This research was supported by NIH grants GM055360 to SLB and U54HD068157 to RGM. JMB was supported by the T32 Genetics Training Grant at the University of Pennsylvania (GM008216).

The STA-PUT protocol involves three stages: 1) Set up of the apparatus and reagents, 2) Preparation of cell suspension from whole testes, and 3) Cell loading, sedimentation, and fraction collection. When performed by a team of two researchers, the protocol takes eight hours on average.
1. Setting up the STA-PUT Apparatus (Figure 1)
***STA-PUT apparatus should be placed in a 4&#176;C large refrigerator or a cold room that can also accommodate a fraction collector, if that method of collection is preferred.
<ol>
	<li>The night before (or at least a few hours before) you perform the method, wash all equipment (especially the glassware and tubing) and sterilize with 70% ethanol. Let equipment dry completely before assembling the apparatus as illustrated in Figure 1.</li>
	<li>Secure the two 2 L cylinders (Figures 1B and C) and the cell loading chamber (Figure 1A) to the top platform and connect all with two small pieces of tubing with tube clamps. Clamp all tubes closed. Seal the spout on the right-most 2 L cylinder.</li>
	<li>Place a small stir bar in the cell loading chamber (Figure 1A) and a larger stir bar in the left-most 2 L cylinder (Figure 1B) that will contain the 2% BSA.</li>
	<li>Place the 2 L sedimentation chamber on the platform (Figure 1D). Place the metal baffle (Figure 1F) directly on top of the opening in the bottom of the sedimentation chamber (Figure 1D). This is critical, as the baffle prevents vortexing of the liquid and disruption of the cell gradient during fraction collection. Place the lid on top of the sedimentation chamber.</li>
	<li>After applying a very small amount of vacuum grease to the ground glass joint of the three-way stopcock (Figure 1G), clamp the stopcock to the bottom of the sedimentation chamber, connecting the ground glass joints of the stopcock and the sedimentation chamber.</li>
	<li>Connect the cell-loading chamber (Figure 1A) to the right outlet of the stopcock with tubing. Close the stopcock.</li>
	<li>Attach the cell fractionation tubing to the left outlet of the stopcock. The fractionation tubing comprises a piece of tubing with a glass Pasteur pipette connected to the open end. A piece of smaller bore tubing is attached to the narrow end of the glass pipette. The narrow pipette restricts the flow of the cell suspension during fraction collecting. Clamp this small tube at the very bottom.</li>
	<li>Prepare 2 L Krebs (1x) buffer the day of the experiment (Table 1). Then, prepare 550 ml 2% BSA in 1x Krebs, 550 ml 4% BSA in 1x Krebs, and 50 ml 0.5% BSA in 1x Krebs. Filter and cool these solutions to 4 &#176;C.</li>
	<li>Pour the 4% BSA solution in the right 2 L cylinder (Figure 1C) and the 2% BSA solution in the left 2 L cylinder (Figure 1B). Make sure there are no large bubbles in the tubing that connects these cylinders.</li>
	<li>Pour Krebs buffer into the cell loading chamber and fill the tubing connecting this chamber with the sedimentation chamber, making sure there are no large bubbles, as these could disturb the gradient. Squeezing or flicking the tubing gently helps to remove the bubbles. Make sure all of the Krebs buffer is in the tube and not in the cell chamber.</li>
	<li>Allow a small amount of Krebs buffer to flow into the sedimentation chamber and then drain almost all the buffer into the cell fractionation tubing in order to fill the tube and remove any large bubbles.</li>
	<li>Place the fraction collector directly under the sedimentation chamber. Make sure that all the fraction tubes are in place and covered with plastic wrap to prevent contamination.</li>
</ol>
2. Isolating Spermatogenic Cells from Whole Testes
<ol>
	<li>Dissect testes from ca12 adult mice (preferably at least eight weeks old) and place in Krebs buffer at room temperature to wash off any contaminating material. This protocol being demonstrated involves the use of laboratory mice and was executed in compliance with the guidelines of the University of Pennsylvania Institutional Animal Care and Use Committee.</li>
