This work was supported by grants from Academy of Finland [315948, 314387 to N.K.]; Sigrid Jus&#233;lius Foundation [to N.K., J.T.]; Emil Aaltonen Foundation [to J.-A.M., T.L.]; Turku Doctoral Programme of Molecular Medicine [S.C.-M., O.O.].

Maintenance of laboratory mice and all animal experiments were done in accordance with the relevant guidelines and regulations for the care and use of laboratory animals in University of Turku.
1. Preparation of seminiferous tubules for microdissection
<ol>
	<li>Sacrifice an adult male mouse (&#8805;8 weeks old, testis 80&#8211;120 mg depending on the strain and age) via CO2 asphyxiation followed by cervical dislocation. 
	NOTE: The mouse should be sexually mature, and preferably at least 8 weeks old. Transillumination pattern of juvenile mice differs from adult because the wave of the seminiferous epithelium has not yet been fully established, and the timing of the first wave of spermatogenesis is distinct21,22. Lack of elongating spermatids in &#60;4-week-old male mice precludes their use for transillumination-assisted microdissection. All mouse strains that have normal spermatogenesis can be used.</li>
	<li>Spray the ventral abdomen with 70% ethanol. Open the abdominopelvic cavity using sterile scissors, making a V-shaped opening.</li>
	<li>Pulling on the epididymal fat pad with sterile forceps, locate the testes, dissect them using scissors and place them on a sterile 100-mm Petri dish containing PBS. 
	NOTE: To maintain sterility, make sure all labware and surgical tools are sterile.</li>
	<li>Using fine-tipped scissors decapsulate the testes by cutting a slit in the tunica albuginea, the thick fibrous sheet encapsulating the testis. Then tear the tunica open using a pair of forceps. Force the tubules out by pressing with forceps and discard the tunica. 
	NOTE: While discarding the tunica, it might be beneficial for some downstream applications that arteria testicularis is removed together with the tunica. Avoid damaging the seminiferous tubules.</li>
	<li>Move the seminiferous tubules to a new Petri dish and pour enough sterile PBS to cover the bottom of the Petri dish. Next, gently pull the tubules apart but avoid damaging the tubules. 
	NOTE: Too much mechanical stress will impinge on the transillumination pattern and affect the viability of the tissue and its cellular architecture. The tubules can also be processed for whole-mount immunostainings from this point without staging (3B). Sometimes it is sufficient to define the stage retrospectively by including antibodies against proteins expressed in differentiating spermatogonia, such as SALL4, c-KIT and DNMT3A18,23. The density of spermatogonia is a relatively reliable stage indicator (Figure 2).</li>
</ol>
2. Transillumination-assisted microdissection
<ol>
	<li>Place the Petri dish under a dissection microscope firmly by taping it to the stage. 
	NOTE: It is important to tape the Petri dish well to prevent its movement which could cause mixing of the collected staged seminiferous tubule segments.</li>
	<li>To reveal the light absorption pattern of seminiferous tubules under focus, make sure that the sample is illuminated from below and the light passes through the sample, i.e., it is transilluminated. 
	NOTE: The amount of light absorbed/scattered is relative to the level of chromatin condensation in elongating spermatids and their bundling inside the seminiferous tubule: the more condensed, the more light is absorbed, i.e., appears darker.</li>
	<li>Get acquainted with the light absorption pattern of different stages as described in Figure 2, Figure 5A and Figure S1 by carefully moving bundles of tubules using fine forceps. 
	NOTE: The stages always follow each other in a logical order, forming the wave of seminiferous epithelium. However, it is important to know that the direction of the spermatogenic wave occasionally reverses and then reverts back again (also known as modulations4,9), sometimes complicating the procedure. Also, the length of each stage, in terms of how many mm of tubule, varies considerably.</li>
	<li>Carefully lift the tubule of interest using forceps with a hooked tip, and then cut a segment of appropriate length using microdissection scissors (see Supplementary Video 1). A hook at the tip of the forceps makes lifting and holding a tubule easier and helps to avoid squeezing it. 
	NOTE: The length of the segments to be cut depends on downstream applications. For the collection of pooled tubule pieces of a specific stage for protein or RNA analysis12,13 (II&#8211;V, VII&#8211;VIII and IX&#8211;XI, Figure 5B) the length is typically 2&#8211;5 mm. When standard phenol-chloroform extraction is used, around 200 ng of RNA can be derived from 1 mm of tubule. For whole-mount staining of staged tubule segments, the length of the segments should be &#62;5 mm. For squash preparations the length of segments should not exceed 1&#8211;2 mm because the cells in the middle of the segment may fail to exit if too long. Use a mm scale under the Petri dish for an accurate measurement of the tubule length.</li>
</ol>
3. Immunostaining of different preparations
<ol>
	<li>Squash preparation: stage verification and immunostaining 
	NOTE: Stage-specific tubule pieces can be squashed on a microscope slide with a cover glass to perform morphological analysis of living cells by phase-contrast microscopy and subsequent immunostaining. A beginner is recommended to use this approach to verify the stages when getting acquainted with the transillumination-assisted microdissection method.
	<ol>
		<li>Collect the segment in a volume of 10 &#956;L using a pipette and move it onto a microscope slide.</li>
		<li>Squash the tubule by placing a cover glass (20 mm x 20 mm) carefully on the tubule. As a result, cells will flow out the tubule and form a live-cell monolayer. Place a filter paper on the edge of the cover glass to facilitate the spreading of cells. Avoid squashing the cells too much to keep them alive.</li>
