This work was supported by the CU Boulder Summer Underground Research Opportunities Program (UROP) grant (C.B.), the NSF Graduate Research Fellowship (L.S.), the Schmidt Science Fellowship (C.L.), the University of Colorado Research &#38; Innovation Seed Grant Program (2020 award to V.F., S.C., and K.C.), and the Anschutz Boulder Nexus Seed Grant at the University of Colorado (to V.F. and K.C.). Special acknowledgment goes to Dr. Tyler Tuttle for help with the loading chamber design as well as the Calve lab members for helpful discussions.

All animal experiments and procedures were performed according to protocol #2705, approved by the Animal Care and Use Committee of the University of Colorado Boulder. Six week old female C57BL/6J mice were used for the present study. The animals were obtained from a commercial source (see Table of Materials).
1. Animal preparation
<ol>
	<li>Euthanize the animal following the institutionally approved method. 
	NOTE: The present study used CO2 inhalation in alignment with the American Veterinary Medical Association&#39;s guidelines (a displacement rate of 30% to 70% of the chamber volume with CO2 per minute), followed by cervical dislocation, to ensure successful euthanization.
	<ol>
		<li>Work under a hood, if possible, to minimize the spread of mice allergens. Once the mouse stops moving and breathing, allow 2 min or more to verify the lack of response. 
		NOTE: If the mouse is pregnant or post-partum, the pups must be individually euthanized. Pups E15.5 and older must be decapitated during the dissection.</li>
	</ol>
	</li>
	<li>Prepare the dissection setup with a dissection pad, an 11-blade scalpel, curved thin sharp scissors, two pairs of forceps, curved forceps, 5-0 polyglactin suture, a dissecting microscope, and six pins (Figure 1, see Table of Materials).</li>
	<li>Place the mouse on the pad, and pin the forelimbs down (Figure 2A). Make an incision of approximately 1-1.5 cm in the abdomen with scissors (Figure 2B). Gently use the scissors to separate the skin at the cranial, caudal, and lateral sides of the incision (Figure 2C, D).</li>
	<li>Flip the mouse to its dorsal side, and gently peel back the skin toward the hindlimbs to remove the skin away from the dissection site (Figure 2E-H).</li>
	<li>Pin the mouse at the limbs (Figure 2I), and make an incision of approximately 1 cm into the abdomen from the thorax to the pelvis (Figure 2J). 
	NOTE: Ensure not to damage the underlying organs.</li>
	<li>Gently push the organs toward the thorax to clear the field of view (Figure 2K). 
	NOTE: Irrigate the tissues with 1x PBS to maintain hydration.</li>
	<li>Clear all of the fat tissue from the pelvic floor (Figure 2L-N). 
	&#8203;NOTE: Use forceps to gently pull and clear fat off the organs and tissues of interest.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65074/65074fig01.jpg" /> 
Figure 1: A clean workspace with all the tools needed to perform the dissections. <a href="https://www.jove.com/files/ftp_upload/65074/65074fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65074/65074fig02.jpg" /> 
Figure 2: Removal of the skin and opening of the pelvic and thoracic cavities of the mouse. (A) Pinning down all the limbs. (B) Initial incision. (C) Separating the skin from underlying fascia using scissors. (D) Cutting of the skin and preparing for removal. (E-G) Pulling the skin off by going around the mouse. (H) Completely removing the skin from the dorsal side. (I) Complete removal of the skin from the torso, and re-pinning of the mouse limbs. (J) Opening of the abdomen. (K) View of the open abdomen. (L) Moving the organs out of the field of view. (M) Removing the fat. (N) View of the cleared pelvic floor. <a href="https://www.jove.com/files/ftp_upload/65074/65074fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. USL harvesting
