This research was funded by the European Union&#39;s Horizon 2020 research and innovation programs under the Marie Sklodowska-Curie grant agreement No 722609 and 764474, NWO ZonMw (MKMD 40-42600-98-13007). This research was supported by BioSPX. WJ-D received funding from the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) TRR219-project ID 322900939 and project ID 403041552

1. Cell seeding, maintenance, and calcification induction
<ol>
	<li>For culturing primary cells, use a laminar airflow cabinet, gloves, and sterile equipment. Disinfect hands and workspace before and after conducting any work. Treat all primary cells and culture media as a potential biohazard, unless proven otherwise. Preferably autoclave surplus cells and media before disposal. Do not chemically inactivate and autoclave since this will liberate toxic fumes.</li>
	<li>Culture hVSMC on uncoated cell culture plates.</li>
	<li>Routinely maintain hVSMC in growth medium consisting of M199 medium supplemented with 10%-20% FBS and 1% Pen/Strep and incubate at 37 &#176;C and 5% CO2.</li>
	<li>Split the cells upon 70%-90% confluency.
	<ol>
		<li>To split, wash the hVSMCs 2x with PBS. Add trypsin and incubate for 2-5 min at 37 &#176;C. Check for the detachment of cells under the microscope.</li>
		<li>Inhibit the trypsin reaction by adding serum-containing medium. Centrifuge the cells at 350 x g for 4 min and resuspend the pellet in maintenance medium. hVSMCs follow a 1:2 splitting scheme.</li>
	</ol>
	</li>
	<li>To start the calcification experiment, prepare the cells following instructions for a split (Step 1.4.1). Upon resuspension, count the cells and seed into a 48-well plate at a density 10-15 x 103 cells/cm2. Seed the cells avoiding the use of the outer wells, as recommended in Supplemental Figure 1.</li>
	<li>Let the cells adhere and recover for 24 h (overnight) while incubating at 37 &#176;C and 5% CO2.</li>
	<li>Gently wash the cells 2x with PBS, carefully aspirating all the remaining PBS after the 2nd wash.</li>
	<li>Gently add calcification medium and further incubate at 37 &#176;C and 5% CO2. Calcification medium must contain a calcification stimulus, fluorescently labeled fetuin-A (e.g., with red fluorescent protein [RFP]) (1 &#181;g/mL), and HOECHST 33342 (0.1 &#181;g/mL) nuclear stain or similar live fluorescent nuclear stain. Do not use DAPI, as this is non-permeable.</li>
	<li>Check daily with a light microscope until calcification occurs. 
	&#8203;NOTE: For notes on the cell culture and maintenance of hVMSCs, please see Supplemental File 1.</li>
</ol>
2. Calcification detection via imaging
NOTE: The following protocol provides the general steps to be taken in preparation, imaging, and data analysis. Screenshots supporting the instructions for each step using an automated imaging platform and corresponding image analysis software (see Table of Materials for details) are provided in Supplemental File 2 and Supplemental File 3. Other imaging instruments and image processing tools may be used to apply this protocol. However, repeated imaging at the same location in each well is crucial for meaningful data acquisition. Creating a protocol to image calcification and re-use at every imaging step is necessary for obtaining reproducible results. The first time applying the method, follow the steps below to prepare before the imaging.
<ol>
	<li>Protocol setup
	<ol>
		<li>Open the software, and then select Protocols and Create New.</li>
		<li>Select Procedure and click Set Temperature. In the pop-up window, set the temperature to 37 &#176;C and the gradient to 1 &#176;C, and click OK. Select the plate type of choice from the dropdown menu. Add the imaging step by clicking Image. Select the inverted imager, and click OK.</li>
		<li>Add three imaging channels and set them to DAPI (377, 447 nm), RFP (531, 593 nm), and brightfield. Tick the Montage box and set the desired number and location of the images. 2 x 2 for a 48-well plate is a common choice. Change the overlap to represent a better coverage of each well. Select which wells are to be imaged. A new window will open where the wells of interest can be selected.</li>
		<li>Click Focus Options to set the focus mode. A new window will open. Set each channel individually. Commonly, wells are autofocused on the &#34;DAPI&#34; channel. Set all other channels to Fixed Focal Height From First Channel and set by clicking OK. Close the window.</li>
		<li>Start the pre-setting data reduction steps by clicking Data Reduction and select Image Preprocessing. Setting up this step reduces the fluorescence background. Accept the default settings by selecting OK. A new set of images will be created with the prefix &#34;TSF&#34;. The original images will be preserved.</li>
		<li>Set up a step to count the cells by selecting Cellular Analysis. Select TSF DAPI images from the Channel dropdown menu. Set further details after imaging has been performed. This step can also be considered data reduction; always make sure to keep the original images.</li>
		<li>Prepare the analysis of the RFP (calcification) signal by selecting Statistics. In the pop-up window, label Step and select TSF RFP as the input channel. Tick the Upper Value and Lower Value boxes. Tick the box for Total Area in the lower list. Select None in the Color Effect column. In the pop-up window, click Custom and tick the Background box. This will color code the results from low-high for easy assessment. Press OK.</li>