	<li>Add 45 mg collagenase to 50 ml Krebs buffer in a conical tube right before you intend to add the seminiferous tubules. Allow this solution to heat up to 33 &#176;C for a few minutes. Split evenly into two 50 ml conical tubes.</li>
	<li>Decapsulate the testes in a separate plate containing 8 ml Krebs buffer, discarding the tunica albuginea and releasing the seminiferous tubules. The tunica albuginea is the thin membrane that surrounds the seminiferous tubules. The tubules should readily detach from the tunica albuginea. To do this, make an incision in the tunica albuginea, hold this membrane with a pair of forceps, and push the tubules out of and away from the membrane with another pair of forceps.</li>
	<li>Add 4 ml Krebs buffer containing the tubules to each conical tube containing 25 ml of collagenase solution and incubate, shaking, at 33 &#176;C for 10 min. At the end, the tubules should have a "spaghetti-like" appearance.</li>
	<li>Allow the tubules to settle for ~5 min to the bottom of the tube. Pour out the supernatant and wash 2x in 25 ml Krebs buffer (at room temperature), allowing the tubules to settle to the bottom of the tube each time. Leave ~5 ml Krebs buffer in each tube.</li>
	<li>While washing, add 30 mg trypsin to 50 ml Krebs buffer in a falcon tube right before you intend to add the seminiferous tubules. Allow this solution to heat up to 33 &#176;C for a few minutes. Split evenly into two 50 ml conical tubes.</li>
	<li>Add 25 ml trypsin solution to each of the two tubes containing the tubules. Add 3 &#956;g DNAse (1 &#956;g/10ml) to each tube to prevent cells from clumping. Incubate, shaking, at 33 &#176;C for 10 min.</li>
	<li>Use a wide bore pipette to agitate the solution containing the tubules, pipetting them in and out approximately 10x. The solution should begin to look more like a single cell suspension.</li>
	<li>Incubate, shaking, at 33 &#176;C for an additional 10 min. Use a wide bore pipette to disperse the tubules into a single cell suspension, pipetting them in and out approximately 25x. If you still see a lot of tubules or cell clumps, disperse more with the pipette and/or add another 3 &#956;g DNAse to each tube (double initial concentration).</li>
	<li>Filter the single cell suspension through a 100 &#956;m mesh cell strainer (one for each tube of 30 ml cell suspension). Combine the cell suspensions and count the total number of cells (should be between 7-8 x 108 cells). ***DO NOT LOAD MORE THAN 800,000,000 cells onto the gradient.</li>
	<li>Based on the cell count obtained in step 2.10, pellet the appropriate volume of filtered cells at 450 rcf for 5 min and wash with 30 ml Krebs buffer. Repeat. ***Usually 22 testes will yield more than 800,000,000 cells, but do not load more than this number of cells.</li>
	<li>Carefully resuspend cells in 25 ml 0.5% BSA containing between 2.5 and 5 &#956;g DNAse (depending on the degree of cell clumping) without creating bubbles. Make sure the cells do not clump. Filter the single cell suspension through a 100 &#956;m mesh cell strainer. The cells are now ready to load. Mix well before loading by inverting the tube a few times.</li>
</ol>
3. Cell Loading and Sedimentation
<ol>
	<li>Make sure that the stopcock is closed, but in the position that will allow liquid to flow from the cell chamber (Figure 1A) into the sedimentation chamber (Figure 1D).</li>
	<li>Turn both stir bar plates on to a low setting (approximately 70 rpm) to allow the stir bars to move continuously, but slowly.</li>
	<li>Pour the cell suspension into the cell chamber (Figure 1A) and open the stopcock so that the cells flow slowly into the sedimentation chamber at a rate of 10 ml/min (Figure 1D). Close the stopcock. Flow rate can be determined by marking volume intervals on the sedimentation chamber.</li>
	<li>Pour 5 ml of 0.5% BSA solution into the cell chamber and drain into the sedimentation chamber at a rate of 10 ml/min. Close the stopcock. This step will wash the cells out of the cell chamber. Repeat 4x.</li>
	<li>Prepare the gradient: Open the clamps between the cell chamber (Figure 1A) and the two 2 L cylinders (Figure 1B and C), and begin to drain the liquid into the sedimentation chamber at a rate of 40 ml/min (Figure 1D) immediately. Adjust the flow rate so it takes approximately 20-30 min to load the BSA gradient into the sedimentation chamber. A thin, undisturbed layer of cells lying on top of the BSA gradient should be visible. Once most of the BSA is loaded, close the stopcock and turn it to the position that will allow liquid to drain from the sedimentation chamber into the fractionation tube.</li>
