This work was supported by NIGMS (R01GM11755) to P.W.J. and by a training grant fellowship from the National Cancer Institute (NIH) (CA009110) to S.R.W. and J.H. 
&#160;

The use of mice was approved by the Institutional Animal Care and Use Committee of Johns Hopkins University. Experiments were performed on juvenile (20 - 26 days post-partum, dpp) C57BL/6J mice, taking advantage of the semi-synchronous first wave of spermatogenesis. However, this technique can also be performed using adult mice.
1. Dissection and isolation of mouse seminiferous tubules
<ol>
	<li>Prepare the fix/lysis solution and antibody dilution buffer (ADB) described in Table 1 and Table 2.</li>
	<li>Prepare one 35-mm Petri dish with 3 mL of 1x phosphate buffered saline (1x PBS, pH 7.4), and another 35-mm dish with 2 mL of fix/lysis solution per mouse.</li>
	<li>Sacrifice the mouse via cervical dislocation or CO2 asphyxiation and spray the ventral abdomen with 70% ethanol. Open the abdominopelvic cavity using sterile scissors, making a V-shaped opening. Then remove the testes by pulling on the epididymal fat pad using forceps, and avoid disturbing the tunica albuginea. Place the testes in a 35-mm Petri dish containing 1x PBS. 
	NOTE: Utilize the first wave of spermatogenesis in order to observe an enriched cell population of interest14. Specific stages of spermatogenesis are enriched at different mouse ages (Table 3).</li>
	<li>Remove the testicular tunica albuginea by puncturing the tissue with sharp tipped forceps and collect the loose seminiferous tubules. Transfer the tubules to a new 35-mm Petri dish containing 2 mL of fix/lysis solution.</li>
	<li>Incubate the tubules in the fix/lysis solution for 5 min at room temperature.</li>
</ol>
2. Preparation of seminiferous tubules
<ol>
	<li>Prepare a poly-L-lysine coated glass slide by outlining the edges of the slide with a liquid blocker pen.</li>
	<li>Apply 100 &#181;L of fix/lysis solution to the center of the glass slide.</li>
	<li>Using sterile forceps and scissors gently tease apart the seminiferous tubules. Cutting out long individual tubule segments approximately 20 mm in length. 
	NOTE: To visualize structures that are sensitive to prolonged fixative exposure, such as the centrosomes, Separate the tubules first in the 1x PBS, then directly mince the tubules on the surface of the poly-lysine slides coated in 100 &#181;L of fix-lysis solution.</li>
	<li>Transfer five 20 mm long seminiferous tubule segments to the prepared poly-L-lysine coated glass slide containing fix/lysis solution.</li>
	<li>Using sterile scissors mince seminiferous tubules into 1.5 to 3.0 mm segments.</li>
	<li>Using sterile forceps arrange the tubule segments on the glass slide so that no tissue overlaps, and the tubules are distributed evenly. 
	NOTE: The optimal number of tubule segments on a glass slide is between 20 and 40.</li>
</ol>
3. Squashing of the seminiferous tubules
<ol>
	<li>Transfer the glass slide containing dispersed seminiferous tubule segments onto a benchtop. Remove excess liquid using a laboratory wipe to avoid losing tissue during the squash step.</li>
	<li>Apply a coverslip (22 x 60 - 1.5 mm) on top of the glass slide and apply pressure with the heel of the palm for 10 - 20 seconds. It is important to apply enough force to disperse spermatocytes from the seminiferous tubules. 
	NOTE: Observe the tubules using a dissection microscope in order to optimize the squashing step so that all tubule segments are disrupted.</li>
	<li>Using slide forceps, immediately flash freeze the glass slide in a small dewar of liquid N2 for 15 seconds, or until the liquid ceases to bubble. If immunolabeling immediately, remove the coverslip with a straight edge razor blade, fine tip forceps or 21-gauge needle. 
