The authors wish to thank Dr. Leonid Volkov and &#201;ric Bouchard for their technical advice regarding epifluorescence microscopy.
Financial support 
Funded by the Canadian Institutes of Health Research (grant #MOP-93781) to G.B.

Animal care was in accordance with the Universit&#233; de Sherbrooke animal care and use committee.

1. Tube Preparation
<ol>
	<li>The day before cell sorting, add 1-2 ml of heat-inactivated fetal bovine serum (FBS) to 5 ml polypropylene round bottom tubes, and to 15 ml and 50 ml polypropylene conical tubes. 
	Critical step: Ensure that every tube used in the protocol is coated. 
	Note: FBS coating prevents germ cells from sticking to tube walls.</li>
	<li>Slowly mix O/N by inversion at 4&#160;&#176;C to coat the tubes uniformly using a rotator.</li>
	<li>The next day, remove FBS from coated tubes by decanting.
	<ol>
		<li>*Critical step: Ensure that the 5 ml polypropylene round bottom tubes and the 15 and 50 ml tubes used for cell preparation (protocol steps 2.1 to 2.11) are completely emptied of FBS to prevent fanning (i.e., imprecise deviation of the side streams). Use a P200 pipet to remove the FBS. 
		Note: Residual FBS in tubes prior to sorting may lead to a loss of cells or contamination of other tubes used during cell sorting,&#160;</li>
		<li>Leave a small residual volume of FBS (about 100-200 &#181;l) in the 5 ml polypropylene round bottom tubes used for collecting purified cells.</li>
	</ol>
	</li>
</ol>
2. Cell Preparation
<ol>
	<li>Sacrifice one male mouse under anesthesia using O2/isoflurane (induction at 5% then maintained at 2%) followed by CO2 asphyxiation.</li>
	<li>Excise and decapsulate both testes and mince with small scissors in 500 &#181;l of sorting buffer into a 1.5 ml tube.</li>
	<li>Flush germ cells from seminiferous tubules by gentle up and down pipetting in the 1.5 ml tube, first with a truncated 100-1,000 &#181;l tip, and then with an intact 100-1,000 &#181;l tip.</li>
	<li>Remove debris and clumps by one filtration step using a 40 &#181;m cell strainer and collect filtrate into the 50 ml conical tube previously coated with FBS in step 1.</li>
	<li>Wash the filter once with sorting buffer up to a total volume of 3 ml and transfer filtrate into the 15 ml conical tube previously coated in step 1.</li>
	<li>Add 16 &#181;l of 50 mM EDTA pH 8 per milliliter of cell suspension (48 &#181;l for 3 ml) so as to obtain a final concentration of 0.8 mM EDTA.</li>
	<li>To fix germ cells, slowly add 3 volumes of ice-cold 100% ethanol using a 10 ml pipet with gentle agitation (vortex at low speed), which will produce a milky suspension. 
	Critical step: Slowly adding ethanol is crucial and important to preserve the integrity of the cell preparation. We noted that rapid addition of ethanol causes cell lysis resulting in a major decrease of sorting efficiency. For a 3 ml cell suspension, 9 ml of ethanol should be added in approximately 1 min.</li>
	<li>Incubate cells on ice for 15 min with occasional mixing by inversion.</li>
	<li>Centrifuge cell suspension at 800 x g for 5 min and remove the clear supernatant.</li>
	<li>Gently resuspend the fixed germ cells in 2 ml of sorting buffer.</li>
	<li>Add 4 &#181;l of SYTO16 DNA dye (1 mM solution in DMSO) and incubate for at least 30 min (protected from light). 
	Note: Best sorting results are obtained using SYTO16 since it is a permeable DNA dye that is easily incorporated into the highly compacted nuclei of spermatids.</li>
</ol>
3. Cell Sorter Set Up
Note: Here, a 4-Laser (405 nm - violet, 488 nm - blue, 561 nm - yellow-green, 633 nm - red) 20-parameter BDFACSAria III cell sorter is used. The BD FACSDiva 6.1.3 software is used to visualize and analyze the data. The settings may vary depending on the type of sorter used.
<ol>
	<li>Sterilize the FACS sorter by running 70% ethanol as sheath fluid during the fluidics shutdown procedure (&#34;Cytometer&#34; menu in the software) prior to sorting. Select &#34;Cytometer&#34; in the toolbar of the software and choose &#34;Fluidic shutdown&#34;. Follow the steps mentioned by the processes.</li>
	<li>Prepare and filter 1x PBS with a 0.22 &#181;m filter so as to eliminate any crystals and contaminants. Fill the sheath tank to the upper weld line on the inside of the tank.</li>
	<li>Start the software and log in. Start the FACS sorter to warm up the lasers for at least 30 min prior to sorting. Run two fluidic startup processes to flush out the ethanol from the fluidics. Select &#34;Cytometer&#34; in the toolbar of the software and choose &#34;Fluidic startup&#34;. Follow the steps mentioned by the processes. 
