrogen. Samples were warmed in a series of three consequent warming solutions. The main advantages of this technique are (1) the lack of spermatozoa after digestion of warmed testes thus facilitating downstream manipulations; (2) ultra-rapid cooling enabling the optimal exposure of tissues to liquid nitrogen therefore maximizing the cooling and reducing the required concentration of cryoprotectants, thereby reducing their toxicity; (3) synchronous exposure of several testes to cryoprotectants and liquid nitrogen; and (4) repeatability demonstrated by obtaining viability of above 50% in five different zebrafish strains.

On average, the number of early-stage germ cells isolated from a single fresh zebrafish testis varied between 40,000 and 200,000 cells depending on the size of the fish. When digesting fresh zebrafish testes in all 5 strains, early-germ cells were not the only cells present in the cell suspensions (Figure 2). Beside the early-stage germ cells, numerous spermatozoa were found as well. On the other hand, there were far less spermatozoa after digestion of cryopr...

Watch this Scientific Journal Video about Cryopreservation of Zebrafish Spermatogonia by Whole Testes Needle Immersed Ultra-Rapid Cooling at JoVE.com

Cryopreservation of Zebrafish Spermatogonia by Whole Testes Needle Immersed Ultra-Rapid Cooling

The main purpose of this study was to adapt the needle immersed vitrification (NIV) procedure to cryopreserve whole zebrafish testes. Additionally, the repeatability of the method in five different zebrafish strains was tested.

Current trends in science and biotechnology lead to creation of thousands of new lines in model organisms thereby leading to the necessity for new methods ...

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Research

JoVE Journal

Biology

8.2K Views.  Szent István University. The main purpose of this study was to adapt the needle immersed vitrification (NIV) procedure to cryopreserve whole zebrafish testes. Additionally, the repeatability of the method in five different zebrafish strains was tested.

Video: Cryopreservation of Zebrafish Spermatogonia by Whole Testes Needle Immersed Ultra-Rapid Cooling

Transplantation of Whole Kidney Marrow in Adult Zebrafish

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an organism. The functional definition of a HSC is a transplanted cell that has the ability to reconstitute all the blood lineages of an irradiated recipient long term. This designation was established by decades of seminal work in mammalian systems. Using hematopoietic cell transplantation (HCT) and reverse genetic manipulations in the mouse, the underlying regulatory factors of HSC biology are beginning to be unveiled, but are still largely under-explored. Recently, the zebrafish has emerged as a powerful genetic model to study vertebrate hematopoiesis. Establishing HCT in zebrafish will allow scientists to utilize the large-scale genetic and chemical screening methodologies available in zebrafish to reveal novel mechanisms underlying HSC regulation. In this article, we demonstrate a method to perform HCT in adult zebrafish. We show the dissection and preparation of zebrafish whole kidney marrow, the site of adult hematopoiesis in the zebrafish, and the introduction of these donor cells into the circulation of irradiated recipient fish via intracardiac injection. Additionally, we describe the post-transplant care of fish in an "ICU" to increase their long-term health. In general, gentle care of the fish before, during, and after the transplant is critical to increase the number of fish that will survive more than one month following the procedure, which is essential for assessment of long term (&lt;3 month) engraftment. The experimental data used to establish this protocol will be published elsewhere. The establishment of this protocol will allow for the merger of large-scale zebrafish genetics and transplant biology.

In this article, we demonstrate a method to perform HCT in adult zebrafish.

Hematopoietic stem cells (HSC) are a rare population of pluripotent cells that maintain all the differentiated blood lineages throughout the life of an ...

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. The protocol is a modified version of the standard in situ hybridization using alkaline phosphatase and substrates such as NBT/BCIP and Fast Red 1,2. This protocol utilizes standard digoxygenin and fluorescein labeled probes along with tyramide signal amplification (TSA) 3. The commercially available TSA kits allow flexible experimental design as fluorescence emission from green to far-red can be used in combination with various nuclear stains, such as propidium iodide, or fluorescence immunohistochemistry for proteins. TSA produces a reactive fluorescent substrate that quickly covalently binds to moieties, typically tyrosine residues, in the immediate vicinity of the labeled antisense riboprobe. The resulting staining patterns are high resolution in that subcellular localization of the mRNA can be observed using laser scanning confocal microscopy 3,4. One can observe nascent transcripts at the chromosomal loci, distinguish nuclear and cytoplasmic staining and visualize other patterns such as cortical localization of mRNA. Studies in Drosophila indicate that roughly 70% of mRNAs exhibit specific patterns of subcellular localization that frequently correlate with the function of the encoded protein 5. When combined with computer-aided reconstruction of 3D confocal datasets, our protocol allows the detailed analysis of mRNA distribution with sub-cellular resolution in whole vertebrate embryos.

