This work was supported by grants from the National Heart, Lung, and Blood Institute of the National Institutes of Health (NIH) (1R01HL160740 and 5 T32GM132054-04) and the Florida Heart Research Foundation.&#160;We would like to thank Kassandra Gomez for her help synthesizing and imaging the hydrogels.

The in vivo work was approved and overseen by the Institutional Animal Care and Use Committee (IACUC) at Florida International University and conformed to current National Institutes of Health (NIH) guidelines. For this protocol, the procedure does not differ depending on the strain, weight, age, and sex of the mouse. The type of calcification being studied, diets, and treatments may alter the length of the study and the weight of the mouse used and may be dependent on a specific strain and sex of mouse used in the study. For this protocol, both male and female C57BL/6J mice were used, and they were fed a calcification diet. The mice were sacrificed between 20 weeks and 24 weeks of age.
1. Isolation and excision of the aorta
<ol>
	<li>Fluorescent markers
	<ol>
		<li>Forty-eight hours before euthanasia, inject the mice with 20 &#181;M OsteoSense 680, a fluorescent imaging agent, at the recommended dose of 100 &#181;L per 25 g via an intravenous injection.</li>
		<li>Under the fume hood, place a Styrofoam board with four t-pins, a bucket lined with a paper towel, a 50 mL tube stuffed with a paper towel, a 1.5 mL microcentrifuge tube, dissection tools (see the Table of Materials), and ethanol in a spray bottle. Place beakers with PBS on ice.</li>
	</ol>
	</li>
	<li>Blood draw
	<ol>
		<li>Add 2 mL of isoflurane into a 50 mL conical tube stuffed with the paper towel. Place the tube in the bucket, and close it with its lid. 
		CAUTION: Isoflurane should be handled in a well-ventilated space with no recirculation of exhaust air due to the toxicity of short-term exposure.</li>
		<li>Transfer one mouse into the bucket at a time. Once sedated, place the mouse on the paper towel in the dorsal position. Holding the mouse with the palm of the non-dominant hand, place the ears back using the thumb and pointer finger.</li>
		<li>Using the tweezers, excise one of the eyes. Hold the mouse above the 1.5 mL tube to begin blood collection, during which 1 mL of blood should be collected. Once the blood stops draining, place the mouse back in the bucket for further sedation.</li>
	</ol>
	</li>
	<li>Euthanasia
	<ol>
		<li>Place a paper towel on the dissection mat. Once the mouse is sedated, place the mouse on the mat in a supine position. Secure the mouse in the supine position using a t-pin to hold down each limb.</li>
		<li>Spray down the abdomen of the mouse with alcohol. Using the tweezers, lift the skin at the chest region. With the scissors, cut down the medial line into the thoracic cavity. Once the sternum has been cut down the center, cut the diaphragm on either side of the ribcage.</li>
		<li>If necessary, cut away the frontal portion of the ribcage to expose the heart and lungs.</li>
	</ol>
	</li>
	<li>Organ removal
	<ol>
		<li>Using tweezers, lift the skin and make an incision down the midline. Remove any excess skin or fat. Remove the reproductive organs and bladder, as well as any fat outside of the midline.</li>
		<li>Move the gastrointestinal (GI) organs to the right-hand side, and follow the intestines in the midline. Once found, lift the midline, and excise the GI tract up to the stomach.</li>
		<li>Excise the kidneys by lifting them away from the midline. Cut as close to the kidney as possible.</li>
		<li>Next, begin the removal of the liver by lifting the lobes and cutting away from the midline.</li>
		<li>Perfuse the heart and aorta. Using a 10 mL syringe, inject cold PBS into the right ventricle. Wait for the lungs to inflate and turn white. Remove the lungs, as well as any excess diaphragm or ribcage.</li>
	</ol>
	</li>
	<li>Bone removal
	<ol>
		<li>Open a pair of scissors wide, and place them at the lower region of the mouse above the tail. Cut downward, and begin lifting the spine from the skin. Remove the fat from the sides of the midline. Once the skin is detached, excise the spine and intact organs by cutting right above the heart. If transporting to a stereoscope, place the spine and intact organs in a beaker of PBS in the ice bucket.</li>
		<li>Move the spine and organs into the dissecting dish. Fill with ice-cold PBS. Pin down the spine and ribcage using needles.</li>
		<li>Under the dissection microscope, use tweezers to lift the heart carefully, and begin cutting above the spine and underneath the aorta. Continue this until the bottom of the spine is reached. Remove the spine from the dissecting dish, and pin down the heart and any extraneous fat or muscle.</li>
	</ol>
	</li>
	<li>Microdissection
	<ol>
		<li>Starting from the area right below the heart, identify the three tubular structures: the esophagus, vena cava, and aorta. Remove the esophagus and vena cava to have a clear visual field of the aorta.</li>
		<li>Using the microdissection forceps and scissors, begin lifting the adipose tissue and cutting as close to the aorta as possible. Be sure not to cut the aorta. Continue this down the medial line until all the aorta as well as the femoral arteries are exposed.</li>
		<li>At the heart, remove all the adipose tissue, exposing the three branches at the aortic arch. Remove the aortic root from the left ventricle by inserting the microdissection scissors into the left ventricle and cutting the muscle surrounding the aorta.</li>
		<li>Once the aorta is isolated and cleaned, image the aorta using a near-infrared scanner to visualize vascular calcification.</li>
		<li>Use the custom MATLAB script (Supplemental File 1) to quantify the total signal of the calcium tracer, which is normalized to the total scanned aorta area. The MATLAB script functions using the Bioinformatics toolbox add-in. MATLAB is first directed to the location of the TIFF files from the near-infrared scan of the aortas, the individual files are then opened, and the pixel intensity values are extracted from the TIFF files.
		<ol>
			<li>Once the image is presented to the user with a color map, select the aorta with the greatest calcification as the scale maximum.</li>
			<li>When MATLAB asks &#34;How many specimens are in the image?&#34; in the command window, input the number of aortas in the current image, and select each aorta one by one. Once an aorta is selected, MATLAB will create a binary image with a mask of the total area of the aorta and another binary image with a mask of the calcified area of the aorta. The values from these masked images are then used to determine the total calcified area, the total area of the aorta, the percentage area positive for calcification, and the mean intensity of the calcified area.</li>