	Note: This restores fluidics with normal sheath fluid (1x PBS).</li>
	<li>Clean the sorting block and the deflection plates with distilled water and dry them thoroughly. Sonicate a 100 &#181;m nozzle placed in a tube of distilled water for 2 min in a sonication bath. Wipe the nozzle thoroughly.</li>
	<li>Insert the 100 &#181;m nozzle into the flow cell and turn the nozzle-locking lever clockwise to the 12 hr position. Set the sheath pressure to 20 psi.</li>
	<li>Turn on the stream and adjust the amplitude to obtain target values for frequency (around 30), Drop 1 (around 150) and Gap (around 12) and to establish a stable droplet breakoff pattern (few satellite drops). Turn off attenuation.</li>
	<li>Make sure that the stream is straight and hits the center of the waste aspirator by changing the angle of the sort block. Note: A 130 &#181;m nozzle at 10 psi was also tested and no obvious difference was found.</li>
	<li>Set laser delay to 0 for the blue, -77.39 for the red, 37.33 for the violet and -39.85 for the yellow-green lasers.</li>
	<li>Set areas scaling to 1.14 for the blue, 1.0 for the red, 0.75 for the violet and 0.96 for the yellow-green lasers.</li>
	<li>Using a Test Sort (in the &#34;Side Stream&#34; window of the software), make sure that side streams are not fanning. In the &#34;Side Stream&#34; window, adjust the voltage sliders of each side stream so that they hit the middle of test collection tube. If the center stream is not tight, adjust the 2nd, 3rd, and 4th drop settings to constrain the center stream.</li>
	<li>Start the Cytometer Setup and Tracking (CS&#38;T) application in the software (&#34;Cytometer&#34; menu).</li>
	<li>Use the cytometer setup and tracking beads at 1 drop per 350 &#181;l to automate the characterization and tracking of the cytometer performance using manufacturer&#39;s instructions.</li>
	<li>Quit the CS&#38;T application. Apply CS&#38;T setting, make sure the stream parameters still allow a stable droplet breakoff pattern, and turn on the Sweet Spot button as described in manufacturer&#39;s instructions (in the &#34;Breakoff window&#34;).</li>
	<li>Optimize the Drop Delay value using fluorescent beads (see Table of Materials).
	<ol>
		<li>Install the collection tubes in the holder. Open the sort block door. In the &#34;Side Stream&#34; window, turn on voltage then select &#34;Test sort&#34; and click the Aspirator Drawer button to open the drawer and have access to the collection tubes.</li>
		<li>Optimize the side streams so they go into the center of the tube. Adjust values and sliders as needed.</li>
		<li>Close the sort block door and make sure to adjust the micrometer dial to obtain the brightest bead spot at the center. Unselect waste drawer, test sort and voltage. Remove the collection tubes.</li>
		<li>Optimize the drop delay according to manufacturers&#39; instructions.
		<ol>
			<li>Briefly, shake bead vial vigorously and dilute 1 drop of beads in 0.5 ml of PBS.</li>
			<li>Using manufacturers&#39; instructions, set the drop delay using the Drop Delay experiment template available in the software. Alternatively, use the Auto Drop Delay feature to set the drop delay.</li>
			<li>Load the tube of fluorescent beads and adjust the flow rate until the threshold rate is ~3,000 to 4,000 events/second. Set the sort precision to &#34;Initial&#34; and finally to &#34;Fine tune&#34; in the &#34;Sort layout&#34; window. Select the optical filter and sort beads. Adjust the Drop Delay until ~100% are sorted to the left. 
			Note: This step is critical to make sure that cells of interest sort into a side stream. The beads and the optical filter are used to achieve close to a 100% droplet deviation.&#160;</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Place a FSC 1.5 ND (neutral density) filter at the left end of the FSC detector block. Set the window extension to 2.00 &#181;s and the FSC Area scaling to 1.00 (in the &#34;Laser&#34; tab in the &#34;Cytometer&#34; window).</li>
	<li>Before loading sample tube, place a sample filter line of 50 &#181;m at the end of the sample line to avoid clumping. To do so, select &#34;Change Sample Filter&#34; in the &#34;Cytometer&#34; menu.</li>
	<li>Test the composition of the sample fluid to achieve good separation without fanning between side stream and to prevent the cells from sticking to the polypropylene tubes.