	<li>Turn off the stir plates.</li>
	<li>Allow the cells to sediment for one hour and 45 minutes. DO NOT DISTURB THE APPARATUS DURING THIS TIME. Any mechanical agitation could disturb the gradient.</li>
</ol>
4. Fractionation and Analysis of Fractions
***As you perform the following steps, be careful not to disturb the gradient. If the BSA gradient is disturbed, the procedure will not work!
<ol>
	<li>Once the cells have sedimented, slowly drain 50 ml from the sedimentation chamber into a 50 ml conical tube and set aside. This fraction usually contains unwanted clumps of cells and debris.</li>
	<li>Attach the fraction tube to the fraction collector. ***It is also possible to collect the fractions by hand if a fraction collector is not available.</li>
	<li>Collect fractions: Adjust the flow rate with the stopcock so that 10 ml (filling one collection tube) is collected every 45 sec.</li>
	<li>Cap the fraction collection tubes and spin for 5 min at 450 rcf at 4 &#176;C. Pour off supernatant gently, resuspend cells in remaining liquid, and keep cells on ice.</li>
	<li>Once all fractions are collected, microscopically analyze the fractions for different cell populations. Different cell types can be distinguished based on cell size and nuclear morphology8,15 . Figure 2 illustrates "representative results" for the three pooled fractions that are enriched for (a) somatic and meiotic cells and spermatogonia, (b) round spermatids, and (c) elongating and condensing spermatids.
	<ul>
		<li>Type A spermatogonia are 12-14&#956;m ca. in diameter and have round nuclei that are homogenous in chromatin composition.</li>
		<li>Type B spermatogonia are 8-9&#956;m ca. in diameter and have round nuclei that show more dense heterochromatin along the nuclear periphery.</li>
		<li>Pre-leptotene spermatocytes are 7.5-8.2&#956;m ca. in diameter, have less cytoplasm than Type B spermatogonia, and have nuclei that look similar to those in Type B spermatogonia.</li>
		<li>Leptotene primary spermatocytes are 8-10 &#956;m ca. in diameter and have nuclei that look similar to those of pre-leptotene spermatocytes, but seem to gain more dense chromatin at the nuclear membrane.</li>
		<li>Zygotene primary spermatocytes are 10-12 &#956;m ca. in diameter and have nuclei that look similar to those of leptotene primary spermatocytes.</li>
		<li>Pachytene spermatocytes are anywhere from 12-18 &#956;m ca. in diameter and consist of a thin rim of cytoplasm surrounding a large nucleus. More clumps of dense chromatin can be seen in the nucleus.</li>
		<li>Round spermatids are 10 &#956;m ca. in diameter and have a round nucleus with a densely-staining chromocenter in the middle.</li>
		<li>Residual bodies are ~6.5 &#956;m in diameter, are round, and lack a nucleus.</li>
		<li>Elongating and condensing spermatids are similar in size to round spermatids, but have a unique, sickle-shaped nucleus.</li>
	</ul>
	<ol>
		<li>Start with fraction 15 and analyze every five fractions up to 85. Usually, fractions below 15 will be too mixed and clumpy, and fractions above 85 will contain few to no usable cells.</li>
		<li>To analyze a fraction, take 5uL of liquid from the fraction collection tube (once cells have been resuspended after centrifugation) and add to 5uL of 8% formaldehyde in Krebs buffer. Allow the fixed cells to sit at room temperature for five minutes.</li>
		<li>Add 5uL of 0.1% Triton and DAPI (5 &#956;/ml Krebs buffer) to the fixed cells. Allow the cells to sit at room temperature for 5 min.</li>
		<li>Place 10 &#956;l of the resulting solution onto a slide, cover with a cover slip, and analyze with a fluorescence microscope to determine the purity of each fraction.</li>
		<li>Combine fractions that are similar in size and nuclear morphology (see "Representative Results") to create the following populations of cells: meiotic and somatic diploid cells, round spermatids, and condensing/elongating spermatids.</li>
	</ol>
	</li>
</ol>