			<li>To select an aorta, make a box around the aorta of interest on the most recent figure on the screen by selecting four corners with one left click and creating a box around the image. To close the box, double-click on the first corner. 
			&#8203;NOTE: The threshold may be different depending on the model of calcification used. It is up to the user to determine what value determines a positive signal for vascular calcification. Once the threshold is set for an experiment, it must remain the same throughout the duration of the experiment.</li>
		</ol>
		</li>
		<li>Once the quantitative data are extracted, perform a one-way ANOVA with Tukey&#39;s post-hoc test to show significance between the groups.</li>
	</ol>
	</li>
</ol>
2. Isolating and extracting EVs from aortas
NOTE: Once the aortas have been isolated and removed, EVs can be extracted from the tissue. Using a digestive solution and multiple centrifuge cycles, EVs can be collected and used for many different techniques, including calcification assays, gel electrophoresis, and immunoblotting13. The protocol for isolating and extracting EVs is as follows:
<ol>
	<li>Immediately after scanning the aortas, pool two to three aortas to yield a sufficient protein concentration.</li>
	<li>Prepare a digestive solution containing 0.25 M sucrose, 0.12 M sodium chloride (NaCl), 0.01 M potassium chloride (KCl), 0.02 M Tris hydrochloride, and 600 U/mL collagenase13.</li>
	<li>Incubate two to three pooled aortas in 1.5 mL of digestive solution for 2 h at 37 &#176;C. Collect the solution. Centrifuge the solution at 1,000 &#215; g for 15 min to remove cell debris.</li>
	<li>Centrifuge the supernatant for an additional 30 min at 33,000 &#215; g to remove microvesicles. Collect and assess the supernatant for calcification potential (see section 3). 
	NOTE: Follow the subsequent steps if gel electrophoresis and protein immunoblotting of the EVs are to be performed.</li>
	<li>Ultracentrifuge the supernatant at 100,000 &#215; g for 1 h to isolate the EVs of interest.</li>
	<li>While the supernatant is undergoing ultracentrifugation, prepare RIPA lysis and extraction buffer (see the Table of Materials) by adding the protease inhibitor and vortexing until dissolved.</li>
	<li>Once the ultracentrifugation is complete, aspirate the supernatant, leaving the pellet, which contains the EVs. Suspend the pellet in the prepared RIPA lysis and protease inhibitor mixture. The samples are ready for gel electrophoresis and western blotting.</li>
</ol>
3. Assessing the calcification potential of extracellular vesicles with light scattering absorbance
NOTE: To measure the real-time mineral formation of EVs, we used an assay originally developed to study mineral formation from growth plate cartilage EVs from cell culture14. Since calcium phosphate deposition is the hallmark of calcification, an increase in light scattering absorbance as the result of calcium phosphate compound formation is indicative of the calcification potential14. In vitro VSMC models are convenient for measuring the calcification potential of EVs. In this technique, a plate reader with a short wavelength filter can quantify the in vitro calcification of EVs. The absorbency reading is recorded at 340 nm, and a higher absorbance is indicative of more calcium phosphate mineral formation. The protocol for the light scattering absorbance assay is as follows:
<ol>
	<li>Centrifuge the conditioned collected medium of VSMCs at 1,000 &#215; g for 5 min to remove cell debris14.</li>
	<li>Centrifuge the supernatant at 33,000 &#215; g for 30 min, collect the supernatant, and transfer it into a new tube. 
	NOTE: The presence of EVs in the collected supernatant can be confirmed through dynamic light scattering (DLS) or nanoparticle tracking analysis (NTA)</li>
	<li>Prepare a 300 mM sodium phosphate monobasic (NaH2PO4) sterile solution. Filter the solution to ensure sterile conditions if necessary.</li>
	<li>Add 1% (v/v) of 300 mM NaH2PO4 to the EV samples, and gently pipette the solution to mix well. Transfer 200 &#181;L of the mixture solution to a 96-well plate. Incubate the 96-well plate in the microplate reader at 37 &#176;C.</li>
	<li>Set the plate reader to record the absorbance at a wavelength of 340 nm every 1 min.</li>
</ol>
4. Assessing the calcification mineral formation of extracellular vesicles with collagen hydrogels
NOTE: The aggregation of EVs and the formation of microcalcifications are observed through collagen hydrogels. These hydrogels act as a scaffold that mimics the collagen density observed in vivo15. This demonstrates the effect of collagen on calcification growth. The protocol for assessing the mineral formation of EVs in hydrogels is as follows:
<ol>
	<li>Day 1
	<ol>
		<li>Prepare a 2.5 mg/mL collagen solution in DMEM. Slowly add 5 M sodium hydroxide to the collagen/DMEM mixture until the color changes to red. Check that the pH of the solution is approximately 7. 
		NOTE: If the solution turns purple and becomes too basic, restart the process.</li>
		<li>Pipette 300 &#181;L of the 2.5 mg/mL collagen solution with a pH of 7 into each well of an 8-well chamber glass. Incubate at 37 &#176;C and 5% CO2 controlled humidity for 72 h.</li>
	</ol>
	</li>
	<li>Day 4
	<ol>
		<li>Add 300 &#181;L of DMEM as a control into the wells.</li>
		<li>Add 300 &#181;L of the EV medium samples collected previously to the remaining wells.</li>
		<li>Incubate at 37 &#176;C and 5% CO2 controlled humidity for 6 days.</li>
	</ol>
	</li>
	<li>Day 10
	<ol>
		<li>Prepare 2.5 &#181;L of Osteosense 680EX with 22.5 &#181;L of DMEM.</li>
		<li>Pipette 2.5 &#181;L of the Osteosense and DMEM mixture into each well. Incubate at 37 &#176;C and 5% CO2 controlled humidity for 24 h.</li>
	</ol>
	</li>
	<li>Day 11
	<ol>
		<li>Image the hydrogels with filters appropriate for Osteosense fluorescence. Image them through the bottom of a coverglass chamber using an inverted microscope or from the top with a long working distance objective.</li>
		<li>Assess the number of calcifications and the calcification size in the collected images using the Particle Analysis options available in ImageJ.
		<ol>
			<li>Use the Image | Adjust | Threshold command to binarize the images such that only the Osteosense signal appears white. Use the Analyze | Analyze particles command to obtain information for each calcification in the image. Be sure to use the same threshold parameters for each image analyzed for consistency.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

