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

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

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanistic+insights+into+the+regulation+of+the+spermatogonial+stem+cell+niche.">Mechanistic insights into the regulation of the spermatogonial stem cell niche.</a> Cell Cycle. 5, 1164-1170 (2006).</li><li>Dores, C., Alpaugh, W., Dobrinski, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=From+in+vitro+culture+to+in+vivo+models+to+study+testis+development+and+spermatogenesis.">From in vitro culture to in vivo models to study testis development and spermatogenesis.</a> Cell Tissue Res. 349, 691-702 (2012).</li><li>Hayashi, K., Ohta, H., Kurimoto, K., Aramaki, S., Saitou, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reconstitution+of+the+mouse+germ+cell+specification+pathway+in+culture+by+pluripotent+stem+cells.">Reconstitution of the mouse germ cell specification pathway in culture by pluripotent stem cells.</a> Cell. 146, 519-532 (2011).</li><li>Easley, C. 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The authors declare that they have no competing financial interests. This protocol being demonstrated involves the use of laboratory mice and was executed in compliance with all relevant guidelines, regulations and regulatory agencies. Animals used in this protocol are maintained under the guidance and approval of the University of Pennsylvania Institutional Animal Care and Use Committee (IACUC protocol #804284).

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

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

<table border="0" cellpadding="0" cellspacing="0" width="856">
	<tbody>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">BSA</td>
			<td style="width:109px;">Affymetrix/USB</td>
			<td style="width:255px;">10857 100MG</td>
			<td style="width:183px;">Alternate can be used.</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:311px;">0.5%, 2%, and 4% BSA solutions</td>
			<td colspan="2" style="width:364px;">Dissolve 0.25 g, 11 g, and 22 g (respectively) in total volumes of 50 ml, 550 ml, and 550 ml of 1x KREBS (respectively).</td>
			<td style="width:183px;">To be made day of the STA-PUT procedure. Filter sterilize.</td>
		</tr>
		<tr height="91">
			<td height="91" style="height:91px;width:311px;">KREBS (10x)</td>
			<td colspan="2" style="width:364px;">3.26 g KH2PO4 + 139.5 g NaCl +5.89 g MgSO4/7H20 + 40 g Dextrose + 3.78 g CaCl2/2H20 + 7.12 g KCl, bring to 2 L with ddH20</td>
			<td style="width:183px;">Autoclave/filter and store at 4 &#176;C for several months.</td>
		</tr>
		<tr height="64">
			<td height="64" style="height:64px;width:311px;">KREBS (1x)</td>
			<td colspan="2" style="width:364px;">Dissolve 4.24 g NaHCO3 in 100 ml ddH20. Add 200 ml 10x KREBS. Bring to 2 L with ddH20.</td>
			<td style="width:183px;">To be made day of the STA-PUT procedure. Filter sterilize.</td>
		</tr>
		<tr height="17">
			<td height="35" rowspan="2" style="height:35px;width:311px;">Collagenase</td>
			<td rowspan="2" style="width:109px;">Sigma</td>
			<td rowspan="2" style="width:255px;">C9891-1G</td>
			<td rowspan="2" style="width:183px;"> </td>
		</tr>
		<tr height="18">
		</tr>
		<tr height="17">
			<td height="35" rowspan="2" style="height:35px;width:311px;">DNAse</td>
			<td rowspan="2" style="width:109px;">Sigma</td>
			<td rowspan="2" style="width:255px;">DNEP-5MG</td>
			<td rowspan="2" style="width:183px;"> </td>
		</tr>
		<tr height="18">
		</tr>
		<tr height="17">
			<td height="35" rowspan="2" style="height:35px;width:311px;">Trypsin</td>
			<td rowspan="2" style="width:109px;">Sigma</td>
			<td rowspan="2" style="width:255px;">T9201-1G</td>
			<td rowspan="2" style="width:183px;"> </td>
		</tr>
		<tr height="18">
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">DAPI</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Triton-X100</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Formaldehyde (~37%)</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
		<tr height="17">
			<td height="17" style="height:17px;"> </td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:311px;">TABLE 2: EQUIPMENT</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="18">
			<td height="18" style="height:18px;"> </td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Equipment</td>
			<td style="width:109px;">Company</td>
			<td style="width:255px;">Catalog #</td>
			<td style="width:183px;">Comments</td>
		</tr>
		<tr height="169">
			<td height="169" style="height:169px;width:311px;">Complete STA-PUT apparatus</td>
			<td style="width:109px;">ProScience Glass Shop</td>
			<td style="width:255px;">STA-PUT (this procedure uses the standard sedimentation chamber: Cat. No. 56700-500)</td>
			<td style="width:183px;">Includes all glassware and equipment for the apparatus. Alternate glassware is available for performing STA-PUT with smaller cell numbers.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">10 &#956;m mesh filter</td>
			<td style="width:109px;">Fisherbrand</td>
			<td style="width:255px;">22363549</td>
			<td style="width:183px;"> </td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Mouse dissection tools</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
		<tr height="21">
			<td height="43" rowspan="2" style="height:43px;width:311px;">14 ml round bottom fraction collection tubes with caps</td>
			<td colspan="3" style="width:547px;">Preference of researcher </td>
		</tr>
		<tr height="22">
			<td colspan="3" height="22" style="height:22px;width:547px;">Need approximately 100 tubes</td>
		</tr>
		<tr height="43">
			<td height="43" style="height:43px;width:311px;">Fraction collector</td>
			<td style="width:109px;">GE Healthcare Life Sciences</td>
			<td style="width:255px;">Model: Frac-920, Product code: 18-1177-40</td>
			<td style="width:183px;">Alternate can be used.</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Microscope slides, cover glass</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:311px;">Light/Fluorescent Microscope</td>
			<td colspan="3" style="width:547px;">Preference of researcher</td>
		</tr>
	</tbody>
</table>