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The authors declare that they have no competing financial interests. This protocol being demonstrated involves the use of laboratory mice and was executed in compliance with all relevant guidelines, regulations and regulatory agencies. Animals used in this protocol are maintained under the guidance and approval of the University of Pennsylvania Institutional Animal Care and Use Committee (IACUC protocol #804284).

Those who study spermatogenesis rely on animal models for spermatogenic cell samples, as a reliable cell culture system does not yet exist for generating all spermatogenic cell types3. Although spermatogenic cells are readily collected from whole testes, only a mixed population results. This poses a problem for those who wish to study specific subtypes of these cells, such as meiotic cells, round spermatids, and condensing spermatids. Three different methods are currently used to separate these subtypes of ...

Mammalian spermatogenesis is a complex differentiation process that occurs in several stages in the seminiferous tubules of the testes1. Briefly, stem-like spermatogonia that reside near the epithelium of the seminiferous tubule divide and differentiate into spermatocytes, which then undergo meiotic divisions. After meiosis is complete, the resulting haploid cells, or round spermatids, undergo spermiogenesis, a differentiation process that involves the shedding of cytoplasm and compaction of the nucleus. Spermatids gradually develop a flagellum and undergo elongation and condensation of the nucleus, producing elongating and then condensing spermatids, respectively. The end products are spermatozoa, which are released into the lumen of the seminiferous tubule and ultimately into the epididymis where they mature further.
Because the process of spermatogenesis relies on special hormonal and molecular conditions in the testes, a reliable in vitro culture system for the entire process of spermatogenesis has not yet been developed2,3. Culture methods have been developed for creating "primordial germ cell-like cells" and haploid, "round spermatid-like cells" from stem cells, but these methods are not yet able to generate large numbers of these cells and fail to produce later spermatogenic cell types4,5. Fortunately, the spermatogenic cell types differ significantly in size, which allows for a single-cell suspension obtained from whole testes to be separated with a liquid gradient. The STA-PUT method, demonstrated here, uses a linear BSA gradient and simple sedimentation to separate spermatogenic cells based on size and mass6-9.
The STA-PUT method has several advantages over the other two most widely used methods to separate spermatogenic cell types: FACS and elutriation10-13. The STA-PUT apparatus requires only several pieces of specialized glassware assembled in a cold room or large refrigerator. Thus, it is less expensive than using a cell sorter or an elutriator. The STA-PUT method yields higher amounts of cells per cell type and testis than can be sorted by FACS in a comparable time frame, although the purity of each cell population is not as high as those obtained with FACS11. Cell sorting utilizing magnetic beads (magnetic activated cell sorting, MACS) has recently been successfully employed for enrichment of spermatogonia from a mixed testicular cell population, but it is currently unsuitable for separating spermatocytes or spermatids due to lack of knowledge of appropriate surface markers14. An additional advantage of the STA-PUT method over FACS or MACS is the ability to isolate viable cells suitable for subsequent culture because, in contrast to most FACS protocols, it does not require any DNA or other types of staining. For studies that require large yields of spermatogenic cells types at ~90% purity, the STA-PUT is an ideal method.

<table><tbody><tr><td>BSA</td><td>Affymetrix/USB</td><td>10857 100MG</td><td>Alternate can be used.</td></tr><tr><td>0.5%, 2%, and 4% BSA solutions</td><td>Dissolve 0.25 g, 11 g, and 22 g (respectively) in total volumes of 50 ml, 550 ml, and 550 ml of 1x KREBS (respectively).</td><td>To be made day of the STA-PUT procedure. Filter sterilize.</td><td></td></tr><tr><td>KREBS (10x)</td><td>3.26 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; + 139.5 g NaCl +5.89 g MgSO&lt;sub&gt;4&lt;/sub&gt;/7H&lt;sub&gt;2&lt;/sub&gt;0 + 40 g Dextrose + 3.78 g CaCl&lt;sub&gt;2&lt;/sub&gt;/2H&lt;sub&gt;2&lt;/sub&gt;0 + 7.12 g KCl, bring to 2 L with ddH&lt;sub&gt;2&lt;/sub&gt;0</td><td>Autoclave/filter and store at 4 &amp;deg;C for several months.</td><td></td></tr><tr><td>KREBS (1x)</td><td>Dissolve 4.24 g NaHCO&lt;sub&gt;3 &lt;/sub&gt;in 100 ml ddH&lt;sub&gt;2&lt;/sub&gt;0. Add 200 ml 10x KREBS. Bring to 2 L with ddH&lt;sub&gt;2&lt;/sub&gt;0.</td><td>To be made day of the STA-PUT procedure. Filter sterilize.</td><td></td></tr><tr><td>Collagenase</td><td>Sigma</td><td>C9891-1G</td><td> </td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>DNAse</td><td>Sigma</td><td>DNEP-5MG</td><td> </td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>Trypsin</td><td>Sigma</td><td>T9201-1G</td><td> </td></tr><tr><td></td><td></td><td></td><td></td></tr><tr><td>DAPI</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td>Triton-X100</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td>Formaldehyde (~37%)</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td> </td><td></td><td></td><td></td></tr><tr><td>TABLE 2: EQUIPMENT</td><td></td><td></td><td></td></tr><tr><td> </td><td></td><td></td><td></td></tr><tr><td>Equipment</td><td>Company</td><td>Catalog #</td><td>Comments</td></tr><tr><td>Complete STA-PUT apparatus</td><td>ProScience Glass Shop</td><td>STA-PUT (this procedure uses the standard sedimentation chamber: Cat. No. 56700-500)</td><td>Includes all glassware and equipment for the apparatus. Alternate glassware is available for performing STA-PUT with smaller cell numbers.</td></tr><tr><td>10 &amp;mu;m mesh filter</td><td>Fisherbrand</td><td>22363549</td><td> </td></tr><tr><td>Mouse dissection tools</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td>14 ml round bottom fraction collection tubes with caps</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td>Need approximately 100 tubes</td><td></td><td></td><td></td></tr><tr><td>Fraction collector</td><td>GE Healthcare Life Sciences</td><td>Model: Frac-920, Product code: 18-1177-40</td><td>Alternate can be used.</td></tr><tr><td>Microscope slides, cover glass</td><td>Preference of researcher</td><td></td><td></td></tr><tr><td>Light/Fluorescent Microscope</td><td>Preference of researcher</td><td></td><td></td></tr></tbody></table>