		<li>Monitor cell spreading under a microscope. Use a phase-contrast microscope at 40x objective to verify stage recognition by examining the cell types present (Figure 2, Figure S2).</li>
		<li>Once the cells have spread to form a round monolayer from both ends of the tubule, dip the slide into a container containing liquid nitrogen while holding it with forceps. Keep it submerged for 10 s. Alternatively place the slide on a dry ice plate for freezing.</li>
		<li>Remove the cover glass by flipping it off using a scalpel.</li>
		<li>Without a delay, proceed with the fixation and quickly place the slide in a container with 90% ethanol for 2&#8211;5 min. 
		NOTE: Make sure the squash preparation does not thaw before placing it to 90% ethanol. Other fixatives can also be used, such as acetone, for 10 min.</li>
		<li>Air-dry and store at room temperature (RT) (up to some days) or at -80 &#176;C (long-term).</li>
		<li>For immunostaining, post-fix the samples in 4% paraformaldehyde (PFA) for 10 min at RT.</li>
		<li>Rinse in PBS and permeabilize with 0.1% Triton X-100 in PBS for 5 min.</li>
		<li>Rinse in PBS and draw a grease ring around each squash sample.</li>
		<li>Add 50&#8211;100 &#956;L of 10% BSA (bovine serum albumin) in 0.1% Tween in PBS (PBST) inside the grease ring and block samples for 30 min at RT.</li>
		<li>Remove the BSA solution and incubate with a primary antibody diluted in 10% BSA in PBST for 1 h at RT.</li>
		<li>Wash 3x for 5 min with PBST.</li>
		<li>Incubate with a secondary antibody diluted in 10% BSA in PBST. 
		NOTE: To stain the acrosomes, the samples can be incubated with Rhodamine-labeled Peanut agglutinin antibody (PNA, 1:1000) in 10% BSA in PBST for 1 h at RT (Figure S3) instead of specific primary and secondary antibodies.</li>
		<li>Wash 3x for 5 min each with PBST, rinse with PBS and mount with a mountant containing DAPI.</li>
	</ol>
	</li>
	<li>Whole-mount immunostaining of seminiferous tubules 
	NOTE: The protocol below describes whole-mount staining for staged (from step 2.4) tubule segments. If a researcher wants to perform whole-mount staining without staging (from step 1.5), pay attention to notes in 3.2.1 and 3.2.7.
	<ol>
		<li>Using a pipette, transfer the tubule segments (from step 2.4) in ice-cold PBS into a 15 mL conical tube and allow them to sediment on ice. 
		NOTE: If using unstaged tubules from 1.5., separate the tubules in a Petri dish by pipetting up and back onto a tilted dish several times. Use a 1-mL pipette with a cut tip. This step is intended to open up the structure of the tissue. However, avoid too much pipetting as it might damage the tubules. Sedimentation of tubules will take some tens of seconds. Small tubule fragments, interstitial cells and cell debris stay in the supernatant.</li>
		<li>Carefully remove the supernatant (SN) by pipetting or with an aspirator. Add 10 mL of ice-cold PBS and mix by inversion.</li>
		<li>Allow to sediment and then remove SN as before.</li>
		<li>Add 5 mL of 4% PFA and fix for 5 h on a rotating table (20&#8211;30 rpm) at +4 &#176;C. 
		NOTE: Fixation time depends on the proteins of interest and their subcellular localization. For nuclear and cytoplasmic proteins, a 2 h fixation is typically sufficient, however, membrane markers, such as GFR&#945;1 (GDNF family receptor alpha 1; Figure 6A,B), benefit from a longer fixation, up to 6 h.</li>
		<li>Allow to sediment, remove SN (PFA) as before, and rinse briefly by adding 10 mL of PBS and inverting the tube.</li>
		<li>Allow to sediment, remove SN as before, and repeat the PBS washing step three times for at least 10 min each on a rotating table at +4 &#176;C and proceed with staining or store at +4 &#176;C. 
		NOTE: If the working conditions are sterile and clean the samples can be stored and used for at least some weeks. Alternatively, add Sodium Azide to a final concentration of 0.02% (w/v) from a 2% stock solution to help preserve the tubules prior to storing at +4 &#176;C.</li>
		<li>Using a 1-mL pipette, move 10&#8211;20 fixed tubule segments to a 2-mL round-bottom tube. Allow to sediment and remove SN. 
		NOTE: If working with long tubules that have not been staged, pour the tubules onto a Petri dish and using microdissection scissors and forceps cut segments of around 5&#8211;20 mm. Too long segments will tangle during the staining procedure, whereas too short segments are easily lost.</li>
		<li>Add 1 mL of 2% BSA + 10% FBS (fetal bovine serum) in 0.3% Triton X-100 in PBS (PBSX). Block for at least 1 h on a rotating table (20&#8211;30 rpm) at RT.</li>
		<li>Rinse with 1 mL of PBSX, remove SN by pipetting and add 250 &#956;L of primary antibody diluted in 1% BSA in PBSX (1:100&#8211;1:2000 dilution). Incubate for 2 h at RT or overnight at +4 &#176;C on a rotating table (20&#8211;30 rpm).</li>
		<li>Remove the antibody solution by pipetting and rinse the tubules with 1 mL of PBSX as above. Wash three times for 1 h in PBSX on a rotating table (20&#8211;30 rpm) at RT. 
		NOTE: After this first wash the sample can be left overnight at +4 &#176;C if necessary.</li>
		<li>Remove SN and add 250 &#956;L of secondary antibody diluted in 1% BSA in PBSX (typically 1:500 dilution of a fluorescent-labeled antibody). Cover in foil and incubate on a rotating table (20&#8211;30 rpm) at RT for 1 h.</li>
		<li>Repeat 3.2.10.</li>
		<li>Finally remove SN and pour the tubules to a microscope slide. Gently separate and arrange tubules in linear strips using gel-loading tips. Drain off excess buffer and add mounting medium and a coverslip. 
		NOTE: Avoid drying of the tubules while arranging them. Counter-staining of the nuclei with DAPI is not necessary in most cases. Seal off the edge of the coverslip with nail polish to prevent sample drying and deterioration. Slides can be stored for 1&#8211;2 weeks at +4 &#176;C before imaging.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have nothing to disclose.