<ol>
	<li>Cut the uterine horns from the ovaries (Figure 3B), pull away from the field of view, and cut off at the cervical connection (Figure 3C). 
	NOTE: The uterine horns can be identified following the schematic in Figure 3A. Irrigate the tissues with 1x PBS to maintain hydration.</li>
	<li>Cut the ureters away from the bladder connection (Figure 3D). 
	NOTE: This is to avoid confusion with the USL.</li>
	<li>Cut the colon as close to the cervix as possible (Figure 3E,F). 
	NOTE: Irrigate the tissues with 1x PBS to maintain hydration.</li>
	<li>Place the mouse along with the dissection pad under the dissecting scope to visualize the USLs (Figure 3G).</li>
	<li>Gently use the forceps to clean the surrounding fat from the USLs. 
	NOTE: Use a second pair of forceps to hold the cervix up at a small angle to enhance the visualization of where the USL intersects with the cervix. Irrigate the tissues with 1x PBS to maintain hydration.</li>
	<li>Tie a 5-0 polyglactin suture around the cervical end of both USLs (Figure 4B, C). 
	NOTE: The USLs can be identified using the schematic and magnification images (Figure 4I-K)</li>
	<li>In this study, one USL is used for morphological or biochemical analyses (i.e., Raman microscopy, immunohistochemistry, histology). Cut the cervical end of the USL, leaving a piece of the cervix attached, and cut a piece of muscle from the bottom of the USL (Figure 4D). Place the dissected tissue in a bath with 1x PBS to keep the tissue hydrated (Figure 4G, H).</li>
	<li>Use the remaining USL for mechanical testing and imaging. Cut the cervical end of the USL, leaving a piece of cervix attached, to facilitate the mechanical setup (Figure 4D). 
	NOTE: The cervical tissue will act as an anchor to secure the USL during the mechanical test.</li>
	<li>Once all the tissues of interest are harvested (steps 3-5), dislocate the femurs from the pelvis (Figure 4E). 
	NOTE: One should hear a faint clicking sound when the femoral head is disarticulated from the acetabular cup.</li>
	<li>Cut the pelvic bone from the distal and proximal ends of the pelvic bone, leaving about 10 mm of total tissue (Figure 4F). Place the dissected tissue in 1x PBS.</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65074/65074fig03.jpg" /> 
Figure 3: Cleared pelvic floor for USL dissection. (A) Schematic of the anatomy. (B) Cutting the uterine horns at the ovarian connection. (C) Cutting off uterine horns. (D) Cutting of the ureters. (E) Cutting of the colon. (F) A clear view of the rectum and USLs. (G) Placing the mouse and dissection pad under the dissecting scope. <a href="https://www.jove.com/files/ftp_upload/65074/65074fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65074/65074fig04.jpg" /> 
Figure 4: View of the USL and surrounding tissues and dissection of the USLs. (A) Schematic of anatomical landmarks surrounding the USL. (B) Tying a suture around the cervical ends. (C) Slicing off the cervical ends of the USL. (D) Slicing off the USL to be used for the biochemical analyses at the sacral connection. (E) Cutting of the femurs from the pelvic bone. (F) Cutting off the proximal end of the pelvis. (G) Dissecting the USL in a 35 mm Petri dish. (H) The USL with the attached pelvis in a 35 mm petri dish. (I) The USL and rectum at 0.75x magnification. (J) Removing fat from the USL. (K) Cleaning of the USLs at 1.0x magnification. Scale bar = 2 mm. <a href="https://www.jove.com/files/ftp_upload/65074/65074fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Bladder harvesting
<ol>
	<li>After the fat is cleared, hold the bladder with the forceps, and gently lift it at an angle of approximately 40&#176; (Figure 5A).</li>
	<li>With the scissors, slice off the bladder from the distal side, right above the cervix (Figure 5B).</li>