separation of spermatogenic cell types using sta-put velocity sedimentation

Mammalian spermatogenesis is a complex differentiation process that occurs in several stages in the seminiferous tubules of the testes. Currently, there is no reliable cell culture system allowing for spermatogenic differentiation in vitro, and most biological studies of spermatogenic cells require tissue harvest from animal models like the mouse and rat. Because the testis contains numerous cell types - both non-spermatogenic (Leydig, Sertoli, myeloid, and epithelial cells) and spermatogenic (spermatogonia, spermatocytes, round spermatids, condensing spermatids and spermatozoa) - studies of the biological mechanisms involved in spermatogenesis require the isolation and enrichment of these different cell types. The STA-PUT method allows for the separation of a heterogeneous population of cells - in this case, from the testes - through a linear BSA gradient. Individual cell types sediment with different sedimentation velocity according to cell size, and fractions enriched for different cell types can be collected and utilized in further analyses. While the STA-PUT method does not result in highly pure fractions of cell types, e.g. as can be obtained with certain cell sorting methods, it does provide a much higher yield of total cells in each fraction (~1 x 108 cells/spermatogenic cell type from a starting population of 7-8 x 108 cells). This high yield method requires only specialized glassware and can be performed in any cold room or large refrigerator, making it an ideal method for labs that have limited access to specialized equipment like a fluorescence activated cell sorter (FACS) or elutriator.

The ideal result from the STA-PUT procedure is a fairly noticeable separation of cells from the testes based on cell size and density. While cells isolated from the testes are sedimenting through the BSA gradient, several distinct bands of cells can be observed. Any clumps of cells tend to sink to the bottom of the gradient and will not contaminate the other fractions. A little further up the gradient will be the large somatic and meiotic cells. Farther up the gradient still will be smaller round spermatids. At the ...

Watch this Scientific Journal Video about Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation at JoVE.com

Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation

The STA-PUT method allows for the separation of different populations of spermatogenic cells based on size and density.

Mammalian spermatogenesis is a complex  differentiation process that occurs in several stages in the seminiferous  tubules of the testes. Currently, there ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

25.7K Views.  University of Pennsylvania. The STA-PUT method allows for the separation of different populations of spermatogenic cells based on size and density.

Video: Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation

Step-specific Sorting of Mouse Spermatids by Flow Cytometry

The differentiation of mouse spermatids is one critical process for the production of a functional male gamete with an intact genome to be transmitted to the next generation. So far, molecular studies of this morphological transition have been hampered by the lack of a method allowing adequate separation of these important steps of spermatid differentiation for subsequent analyses. Earlier attempts at proper gating of these cells using flow cytometry may have been difficult because of a peculiar increase in DNA fluorescence in spermatids undergoing chromatin remodeling. Based on this observation, we provide details of a simple flow cytometry scheme, allowing reproducible purification of four populations of mouse spermatids fixed with ethanol, each representing a different state in the nuclear remodeling process. Population enrichment is confirmed using step-specific markers and morphological criterions. The purified spermatids can be used for genomic and proteomic analyses.