separation of spermatogenic cell types using sta-put velocity sedimentation

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

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

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

Separation of Spermatogenic Cell Types Using STA-PUT Velocity Sedimentation

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

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

separation-spermatogenic-cell-types-using-sta-put-velocity

Research

JoVE Journal

Biology

25.5K Views.  University of Pennsylvania. The STA-PUT method allows for the separation of different populations of spermatogenic cells based on size and density.

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

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

Pomiar egzocytozy w Neurony Korzystanie FM etykietowania

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Electrophoretic Separation of Proteins

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to investigate subunit compositions, and to verify homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in a polyacrylamide gel matrix; pore size decreases with increasing acrylamide concentration. The combination of pore size and protein charge, size, and shape determines the migration rate of the protein. In this unit, the standard Laemmli method is described for discontinuous gel electrophoresis under denaturing conditions, i.e., in the presence of sodium dodecyl sulfate (SDS). 

In this video, we demonstrate a method for electrophoretic separation of proteins using poly-acrylimide gel electrophoresis (PAGE).

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to ...

Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis

The unicellular green alga Chlamydomonas reinhardtii (Chlamydomonas) has become a popular organism for research in diverse areas of cell biology and genetics because of its simple life cycle, ease of growth and manipulation for genetic analysis, genomic resources, and transformability of the nucleus and both organelles. Mating strains is a common practice when genetic approaches are used in Chlamydomonass, to create vegetative diploids for analysis of dominance, or following tetrad dissection to ascertain nuclear vs. organellar inheritance, to test allelism, to analyze epistasy, or to generate populations for the purpose of map-based cloning. Additionally, genetic crosses are routinely used to combine organellar genotypes with particular nuclear genotypes. Here we demonstrate standard methods for gametogenesis, mating, zygote germination and tetrad separation. This protocol consists of an easy-to-follow series of steps that will make genetic approaches amenable to scientists who are less familiar with Chlamydomonas. Key parameters and trouble spots are explained. Finally, resources for further information and alternative methods are provided.

Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis

Mating and tetrad separation are required for genetic analysis in Chlamydomonas reinhardtii. Here we demonstrate standard methods for gametogenesis, mating, zygote germination and tetrad dissection. This protocol consists of an easy-to-follow series of steps that will make genetic approaches amenable to scientists who are less familiar with Chlamydomonas.

The unicellular green alga Chlamydomonas reinhardtii (Chlamydomonas) has become a popular organism for research in diverse areas of cell biology and ...