	<ol>
		<li>Load the sample tube, turn on the deflection plates and select &#34;Test sort&#34; with the drawer closed. Look at side streams if separation occurs without too much fanning. 
		Note: Fanning is likely due to a high protein concentration in the sample buffer.</li>
	</ol>
	</li>
	<li>Load the sample tube into the FACS instrument. Select a sample agitation of 300 rpm and a sample temperature of 4 &#176;C. Select &#34;Acquire Data&#34; in the &#34;Acquisition Dashboard&#34;.</li>
	<li>If necessary, dilute the sample to achieve an appropriate event rate to a maximum of 5,000 events/sec (at a maximum flow rate of 3.0). Strictly respect these limits so as to maintain the purity of the sorted cells.</li>
	<li>To analyze and sort the cells, optimize the FSC threshold value to eliminate debris without interfering with the population of interest. In logarithmic scale, adjust the voltage of the photo-multiplier tube (PMT) detector with the 530/30 filter in order to make sure that the fluorescence signal of the total population fits within the scale, and gate on the positive population (Figure 1, Gate 1).</li>
	<li>Produce a Forward Scatter-area (FSC-A)/Side Scatter-area (SSC-A) plot of the SYTO16 positive population and adjust the FSC-A to around 110 V in linear scale, and the SSC-A to around 150 V in logarithmic scale to visualize all the events while excluding very large cells and cell aggregates.</li>
	<li>In the &#34;Sort Layout&#34; window, set the &#34;Sort Precision Mode&#34; to &#34;4-way purity&#34; (Yield mask: 0, Purity Mask: 32, Phase Mask: 0). Select &#34;Continuous&#34; from the &#34;Target Events&#34; menu for continuous sorting.</li>
	<li>Add populations to sort in the field corresponding to the tubes (in the &#34;Sort Layout&#34; window). Place populations with higher frequencies in the middle and those with lower frequencies at the ends. Set the voltage plates to 2,500 V. During sorting, keep the sample collection tubes on ice.&#160;&#160;&#160;&#160;</li>
</ol>
4. Cell Sorting
<ol>
	<li>Immediately before sorting, filter cells using a 50 &#181;m filter in a previously coated 5 ml polypropylene round bottom tube.</li>
	<li>Wash the filter extensively with 1-2 ml of sorting buffer.</li>
	<li>Sort fixed germ cells using a 488 nm laser-equipped cell sorter following the gating scheme detailed in Figure 1. Collect purified fractions in 5 ml polypropylene round bottom tubes previously coated in section 1 on ice.
	<ol>
		<li>Make sure that 100-200 &#181;l of FBS is kept in the collection tubes to avoid sticking of the sorted cells to the tube wall.</li>
		<li>Make sure that the concentration of SYTO16 is saturating to avoid fluorescence fluctuations during the sorting period. SYTO16 is saturating when there is no further variation of cell staining intensity at the Alexa Fluor 488 parameter (DNA staining) in the sorting graphics. If necessary, add 1 &#181;l of SYTO16 to the sorting tube to reach saturation.</li>
		<li>Display the total events on a graphic displaying the following parameters: number of events vs Alexa Fluor 488-A.</li>
		<li>Select the positive DNA staining cells with Gate 1, as shown in Figure 1.
		<ol>
			<li>Display cells from gate 1 on a graphic using SSC-A vs FSC-A as parameters
			<ol>
				<li>Select the cells with Gate 2, as showed in Figure 1.</li>
				<li>Display cells from Gate 2 on a graphic using FSC-A vs Alexa Fluor 488-A as parameters.</li>
				<li>Select cells with Gate 5 and Gate 6 on Gate 2 graphic, as showed in Figure 1.</li>
				<li>Display separately Gate 5 and Gate 6 on graphics using Alexa Fluor 488-W vs Alexa Fluor 488-A as parameters.</li>
				<li>Select spermiogenesis steps 1-9 spermatids on the Gate 5 graphic and spermiogenesis steps 10-12 spermatids on the Gate 6 graphics, as showed in Figure 1.</li>
			</ol>
			</li>
			<li>Display cells from gate 1 on a graphic using FSC-A vs Alexa Fluor 488-A as parameters
			<ol>
				<li>Select cells with Gate 3 and Gate 4, as showed in Figure 1.</li>
				<li>Display separately Gate 3 and Gate 4 on graphics using Alexa Fluor 488-W vs Alexa Fluor 488-A as parameters.</li>
				<li>Select spermiogenesis steps 13-14 spermatids on the Gate 3 graphic and spermiogenesis steps 15-16 spermatids on the Gate 4 graphic, as showed in Figure 1.&#160;&#160;&#160;&#160;</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors declare no competing financial interests.