	<li>Place the tissue in a bath with 1x PBS to keep the tissue hydrated (Figure 5H).</li>
</ol>
4. Rectum harvesting
<ol>
	<li>Once the USLs are disconnected from the cervix and the bladder is dissected, lift the cervix at an angle of approximately 40&#176; with the forceps. There is the rectovaginal fascia that connects the rectum and the cervix. With the scalpel, gently cut this connection (Figure 5C, D).</li>
	<li>Cut the pubic bone at the pubic symphysis using scissors. Gently widen the workspace to increase visual access to the tissue insertions.</li>
	<li>With the forceps, gently pull the rectum toward the thorax, and use the scissors to follow the rectum from its posterior side to the anus. Cut the rectum at the anus (Figure 5E).</li>
	<li>Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5I).</li>
</ol>
5. Cervix-vagina complex harvesting
<ol>
	<li>After the USLs are removed from the cervix, use the forceps to hold the cervix. Cut the cervix as close to the vulva as possible using scissors (Figure 5F, G). 
	NOTE: Ensure to cut the pubic symphysis to see the distal end of the vagina visually.</li>
	<li>Place the tissue in 1x PBS to keep the tissue hydrated (Figure 5J).</li>
</ol>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65074/65074fig05.jpg" /> 
Figure 5: Bladder, rectum, and cervix/vagina dissections. (A) Holding the bladder at an angle. (B) Cutting off the bladder. (C) Cutting the tendon connecting the cervix and rectum. (D) The tendon at 1.0x magnification. (E) Cutting the rectum. (F) Holding onto the cervix with forceps. (G) Cutting at the distal end of the vagina. (H) The bladder in a 35 mm Petri dish. (I) The rectum in a 35 mm Petri dish. (J) The cervix-vagina tissue complex in a 35 mm Petri dish. <a href="https://www.jove.com/files/ftp_upload/65074/65074fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Sample preparation for tissue characterization
<ol>
	<li>Mechanical and visual analyses of the USL
	<ol>
		<li>Place the USL with the pelvic attachment over a T-shaped wall within a custom staining well to ensure full immersion in the staining solution (CAD drawings of the well can be found in Supplementary Coding File 1 and Supplementary Coding File 2). 
		NOTE: Use suture and forceps to help with the placement.</li>
		<li>Dilute a commercially available dye that stains free amine groups (5 &#181;L, see Table of Materials) in 2.5 mL of 1x PBS, add the solution to the custom staining well, and stain the tissue for 2 h on a rocker at 4 &#176;C. 
		NOTE: Vortex the solution before adding it to the staining well.</li>
		<li>During the last 15 min of staining, add 2.5 &#181;L of a commercially available dead cell nuclei stain (see Table of Materials) to the solution. 
		NOTE: Vortex the solution prior to adding to the staining well.</li>
	</ol>
	</li>
	<li>Raman analysis of the USLs
	<ol>
		<li>Pin the USL in a straight line on a polydimethylsiloxane (PDMS) block that is contained in a custom well. 
		NOTE: The PDMS is used as a soft substrate to enable the pinning of the sample in the desired configuration. Blocks of varying dimensions can be made by mixing the two components following the manufacturer&#39;s instructions (see Table of Materials), casting in a Petri dish, and, after polymerization, cutting the PDMS into the required geometry with a scalpel blade.</li>
		<li>Pin with insect pins at the suture loop and at the pelvic muscle. Hydrate the tissue with 1x PBS.</li>
	</ol>
	</li>
	<li>Remaining tissues
	<ol>
		<li>Snap-freeze the remaining tissues with liquid nitrogen or in an appropriate embedding compound, depending on the desired analyses.</li>
		<li>Save the tissues at &#8722;80 &#176;C until subsequent analyses (e.g., immunohistochemical or biochemical assays).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