Two Types of Assays for Detecting Frog Sperm Chemoattraction

Sperm chemoattraction in invertebrates can be sufficiently robust that one can place a pipette containing the attractive peptide into a sperm suspension and microscopically visualize sperm accumulation around the pipette1. Sperm chemoattraction in vertebrates such as frogs, rodents and humans is more difficult to detect and requires quantitative assays. Such assays are of two major types - assays that quantitate sperm movement to a source of chemoattractant, so-called sperm accumulation assays, and assays that actually track the swimming trajectories of individual sperm.
Sperm accumulation assays are relatively rapid allowing tens or hundreds of assays to be done in a single day, thereby allowing dose response curves and time courses to be carried out relatively rapidly. These types of assays have been used extensively to characterize many well established chemoattraction systems - for example, neutrophil chemotaxis to bacterial peptides and sperm chemotaxis to follicular fluid. Sperm tracking assays can be more labor intensive but offer additional data on how chemoattractancts actually alter the swimming paths that sperm take. This type of assay is needed to demonstrate the orientation of sperm movement relative to the chemoattrractant gradient axis and to visualize characteristic turns or changes in orientation that bring the sperm closer to the egg.
Here we describe methods used for each of these two types of assays. The sperm accumulation assay utilized is called a "two-chamber" assay. Amphibian sperm are placed in a tissue culture plate insert with a polycarbonate filter floor having 12 &mu;m diameter pores. Inserts with sperm are placed into tissue culture plate wells containing buffer and a chemoatttractant carefully pipetted into the bottom well where the floor meets the wall (see Fig. 1). After incubation, the top insert containing the sperm reservoir is carefully removed, and sperm in the bottom chamber that have passed through the membrane are removed, pelleted and then counted by hemocytometer or flow cytometer.
The sperm tracking assay utilizes a Zigmond chamber originally developed for observing neutrophil chemotaxis and modified for observation of sperm by Giojalas and coworkers2,3. The chamber consists of a thick glass slide into which two vertical troughs have been machined. These are separated by a 1 mm wide observation platform. After application of a cover glass, sperm are loaded into one trough, the chemoattractant agent into the other and movement of individual sperm visualized by video microscopy. Video footage is then analyzed using software to identify two-dimensional cell movements in the x-y plane as a function of time (xyt data sets) that form the trajectory of each sperm.

Eggs and the extracellular coatings around eggs frequently release peptides, proteins and small molecules that communicate with sperm to guide them to the egg thereby promoting fertilization. Using frog sperm we describe and compare two classes of assays used to detect sperm chemoattraction &#x2013; sperm accumulation assays and sperm tracking assays.

Sperm chemoattraction in invertebrates can be sufficiently robust that one can place a pipette containing the attractive peptide into a sperm suspension ...

Agarose Gel Electrophoresis for the Separation of DNA Fragments

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from the seaweed genera Gelidium and Gracilaria, and consists of repeated agarobiose (L- and D-galactose) subunits2. During gelation, agarose polymers associate non-covalently and form a network of bundles whose pore sizes determine a gel&#39;s molecular sieving properties. The use of agarose gel electrophoresis revolutionized the separation of DNA. Prior to the adoption of agarose gels, DNA was primarily separated using sucrose density gradient centrifugation, which only provided an approximation of size. To separate DNA using agarose gel electrophoresis, the DNA is loaded into pre-cast wells in the gel and a current applied. The phosphate backbone of the DNA (and RNA) molecule is negatively charged, therefore when placed in an electric field, DNA fragments will migrate to the positively charged anode. Because DNA has a uniform mass/charge ratio, DNA molecules are separated by size within an agarose gel in a pattern such that the distance traveled is inversely proportional to the log of its molecular weight3. The leading model for DNA movement through an agarose gel is &#34;biased reptation&#34;, whereby the leading edge moves forward and pulls the rest of the molecule along4. The rate of migration of a DNA molecule through a gel is determined by the following: 1) size of DNA molecule; 2) agarose concentration; 3) DNA conformation5; 4) voltage applied, 5) presence of ethidium bromide, 6) type of agarose and 7) electrophoresis buffer. After separation, the DNA molecules can be visualized under uv light after staining with an appropriate dye. By following this protocol, students should be able to: 
 
1. Understand the mechanism by which DNA fragments are separated within a gel matrix 
2. Understand how conformation of the DNA molecule will determine its mobility through a gel matrix 
3. Identify an agarose solution of appropriate concentration for their needs 
4. Prepare an agarose gel for electrophoresis of DNA samples 
5. Set up the gel electrophoresis apparatus and power supply 
6. Select an appropriate voltage for the separation of DNA fragments 
7. Understand the mechanism by which ethidium bromide allows for the visualization of DNA bands 
8. Determine the sizes of separated DNA fragments 
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A basic protocol for the separation of DNA fragments using agarose gel electrophoresis is described.