The transillumination-assisted microdissection method that we have described above enables a stage-oriented approach for the study of spermatogenesis. Spermatogenesis is a highly synchronized process, and all key steps during the spermatogenic differentiation are regulated and executed in a stage-dependent manner, such as differentiation commitment (at stages VII-VIII), onset of meiosis (VII-VIII), meiotic divisions (XII), onset of spermatid elongation (VIII) and spermiation (VIII)1,26,27. The stage-oriented analysis provides a powerful tool to study these particular events that are restricted to specific steps of spermatogenesis and therefore found only at defined stages of the seminiferous epithelial cycle. Mastering the method takes some practice and use of a good quality dissection microscope and proper illuminating conditions are key to success. Implementing this method as a part of the everyday toolkit has a capacity to greatly improve the impact and biological relevance of research on male reproductive functions by allowing more accurate dissection of molecular events during spermatogenesis.
All WT mouse strains that we have studied display a similar transillumination pattern and exhibit conserved cell associations at stages of the seminiferous epithelial cycle. On the condition that spermiogenic differentiation of germ cells is not grossly different from WT mice, the same also applies to all the knock-out mouse models that we have studied. Moreover, it can be applied to other species that exhibit longitudinal segmental arrangement of stages of the seminiferous epithelial cycle7. However, species with non-segmental stages (such as human) cannot be used. Given the essential role of chromatin condensation in elongating spermatids in defining the transillumination pattern, it is clear that any misregulation of this process will inevitably impinge on the implementation of this method. In juvenile mice and young adults (5-6 weeks) the transillumination pattern has not yet been fully established, and, therefore, only mice older than 8 weeks should be used. It is also important to bear in mind that squeezing and pulling the tubules will inevitably impinge on the transillumination pattern because it distorts the cellular architecture within the seminiferous epithelium.
The isolated seminiferous tubule segments can also be cultured enabling ex vivo observation and manipulation of spermatogenesis-coupled processes, including meiosis. To ensure viability of the tissue and to prevent RNA and protein degradation, the samples should be collected and processed no more than 2 hours after sacrificing the mouse. For ex vivo culture of seminiferous tubules, the time from sacrifice to onset of culture should not exceed 1 hour. The integrity of tubule fragments can typically be maintained up to 72 hours in vitro if harvested properly.
The stage of the seminiferous epithelial cycle can be verified and even more accurately defined using phase-contrast microcopy of squash preparations16. Microscopy is performed on living cells, which provides an additional dimension into the analysis and allows observation of organelle or cell movements in specific stages of spermatogenesis28,29,30. Phase-contrast microscopy provides exact staging for subsequent immunostaining, which enables very detailed analysis of protein expression and localization dynamics during spermatogenesis, including stage-specific changes.
While cells are released from the epithelial context in squash preparations, whole-mount immunostainings of tubule segments enable the study of spermatogenic cells in their physiological environment. Therefore, whole-mount preparations may provide a better visualization of seminiferous tubule architecture and its intercellular contacts than immunostaining on cross-sections. Importantly, transillumination-assisted staging of the tubule segments prior to immunostaining makes the approach even more powerful by including information on the specific stage of a given segment. Whole-mount staining is a particularly useful tool for the study of cells at the periphery of seminiferous tubules, such as peritubular myoid cells, peritubular macrophages and spermatogonia, but might also open novel insights into research on meiotic and postmeiotic germ cells.