<ol><li>Bakhshian Nik, A., Hutcheson, J. D., Aikawa, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Extracellular+vesicles+as+mediators+of+cardiovascular+calcification%5BTitle%5D)+AND+%22Frontiers+in+Cardiovascular+Medicine%22%5BJournal%5D)">Extracellular vesicles as mediators of cardiovascular calcification.</a> Frontiers in Cardiovascular Medicine. 4, 78(2017).</li>
<li>Ho, C. Y., Shanahan, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Medial+arterial+calcification%3A+an+overlooked+player+in+peripheral+arterial+disease%5BTitle%5D)+AND+%22Arteriosclerosis%2C+Thrombosis%2C+and+Vascular+Biology%22%5BJournal%5D)">Medial arterial calcification: an overlooked player in peripheral arterial disease.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 36 (8), 1475-1482 (2016).</li>
<li>Marinelli, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Diagnosis+of+arterial+media+calcification+in+chronic+kidney+disease%5BTitle%5D)+AND+%22Cardiorenal+Medicine%22%5BJournal%5D)">Diagnosis of arterial media calcification in chronic kidney disease.</a> Cardiorenal Medicine. 3 (2), 89-95 (2013).</li>
<li>Moe, S. M., Chen, N. X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mechanisms+of+vascular+calcification+in+chronic+kidney+disease%5BTitle%5D)+AND+%22Journal+of+the+American+Society+of+Nephrology%22%5BJournal%5D)">Mechanisms of vascular calcification in chronic kidney disease.</a> Journal of the American Society of Nephrology. 19 (2), 213-216 (2008).</li>
<li>Mir, B., Goettsch, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Extracellular+vesicles+as+delivery+vehicles+of+specific+cellular+cargo%5BTitle%5D)+AND+%22Cells%22%5BJournal%5D)">Extracellular vesicles as delivery vehicles of specific cellular cargo.</a> Cells. 9 (7), 1601(2020).</li>
<li>Ruiz, J. L., Hutcheson, J. D., Aikawa, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Cardiovascular+calcification%3A+Current+controversies+and+novel+concepts%5BTitle%5D)+AND+%22Cardiovascular+Pathology%22%5BJournal%5D)">Cardiovascular calcification: Current controversies and novel concepts.</a> Cardiovascular Pathology. 24 (4), 207-212 (2015).</li>
<li>New, S. E., Aikawa, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Role+of+extracellular+vesicles+in+de+novo+mineralization%3A+An+additional+novel+mechanism+of+cardiovascular+calcification%5BTitle%5D)+AND+%22Arteriosclerosis%2C+Thrombosis%2C+and+Vascular+Biology%22%5BJournal%5D)">Role of extracellular vesicles in de novo mineralization: An additional novel mechanism of cardiovascular calcification.</a> Arteriosclerosis, Thrombosis, and Vascular Biology. 33 (8), 1753-1758 (2013).</li>
<li>Aikawa, E., Hutcheson, J. D. <a target="_blank" href="http://scholar.google.com/scholar?hl=en&safe=off&q=author%3aAikawa%2C+E+%22Cardiovascular+Calcification+and+Bone+Mineralization%22">Cardiovascular Calcification and Bone Mineralization</a>. , Humana Press. Cham, Switzerland. (2020).</li>
<li>Johnson, T. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+P-407-induced+murine+model+of+dose-controlled+hyperlipidemia+and+atherosclerosis%3A+A+review+of+findings+to+date%5BTitle%5D)+AND+%22Journal+of+Cardiovascular+Pharmacology%22%5BJournal%5D)">The P-407-induced murine model of dose-controlled hyperlipidemia and atherosclerosis: A review of findings to date.</a> Journal of Cardiovascular Pharmacology. 43 (4), 595-606 (2004).</li>
<li>Shobeiri, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Vascular+calcification+in+animal+models+of+CKD%3A+A+review%5BTitle%5D)+AND+%22American+Journal+of+Nephrology%22%5BJournal%5D)">Vascular calcification in animal models of CKD: A review.</a> American Journal of Nephrology. 31 (6), 471-481 (2010).</li>
<li>El-Abbadi, M. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Phosphate+feeding+induces+arterial+medial+calcification+in+uremic+mice%3A+role+of+serum+phosphorus%2C+fibroblast+growth+factor-23%2C+and+osteopontin%5BTitle%5D)+AND+%22Kidney+International%22%5BJournal%5D)">Phosphate feeding induces arterial medial calcification in uremic mice: role of serum phosphorus, fibroblast growth factor-23, and osteopontin.</a> Kidney International. 75 (12), 1297-1307 (2009).</li>
<li>Veseli, B. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Animal+models+of+atherosclerosis%5BTitle%5D)+AND+%22European+Journal+of+Pharmacology%22%5BJournal%5D)">Animal models of atherosclerosis.</a> European Journal of Pharmacology. 816, 3-13 (2017).</li>
<li>Chen, N. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transglutaminase+2+accelerates+vascular+calcification+in+chronic+kidney+disease%5BTitle%5D)+AND+%22American+Journal+of+Nephrology%22%5BJournal%5D)">Transglutaminase 2 accelerates vascular calcification in chronic kidney disease.</a> American Journal of Nephrology. 37 (3), 191-198 (2013).</li>
<li>Wu, L. N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Physicochemical+characterization+of+the+nucleational+core+of+matrix+vesicles%5BTitle%5D)+AND+%22Journal+of+Biological+Chemistry%22%5BJournal%5D)">Physicochemical characterization of the nucleational core of matrix vesicles.</a> Journal of Biological Chemistry. 272 (7), 4404-4411 (1997).</li>
<li>Hutcheson, J. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Genesis+and+growth+of+extracellular-vesicle-derived+microcalcification+in+atherosclerotic+plaques%5BTitle%5D)+AND+%22Nature+Materials%22%5BJournal%5D)">Genesis and growth of extracellular-vesicle-derived microcalcification in atherosclerotic plaques.</a> Nature Materials. 15 (3), 335-343 (2016).</li>
</ol>