The effect of structural damage on female reproductive tissues is understudied, and an easily accessible animal model for POP research is needed. The mouse is a cost-effective model that can mimic human reproductive studies16. Due to the growing interest in the study of the female reproductive system, there is a need for methods that aid the study of these tissues. To address this need, in this work, a method is established to dissect and prepare murine pelvic floor tissues for structural and func...

Approximately 50% of women are affected by pelvic organ prolapse (POP)1,2. About 11% of these women fit the criteria for undergoing surgical repair, which has a poor success rate (~30%)3,4. POP is characterized by the descent of any or all of the pelvic organs (i.e., bladder, uterus, cervix, and rectum) from their natural position due to the failure of the USL and the pelvic floor muscles to provide adequate support5. This condition involves anatomical dysfunction and disruption of the connective tissue, as well as neuromuscular injury, in addition to predisposing factors3,6. POP is associated with multiple factors such as age, weight, parity, and delivery type (i.e., vaginal or caesarian births). These factors are thought to affect the mechanical integrity of all the pelvic floor tissues, with pregnancy and parity thought to be the main drivers of POP5,7,8. 
The uterosacral ligaments (USLs) are important supportive structures for the uterus, cervix, and vagina and tether the cervix to the sacrum4. Damage to the USLs puts women at increased risk of developing POP. It is believed that pregnancy and childbirth impose additional strain on the USL, which potentially induces injury and increases the chances of POP. The USL is a complex tissue composed of smooth muscle cells, blood vessels, and lymphatics distributed heterogeneously along the ligament, which can be divided into three distinct sections: cervical, intermediate, and sacral region9. The mechanical integrity of the USL is derived from extracellular matrix (ECM) components like collagens, elastin, and proteoglycans5,9,10. Type I collagen fibers are known to be a major load-bearing tensile component of ligamentous tissues and are, therefore, likely involved in USL failure and POP11. 
There is a lack of knowledge regarding the causes, prevalence, and effects of POP in women. The development of an appropriate animal model of POP is necessary to advance our understanding of the female pelvic floor. Mice and humans have similar anatomical landmarks within the pelvis, such as the ureters, rectum, bladder, ovaries, and round ligaments9, as well as similar intersection points of the USL with the uterus, cervix, and sacrum. Further, mice offer ease of genetic manipulation and have the potential to be an easily accessible, cost-effective model for the study of POP9. 
This study developed a method to access and isolate the USL and the different pelvic floor tissues from nulliparous (i.e., never pregnant) mice. The extracted USLs were subjected to enzymatic digestion (i.e., to remove collagens and glycosaminoglycans), tested to determine the mechanical response under tensile loading, and evaluated for biochemical composition in a proof-of-concept study. The ability to isolate intact tissues will facilitate further mechanical and biochemical characterizations of the pelvic floor components, which is a crucial first step toward improving our understanding of the injury risks related to childbirth, pregnancy, and POP.

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		<tr>
			<td>11 Blade</td>
			<td>Fisher</td>
			<td>3120030</td>
			<td>Removable blade</td>
		</tr>
		<tr>
			<td>1x PBS</td>
			<td>Fisher</td>
			<td>BP399-1</td>
			<td>Diluted from 10x concentration</td>
		</tr>
		<tr>
			<td>Chondroitinase ABC</td>
			<td>Sigma</td>
			<td>C3667-10UN</td>
			<td>Enzyme&#160;</td>
		</tr>
		<tr>
			<td>Collagenase Type I</td>
			<td>Worthington Biochemical</td>
			<td>LS004194</td>
			<td>Enzyme&#160;</td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Leica</td>
			<td>STELLARIS 5</td>
			<td>Upright configuration</td>
		</tr>
		<tr>
			<td>Dissection Microscope</td>
			<td>Leica</td>
			<td>S9E</td>
			<td>With camera</td>
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		<tr>
			<td>Dumont #5 Forceps</td>
			<td>Fisher</td>
			<td>NC9626652</td>
			<td>Thin tip</td>
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		<tr>
			<td>Female C57BL/6J mice</td>
			<td>Jackson Laboratory</td>
			<td>strain #: 000664</td>
			<td></td>
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		<tr>
			<td>FemtoTools Micromanipulator</td>
			<td>FemtoTools</td>
			<td>FT-RS1002</td>
			<td>100 mN load cell</td>
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		<tr>
			<td>FST Curved Forceps</td>
			<td>Fisher</td>
			<td>NC9639443</td>
			<td>Curved tip</td>
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		<tr>
			<td>FST Sharp 9 mm Scissors&#160;</td>
			<td>Fisher</td>
			<td>NC9639443</td>
			<td>Dissection scissors</td>
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		<tr>
			<td>Ghost Dye 780&#160;</td>
			<td>Tonbo</td>
			<td>13-0865-T500</td>
			<td>Free amine stain</td>
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		<tr>
			<td>Kimwipes</td>
			<td>Fisher</td>
			<td>06-666</td>
			<td>Box of 50 wipes</td>
		</tr>
		<tr>
			<td>OCT</td>
			<td>Tissue Tek</td>
			<td>4583</td>
			<td>Used for tissue preservation</td>
		</tr>
		<tr>
			<td>PDMS</td>
			<td>Thermo Fisher</td>
			<td>044764.AK</td>
			<td>Follow manufacturer&#39;s instructions</td>
		</tr>
		<tr>
			<td>Petri Dishes 35 mm</td>
			<td>Fisher</td>
			<td>FB0875711A</td>
			<td>Used for dissected tissue</td>
		</tr>
		<tr>
			<td>Polyglactin 5-0 Suture</td>
			<td>Veter.Sut</td>
			<td>VS385VL</td>
			<td>With needle</td>
		</tr>
		<tr>
			<td>Renishaw InVia Raman Microscope</td>
			<td>Renishaw</td>
			<td>PN192(EN)-02-A</td>
			<td>With confocal objectives</td>
		</tr>
		<tr>
			<td>Rocking Platform</td>
			<td>VWR</td>
			<td>10127-876</td>
			<td>2 tier platform</td>
		</tr>
		<tr>
			<td>Surgical Gloves</td>
			<td>Fisher</td>
			<td>52818</td>
			<td>For dissection&#160;</td>
		</tr>
		<tr>
			<td>Sytox</td>
			<td>Thermo Fisher</td>
			<td>S11381</td>
			<td>Nuclear stain&#160;</td>
		</tr>
		<tr>
			<td>T-pins</td>
			<td>Fisher</td>
			<td>S99385</td>
			<td>For dissection&#160;</td>
		</tr>
		<tr>
			<td>Transfer Pipets</td>
			<td>Fisher</td>
			<td>13-711-7M</td>
			<td>For dissection&#160;</td>
		</tr>
		<tr>
			<td>Underpads</td>
			<td>Fisher</td>
			<td>22037950</td>
			<td>To cover dissection pad</td>
		</tr>
	</tbody>
</table>