We describe a sorting strategy for mouse spermatids using flow cytometry. Spermatids are sorted into four highly pure populations, including round (spermiogenesis steps 1-9), early elongating (spermiogenesis steps 10-12), late elongating (spermiogenesis steps 13-14) and elongated spermatids (spermiogenesis steps 15-16). DNA staining, size and granulosity are used as selection parameters.

The differentiation of mouse spermatids is one critical process for the production of a functional male gamete with an intact genome to be transmitted to ...

Developmental Biology

A Standardized Approach for Multispecies Purification of Mammalian Male Germ Cells by Mechanical Tissue Dissociation and Flow Cytometry

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. Currently, it allows the discrimination of up to 9 murine germ cell populations with high yield and purity. This high-resolution in discrimination and purification is possible due to unique changes in chromatin structure and quantity throughout spermatogenesis. These patterns can be captured by flow cytometry of male germ cells stained with fluorescent DNA-binding dyes such as Hoechst-33342 (Hoechst). Herein is a detailed description of a recently developed protocol to isolate mammalian testicular germ cells. Briefly, single cell suspensions are generated from testicular tissue by mechanical dissociation, double stained with Hoechst and propidium iodide (PI) and processed by flow cytometry. A serial gating strategy, including the selection of live cells (PI negative) with different DNA content (Hoechst intensity), is used during FACS sorting to discriminate up to 5 germ cell types. These include, with corresponding average purities (determined by microscopy evaluation): spermatogonia (66%), primary (71%) and secondary (85%) spermatocytes, and spermatids (90%), further separated into round (93%) and elongating (87%) subpopulations. Execution of the entire workflow is straightforward, allows the isolation of 4 cell types simultaneously with the appropriate FACS machine, and can be performed in less than 2 h. As reduced processing time is crucial to preserve the physiology of ex vivo cells, this method is ideal for downstream high-throughput studies of male germ cell biology. Moreover, a standardized protocol for multispecies purification of mammalian germ cells eliminates methodological sources of variables and allows a single set of reagents to be used for different animal models.

This work describes the standardization of a method to obtain purified germ cell populations from testicular tissue of different mammalian species. It is a straightforward protocol that combines mechanical testis dissociation, staining with Hoechst-33342 and propidium iodide, and FACS sorting, with wide applications in comparative studies of male reproductive biology.

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. ...

A Seminiferous Tubule Squash Technique for the Cytological Analysis of Spermatogenesis Using the Mouse Model

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during meiosis can lead to infertility, pregnancy loss or genetic defects. Commencing at the onset of puberty and continuing throughout adulthood, continuous semi-synchronous waves of spermatocytes undergo spermatogenesis and ultimately form haploid sperm. The first wave of mouse spermatocytes undergoing meiotic initiation appear at day 10 post-partum (10 dpp) and are released into the lumen of seminiferous tubules as mature sperm at 35 dpp. Therefore, it is advantageous to utilize mice within this developmental time-window in order to obtain highly enriched populations of interest. Analysis of rare cell stages is more difficult in older mice due to the contribution of successive spermatogenic waves, which increase the diversity of the cellular populations within the tubules. The method described here is an easily implemented technique for the cytological evaluation of the cells found within the seminiferous tubules of mice, including spermatogonia, spermatocytes, and spermatids. The tubule squash technique maintains the integrity of isolated male germ cells and allows examination of cellular structures that are not easily visualized with other techniques. To demonstrate the possible applications of this tubule squash technique, spindle assembly was monitored in spermatocytes progressing through the prophase to metaphase I transition (G2/MI transition). In addition, centrosome duplication, meiotic sex chromosome inactivation (MSCI), and chromosome bouquet formation were assessed as examples of the cytological structures that can be observed using this tubule squash method. This technique can be used to pinpoint specific defects during spermatogenesis that are caused by mutation or exogenous perturbation, and thus, contributes to our molecular understanding of spermatogenesis.

The goal of this tubule squash technique is to rapidly assess cytological features of developing mouse spermatocytes while preserving cellular integrity. This method allows for the study of all stages of spermatogenesis, and can be easily implemented alongside other biochemical and molecular biological approaches for the study of mouse meiosis.

Meiotic progression in males is a process that requires the concerted action of a number of highly regulated cellular events. Errors occurring during ...