Agarose gel electrophoresis is the most effective way of separating DNA fragments of varying sizes ranging from 100 bp to 25 kb1. Agarose is isolated from ...

A Novel Bayesian Change-point Algorithm for Genome-wide Analysis of Diverse ChIPseq Data Types

ChIPseq is a widely used technique for investigating protein-DNA interactions. Read density profiles are generated by using next-sequencing of protein-bound DNA and aligning the short reads to a reference genome. Enriched regions are revealed as peaks, which often differ dramatically in shape, depending on the target protein1. For example, transcription factors often bind in a site- and sequence-specific manner and tend to produce punctate peaks, while histone modifications are more pervasive and are characterized by broad, diffuse islands of enrichment2. Reliably identifying these regions was the focus of our work.
Algorithms for analyzing ChIPseq data have employed various methodologies, from heuristics3-5 to more rigorous statistical models, e.g. Hidden Markov Models (HMMs)6-8. We sought a solution that minimized the necessity for difficult-to-define, ad hoc parameters that often compromise resolution and lessen the intuitive usability of the tool. With respect to HMM-based methods, we aimed to curtail parameter estimation procedures and simple, finite state classifications that are often utilized.
Additionally, conventional ChIPseq data analysis involves categorization of the expected read density profiles as either punctate or diffuse followed by subsequent application of the appropriate tool. We further aimed to replace the need for these two distinct models with a single, more versatile model, which can capably address the entire spectrum of data types.
To meet these objectives, we first constructed a statistical framework that naturally modeled ChIPseq data structures using a cutting edge advance in HMMs9, which utilizes only explicit formulas-an innovation crucial to its performance advantages. More sophisticated then heuristic models, our HMM accommodates infinite hidden states through a Bayesian model. We applied it to identifying reasonable change points in read density, which further define segments of enrichment. Our analysis revealed how our Bayesian Change Point (BCP) algorithm had a reduced computational complexity-evidenced by an abridged run time and memory footprint. The BCP algorithm was successfully applied to both punctate peak and diffuse island identification with robust accuracy and limited user-defined parameters. This illustrated both its versatility and ease of use. Consequently, we believe it can be implemented readily across broad ranges of data types and end users in a manner that is easily compared and contrasted, making it a great tool for ChIPseq data analysis that can aid in collaboration and corroboration between research groups. Here, we demonstrate the application of BCP to existing transcription factor10,11 and epigenetic data12 to illustrate its usefulness.

Our Bayesian Change Point (BCP) algorithm builds on state-of-the-art advances in modeling change-points via Hidden Markov Models and applies them to chromatin immunoprecipitation sequencing (ChIPseq) data analysis. BCP performs well in both broad and punctate data types, but excels in accurately identifying robust, reproducible islands of diffuse histone enrichment.

ChIPseq is a widely used technique for investigating protein-DNA interactions. Read density profiles are generated by using next-sequencing of ...

Separation of Mouse Embryonic Facial Ectoderm and Mesenchyme

Orofacial clefts are the most frequent craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal facial development3. In human and mouse, initial growth and patterning of the face relies on several small buds of tissue, the facial prominences4,5. The face is derived from six main prominences: paired frontal nasal processes (FNP), maxillary prominences (MxP) and mandibular prominences (MdP). These prominences consist of swellings of mesenchyme that are encased in an overlying epithelium. Studies in multiple species have shown that signaling crosstalk between facial ectoderm and mesenchyme is critical for shaping the face6. Yet, mechanistic details concerning the genes involved in these signaling relays are lacking. One way to gain a comprehensive understanding of gene expression, transcription factor binding, and chromatin marks associated with the developing facial ectoderm and mesenchyme is to isolate and characterize the separated tissue compartments.
Here we present a method for separating facial ectoderm and mesenchyme at embryonic day (E) 10.5, a critical developmental stage in mouse facial formation that precedes fusion of the prominences. Our method is adapted from the approach we have previously used for dissecting facial prominences7. In this earlier study we had employed inbred C57BL/6 mice as this strain has become a standard for genetics, genomics and facial morphology8. Here, though, due to the more limited quantities of tissue available, we have utilized the outbred CD-1 strain that is cheaper to purchase, more robust for husbandry, and tending to produce more embryos (12-18) per litter than any inbred mouse strain8. Following embryo isolation, neutral protease Dispase II was used to treat the whole embryo. Then, the facial prominences were dissected out, and the facial ectoderm was separated from the mesenchyme. This method keeps both the facial ectoderm and mesenchyme intact. The samples obtained using this methodology can be used for techniques including protein detection, chromatin immunoprecipitation (ChIP) assay, microarray studies, and RNA-seq.