isolation and characterization of the murine uterosacral ligaments and pelvic floor organs

Pelvic organ prolapse (POP) is a condition that affects the integrity, structure, and mechanical support of the pelvic floor. The organs in the pelvic floor are supported by different anatomical structures, including muscles, ligaments, and pelvic fascia. The uterosacral ligament (USL) is a critical load-bearing structure, and injury to the USL results in a higher risk of developing POP. The present protocol describes the dissection of murine USLs and the pelvic floor organs alongside the acquisition of unique data on the USL biochemical composition and function using Raman spectroscopy and the evaluation of mechanical behavior. Mice are an invaluable model for preclinical research, but dissecting the murine USL is a difficult and intricate process. This procedure presents an approach to guide the dissection of murine pelvic floor tissues, including the USL, to enable multiple assessments and characterization. This work aims to aid the dissection of pelvic floor tissues by basic scientists and engineers, thus expanding the accessibility of research on the USL and pelvic floor conditions and the preclinical study of women's health using mouse models.

Each step of the dissection of a wild-type mouse is detailed in the associated video and figures related to the protocol. For this study, 6 week old female C57BL/6J mice were used (Supplementary Table 1). Three sample groups with USLs treated with different enzymes were analyzed: control (no treatment), collagenase-treated, and chondroitinase-treated groups. The smooth muscle, nerves, and lymphatics in the USL are surrounded by an ECM rich in fibrillar collagens and glycosaminoglycans (GAGs)<sup class="x...

Watch this Scientific Journal Video about Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs at JoVE.com

Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs

This article presents a detailed protocol for dissecting uterosacral ligaments and other pelvic floor tissues, including the cervix, rectum, and bladder in mice, to expand the study of female reproductive tissues.

Pelvic organ prolapse (POP) is a condition that affects the integrity, structure, and mechanical support of the pelvic floor. The organs in the pelvic ...

isolation-characterization-murine-uterosacral-ligaments-pelvic-floor

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Biology

1.1K Views.  University of Colorado Boulder. This article presents a detailed protocol for dissecting uterosacral ligaments and other pelvic floor tissues, including the cervix, rectum, and bladder in mice, to expand the study of female reproductive tissues.

Video: Isolation and Characterization of the Murine Uterosacral Ligaments and Pelvic Floor Organs