Differentiation of male germ cells from diploid spermatogonia to mature haploid spermatozoa, i.e., spermatogenesis, is a complex process that takes place in the epithelium of seminiferous tubules in the testes of a sexually-mature individual1. Mitotic descendants of A1 spermatogonia first divide five times to expand the differentiation-committed population, then enter meiosis as spermatocytes that ultimately give rise to haploid spermatids. Differentiation of round spermatids into spermatozoa, i.e., spermiogenesis, involves complex changes in cellular morphology, including nuclear compaction and construction of sperm-specific structures such as the acrosome and the flagellum. In mouse, the whole process of spermatogenesis takes 35 days to complete2,3.
At any given locale, the seminiferous epithelium hosts up to five cohorts of differentiating germ cells plus the germline stem/progenitor cells and the somatic Sertoli cells1. Differentiating germ cells form concentric layers the composition of which is predictable, and haploid cells at a given step of development always associate with certain types of spermatocytes and spermatogonia4,5. Therefore, any cross-section of a tubule hosts cohorts of germ cells of a constant composition. These specific cell associations are defined as the stages of the seminiferous epithelium. Stages per se do not present stagnant check-point-like states but continually develop as the differentiation of germ cell cohorts progresses in synchrony1,2,6. In mice, there are 12 stages (I-XII)2 that are arranged in a segmental fashion along the longitudinal axis of the seminiferous tubule, and they follow each other in a logical order thus forming the wave of seminiferous epithelium, or spermatogenic wave7,8,9 (Figure 1). Completion of spermatogenesis takes four cycles, and hierarchical layers or cohorts of differentiating germ cells&#160;within any seminiferous tubule cross-section are temporally one seminiferous cycle apart from one another. The length of the cycle is species-dependent and in the mouse each cycle takes 8.6 days10.
The stages can be identified on the basis of the cellular composition and organization of the seminiferous epithelium on histological testis sections5 (Figure 1 and Figure 2). However, histological analysis is laborious, time-consuming and requires fixing and staining, and cannot, therefore, be applied to&#160;live tissue. Importantly, staging can also be performed on living tissue under a dissection microscope by taking advantage of distinct light absorption/scatter patterns exhibited by different stages of the cycle (Figure 2). The ability of each stage to absorb and scatter light is relative to the level of chromatin condensation of late post-meiotic spermatids that any given stage hosts and the packing of these cells in bundles7,11. Spermatid differentiation, i.e., spermiogenesis, is further divided into 16 developmental steps, including 8 steps of round spermatid (step 1-8) and 8 steps of elongating spermatid (steps 9-16) differentiation (Figure 1). Step 9-11 elongating spermatids (stage IX-XI) display only a low level of chromatin condensation resulting in low amount of light being absorbed. Chromatin condensation begins in step 11 spermatids (stage XI), and step 15-16 elongating spermatids (stage IV-VIII) contain fully condensed chromatin, and therefore&#160;exhibit maximal light absorption (Figure 3). Chromatin needs to be condensed in order to be tightly packed into the sperm head. Additional factors that contribute to light absorption pattern are location of elongating spermatids within the epithelium (basal vs. apical) and bundling of elongating spermatids (pronounced in stages II-V)11 (Figure 3). Bundles are seen as spots in the middle of the tubules and as stripes on the edges under a dissection microscope and the more condensed the chromatin, the darker the spot/stripe11.
This article describes the use of transillumination-assisted microdissection method for the isolation of seminiferous tubule segments representing specific stages of the seminiferous epithelial cycle. Once isolated, staged tubule segments can be subject to various downstream analyses, including biochemical RNA and protein analyses12,13,14,15, flow cytometry16, ex vivo tubule culture17 and immunostaining18. Here we also provide detailed downstream protocols to prepare squashed monolayers of staged tubule segments for live cell morphological analysis and subsequent immunostaining, as well as whole-mount immunostainings of tubule segments. The workflow in a nutshell in described in Figure 4.
The transillumination-assisted microdissection method allows accurate identification and isolation of germ cells at specific steps of differentiation thanks to the synchronized cellular composition of the stages. Importantly, it also enables the study of stage-dependent events during spermatogenesis on live tissue. Given the lack of scalable in vitro models for spermatogenesis, this method also has a unique advantage of allowing targeted short-term developmental and toxicological studies on stage-specific tubule segments ex vivo12,17. While we describe the method here for the mouse, the same procedure can be applied to any mammalian species with longitudinal and segmental arrangement of seminiferous epithelial stages, such as the rat4,7,15,19,20.