Zebrafish Whole Mount High-Resolution Double Fluorescent In Situ Hybridization

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology. Here, we present a high-resolution double fluorescent in situ hybridization protocol for analyzing the precise expression pattern of a single gene and for determining the overlap of the expression domains of two genes. We include a propidium iodide nuclear counter-stain to highlight tissue organization.

Whole mount in situ hybridization is one of the most widely used techniques in developmental biology.  Here, we present a high-resolution double ...

Generating Chimeric Zebrafish Embryos by Transplantation

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an organism that contains cells from more than one animal; mosaics are one type of chimera in which cells from more than one genotype are mixed, usually wild-type and mutant. In the zebrafish, chimeras can be readily made by transplantation of cells from a donor embryo into a host embryo at the appropriate embryonic stage. Labeled donor cells are generated by injection of a lineage marker, such as a fluorescent dye, into the one-cell stage embryo. Labeled donor cells are removed from donor embryos and introduced into unlabeled host embryos using an oil-controlled glass pipette mounted on either a compound or dissecting microscope. Donor cells can in some cases be targeted to a specific region or tissue of the developing blastula or gastrula stage host embryo by choosing a transplantation site in the host embryo based on well-established fate maps.

A step-by-step guide to generating targeted chimeric zebrafish embryos by transplantation at the blastula or gastrula stage.

One of the most powerful tools used to gain insight into complex developmental processes is the analysis of chimeric embryos. A chimera is defined as an ...

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to the original protocol that both streamline the procedure and increase sample uniformity.  First, we normalize all sperm volumes using freezing media that does not contain the cryoprotectant. Second, cryopreserved sperm are stored in cryovials instead of capillary tubes.  The rates of sperm freezing and thawing (&delta;&deg;C/time) are probably the two most critical variables to control in this procedure.  For this reason, do not substitute different tubes for those specified.  Working in teams of 2 it is possible to freeze the sperm of 100 males per team in ~2 hrs.  Sperm cryopreserved using this protocol has an average of 25% fertility (measured as the number of viable embryos generated in an in vitro fertilization divided by the total number of eggs fertilized) and this percent fertility is stable over many years.

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a high-throughput sperm cryopreservation protocol for zebrafish. Sperm cryopreserved using this protocol has an average of 25% fertility in subsequent vitro fertilization and is stable over many years.

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to ...

Making Gynogenetic Diploid Zebrafish by Early Pressure

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from heterozygous females allow F1 mutagenized females to be screened directly for deleterious mutations in an accelerated forward genetic screen. Streisinger et al.1,2 described methods for making gynogenetic (homozygous) diploid zebrafish by activating zebrafish eggs with ultraviolet light-inactivated sperm and preventing either the second meiotic or the first zygotic cell division using physical treatments (heat or pressure) that deploymerize microtubules. The "early pressure" (EP) method blocks the meiosis II, which occurs shortly after fertilization. The EP method produces a high percentage of viable embryos that can develop to fertile adults of either sex. The method generates embryos that are homozygous at all loci except those that were separated from their centromere by recombination during meiosis I. Homozygous mutations are detected in EP clutches at between 50% for centromeric loci and less than 1% for telomeric loci. This method is reproduced verbatim from the Zebrafish Book3.

This is a method for generating gynogenetic diploid zebrafish embryos (embryos whose only genetic contribution comes from the mother) by blocking the second meiotic division immediately after fertilization with ultraviolet light-inactivated sperm. EP embryos are not fully homozygous due to recombination during the first meiotic division, however they are homozygous at all loci that have not been separated from their centromere by recombination.