Isolation and Characterization of RNA-Containing Exosomes

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of endocytic origin were first proposed to function as a way for reticulocytes to eradicate the transferrin receptor while maturing into erythrocytes1, and were later named exosomes. Exosomes are formed by inward budding of late endosomes, producing multivesicular bodies (MVBs), and are released into the environment by fusion of the MVBs with the plasma membrane2. Since the first discovery of exosomes, a wide range of cells have been shown to release these vesicles. Exosomes have also been detected in several biological fluids, including plasma, nasal lavage fluid, saliva and breast milk3-6. Furthermore, it has been demonstrated that the content and function of exosomes depends on the originating cell and the conditions under which they are produced. A variety of functions have been demonstrated for exosomes, such as induction of tolerance against allergen7,8, eradication of established tumors in mice9, inhibition and activation of natural killer cells10-12, promotion of differentiation into T regulatory cells13, stimulation of T cell proliferation14 and induction of T cell apoptosis15. Year 2007 we demonstrated that exosomes released from mast cells contain messenger RNA (mRNA) and microRNA (miRNA), and that the RNA can be shuttled from one cell to another via exosomes. In the recipient cells, the mRNA shuttled by exosomes was shown to be translated into protein, suggesting a regulatory function of the transferred RNA16. Further, we have also shown that exosomes derived from cells grown under oxidative stress can induce tolerance against further stress in recipient cells and thus suggest a biological function of the exosomal shuttle RNA17. Cell culture media and biological fluids contain a mixture of vesicles and shed fragments. A high quality isolation method for exosomes, followed by characterization and identification of the exosomes and their content, is therefore crucial to distinguish exosomes from other vesicles and particles. Here, we present a method for the isolation of exosomes from both cell culture medium and body fluids. This isolation method is based on repeated centrifugation and filtration steps, followed by a final ultracentrifugation step in which the exosomes are pelleted. Important methods to identify the exosomes and characterize the exosomal morphology and protein content are highlighted, including electron microscopy, flow cytometry and Western blot. The purification of the total exosomal RNA is based on spin column chromatography and the exosomal RNA yield and size distribution is analyzed using a Bioanalyzer.

This paper demonstrates methods for the isolation, purification and detection of exosomes, as well as techniques for analysis of their molecular content. These methods are adaptable for exosome isolation from both cell culture media and biological fluids, and can beyond analysis of molecular content also be useful in functional studies.

The field of exosome research is rapidly expanding, with a dramatic increase in publications in recent years. These small vesicles (30-100 nm) of ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of ...

Isolation of Murine Valve Endothelial Cells

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and surrounded by a monolayer of valve endothelial cells (VECs). VECs play essential roles in establishing the valve structures during embryonic development, and are important for maintaining life-long valve integrity and function. In contrast to a continuous endothelium over the surface of healthy valve leaflets, VEC disruption is commonly observed in malfunctioning valves and is associated with pathological processes that promote valve disease and dysfunction. Despite the clinical relevance, focused studies determining the contribution of VECs to development and disease processes are limited. The isolation of VECs from animal models would allow for cell-specific experimentation. VECs have been isolated from large animal adult models but due to their small population size, fragileness, and lack of specific markers, no reports of VEC isolations in embryos or adult small animal models have been reported. Here we describe a novel method that allows for the direct isolation of VECs from mice at embryonic and adult stages. Utilizing the Tie2-GFP reporter model that labels all endothelial cells with Green Fluorescent Protein (GFP), we have been successful in isolating GFP-positive (and negative) cells from the semilunar and atrioventricular valve regions using fluorescence activated cell sorting (FACS). Isolated GFP-positive VECs are enriched for endothelial markers, including CD31 and von Willebrand Factor (vWF), and retain endothelial cell expression when cultured; while, GFP-negative cells exhibit molecular profiles and cell shapes consistent with VIC phenotypes. The ability to isolate embryonic and adult murine VECs allows for previously unattainable molecular and functional studies to be carried out on a specific valve cell population, which will greatly improve our understanding of valve development and disease mechanisms.

The ability to isolate heart valve endothelial cells (VECs) is critical for understanding mechanisms of valve development, maintenance, and disease. Here we describe the isolation of VECs from embryonic and adult Tie2-GFP mice using FACS that will allow for studies determining the contribution of VECs in developmental and disease processes.

Normal valve structures consist of stratified layers of specialized extracellular matrix (ECM) interspersed with valve interstitial cells (VICs) and ...

Isolation of Murine Coronary Vascular Smooth Muscle Cells

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs from the coronary circulation. Hearts with intact aortic arches were removed and perfused via retrograde Langendorff with digestion solution containing 300 Units/ml of collagenase type II, 0.1 mg/ml soybean trypsin inhibitor and 1 M CaCl2. The perfusates were collected at 15 min intervals for 90 min, pelleted by centrifugation, resuspended in plating media, and plated on tissue culture dishes. VSMCs were characterized by presence of SM22&#945;, &#945;-SMA, and vimentin. One of the main advantages of using this technique is the ability to isolate VSMCs from the coronary circulation of mice. Although the small number of cells obtained can limit some of the applications for which the cells can be utilized, isolated coronary VSMCs can be used in a variety of well-established cell culture techniques and assays. Studies investigating VSMCs from genetically modified mice can provide further information about structure-function and signaling processes associated with vascular pathologies.