The authors have no conflicts of interest to disclose.

When performing the protocol, it is important to note the critical steps for obtaining successful results. During the isolation of the murine aorta, it is vital that the perfusion is performed properly. When injecting the PBS, care must be taken not to puncture the right ventricle. This would cause the liquid to leak directly out of the ventricle and fail to circulate through the lungs, leaving blood within the aorta. Once the perfusion has been conducted properly and microdissection has begun, all the adipose and fatty ...

Calcification is the most significant predictor of cardiovascular disease mortality and morbidity1. Calcification alters the arterial wall mechanics due to the buildup of calcium and phosphate minerals2. In atherosclerosis, calcification can exacerbate local stress and result in plaque rupture, which is the leading cause of heart attacks. Medial calcification-often resulting from chronic kidney disease-is more widespread and leads to significant arterial stiffening, dysfunction, and cardiac overload2,3. Currently, there are no therapeutic options for the treatment or prevention of vascular calcification.
Vascular smooth muscle cells (VSMCs) adopt an osteoblast-like phenotype and release calcifying extracellular vesicles (EVs) that nucleate nascent minerals, thus driving calcification4,5,6. This process resembles the physiological mineralization of osteoblasts in bone7. Although the endpoint of mineralization is similar in the vascular wall and bone matrix, the mechanisms by which the calcifying EVs originate differ in the two tissues8. There are many types of models that are used to study vascular calcification. In vitro, cell culture models mimic the osteogenic transition of VSMCs and subsequent mineral formation with specialized media.
When studying in vivo calcification, the model used depends upon the type of calcification being studied. Hyperlipidemic mouse models are often used to study atherosclerotic calcification, which appears more focal within lipid-rich plaques9. In contrast, medial calcification is more widespread throughout the vasculature and is often studied using chronic kidney disease models that employ an adenine-rich diet regimen to induce renal failure or surgical techniques to remove significant portions of the kidneys10,11. Aggressive models of vascular calcification have used a combination of both hyperlipidemic and chronic kidney disease models12. This protocol provides a method for assessing vascular calcification in murine aortas for both medial and atherosclerotic calcification, extracting EVs from the aortic wall, and observing the mineralization potential in the EVs obtained from in vitro cell culture models. Future studies can use these procedures in mechanistic analyses of vascular calcification and to assess potential therapeutic interventions.