Heterozygosity in diploid eukaryotes often makes genetic studies cumbersome. Methods that produce viable homozygous diploid offspring directly from ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Cryopreservation of Mouse Embryos by Ethylene Glycol-Based Vitrification

Cryopreservation of mouse embryos is a technological basis that supports biomedical sciences, because many strains of mice have been produced by genetic modifications and the number is consistently increasing year by year. Its technical development started with slow freezing methods in the 1970s1, then followed by vitrification methods developed in the late 1980s2. Generally, the latter technique is advantageous in its quickness, simplicity, and high survivability of recovered embryos. However, the cryoprotectants contained are highly toxic and may affect subsequent embryo development. Therefore, the technique was not applicable to certain strains of mice, even when the solutions are cooled to 4&deg;C to mitigate the toxic effect during embryo handling. At the RIKEN BioResource Center, more than 5000 mouse strains with different genetic backgrounds and phenotypes are maintained3, and therefore we have optimized a vitrification technique with which we can cryopreserve embryos from many different strains of mice, with the benefits of high embryo survival after vitrifying and thawing (or liquefying, more precisely) at the ambient temperature4.
Here, we present a vitrification method for mouse embryos that has been successfully used at our center. The cryopreservation solution contains ethylene glycol instead of DMSO to minimize the toxicity to embryos5. It also contains Ficoll and sucrose for prevention of devitrification and osmotic adjustment, respectively. Embryos can be handled at room temperature and transferred into liquid nitrogen within 5 min. Because the original method was optimized for plastic straws as containers, we have slightly modified the protocol for cryotubes, which are more easily accessible in laboratories and more resistant to physical damages. We also describe the procedure of thawing vitrified embryos in detail because it is a critical step for efficient recovery of live mice. These methodologies would be helpful to researchers and technicians who need preservation of mouse strains for later use in a safe and cost-effective manner.

An ethylene glycol-based vitrification method for mouse embryos is described. It is advantageous to other methods in its simplicity and low embryonic toxicity, and therefore can be broadly applicable to many strains of mice, including inbred and gene-modified mice.

Cryopreservation of mouse embryos is a technological basis that supports biomedical sciences, because many strains of mice have been produced by genetic ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

Imaging Centrosomes in Fly Testes

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell differentiation. Centrosomes are essential to determine the cell division axis during mitosis and to nucleate cilia during interphase. The identity of the proteins that mediate these dynamic changes remains only partially known, and the function of many of the proteins that have been implicated in these processes is still rudimentary. Recent work has shown that Drosophila spermatogenesis provides a powerful system to identify new proteins critical for centrosome function and formation as well as to gain insight into the particular function of known players in centrosome-related processes. Drosophila is an established genetic model organism where mutants in centrosomal genes can be readily obtained and easily analyzed. Furthermore, recent advances in the sensitivity and resolution of light microscopy and the development of robust genetically tagged centrosomal markers have transformed the ability to use Drosophila testes as a simple and accessible model system to study centrosomes. This paper describes the use of genetically-tagged centrosomal markers to perform genetic screens for new centrosomal mutants and to gain insight into the specific function of newly identified genes.

Imaging of centrosomal proteins during Drosophila spermatogenesis is a powerful method to identify new proteins critical for centrosome biology as well as to elucidate the particular function of known players in this process.

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell ...

Analysis of Apoptosis in Zebrafish Embryos by Whole-mount Immunofluorescence to Detect Activated Caspase 3

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis in zebrafish embryos. The whole-mount analysis provides spatial information in regard to tissue specificity of apoptosing cells, although sectioning and/or colabeling is ultimately required to pinpoint the exact cell types undergoing apoptosis. The whole-mount Casp3 assay is optimized for analysis of fixed embryos between the 4-cell stage and 32 hr-post-fertilization and is useful for a number of applications, including analysis of zebrafish mutants and morphants, overexpression of mutant and wild-type mRNAs, and exposure to chemicals. Compared to acridine orange staining, which can identify apoptotic cells in live embryos in a matter of hours, Casp3 and TUNEL assays take considerably longer to complete (2-4 days). However, because of the dynamic nature of apoptotic cell formation and clearance, analysis of fixed embryos ensures accurate comparison of apoptotic cells across multiple samples at specific time points. We have also found the Casp3 assay to be superior to analysis of apoptotic cells by the whole-mount TUNEL assay in regard to cost and reliability. Overall, the Casp3 assay represents a robust, highly reproducible assay in which to analyze apoptotic cells in early zebrafish embryos.

Certain genetic perturbations or exposure to toxins can disrupt normal developmental processes leading to death of specific cell types. The analysis of activated Caspase 3 by whole-mount immunofluorescence in zebrafish embryos reveals stage- and tissue-specific localization of cells specifically undergoing apoptosis.

Whole-mount immunofluorescence to detect activated Caspase 3 (Casp3 assay) is useful to identify cells undergoing either intrinsic or extrinsic apoptosis ...