Enrichment of Pachytene Spermatocytes and Spermatids from Mouse Testes Using Standard Laboratory Equipment

To characterize each step of spermatogenesis, researchers must separate different subpopulations of germ cells from testes. However, isolating discrete populations is challenging, because the adult testis contains a complex mix of germ cells from all steps of spermatogenesis along with certain populations of somatic cells. Over the past few decades, different techniques such as centrifugal elutriation, fluorescence-activated cell sorting (FACS), and STA-PUT have been successfully applied to the isolation of germ cells. A drawback is that they all require dedicated devices and specialized training. Following principles underlying the STA-PUT method, a simple protocol has been developed for the isolation of pachytene spermatocytes, round spermatids, and elongating spermatids from mouse testes. After preparing a single cell suspension of testicular cells, specific cell populations are enriched by gravity sedimentation through a discontinuous bovine serum albumin (BSA) density gradient. The cell fractions are then manually collected and microscopically analysed. This modified density gradient for round spermatids (MDR) sedimentation protocol can be widely applied, because it requires only standard laboratory equipment. Furthermore, the protocol requires minimal starting materials, reducing its cost and use of laboratory animals.

Presented here is a protocol for enriching pachytene spermatocytes, round spermatids, and elongating spermatids from adult mouse testes using a discontinuous bovine serum albumin density gradient with standard laboratory equipment.

To characterize each step of spermatogenesis, researchers must separate different subpopulations of germ cells from testes. However, isolating discrete ...

Isolation of Murine Spermatogenic Cells using a Violet-Excited Cell-Permeable DNA Binding Dye

Isolation of meiotic spermatocytes is essential to investigate molecular mechanisms underlying meiosis and spermatogenesis. Although there are established cell isolation protocols using Hoechst 33342 staining in combination with fluorescence-activated cell sorting, it requires cell sorters equipped with an ultraviolet laser. Here we describe a cell isolation protocol using the DyeCycle Violet (DCV) stain, a low cytotoxicity DNA binding dye structurally similar to Hoechst 33342. DCV can be excited by both ultraviolet and violet lasers, which improves the flexibility of equipment choice, including a cell sorter not equipped with an ultraviolet laser. Using this protocol, one can isolate three live-cell subpopulations in meiotic prophase I, including leptotene/zygotene, pachytene, and diplotene spermatocytes, as well as post-meiotic round spermatids. We also describe a protocol to prepare single-cell suspension from mouse testes. Overall, the procedure requires a short time to complete (4-5 hours depending on the number of needed cells), which facilitates many downstream applications.

Here we present a simple and efficient method to isolate live meiotic and post-meiotic germ cells from adult mouse testes. Using a low-cytotoxicity, violet-excited DNA binding dye and fluorescence-activated cell sorting, one can isolate highly enriched spermatogenic cell populations for many downstream applications.

Isolation of meiotic spermatocytes is essential to investigate molecular mechanisms underlying meiosis and spermatogenesis. Although there are established ...

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Murine Round Spermatid Injection Procedure to Study Embryonic Development

Mouse Round Spermatid Injection

Round spermatids, characterized by their haploid genetic content, represent the precursor cells to mature spermatozoa. Through the innovative technique of round spermatid injection (ROSI), oocytes can be successfully fertilized and developed into viable fetuses. In a groundbreaking milestone achieved in 1995, the first mouse fetus was born through ROSI technology. ROSI has since emerged as a pivotal tool for unraveling the intricate mechanisms governing embryonic development and holds significant potential in various applications, including the acceleration of mouse generation and the production of genetically modified mice. In 1996, a milestone was reached when the first human fetus was born through ROSI technology. However, the clinical applications of this method have shown a fluctuating pattern of success and failure. To date, ROSI technology has not found widespread application in clinical practice, primarily due to its low birth efficiency and insufficient validation of fetal safety. This article provides a comprehensive account of the precise methods of performing ROSI in mice, aiming to shed new light on basic research and its potential clinical applications.

Here, we introduce a method for performing round spermatid injection (ROSI) in mice, a technique with promising clinical applications and utility for investigating the mechanisms underlying embryonic development.

Round spermatids, characterized by their haploid genetic content, represent the precursor cells to mature spermatozoa. Through the innovative technique of ...