<table><tbody><tr><td>8-well chambered coverglass</td><td>Thermo Scientific</td><td>155409PK</td><td></td></tr><tr><td>10 mL Syringe</td><td>BD</td><td>302995</td><td></td></tr><tr><td>20 G 1 inch Needle</td><td>BD</td><td>305175</td><td></td></tr><tr><td>Collagen, High Concentraion, Rat Tail</td><td>Corning</td><td>354249</td><td></td></tr><tr><td>Collagenase</td><td>Worthington Biochemical</td><td>LS004174</td><td></td></tr><tr><td>Curved Forceps</td><td>Roboz Surgical Instrument</td><td>RS-8254</td><td></td></tr><tr><td>Dissection Dish</td><td>Living Systems Instrumentation</td><td>DD-90-S</td><td></td></tr><tr><td>Dissection Pan and Wax</td><td>United Scientific Supplies</td><td>DSPA01-W</td><td></td></tr><tr><td>DMEM</td><td>Cytiva</td><td>SH30022.FS</td><td></td></tr><tr><td>Isoflurane</td><td>Sigma-Aldrich</td><td>26675-46-7</td><td></td></tr><tr><td>LI-COR Odyssey</td><td>LI-COR</td><td>DLx</td><td></td></tr><tr><td>Micro Dissecting Curved Scissors (24 mm Blade)&amp;nbsp;</td><td>Roboz Surgical Instrument</td><td>RS-5913</td><td></td></tr><tr><td>Micro Dissecting Spring Scissors (13 mm Blade)</td><td>Roboz Surgical Instrument</td><td>RS-5677</td><td></td></tr><tr><td>Micro Dissecting Spring Scissors (5 mm Blade)</td><td>Roboz Surgical Instrument</td><td>RS-5600</td><td></td></tr><tr><td>Micro Dissecting Tweezers (0.10 x 0.06 mm Tip)</td><td>Roboz Surgical Instrument</td><td>RS-4976</td><td></td></tr><tr><td>Optima MAX-TL Ultracentrifuge</td><td>Beckman Coulter</td><td>B11229</td><td></td></tr><tr><td>OsteoSense 680EX</td><td>Perkin Elmer</td><td>NEV10020EX</td><td></td></tr><tr><td>Pierce Protease Inhibitor</td><td>Thermo Scientific</td><td>A32963</td><td></td></tr><tr><td>Potassium Chloride</td><td>Fischer Chemical</td><td>P217</td><td></td></tr><tr><td>RIPA Lysis and Extraction Buffer</td><td>G Biosciences</td><td>786-489</td><td></td></tr><tr><td>Sodium Chloride</td><td>Fischer Chemical</td><td>BP358</td><td></td></tr><tr><td>Sodium Hydroxide</td><td>Thermo Scientific</td><td>A4782602</td><td></td></tr><tr><td>Sodium phosphate monobasic</td><td>Sigma-Aldrich</td><td>S0751</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S7903</td><td></td></tr><tr><td>Synergy HTX Multimode Reader</td><td>Agilent</td><td></td><td></td></tr><tr><td>Tissue culture plate, 96-well</td><td>Thermo Fisher</td><td>167008</td><td></td></tr><tr><td>T-Pins</td><td>United Scientific Supplies</td><td>TPIN02-PK/100</td><td></td></tr><tr><td>Tris Hydrochloride</td><td>Fischer Chemical</td><td>BP153</td><td></td></tr></tbody></table>

analysis of extracellular vesicle-mediated vascular calcification using in vitro and in vivo models

Cardiovascular disease is the leading cause of death in the world, and vascular calcification is the most significant predictor of cardiovascular events; however, there are currently no treatment or therapeutic options for vascular calcification. Calcification begins within specialized extracellular vesicles (EVs), which serve as nucleating foci by aggregating calcium and phosphate ions. This protocol describes methods for obtaining and assessing calcification in murine aortas and analyzing the associated extracted EVs. First, gross dissection of the mouse is performed to collect any relevant organs, such as the kidneys, liver, and lungs. Then, the murine aorta is isolated and excised from the aortic root to the femoral artery. Two to three aortas are then pooled and incubated in a digestive solution before undergoing ultracentrifugation to isolate the EVs of interest. Next, the mineralization potential of the EVs is determined through incubation in a high-phosphate solution and measuring the light absorbance at a wavelength of 340 nm. Finally, collagen hydrogels are used to observe the calcified mineral formation and maturation produced by the EVs in vitro.

Once the aortas are extracted, imaging using a near-infrared optical scanner shows a visual representation of the aorta as well as the vascular calcification (Figure 1). The pixel intensity values within the scanned fluorescent image represent the distribution of calcification and are shown here using a colored heatmap. Quantification methods include identifying a positive threshold and reporting the percentage area of the aorta with values greater than this threshold value and/or reporting ...

Watch this Scientific Journal Video about Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models at JoVE.com

Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

This paper presents the methodology for obtaining and assessing vascular calcification by isolating murine aortas followed by extracting calcified extracellular vesicles to observe the mineralization potential.

Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

Cardiovascular disease is the leading cause of death in the world, and vascular calcification is the most significant predictor of cardiovascular events; ...

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1.6K Views.  Florida International University. Dit protocol biedt een methode die kan worden gebruikt om te helpen bij het beoordelen en in de toekomst helpen bij het ontwikkelen van behandelingen voor vasculaire calcificatie, de belangrijkste voorspeller van hart- en vaatziekten. Het vermogen om EV's te isoleren stelt ons in staat om de mechanismen die leiden tot vasculaire verkalking en de fenotypische verandering van vasculaire gladde spiercellen verder te bestuderen. Deze techniek geeft inzicht in vroege moleculaire mechanismen van va...

Video: Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

Isolation and Characterization of Primary Rat Valve Interstitial Cells: A New Model to Study Aortic Valve Calcification

Calcific aortic valve disease (CAVD) is characterized by the progressive thickening of the aortic valve leaflets. It is a condition frequently found in the elderly and end-stage renal disease (ESRD) patients, who commonly suffer from hyperphosphatemia and hypercalcemia. At present, there are no medication therapies that can stop its progression. The mechanisms that underlie this pathological process remain unclear. The aortic valve leaflet is composed of a thin layer of valve endothelial cells (VECs) on the outer surfaces of the aortic cusps, with valve interstitial cells (VICs) sandwiched between the VECs. The use of a rat model enables the in vitro study of ectopic calcification based on the in vivo physiopathological serum phosphate (Pi) and calcium (Ca) levels of patients who suffer from hyperphosphatemia and hypercalcemia. The described protocol details the isolation of a pure rat VIC population as shown by the expression of VIC markers: alpha-smooth muscle actin (&#945;-SMA) vimentin and tissue growth factor beta (TGF&#946;) 1 and 2, and the absence of cluster of differentiation (CD) 31, a VEC marker. By expanding these VICs, biochemical, genetic, and imaging studies can be performed to study and unravel the key mediators underpinning CAVD.