<table><tbody><tr><td>bovine serum albumin (BSA)</td><td>Sigma</td><td>A9647</td><td></td></tr><tr><td>cover glass 20x20 mm</td><td>Menzel Gl&amp;auml;ser</td><td>11961988</td><td></td></tr><tr><td>Falcon conical tube 15-ml</td><td>Sarstedt</td><td>62.554.502</td><td></td></tr><tr><td>fetal bovine serum (FBS)</td><td>Biowest</td><td>S1810</td><td></td></tr><tr><td>grease pen (ImmEdge)</td><td>Vector Laboratories</td><td>H-4000</td><td></td></tr><tr><td>microscope slide Superfrost Plus</td><td>Thermo Scientific</td><td>22-037-246</td><td></td></tr><tr><td>Parafolmaldehyde (PFA)</td><td>Electron Microscopy Sciences</td><td>15714</td><td></td></tr><tr><td>Petri dish (100-mm)</td><td>Greiner</td><td>664160</td><td></td></tr><tr><td>phosphate-buffered saline (PBS)</td><td>Gibco</td><td>11503387</td><td></td></tr><tr><td>ProLong Diamond Antifade Mountant</td><td>Thermo Fisher</td><td>P36962</td><td></td></tr><tr><td>rhodamine-labelled Peanut agglutinin (PNA)</td><td>Vector Laboratories</td><td>RL-1072</td><td></td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>93443</td><td></td></tr><tr><td>Tween-20</td><td>Sigma</td><td>P2287</td><td></td></tr></tbody></table>

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.