Simple and Efficient Technique for the Preparation of Testicular Cell Suspensions

Mammalian testes are very complex organs that contain over 30 different cell types, including somatic testicular cells and different stages of germline cells. This heterogeneity is an important drawback concerning the study of the bases of mammalian spermatogenesis, as pure or enriched cell populations in certain stages of sperm development are needed for most molecular analyses1.
 Various strategies such as Staput2,3, centrifugal elutriation1, and flow cytometry (FC)4,5 have been employed to obtain enriched or purified testicular cell populations in order to enable differential gene expression studies. 
It is required that cells are in suspension for most enrichment/ purification approaches. Ideally, the cell suspension will be representative of the original tissue, have a high proportion of viable cells and few multinucleates - which tend to form because of the syncytial nature of the seminiferous epithelium6,7 - and lack cell clumps1 . Previous reports had evidenced that testicular cell suspensions prepared by an exclusively mechanical method clumped more easily than trypsinized ones1 . On the other hand, enzymatic treatments with RNAses and/or disaggregating enzymes like trypsin and collagenase lead to specific macromolecules degradation, which is undesirable for certain downstream applications. The ideal process should be as short as possible and involve minimal manipulation, so as to achieve a good preservation of macromolecules of interest such as mRNAs. Current protocols for the preparation of cell suspensions from solid tissues are usually time-consuming, highly operator-dependent, and may selectively damage certain cell types1,8 . 
	The protocol presented here combines the advantages of a highly reproducible and extremely brief mechanical disaggregation with the absence of enzymatic treatment, leading to good quality cell suspensions that can be used for flow cytometric analysis and sorting4, and ulterior gene expression studies9 .

A novel protocol for the mechanical preparation of testicular cell suspensions from rodent material, avoiding enzymes and detergents, is described. The method is very simple, fast, reproducible, and renders good quality cell suspensions, which are suitable for flow sorting and RNA extraction.

Mammalian testes are very complex organs that contain over 30 different cell types, including somatic testicular cells and different stages of germline ...

Sperm Collection of Differential Quality Using Density Gradient Centrifugation

In sexual reproduction, a male gamete or sperm cell fuses with a female gamete to bring about fertilization. However, a large number of sperm cells with fertilizing ability are required to interact with a female gamete to ensure fertilization. As such, the fertilizing ability of individual sperm cells is critical for successful reproduction. Density gradient centrifugation has been utilized for several decades as a reproducible, fast, efficient, effective and extremely adaptable method to collect only high-quality sperm to be used in assisted reproductive technology. The protocols we described herein focus on the utilization of the discontinuous Percoll density gradient centrifugation (PDGC) technique to isolate three distinct populations of rooster sperm by their quality. We were able to collect low-, medium- and high-quality sperm. We also describe reproducible protocols that entail determining fertility potential of sperm by assessing their viability, mobility and penetrability. Collection of sperm by their quality using PDGC technique would be useful to accurately and thoroughly characterize sperm with differential fertility potential.

In this paper, we aim to describe the performance of the density gradient centrifugation technique and its application in sperm physiology research.

In sexual reproduction, a male gamete or sperm cell fuses with a female gamete to bring about fertilization. However, a large number of sperm cells with ...

Isolation of Immune Regulatory Sertoli Cells from Mouse Seminiferous Tubules

To isolate the Sertoli cells, use a new 25-milliliter pipette to resuspend the settled tubules thoroughly in 25 milliliters of PBS. After allowing the tubules to settle for another 12 minutes, use the same pipette to wash the tubules three more times in the same way. Then, transfer the tubules into a new 100-milliliter screw-cap bottle and add the Hyaluronidase-DNase-I solution. Further, digest the tubules in a shaking water bath at 32 Celsius for five minutes.
Check an aliquot of the suspension by light microscope inspection at 100x magnification. Short tubules, tubular aggregates, and released cells should be visible. Allow the tubular aggregates to settle for 10 minutes. Then, use a new 25-milliliter pipette to remove the supernatant and wash the aggregates four times in 25ml of PBS, with 10 minutes of sedimentation between each wash.
The contaminating PTC population will have been reduced. After the last wash, add 20ml of RPMI 1640 medium to the cells, and pass the suspension through an 18G needle mounted on a 20-milliliter syringe 10 times. Next, quickly confirm that all of the aggregates have been dissociated, and filter the cell suspension through a 70-micron cell strainer to obtain a pure single-cell suspension.
Spin down the filtrate, using the 25-milliliter pipette to carefully aspirate the supernatant. Then, resuspend the isolated Sertoli cells in 40ml of serum-free RPMI 1640 medium. Then, count the number of viable cells by trypan-blue exclusion and adjust the cell concentration to 3 x 106 cells per milliliter. Finally, feed 1 milliliter of cells per well in a six-well plate.