	NOTE: For optimal results, immunolabel the slides immediately (see step 4). However, at this point slides can be preserved at -80 &#176;C for up to two weeks. To maximize the lifetime of proteins and cellular structures, immediately store the slides on dry ice or transfer to a -80 &#176;C freezer, and keep the coverslip on the slide. When immunolabeling stored slides, remove the coverslip by immersing the slides in liquid N2 for 15 seconds, as described in step 3.3.</li>
</ol>
4. Immunolabeling of mounted seminiferous tubules
<ol>
	<li>Immerse the slide in 1x PBS, and wash three times for 5 min in a 50-mL Coplin jar containing 1x PBS. 
	NOTE: Never allow the slides to completely dry during immunolabeling.</li>
	<li>Apply 1 mL of antibody dilution buffer onto the glass slide to block for 1 - 2 h in a humidified chamber.</li>
	<li>Remove the antibody dilution buffer and apply 100 &#181;L of primary antibody diluted in antibody dilution buffer in a humidified chamber. 
	NOTE: For best results, incubate primary antibodies overnight at 4 &#176;C. Use smaller volumes of antibody (e.g. 50 &#181;L) by covering the slide with a coverslip or parafilm. Validate and optimize primary antibodies15.</li>
	<li>Rinse slides three times for 5 min in a 50 mL Coplin jar containing 1x PBS. 
	NOTE: To enhance washes, agitate Coplin jars by using a small magnetic stir bar and place on a magnetic stir plate at low speed, or place the Coplin jar on a tabletop mixer at low speed.</li>
	<li>Apply 100 &#181;L of secondary antibody diluted in antibody dilution buffer for 1 to 1.5 h in a humidified chamber at room temperature. Incubate the slides in a dark box to avoid photo bleaching of fluorophore conjugated antibodies. 
	NOTE: Use smaller volumes of antibody (e.g. 50 &#181;L) by covering the slide with a coverslip or parafilm.</li>
	<li>Rinse slides two times for 5 min in a 50-mL Coplin jar containing 1x PBS.</li>
	<li>Mount the slides in mounting medium containing 4&#39;,6-diamidino-2-phenylindole (DAPI, 1.5 &#181;g/mL) and apply coverslips (22 x 60 -&#160;1.5 mm).</li>
	<li>Seal the coverslips to the slides with clear nail polish to preserve the slides and prevent the coverslips from moving.</li>
</ol>
5. Analysis and imaging tubule squash preparations
<ol>
	<li>Use an epifluorescence microscope with automated stage that enables precise Z-axis movement and a high-resolution camera for image capturing. Alternatively, use a laser confocal microscope. 
	NOTE: Use any available hardware and software for this step. For example, images displayed in Figure 1 and Figure 2 were captured using a Zeiss Cell Observer Z1 linked to an ORCA-Flash 4.0 CMOS camera and analyzed with the Zeiss ZEN 2012 blue edition image software.</li>
	<li>Assess the slides using a 20X&#160;objective to determine the quality of the squash preparation. Good quality slides have a monolayer of nuclei that are evenly distributed. 
	NOTE: Some sections of the slide may be better than others, it is useful to note the coordinates of where the best regions of the slide can be found for further analysis using higher magnification.</li>
	<li>Using higher magnification objective (e.g. 63X&#160;or 100X), capture regions of the slide that have a consistent monolayer of nuclei. Once a region is selected, set the upper and lower Z-stacks to ensure that all in-focus light is captured. 
	NOTE: The optimal number of Z-stacks captured between the upper and lower limit is generally designated by the image acquisition software and is different for each objective.</li>
	<li>Use the image processing software to compile an extended depth focus image. The software combines the in-focus light from each Z-stack, and is essential for optimal imaging of tubule squash preparations.</li>
</ol>