The purpose of this protocol is to demonstrate the isolation and culture techniques of murine primary vascular smooth muscle cells (VSMCs) from the coronary circulation. Once VSMCs have been isolated, they can be used for many standard culture techniques.

While the isolation and culture of vascular smooth muscle cells (VSMCs) from large vessels is well established, we sought to isolate and culture VSMCs ...

Isolation and Characterization of Microvesicles from Peripheral Blood

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived microvesicles (MVs, diameter &#62; 100 nm) is a fundamental cellular process that occurs in all living cells. These vesicles transport proteins, lipids and nucleic acids specific for their cell of origin and in vitro studies have highlighted their importance as mediators of intercellular communication. EVs have been successfully isolated from various body fluids and especially EVs in blood have been identified as promising biomarkers for cancer or infectious diseases. In order to allow the study of MV subpopulations in blood, we present a protocol for the standardized isolation and characterization of MVs from peripheral blood samples. MVs are pelleted from EDTA-anticoagulated plasma samples by differential centrifugation and typically possess a diameter of 100 - 600 nm. Due to their larger size, they can easily be studied by flow cytometry, a technique that is routinely used in clinical diagnostics and available in most laboratories. Several examples for quality control assays of the isolated MVs will be given and markers that can be used for the discrimination of different MV subpopulations in blood will be presented.

Extracellular vesicles present in blood have been suggested as novel biomarkers for various diseases. Here, we present a protocol for the isolation of large plasma membrane-derived microvesicles from peripheral blood samples and their subsequent analysis by conventional flow cytometry and Western Blotting.

The release of extracellular vesicles (EVs) including small endosomal-derived exosomes (Exos, diameter &#60; 100 nm) and large plasma membrane-derived ...

Isolation of Myoepithelial Cells from Adult Murine Lacrimal and Submandibular Glands

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ductal epithelial, and myoepithelial cells (MECs). MECs express alpha smooth muscle actin (&#945;SMA) and have a contractile function. They are found in multiple glandular organs and are of ectodermal origin. In addition, the LG contains SMA+ vascular smooth muscle cells of endodermal origin called pericytes: contractile cells that envelop the surface of vascular tubes. A new protocol allows us to isolate both MECs and pericytes from adult murine LGs and submandibular glands (SMGs). The protocol is based on the genetic labeling of MECs and pericytes using the SMACreErt2/+:Rosa26-TdTomatofl/fl mouse strain, followed by preparation of the LG single-cell suspension for fluorescence activated cell sorting (FACS). The protocol allows for the separation of these two cell populations of different origins based on the expression of the epithelial cell adhesion molecule (EpCAM) by MECs, whereas pericytes do not express EpCAM. Isolated cells could be used for cell cultivation or gene expression analysis.

The lacrimal gland (LG) has two cell types expressing &#945;-smooth muscle actin (&#945;SMA): myoepithelial cells (MECs) and pericytes. MECs are of ectodermal origin, found in many glandular tissues, while pericytes are vascular smooth muscle cells of endodermal origin. This protocol isolates MECs and pericytes from murine LGs.

The lacrimal gland (LG) is an exocrine tubuloacinar gland that secretes an aqueous layer of tear film. The LG epithelial tree is comprised of acinar, ...

Isolation and Characterization of Adult Cardiac Fibroblasts and Myofibroblasts

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that mitigate fibrosis. However, fibroblast research has been hindered due to the lack of universally acceptable fibroblast markers to identify quiescent as well as activated fibroblasts. Fibroblasts are a heterogenous cell population, making them difficult to isolate and characterize. The presented protocol describes three different methods to enrich fibroblasts and myofibroblasts from uninjured and injured mouse hearts. Using a standard and reliable protocol to isolate fibroblasts will enable the study of their roles in homeostasis as well as fibrosis modulation.

Obtaining a pure population of fibroblasts is crucial to studying their role in wound repair and fibrosis. Described here is a detailed method to isolate fibroblasts and myofibroblasts from uninjured and injured mouse hearts followed by characterization of their purity and functionality by immunofluorescence, RTPCR, fluorescence-assisted cell sorting, and collagen gel contraction.