Spermatogenic cells have always been challenging to study given the complexity of the seminiferous epithelium, as well as the limited success of in vitro culture. Over the years, many approaches to purify germ cells from various species were developed. Sedimentation techniques using gravity purification with Percoll or bovine serum albumin gradients usually provide a good yield of intact germinal cells, but lack proper definition between some cell types such as meiotic tetraploid cells and spermatids10</sup...

Haploid round spermatids differentiate into spermatozoa by a process called spermiogenesis. This involves many different steps including the acquisition of a flagellum, chromatin and cytoskeleton remodeling, condensation of the nucleus as well as the loss of most of the cytoplasm. These unique cellular events must be finely regulated in order to produce a mature functional gamete with an intact genome suitable for fertilization. Spermiogenesis can hardly be studied in vitro since no reliable cell culture system has so far been able to support progression through the different steps of the process. Moreover, actual in vitro techniques lead to a poor yield1,2. In vivo, proper transitions through the different steps of spermiogenesis are crucial for the natural functional integrity of the male gamete. Successful purification of spermatids according to their differentiation steps has never been accomplished with a level of enrichment sufficient to allow molecular characterization of spermiogenesis. For instance, purification of key steps of the spermatidal differentiation would be especially useful to study the developing acrosome, formation of the midpiece3, cell junction dynamics4, RNA dynamics5, chromatin remodeling process6,7 or genomic stability8. Purification of spermatids has been hampered by their progressive morphological transformation, the lack of known stage-specific external biomarkers, and their peculiar shape and size. 
Although most male germ cells display a direct relationship between DNA staining and ploidy (DNA content), we noticed that such positive correlation is no longer applicable to spermatids. This stems from our early observation that seminiferous tubule sections show variable intensity of DNA staining throughout the different spermiogenesis steps. Although DNA staining is consistent with their haploid set of chromosomes from spermiogenesis steps 1 to 7 (round spermatids), we observed a sharp increase in fluorescence intensity with DAPI or SYTO 16 around the onset of nuclear reorganization and chromatin remodeling (spermiogenesis step 8) reaching a peak at the onset of nuclear condensation (spermiogenesis steps 11-12). Following condensation of the nucleus, DNA staining intensity decreases until spermiation (spermiogenesis step 16). We surmised that this was likely associated with the formation of their peculiar chromatin structure transition where histones are replaced by protamines. We therefore developed a reliable flow cytometry method that allows the separation of spermatids using the variation of DNA intensity of spermatids as a main selection parameter.
A simple flow cytometry approach is described to separate mouse spermatids with high purity (95-100%) based on their apparent DNA content (SYTO16 staining), size and granulosity. Spermatids are separated into four populations; spermiogenesis steps 1-9, 10-12, 13-14 and 15-16. Purified spermatids are suitable for genetic/genomic analysis, as well as proteomic applications as described in a recent publication from our group9.