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The authors have nothing to disclose.

Mice have proven to be a useful model organism for studying the cellular events that govern meiotic progression during spermatogenesis. Further, it is necessary to develop tools catered to the study of spermatogenesis because many events, such as exit from meiotic prophase I, are sexually dimorphic. This protocol describes a seminiferous tubule squash method for visualization and study of different stages of mouse spermatogenesis. This method preserves cellular integrity and thereby allows detailed analyses of nuclear an...

Meiosis is a complex cellular event in which a single round of DNA replication is followed by two successive rounds of cell division. Several meiosis-specific events must be coordinated during the initial stages of meiosis to ensure accurate chromosome segregation. These events include the completion of homologous recombination, co-orientation of sister kinetochores during the first meiotic division, and the stepwise loss of cohesin complexes to resolve chiasmata between homologs. Precise regulation of these processes is necessary to maintain fertility and to prevent chromosome missegregation events that can lead to genetic developmental disorders and spontaneous miscarriage1.
While the key events of meiosis take place in both males and females, significant temporal and mechanistic differences exist between spermatogenesis and oogenesis2. For example, during female meiosis, prophase I occurs during embryonic development and arrests at the dictyate stage until puberty. In contrast, spermatogenesis commences at puberty and progresses in waves throughout adult life without arrest. The differences between male and female meiosis emphasizes the need to develop methods that are specifically catered towards assessing these processes in both spermatocytes and oocytes. Currently, assessing meiotic progression largely relies on the use of chromatin spreads3,4,5. While chromatin spreads are useful for studying meiotic chromosomes, they fail to preserve cellular integrity, preventing evaluation of cellular structures such as spindle microtubules, centrosomes, the nuclear envelope, and telomere attachments. Live imaging and long-term culturing techniques have greatly advanced our understanding of female meiosis; similar approaches to visualize the entire intact cell, however, are less frequently implemented for the study of spermatogenesis6,7. In order to visualize dynamic events throughout male meiosis, we have adapted established tubule squash techniques to rapidly assess the cytological features of developing mouse spermatocytes8,9. The method described here maintains the integrity of the cell, enabling the study of multiple cellular structures during different stages of spermatogenesis.
This tubule squash technique is a whole cell approach, which allows for the assessment of cellular structures via immunofluorescence microscopy. Common histological approaches to visualize meiotic progression in male mice such as haematoxylin and eosin staining of paraffin embedded testes, and immunofluorescent labeling of cryosections allow for a broad overview of meiotic progression. However, these techniques fail to resolve single cells to the extent necessary for detailed analysis of the events occurring throughout meiosis10,11. Alternative techniques to visualize meiotic processes rely on significant chemiosmotic disruption to the spermatocyte to isolate and fix nuclear materials3,4,5. These chemical treatments hinder the observation of cell types other than primary spermatocytes. A recently described method by Namekawa has enabled the research community to preserve the nuclear architecture of isolated spermatocytes, but requires the use of a cytospin and accessories that may not be readily available to some laboratories4. In contrast, the tubule squash technique only requires equipment that is generally standard in most cell biology laboratories.
The tubule squash method described here can be used to visualize the diverse cell types found within the seminiferous tubule, including sertoli cells, spermatogonia, primary and secondary spermatocytes, and spermatids. By coupling this technique with the near-synchronous first wave of spermatogenesis in juvenile mice, it is possible to obtain enriched populations of spermatogenic cells as they progress through meiosis12. This process permits the detailed analysis of processes throughout spermatogenesis, such as early prophase events, the G2/MI and metaphase to anaphase transitions, and spermiogenesis. Furthermore, tubule squash preparations can be used to visualize cytological features of the chromosomes (e.g. interchromatid domains (ICDs) and kinetochores) and centrosomes (centrioles and pericentriolar material/matrices). The squash method can be readily performed in parallel with other experimental approaches, such as chromatin spreads and protein extraction. In addition, this technique has been successfully modified to deposit living spermatogenic cells on slides for direct visualization13.
The method described here involves a whole cell seminiferous tubule squash technique to analyze the G2/MI transition in wild-type C57BL/6J mice. The cytological features of primary spermatocytes entering the first meiotic division were visualized with immunofluorescence microscopy to observe the meiotic spindle. This versatile technique can be easily modified to visualize other meiotic stages and different cell types. The technique is also amenable to alternative visualization strategies, such as DNA and RNA FISH approaches.

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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.

Here, we have used the tubule squash method to visualize transitory cell populations undergoing the prophase to metaphase I (G2/MI) transition, which were enriched by harvesting testes from juvenile wild-type mice undergoing the first wave of spermatogenesis (24 dpp). Figure 1 depicts representative images of the various cell stages that can be visualized using the tubule squash method. Enriched populations of metaphase I spermatocytes were visualized using a...

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

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 ...

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Developmental Biology

14.8K Views.  Johns Hopkins Bloomberg School of Public Health. 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.

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

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 ...

Collection of Post-mating Semen from the Female Reproductive Tract and Measurement of Semen Liquefaction in Mice

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called liquefaction. In this study, we show how to collect the post-ejaculated semen from the female reproductive tract in a mouse model. First, adult female mice in the estrus stage were housed in a male&#39;s cage overnight. The next morning, copulation was confirmed by the presence of copulatory plug at the vaginal opening. Female mice with copulatory plugs were euthanized, and each reproductive tract was collected as a whole (vagina, uterus, oviducts, ovaries), ensuring a closed system to contain the semen. The reproductive tract was placed in a 1.5 mL microcentrifuge tube, and the vagina was cut off to release the semen into the tube. To ensure maximum semen volume for analysis, toothless forceps were used to squeeze the uterine horns from ovarian end to vaginal end expelling remaining semen. The whole reproductive tract was then discarded. The semen-containing tube was briefly spun down. A 25 &#956;L capillary pipette was placed into the tube at a 180&#176; angle (parallel to the tube wall). The amount of time used to fill the capillary tube to the 25 &#956;L line was recorded. Semen from a proven male breeder usually takes approximately 60-180 s to fill a 25 &#956;L capillary tube. This semen collection technique can also be used in other downstream applications such as sperm imaging and motility analysis.