Cardiac fibrosis in response to injury is a physiological response to wound healing. Efforts have been made to study and target fibroblast subtypes that ...

Isolation and Characterization of Intact Phycobilisome in Cyanobacteria

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. Studying the structure and composition of phycobilisome is of great interest to scientists because it reveals the evolution and divergence of photosynthesis in cyanobacteria. This protocol provides a detailed and optimized method to break cyanobacterial cells at low cost by a bead-beater efficiently. The intact phycobilisome can then be isolated from the cell extract by sucrose gradient ultracentrifugation. This method has demonstrated being suitable for both model and non-model cyanobacteria with different cell types. A step-by-step procedure is also provided to confirm the integrity and property of phycobiliproteins by 77K fluorescence spectroscopy and SDS-PAGE stained by zinc sulfate and Coomassie Blue. The isolated phycobilisome can also be subjected to further structural and compositional analyses. Overall, this protocol provides a helpful starting guide that allows researchers unfamiliar with cyanobacteria to quickly isolate and characterize intact phycobilisome.

The current protocol details the isolation of phycobilisomes from cyanobacteria by centrifugation through a discontinuous sucrose density gradient. The fractions of intact phycobilisomes are confirmed by 77K fluorescent emission spectrum and SDS-PAGE analysis. The resulting phycobilisome fractions are suitable for negative staining of TEM and mass spectrometry analysis.

In cyanobacteria, phycobilisome is a vital antenna protein complex that harvests light and transfers energy to photosystem I and II for photochemistry. ...

Isolation and Characterization of Cyanobacterial Extracellular Vesicles

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential biotechnology models. Recent work has demonstrated that both marine and freshwater cyanobacteria produce extracellular vesicles -&#160;small membrane-bound structures released from the outer surface of the microbes. While vesicles likely contribute to diverse biological processes, their specific functional roles in cyanobacterial biology remain largely unknown. To encourage and advance research in this area, a detailed protocol is presented for isolating, concentrating, and purifying cyanobacterial extracellular vesicles. The current work discusses methodologies that have successfully isolated vesicles from large cultures of Prochlorococcus, Synechococcus, and Synechocystis. Methods for quantifying and characterizing vesicle samples from these strains are presented. Approaches for isolating vesicles from aquatic field samples are also described. Finally, typical challenges encountered with cyanobacterial vesicle purification, methodological considerations for different downstream applications, and the trade-offs between approaches are also discussed.

The present protocol providesdetailed descriptions for isolation, concentration,and characterization of extracellular vesicles from cyanobacterial cultures. Approaches for purifying vesicles from cultures at different scales, trade-offs among methodologies, and considerations for working with field samples are also discussed.

Cyanobacteria are a diverse group of photosynthetic, Gram-negative bacteria that play critical roles in global ecosystems and serve as essential ...

Isolation and Characterization of the Immune Cells from Micro-dissected Mouse Choroid Plexuses

The brain is no longer considered as an organ functioning in isolation; accumulating evidence suggests that changes in the peripheral immune system can indirectly shape brain function. At the interface between the brain and the systemic circulation, the choroid plexuses (CP), which constitute the blood-cerebrospinal fluid barrier, have been highlighted as a key site of periphery-to-brain communication. CP produce the cerebrospinal fluid, neurotrophic factors, and signaling molecules that can shape brain homeostasis. CP are also an active immunological niche. In contrast to the brain parenchyma, which is populated mainly by microglia under physiological conditions, the heterogeneity of CP immune cells recapitulates the diversity found in other peripheral organs. The CP immune cell diversity and activity change with aging, stress, and disease and modulate the activity of the CP epithelium, thereby indirectly shaping brain function. The goal of this protocol is to isolate murine CP and identify about 90% of the main immune subsets that populate them. This method is a tool to characterize CP immune cells and understand their function in orchestrating periphery-to-brain communication. The proposed protocol may help decipher how CP immune cells indirectly modulate brain function in health and across various disease conditions.

This study uses flow cytometry and two different gating strategies on isolated perfused mice brain choroid plexuses; this protocol identifies the main immune cell subsets that populate this brain structure.

The brain is no longer considered as an organ functioning in isolation; accumulating evidence suggests that changes in the peripheral immune system can ...