This protocol describes the isolation, culture, and calcification of rat-derived valve interstitial cells, a highly physiological in vitro model of calcific aortic valve disease (CAVD). Exploitation of this rat model facilitates CAVD research in exploring the cell and molecular mechanisms that underlie this complex pathological process.

Calcific aortic valve disease (CAVD) is characterized by the progressive thickening of the aortic valve leaflets. It is a condition frequently found in ...

Developmental Biology

Kinetic Analysis of Vasculogenesis Quantifies Dynamics of Vasculogenesis and Angiogenesis In Vitro

Vasculogenesis is a complex process by which endothelial stem and progenitor cells undergo de novo vessel formation. Quantitative assessment of vasculogenesis has become a central readout of endothelial progenitor cell functionality, and therefore, several attempts have been made to improve both in vitro and in vivo vasculogenesis models. However, standard methods are limited in scope, with static measurements failing to capture many aspects of this highly dynamic process. Therefore, the goal of developing this novel protocol was to assess the kinetics of in vitro vasculogenesis in order to quantitate rates of network formation and stabilization, as well as provide insight into potential mechanisms underlying vascular dysfunction. Application of this protocol is demonstrated using fetal endothelial colony forming cells (ECFCs) exposed to maternal diabetes mellitus. Fetal ECFCs were derived from umbilical cord blood following birth, cultured, and plated in slides containing basement membrane matrix, where they underwent vasculogenesis. Images of the entire slide wells were acquired using time-lapse phase contrast microscopy over 15 hours. Images were analyzed for derivation of quantitative data using an analysis software called Kinetic Analysis of Vasculogenesis (KAV). KAV uses image segmentation followed by skeletonization to analyze network components from stacks of multi-time point phase contrast images to derive ten parameters (9 measured, 1 calculated) of network structure including: closed networks, network areas, nodes, branches, total branch length, average branch length, triple-branched nodes, quad-branched nodes, network structures, and the branch to node ratio. Application of this protocol identified altered rates of vasculogenesis in ECFCs obtained from pregnancies complicated by diabetes mellitus. However, this technique has broad implications beyond the scope reported here. Implementation of this approach will enhance mechanistic assessment and improve functional readouts of vasculogenesis and other biologically important branching processes in numerous cell types or disease states.

Kinetic Analysis of Vasculogenesis Quantifies Dynamics of Vasculogenesis and Angiogenesis In Vitro

Here, we present a protocol for time-lapse imaging and analysis of vasculogenesis in vitro using phase contrast microscopy coupled with the open source software, Kinetic Analysis of Vasculogenesis. This protocol can be applied to quantitatively assess the vasculogenic potential of numerous cell types or experimental conditions that model vascular disease.

Vasculogenesis is a complex process by which endothelial stem and progenitor cells undergo de novo vessel formation. Quantitative assessment of ...

An In Vitro 3D Model and Computational Pipeline to Quantify the Vasculogenic Potential of iPSC-Derived Endothelial Progenitors

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent endothelial progenitors (EPs), which can differentiate into the cell types necessary to assemble mature, functional vasculature, have been derived from both embryonic and induced pluripotent stem cells. However, these cells have not been rigorously evaluated in three-dimensional environments, and a quantitative measure of their vasculogenic potential remains elusive. Here, the generation and isolation of iPSC-EPs via fluorescent-activated cell sorting are first outlined, followed by a description of the encapsulation and culture of iPSC-EPs in collagen hydrogels. This extracellular matrix (ECM)-mimicking microenvironment encourages a robust vasculogenic response; vascular networks form after a week of culture. The creation of a computational pipeline that utilizes open-source software to quantify this vasculogenic response is delineated. This pipeline is specifically designed to preserve the 3D architecture of the capillary plexus to robustly identify the number of branches, branching points, and the total network length with minimal user input.

Endothelial progenitors derived from induced pluripotent stem cells (iPSC-EPs) have the potential to revolutionize cardiovascular disease treatments and to enable the creation of more faithful cardiovascular disease models. Herein, the encapsulation of iPSC-EPs in three-dimensional (3D) collagen microenvironments and a quantitative analysis of these cells&#8217; vasculogenic potential are described.

Induced pluripotent stem cells (iPSCs) are a patient-specific, proliferative cell source that can differentiate into any somatic cell type. Bipotent ...

A Semi-Automated and Reproducible Biological-Based Method to Quantify Calcium Deposition In Vitro

Vascular calcification involves a series of degenerative pathologies, including inflammation, changes to cellular phenotype, cell death, and the absence of calcification inhibitors, that concomitantly lead to a loss of vessel elasticity and function. Vascular calcification is an important contributor to morbidity and mortality in many pathologies, including chronic kidney disease, diabetes mellitus, and atherosclerosis. Current research models to study vascular calcification are limited and are only viable at the late stages of calcification development in vivo. In vitro tools for studying vascular calcification use end-point measurements, increasing the demands on biological material and risking the introduction of variability to research studies. We demonstrate the application of a novel fluorescently labeled probe that binds to in vitro calcification development on human vascular smooth muscle cells and determines the real-time development of in vitro calcification. In this protocol, we describe the application of our newly developed calcification assay, a novel tool in disease modeling that has potential translational applications. We envisage this assay to be relevant in a broader spectrum of mineral deposition research, including applications in bone, cartilage, or dental research.