A protocol for separation of embryo facial ectoderm and mesenchyme is described. We use Dispase II to treat whole embryos first, dissect whole facial prominences out, and then separate the facial ectoderm and mesenchyme.

 Orofacial clefts are the most frequent  craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal ...

Laser-assisted Microdissection (LAM) as a Tool for Transcriptional Profiling of Individual Cell Types

The understanding of developmental processes at the molecular level requires insights into transcriptional regulation, and thus the transcriptome, at the level of individual cell types. While the methods described here are generally applicable to a wide range of species and cell types, our research focuses on plant reproduction. Plant cultivation and seed production is of crucial importance for human and animal nutrition. A detailed understanding of the regulatory networks that govern the formation of the reproductive lineage (germline) and ultimately of seeds is a precondition for the targeted manipulation of plant reproduction. In particular, the engineering of apomixis (asexual reproduction through seeds) into crop plants promises great improvements, as it leads to the formation of clonal seeds that are genetically identical to the mother plant. Consequently, the cell types of the female germline are of major importance for the understanding and engineering of apomixis. However, as the corresponding cells are deeply embedded within the floral tissues, they are very difficult to access for experimental analyses, including cell-type specific transcriptomics. To overcome this limitation, sections of individual cells can be isolated by laser-assisted microdissection (LAM). While LAM in combination with transcriptional profiling allows the identification of genes and pathways active in any cell type with high specificity, establishing a suitable protocol can be challenging. Specifically, the quality of RNA obtained after LAM can be compromised, especially when small, single cells are targeted. To circumvent this problem, we have established a workflow for LAM that reproducibly results in high RNA quality that is well suitable for transcriptomics, as exemplified here by the isolation of cells of the female germline in apomictic Boechera. In this protocol, procedures are described for tissue preparation and LAM, also with regard to RNA extraction and quality control.

Laser-assisted Microdissection LAM as a Tool for Transcriptional Profiling of Individual Cell Types

Here we present a protocol for laser-assisted microdissection of specific plant cell types for transcriptional profiling. While the protocol is suitable for different species and cell types, the focus is on highly inaccessible cells of the female germline important for sexual and apomictic reproduction in the crucifer genus Boechera.

The understanding of developmental processes at the molecular level requires insights into transcriptional regulation, and thus the transcriptome, at the ...

Comparison of Tobacco Host Cell Protein Removal Methods by Blanching Intact Plants or by Heat Treatment of Extracts

Plants not only provide food, feed and raw materials for humans, but have also been developed as an economical production system for biopharmaceutical proteins, such as antibodies, vaccine candidates and enzymes. These must be purified from the plant biomass but chromatography steps are hindered by the high concentrations of host cell proteins (HCPs) in plant extracts. However, most HCPs irreversibly aggregate at temperatures above 60 &#176;C facilitating subsequent purification of the target protein. Here, three methods are presented to achieve the heat precipitation of tobacco HCPs in either intact leaves or extracts. The blanching of intact leaves can easily be incorporated into existing processes but may have a negative impact on subsequent filtration steps. The opposite is true for heat precipitation of leaf extracts in a stirred vessel, which can improve the performance of downstream operations albeit with major changes in process equipment design, such as homogenizer geometry. Finally, a heat exchanger setup is well characterized in terms of heat transfer conditions and easy to scale, but cleaning can be difficult and there may be a negative impact on filter capacity. The design-of-experiments approach can be used to identify the most relevant process parameters affecting HCP removal and product recovery. This facilitates the application of each method in other expression platforms and the identification of the most suitable method for a given purification strategy.

Three heat precipitation methods are presented that effectively remove more than 90% of host cell proteins (HCPs) from tobacco extracts prior to any other purification step. The plant HCPs irreversibly aggregate at temperatures above 60 &#176;C.

Plants not only provide food, feed and raw materials for humans, but have also been developed as an economical production system for biopharmaceutical ...