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

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. ...

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7.3K Views.  University of Turku, Turku, Finland. Dit protocol beschrijft transillumination-ondersteunde microdissectie van segmenten van volwassen muis seminifereuze tubuli die specifieke stadia van seminifereuze epitheelcyclus vertegenwoordigen, en celtypen daarin, en daaropvolgende immunostaining van squashpreparaten en intacte tubulussegmenten.

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

Organ Culture and Whole Mount Immunofluorescence Staining of Mouse Wolffian Ducts

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

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

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

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 ...

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. Whole mount preparations preserve the structure of meiotic nuclei, and importantly, each gonad arm contains all stages of meiosis, organized in a temporal-spatial progression that makes it easy to identify nuclei at different stages. Adult hermaphrodites have two gonad arms, each organized as a closed tube with proliferating germline stem cells at the distal closed end and cellularized oocytes at the proximal open end, which join in the center at the uterus. Dissection releases one or both gonad arms from the body cavity, allowing the entirety of meiosis to be visualized. Here, a common protocol for immunofluorescence against a protein of interest is presented, followed by DAPI staining to mark all chromosomes. Young adults are immobilized in levamisole and quickly dissected using two syringe needles. After germline extrusion, the sample is fixed before undergoing a freeze crack in liquid nitrogen, which helps permeabilize the cuticle and other tissues. The sample can then be dehydrated in ethanol, rehydrated, and incubated with primary and secondary antibodies. DAPI is added to the sample in the mounting medium, which allows reliable visualization of DNA and makes it easy to find animals to image under a fluorescent microscope. This technique is readily adopted by those familiar with handling C. elegans after a few hours spent practicing the dissection method itself. This protocol has been taught to high-schoolers and undergraduates working in a research lab and incorporated into a course-based undergraduate research experience at a liberal arts college.

C. elegans Gonad Dissection and Freeze Crack for Immunofluorescence and DAPI Staining

This is a commonly used method for C. elegans gonad dissection followed by freeze crack, which produces germline samples for immunofluorescence via antibody staining, or for simple DAPI staining to visualize DNA. This protocol has been successful for undergraduates in a research lab and in a course-based undergraduate research experience.

The C. elegans germline makes an excellent model for studying meiosis, in part due to the ease of conducting cytological analyses on dissected animals. ...

Genetics

Analysis of Epididymal Protein Synthesis and Secretion

The mammalian epididymis generates one of the most complex intraluminal fluids of any endocrine gland in order to support the post-testicular maturation and storage of spermatozoa. Such complexity arises due to the combined secretory and absorptive activity of the lining epithelial cells. Here, we describe the techniques for the analysis of epididymal protein synthesis and secretion by focusing on the model protein family of dynamin (DNM) mechanoenzymes; large GTPases that have the potential to regulate bi-directional membrane trafficking events. For the study of protein expression in epididymal tissue, we describe robust methodology for immunofluorescence labeling of target proteins in paraffin-embedded sections and the subsequent detection of the spatial distribution of these proteins via immunofluorescence microscopy. We also describe optimized methodology for the isolation and characterization of exosome like vesicles, known as epididymosomes, which are secreted into the epididymal lumen to participate in intercellular communication with maturing sperm cells. As a complementary approach, we also describe the immunofluorescence detection of target proteins in an SV40-immortalized mouse caput epididymal epithelial (mECap18) cell line. Moreover, we discuss the utility of the mECap18 cell line as a suitable in vitro model with which to explore the regulation of epididymal secretory activity. For this purpose, we describe the culturing requirements for the maintenance of the mECap18 cell line and the use of selective pharmacological inhibition regimens that are capable of influencing their secretory protein profile. The latter are readily assessed via harvesting of conditioned culture medium, concentration of secreted proteins via trichloroacetic acid/acetone precipitation and their subsequent analysis via SDS-PAGE and immunoblotting. We contend that these combined methods are suitable for the analysis of alternative epididymal protein targets as a prelude to determining their functional role in sperm maturation and/or storage.