A Semi-Automated and Reproducible Biological-Based Method to Quantify Calcium Deposition In Vitro

Cardiovascular disease is the leading cause of death worldwide. Vascular calcification contributes substantially to the burden of cardiovascular morbidity and mortality. This protocol describes a simple method to quantify vascular smooth muscle cell-mediated calcium precipitation in vitro by fluorescent imaging.

Vascular calcification involves a series of degenerative pathologies, including inflammation, changes to cellular phenotype, cell death, and the absence ...

Biochemistry

Calcification of Vascular Smooth Muscle Cells and Imaging of Aortic Calcification and Inflammation

Cardiovascular disease is the leading cause of morbidity and mortality in the world. Atherosclerotic plaques, consisting of lipid-laden macrophages and calcification, develop in the coronary arteries, aortic valve, aorta, and peripheral conduit arteries and are the hallmark of cardiovascular disease. In humans, imaging with computed tomography allows for the quantification of vascular calcification; the presence of vascular calcification is a strong predictor of future cardiovascular events. Development of novel therapies in cardiovascular disease relies critically on improving our understanding of the underlying molecular mechanisms of atherosclerosis. Advancing our knowledge of atherosclerotic mechanisms relies on murine and cell-based models. Here, a method for imaging aortic calcification and macrophage infiltration using two spectrally distinct near-infrared fluorescent imaging probes is detailed. Near-infrared fluorescent imaging allows for the ex vivo quantification of calcification and macrophage accumulation in the entire aorta and can be used to further our understanding of the mechanistic relationship between inflammation and calcification in atherosclerosis. Additionally, a method for isolating and culturing animal aortic vascular smooth muscle cells and a protocol for inducing calcification in cultured smooth muscle cells from either murine aortas or from human coronary arteries is described. This in vitro method of modeling vascular calcification can be used to identify and characterize the signaling pathways likely important for the development of vascular disease, in the hopes of discovering novel targets for therapy.

Vascular calcification is an important predictor of and contributor to human cardiovascular disease. This protocol describes methods for inducing calcification of cultured primary vascular smooth muscle cells and for quantifying calcification and macrophage burden in animal aortas using near-infrared fluorescence imaging.

Cardiovascular disease is the leading cause of morbidity and mortality in the world.  Atherosclerotic plaques, consisting of lipid-laden macrophages and ...

Medicine

In Vitro Model of Coronary Angiogenesis

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects in these vessels represent severe health risks such as in atherosclerosis, which can lead to myocardial infarctions and heart failures in patients. Consequently, coronary artery disease is one of the leading causes of death worldwide. Despite its clinical importance, relatively little progress has been made on how to regenerate damaged coronary arteries. Nevertheless, recent progress has been made in understanding the cellular origin and differentiation pathways of coronary vessel development. The advent of tools and technologies that allow researchers to fluorescently label progenitor cells, follow their fate, and visualize progenies in vivo have been instrumental in understanding coronary vessel development. In vivo studies are valuable, but have limitations in terms of speed, accessibility, and flexibility in experimental design. Alternatively, accurate in vitro models of coronary angiogenesis can circumvent these limitations and allow researchers to interrogate important biological questions with speed and flexibility. The lack of appropriate in vitro model systems may have hindered the progress in understanding the cellular and molecular mechanisms of coronary vessel growth. Here, we describe an in vitro culture system to grow coronary vessels from the sinus venosus (SV) and endocardium (Endo), the two progenitor tissues from which many of the coronary vessels arise. We also confirmed that the cultures accurately recapitulate some of the known in vivo mechanisms. For instance, we show that the angiogenic sprouts in culture from SV downregulate COUP-TFII expression similar to what is observed in vivo. In addition, we show that VEGF-A, a well-known angiogenic factor in vivo, robustly stimulates angiogenesis from both the SV and Endo cultures. Collectively, we have devised an accurate in vitro culture model to study coronary angiogenesis.

In vitro models of coronary angiogenesis can be utilized for the discovery of the cellular and molecular mechanisms of coronary angiogenesis. In vitro explant cultures of sinus venosus and endocardium tissues show robust growth in response to VEGF-A and display a similar pattern of COUP-TFII expression as in vivo.

Here, we describe an in vitro culture assay to study coronary angiogenesis. Coronary vessels feed the heart muscle and are of clinical importance. Defects ...

Isolation of Mouse Interstitial Valve Cells to Study the Calcification of the Aortic Valve In Vitro

The calcification of aortic valve cells is the hallmark of aortic stenosis and is associated with valve cusp fibrosis. Valve interstitial cells (VICs) play an important role in the calcification process in aortic stenosis through the activation of their dedifferentiation program to osteoblast-like cells. Mouse VICs are a good in vitro tool for the elucidation of the signaling pathways driving the mineralization of the aortic valve cell. The method described herein, successfully used by these authors, explains how to obtain freshly isolated cells. A two-step collagenase procedure was performed with 1 mg/mL and 4.5 mg/mL. The first step is crucial to remove the endothelial cell layer and avoid any contamination. The second collagenase incubation is to facilitate the migration of VICs from the tissue to the plate. In addition, an immunofluorescence staining procedure for the phenotype characterization of the isolated mouse valve cells is discussed. Furthermore, the calcification assay was performed in vitro by using the calcium reagent measurement procedure and alizarin red staining. The use of mouse valve cell primary culture is essential for testing new pharmacological targets to inhibit cell mineralization in vitro.