Semen liquefaction is required for sperm to be liberated from the seminal gel. This study provides the procedures for collecting semen from the female reproductive tract post-coitus, and measuring semen liquefaction time.

In mice, ejaculated semen is deposited in the uterus. After ejaculation, the semen changes consistency from gel-like to watery, a process called ...

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 ...

Transillumination-Assisted Dissection of Specific Stages of the Mouse Seminiferous Epithelial Cycle for Downstream Immunostaining Analyses

Spermatogenesis is a unique differentiation process that ultimately gives rise to one of the most distinct cell types of the body, the sperm. Differentiation of germ cells takes place in the cytoplasmic pockets of somatic Sertoli cells that host 4 to 5 generations of germ cells simultaneously and coordinate and synchronize their development. Therefore, the composition of germ cell types within a cross-section is constant, and these cell associations are also known as stages (I&#8211;XII) of the seminiferous epithelial cycle. Importantly, stages can also be identified from intact seminiferous tubules based on their differential light absorption/scatter characteristics revealed by transillumination, and the fact that the stages follow each other along the tubule in a numerical order. This article describes a transillumination-assisted microdissection method for the isolation of seminiferous tubule segments representing specific stages of mouse seminiferous epithelial cycle. The light absorption pattern of seminiferous tubules is first inspected under a dissection microscope, and then tubule segments representing specific stages are cut and used for downstream applications. Here we describe immunostaining protocols for stage-specific squash preparations and for intact tubule segments. This method allows a researcher to focus on biological events taking place at specific phases of spermatogenesis, thus providing a unique tool for developmental, toxicological, and cytological studies of spermatogenesis and underlying molecular mechanisms.

This protocol describes transillumination-assisted microdissection of segments of adult mouse seminiferous tubules representing specific stages of seminiferous epithelial cycle, and cell types therein, and subsequent immunostaining of squash preparations and intact tubule segments.

Spermatogenesis is a unique differentiation process that ultimately gives rise to one of the most distinct cell types of the body, the sperm. ...

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 ...

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies showed that microinjection of germ cells from fertile donor mice into the seminiferous tubules of infertile recipient mice results in donor-derived spermatogenesis and sperm production by the recipient animal2. The use of donor males carrying the bacterial &beta;-galactosidase gene allowed identification of donor-derived spermatogenesis and transmission of the donor haplotype to the offspring by recipient animals1. Surprisingly, after transplantation into the lumen of the seminiferous tubules, transplanted germ cells were able to move from the luminal compartment to the basement membrane where spermatogonia are located3. It is generally accepted that only SSCs are able to colonize the niche and re-establish spermatogenesis in the recipient testis. Therefore, germ cell transplantation provides a functional approach to study the stem cell niche in the testis and to characterize putative spermatogonial stem cells. To date, germ cell transplantation is used to elucidate basic stem cell biology, to produce transgenic animals through genetic manipulation of germ cells prior to transplantation4,5, to study Sertoli cell-germ cell interaction6,7, SSC homing and colonization3,8, as well as SSC self-renewal and differentiation9,10.
Germ cell transplantation is also feasible in large species11. In these, the main applications are preservation of fertility, dissemination of elite genetics in animal populations, and generation of transgenic animals as the study of spermatogenesis and SSC biology with this technique is logistically more difficult and expensive than in rodents. Transplantation of germ cells from large species into the seminiferous tubules of mice results in colonization of donor cells and spermatogonial expansion, but not in their full differentiation presumably due to incompatibility of the recipient somatic cell compartment with the germ cells from phylogenetically distant species12. An alternative approach is transplantation of germ cells from large species together with their surrounding somatic compartment. We first reported in 2002, that small fragments of testis tissue from immature males transplanted under the dorsal skin of immunodeficient mice are able to survive and undergo full development with the production of fertilization competent sperm13. Since then testis tissue xenografting has been shown to be successful in many species and emerged as a valuable alternative to study testis development and spermatogenesis of large animals in mice14.