<table><tbody><tr><td>Isoflurane</td><td>ABBOT</td><td>05260-05</td><td>For mouse anesthesia before euthanasia</td></tr><tr><td>Fetal bovine serum</td><td>Wisent</td><td>90150</td><td>For tube coating</td></tr><tr><td>1x PBS</td><td></td><td></td><td></td></tr><tr><td>EDTA</td><td>BioShop</td><td>EDT</td><td>For sorting buffer preparation</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H</td><td>For sorting buffer preparation</td></tr><tr><td>100% Ethanol</td><td>Les alcools de commerce</td><td>092-09-11N</td><td>For cell fixation</td></tr><tr><td>SYTO 16</td><td>Life Technologies</td><td>S7578</td><td>DNA staining</td></tr><tr><td>5 ml polypropylene round bottom tubes</td><td>BD Falcon</td><td>352063</td><td>Sorted cells collection</td></tr><tr><td>15 ml polypropylene conical bottom tubes</td><td>PROgene</td><td>1500</td><td></td></tr><tr><td>50 ml polypropylene conical bottom tubes</td><td>PROgene</td><td>5000</td><td></td></tr><tr><td>TEC4 anaesthetic vaporizer</td><td>Ohmeda</td><td>1160526</td><td>For mouse euthanasia</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; gas tank</td><td>Praxair</td><td>C799117902</td><td>For mouse euthanasia</td></tr><tr><td>O&lt;sub&gt;2&lt;/sub&gt; gas tank</td><td>Praxair</td><td>O254130501</td><td>For mouse euthanasia</td></tr><tr><td>Homemade mouse gas chamber</td><td></td><td></td><td>For mouse euthanasia</td></tr><tr><td>40 &amp;micro;m Falcon cell strainer</td><td>Corning Incorporated</td><td>352340</td><td></td></tr><tr><td>50 &amp;mu;m sample line filters</td><td>BD Biosciences</td><td>649049</td><td></td></tr><tr><td>Vortex mixer</td><td>Labnet international, inc.</td><td>S0200</td><td>For cell fixation</td></tr><tr><td>Dynac centrifuge</td><td>Clay Adams</td><td>101</td><td></td></tr><tr><td>Celltrics 50 &amp;micro;m filters</td><td>Partec</td><td>04-004-2327</td><td></td></tr><tr><td>488 nm laser-euipped cell sorter</td><td>BD Biosciences</td><td>FACSAria III</td><td></td></tr><tr><td>Accudop Fluorescent Beads</td><td>BD Biosciences</td><td>345249</td><td></td></tr><tr><td>Sorting Buffer: 1x PBS, 1 mM EDTA pH 8.0, 25 mM HEPES pH 7.0, 1% FBS</td><td></td><td></td><td>FBS is heat-inactivated. Make fresh solution, 0.22 &amp;mu;m filtered and keep at 4&amp;deg;C.</td></tr></tbody></table>

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.

Gating strategy used with flow cytometry
Figure 1 represents the gating strategy used in flow cytometry to sort four highly pure spermatid populations. Briefly, cells with positive DNA staining (Alexa Fluor 488-A) are first selected with Gate 1. Spermatids from spermiogenesis steps 1-12 are selected (Gate 2) on a dot plot showing the granulosity (SSC-A) vs size (FSC-A) from Gate 1. Then, spermatids from spermiog...

Watch this Scientific Journal Video about Step-specific Sorting of Mouse Spermatids by Flow Cytometry at JoVE.com

Step-specific Sorting of Mouse Spermatids by Flow Cytometry

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 ...

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JoVE Journal

Developmental Biology

10.5K Views.  Université de Sherbrooke. 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.

Video: Step-specific Sorting of Mouse Spermatids by Flow Cytometry

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair cells are aligned in one row of inner hair cells and three rows of outer hair cells that extend along the basal to apical axis of the cochlea. The explant culture technique described here provides an efficient method to isolate and maintain cochlear explants from the embryonic mouse inner ear. Also, the morphology and molecular characteristics of sensory hair cells and nonsensory supporting cells within the cochlear explant cultures resemble those observed in vivo and can be studied within its intrinsic cellular environment. The cochlear explants can serve as important experimental tools for the identification and characterization of molecular and genetic pathways that are involved in cellular specification and patterning. Although transgenic mouse models provide an effective approach for gene expression studies, a considerable number of mouse mutants die during embryonic development thereby hindering the analysis and interpretation of developmental phenotypes. The organ of Corti from mutant mice that die before birth can be cultured so that their in vitro development and responses to different factors can be analyzed. Additionally, we describe a technique for electroporating embryonic cochlear explants ex vivo which can be used to downregulate or overexpress specific gene(s) and analyze their potential endogenous function and test whether specific gene product is necessary or sufficient in a given context to influence mammalian cochlear development1-8.

We present a method that describes isolation and culture of cochlear explants from embryonic mouse inner ear. We also demonstrate a method for gene transfer into cochlear explants via square-wave electroporation. The in vitro explant culture coupled with gene transfer technique enables researchers to study the effects of altering gene expression during development.

Auditory hair cells located within the mouse organ of Corti detect and transmit sound information to the central nervous system. The mechanosensory hair ...

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 ...

Biology

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 ...

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 ...

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 ...

Step-specific Sorting of Mouse Spermatids by Flow Cytometry

Summary

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Chapters in this video

Culture of Embryonic Mouse Cochlear Explants and Gene Transfer by Electroporation

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

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

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

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

Flow Cytometry Purification of Mouse Meiotic Cells

Mouse Round Spermatid Injection

Simple and Efficient Technique for the Preparation of Testicular Cell Suspensions

Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation

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