Here, we report the immunofluorescence localization of dynamin to illustrate the protocols for the detection of proteins in paraffin-embedded mouse epididymal sections and those of an immortalized epididymal cell line (mECap18). We also describe the protocols for the isolation of secretory proteins from both epididymal fluid and conditioned cell media.

The mammalian epididymis generates one of the most complex intraluminal fluids of any endocrine gland in order to support the post-testicular maturation ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

Microdissection and Dissociation of the Murine Oviduct: Individual Segment Identification and Single Cell Isolation

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental treatments. However, because of their small body size, some structures, such as the oviduct with a diameter of 200-400 &#956;m, have proven to be relatively difficult to study except by immunohistochemistry. Recently, immunohistochemical studies have uncovered more complex differences in oviduct segments than were previously recognized; thus, the oviduct is divided into four functional segments with different ratios of seven distinct epithelial cell types. The different embryological origins and ratios of the epithelial cell types likely make the four functional regions differentially susceptible to disease. For example, precursor lesions to serous intraepithelial carcinomas arise from the infundibulum in mouse models and from the corresponding fimbrial region in the human fallopian tube. The protocol described here details a method for microdissection to subdivide the oviduct in such a way to yield a sufficient amount and purity of RNA necessary for downstream analysis such as reverse transcription-quantitative PCR (RT-qPCR) and RNA sequencing (RNAseq). Also described is a mostly non-enzymatic tissue dissociation method appropriate for flow cytometry or single cell RNAseq analysis of fully differentiated oviductal cells. The methods described will facilitate further research utilizing the murine oviduct in the field of reproduction, fertility, cancer, and immunology.

A method for microdissection of the mouse oviduct that allows collection of the individual segments while maintaining RNA integrity is presented. In addition, non-enzymatic oviductal cell dissociation procedure is described. The methods are appropriate for subsequent gene and protein analysis of the functionally different oviductal segments and dissociated oviductal cells.

Mouse model systems are unmatched for the analysis of disease processes because of their genetic manipulability and the low cost of experimental ...

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 ...

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 ...

Biology

In Vivo Microinjection and Electroporation of Mouse Testis

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular transfection technique for testicular mouse cells allows the study of unique processes in spermatogenesis.
The following protocol focuses on transfection of testicular mouse cells with plasmid constructs. Specifically, we used the reporter vector pEGFP-C1, which expresses enhanced green fluorescent protein (eGFP) and also the pDsRed2-N1 vector expressing red fluorescent protein (DsRed2). Both encoded reporter genes were under the control of the human cytomegalovirus immediate-early promoter (CMV).
For performing gene transfer into mouse testes, the reporter plasmid constructs are injected into testes of living mice. To that end, the testis of an anaesthetized animal is exposed and the site of microinjection is prepared. Our preferred place of injection is the efferent duct, with the ultimately connected rete testis as the anatomical transport route of the spermatozoa between the testis and the epididymis. In this way, the filling of the seminiferous tubules after microinjection is excellently managed and controlled due to the use of stained DNA solutions. After observing a sufficient filling of the testis by its colored tubule structure, the organ is electroporated. This enables the transfer of the DNA solution into the testicular&#160;cells. Following 3 days of incubation, the testis is removed and investigated under the microscope for green or red&#160;fluorescence, illustrating transfection success.
Generally, this protocol can be employed for delivering DNA- or RNA- constructs into living mouse testis in order to (over)express or knock down genes, facilitating in vivo gene function analysis. Furthermore, it is suitable for studying reporter constructs or putative gene regulatory elements. Thus, the main advantages of the electroporation technique are fast performance in combination with low effort as well as the moderate technical equipment and skills required compared to alternative techniques.

In Vivo Microinjection and Electroporation of Mouse Testis

This article describes microinjection and electroporation of mouse testis in&#160;vivo as a&#160;transfection technique for testicular mouse cells to study unique processes of spermatogenesis. The presented protocol involves steps of glass capillary preparation, microinjection via the efferent duct, and transfection by electroporation.

This video and article contribution gives a comprehensive description of microinjection and electroporation of mouse testis in vivo. This particular ...

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 ...

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

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