Protocols for germ cell transplantation and testis tissue xenografting are described. Germ cell transplantation results in donor-derived spermatogenesis in recipient testes and represents a functional reconstitution assay for identification of spermatogonial stem cells (SSCs). Testis tissue xenografting reproduces testis development and spermatogenesis of various donor species in recipient mice.

Germ cell transplantation was developed by Dr. Ralph Brinster and colleagues at the University of Pennsylvania in 19941,2. These ground-breaking studies ...

Biology

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 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 ...

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

Preparation of Meiotic Chromosome Spreads from Mouse Spermatocytes

Mammalian meiosis is a dynamic developmental process that occurs in germ cells and can be studied and characterized. Using a method to spread nuclei on the surface of slides (rather than dropping them from a height), we demonstrate an optimized technique on mouse spermatocytes that was first described in 1997. This method is widely used in laboratories to study mammalian meiosis because it yields a plethora of high quality nuclei undergoing substages of prophase I. Seminiferous tubules are first placed in a hypotonic solution to swell spermatocytes. Then spermatocytes are released into a sucrose solution to create a cell suspension, and nuclei are spread onto fixative-soaked glass slides. Following immunostaining, a diversity of proteins germane to meiotic processes can be examined. For example, proteins of the synaptonemal complex, a tripartite structure that connects the chromosome axes/cores of homologs together can be easily visualized. Meiotic recombination proteins, which are involved in repair of DNA double-strand breaks by homologous recombination, can also be immunostained to evaluate progression of prophase I. Here we describe and demonstrate in detail a technique widely used to study mammalian meiosis in spermatocytes from juvenile or adult male mice.

Meiosis is the developmental process by which gametes are formed through a single round of DNA replication and two successive rounds of chromosome segregation. Mammalian meiosis can be examined by utilizing a technique to prepare meiotic chromosome spreads. Here, we demonstrate a method of preparing surface-spread nuclei from mouse spermatocytes.

Mammalian meiosis is a dynamic developmental process that occurs in germ cells and can be studied and characterized. Using a method to spread nuclei on ...

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry

In eukaryotes, meiosis is essential for genome stability and genetic diversity in sexual reproduction. Experimental analyses of spermatocytes in testes are critical for the investigations of spindle assembly and chromosome segregation in male meiotic division. The mouse spermatocyte is an ideal model for mechanistic studies of meiosis, however, the effective methods for the analyses of spermatocytes are lacking. In this article, a practical and efficient method for the in vivo inhibition of kinesin-7 CENP-E in mouse spermatocytes is reported. A detailed procedure for testicular injection of a specific inhibitor GSK923295 through abdominal surgery in 3-week-old mice is presented. Furthermore, described here is a series of protocols for tissue collection and fixation, hematoxylin-eosin staining, immunofluorescence, flow cytometry and transmission electron microscopy. Here we present an in vivo inhibition model via abdominal surgery and testicular injection, that could be a powerful technique to study male meiosis. We also demonstrate that CENP-E inhibition results in chromosome misalignment and metaphase arrest in primary spermatocytes during meiosis I. Our in vivo inhibition method will facilitate mechanistic studies of meiosis, serve as a useful method for genetic modifications of male germ lines, and shed a light on future clinical applications.

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry

This article reports an in vivo inhibition of CENP-E through abdominal surgery and testicular injection of GSK923295, a valuable model for male meiotic division. Using the immunofluorescence, flow cytometry and transmission electron microscopy assays, we show that CENP-E inhibition results in chromosome misalignment and genome instability in mouse spermatocytes.

In eukaryotes, meiosis is essential for genome stability and genetic diversity in sexual reproduction. Experimental analyses of spermatocytes in testes ...

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

Summary

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Chapters in this video

Step-specific Sorting of Mouse Spermatids by Flow Cytometry

Collection of Post-mating Semen from the Female Reproductive Tract and Measurement of Semen Liquefaction in Mice

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

Transillumination-Assisted Dissection of Specific Stages of the Mouse Seminiferous Epithelial Cycle for Downstream Immunostaining Analyses

Mouse Round Spermatid Injection

Germ Cell Transplantation and Testis Tissue Xenografting in Mice

Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Preparation of Meiotic Chromosome Spreads from Mouse Spermatocytes

Functional Assessment of Kinesin-7 CENP-E in Spermatocytes Using In Vivo Inhibition, Immunofluorescence and Flow Cytometry