Isolation of Mouse Interstitial Valve Cells to Study the Calcification of the Aortic Valve In Vitro

This article describes the isolation of mouse aortic valve cells by a two-step collagenase procedure. Isolated mouse valve cells are important for performing different assays, such as this in vitro calcification assay, and for investigating the molecular pathways leading to aortic valve mineralization.

The calcification of aortic valve cells is the hallmark of aortic stenosis and is associated with valve cusp fibrosis. Valve interstitial cells (VICs) ...

Isolation of Human Primary Valve Cells for In vitro Disease Modeling

Calcific aortic valve disease (CAVD) is present in nearly a third of the elderly population. Thickening, stiffening, and calcification of the aortic valve causes aortic stenosis and contributes to heart failure and stroke. Disease pathogenesis is multifactorial, and stresses such as inflammation, extracellular matrix remodeling, turbulent flow, and mechanical stress and strain contribute to the osteogenic differentiation of valve endothelial and valve interstitial cells. However, the precise initiating factors that drive the osteogenic transition of a healthy cell into a calcifying cell are not fully defined. Further, the only current therapy for CAVD-induced aortic stenosis is aortic valve replacement, whereby the native valve is removed (surgical aortic valve replacement, SAVR) or a fully collapsible replacement valve is inserted via a catheter (transcatheter aortic valve replacement, TAVR). These surgical procedures come at a high cost and with serious risks; thus, identifying novel therapeutic targets for drug discovery is imperative. To that end, the present study develops a workflow where surgically removed tissues from patients and donor cadaver tissues are used to create patient-specific primary lines of valvular cells for in vitro disease modeling. This protocol introduces the utilization of a cold storage solution, commonly utilized in organ transplant, to reduce the damage caused by the often-lengthy procurement time between tissue excision and laboratory processing with the benefit of greatly stabilizing cells of the excised tissue. The results of the present study demonstrate that isolated valve cells retain their proliferative capacity and endothelial and interstitial phenotypes in culture upwards of several days after valve removal from the donor. Using these materials allows for the collection of control and CAVD cells, from which both control and disease cell lines are established.

This protocol describes the collection of human aortic valves extracted during surgical aortic valve replacement procedures or from cadaveric tissue, and the subsequent isolation, expansion, and characterization of patient specific primary valve endothelial and interstitial cells. Included are important details regarding the processes needed to ensure cell viability and phenotype specificity.

Calcific aortic valve disease (CAVD) is present in nearly a third of the elderly population. Thickening, stiffening, and calcification of the aortic valve ...

Evaluating Salidroside as a Therapeutic Agent for Vascular Calcification Using Network Pharmacology and Experimental Rat Models

Vascular calcification (VC) is a critical pathological condition associated with significant morbidity and mortality. This study employs a hybrid approach of network pharmacology and molecular biology to delineate the therapeutic mechanisms of salidroside (SAL), an active compound from Rhodiola crenulata, against VC. Through database mining and network analysis, 388 SAL targets intersecting with 2871 VC-associated targets were identified, resulting in 208 common targets. A protein-protein interaction (PPI) network constructed via the String database and topological analysis in Cytoscape 3.9.1 pinpointed 10 key targets, including IL6, TNF, TP53, IL1B, HIF1A, CASP3, and STAT3, among others. The identified genes were concentrated in the lipid and atherosclerosis pathways, indicating that the improvement of VC by SAL may occur through the regulation of abnormal expression of lipid and inflammatory factors. It was also found that SAL inhibits the abnormal expression of inflammatory factors, thereby activating the JAK2/STAT3 pathway to intervene in the progression of VC. The JAK2/STAT3 pathway is a key molecular mechanism by which SAL prevents further deterioration of VC. Functional enrichment analyses revealed the involvement of these targets in inflammatory responses and lipid metabolism, pivotal pathways in VC. In vivo studies in rats demonstrated SAL's efficacy in mitigating dyslipidemia and vascular inflammation, with improved serum lipid profiles and reduced vascular calcium deposition. The mechanistic exploration, grounded in Western blot analysis, demonstrated salidroside's ability to regulate the JAK2/STAT3 signaling pathway, highlighting its potential as a modulator in this critical molecular mechanism and offering a potential therapeutic target for VC. The strength of this research lies in its methodological rigor, integrating computational predictions with in vivo validations. This comprehensive approach establishes a robust framework for exploring the therapeutic mechanisms of natural compounds in combating VC.

This study establishes a rat model of vascular calcification induced by a high-fat diet (HFD) combined with vitamin D3 (VD3). The model was used to evaluate the therapeutic efficacy of salidroside in preventing and treating vascular calcification, providing insights into its potential mechanisms of action through network pharmacology and in vivo experiments.

Vascular calcification (VC) is a critical pathological condition associated with significant morbidity and mortality. This study employs a hybrid approach ...

Von Kossa Staining of Aortic Vascular Smooth Muscle Cells: An In Vitro Technique for Assessment of Arterial Medial Layer Calcification

Source: O&#39;Rourke, C., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/54017">Calcification of Vascular Smooth Muscle Cells and Imaging of Aortic Calcification and Inflammation.</a> J. Vis. Exp. (2016).
This video demonstrates a staining technique for assessing arterial medial calcification of the vascular smooth muscle cells in vitro. The von Kossa method employs a photochemical reaction to deposit metallic silver at the sites of calcification and impart a black coloration.

Source: O&#39;Rourke, C., et al. Calcification of Vascular Smooth Muscle Cells and Imaging of Aortic Calcification and Inflammation. J. Vis. Exp. (2016).
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Analysis of Extracellular Vesicle-Mediated Vascular Calcification Using In Vitro and In Vivo Models

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