		<li>For a fair readout, normalize the RFP signal per cell. In doing so, a total area is divided per cell. To set up this step, press Ratio on the left side. In the pop-up window, select for data input 1 RFP Quantification: Total Area from the drop-down menu. Further select Cell Count as data input 2.</li>
		<li>Finally, select a New Data Set Name and press OK and OK again. Select Color Effect as described previously. In the upper-left corner, select File and save the file as protocol (.prt file type).</li>
	</ol>
	</li>
	<li>Daily after the first calcification occurs
	<ol>
		<li>For every time point, repeat these steps. In the startup software, press Read Now and Existing Protocol.... Select the protocol that has been created in the previous step. The program will ask to save the experiment. This can be done at this step but also at any later step manually by clicking the Save button on the upper left.</li>
		<li>A pop-up window will ask to wait for the system to heat. Turn on the CO2 gas-controller and set it to 5%.</li>
		<li>Once the system has reached the set temperature, a prompt will appear to insert a plate and read. Press Cancel. This is because, for each time point, the exposure of each fluorochrome must be adjusted individually.</li>
		<li>Transport the plate from the incubator to the imager in a safe box that resists breaking and spilling and is in line with local biosafety regulations for transporting live cells, in case of an accident. Place the plate into the plate reader.</li>
		<li>After canceling, click on Procedure. In the pop-up window, select Pre-Set Imaging Step. Adjust the focus and exposure on the DAPI channel first. Untick the Auto Exposure box at all channels. Then, click the Microscope icon next to the DAPI channel. A new window will open.</li>
		<li>Select the well to focus on. Use a well with medium to high calcification for this step. If a signal is already visible, first click Autofocus and then Autoexposure. If not, first increase the exposure and repeat Autofocus and Autoexposure. If desired, the exposure can be adjusted manually by selecting the exposure drop-down menu. Check the settings on multiple wells. Once satisfied, save the settings.</li>
		<li>Adjust the exposure in the same manner for all channels. 
		CAUTION: Do not focus on other channels; only adjust exposure. The focus plain should be the same for all channels.</li>
		<li>Close the procedure window. Select the green Read Now button in the top task bar. Save the experiment as indicated by the software. 
		&#8203;NOTE: The software will now automatically read all the selected wells in the selected settings.</li>
	</ol>
	</li>
</ol>
3. Data analysis
NOTE: For detailed screenshots on how to perform the data analysis using an automated imaging platform and corresponding image analysis software (see Table of Materials for details), please see Supplemental File 4. If using alternative imaging instruments or analysis software, the images should be exported and batch processed ensuring that the exposure, fluorescence threshold, or intensity are adjusted equally for all images in a comparative data set.
<ol>
	<li>While the system reads the plate, all the images are automatically processed in the background. Adjust the settings for cell count by selecting TSF DAPI images to display. Select the well of choice by double-clicking. A new window will pop up. Select one of the images.</li>
	<li>Select the Analysis tab. In the pop-up window, select Cellular Analysis: Cell Count. Click OK. Deselect the RFP and brightfield channels. Tick the Highlight Objects box.</li>
	<li>Click Options to open the menu. Adjust the size and intensity threshold to include all nuclei but exclude debris. Click OK. Click Apply changes in the lower left to also transfer the settings to all other images.</li>
	<li>In a similar manner as before, select a well and image with a medium to high amount of calcification to best adjust the signal-to-noise ratio; click the Analysis tab in the task menu.
	<ol>
		<li>This time select Image Statistics. Deselect the DAPI and brightfield channel. Click Options to open the menu. Tick the box Threshold Outliers. Adjust the threshold&#39;s upper and lower values. Only count the signal if it is above the background and exclude if there are debris/artifacts.</li>
		<li>To set a non-subjective threshold value for the signal above background, select the line tool from the task bar. A window will pop up. Draw a line through a patch of signal, but make sure to also include an area of background. 
		NOTE: A graph will appear in the pop-up window, showing a peak representing the signal above the background line. The 25% peak height value represents the recommended threshold. It is also recommended to measure multiple signal patches to ensure the selection of the right threshold. While selecting patches of signal, selecting regions with very high signal strength may result in a threshold that neglects medium and or lower signals. It is, therefore, recommended to select patches of medium to low signal above background. Examples of too high, too low, and accurate thresholds are shown in Supplemental File 4.</li>
		<li>Once satisfied, click OK and Apply Changes to transfer the settings to all other wells. Check if the threshold is accurately selected in the other wells.</li>
	</ol>
	</li>
	<li>Once the cell count and RFP thresholds are set, export the data. Select the plate of interest. In the drop-down menu, multiple parameters can be selected for export. Of relevance might be cell count, RFP area, and &#34;area/cell&#34; which is the metric used for comparison between groups. Click the Excel button next to the drop-down menu to export the data directly into a spreadsheet. 
	NOTE: In the dropdown menu, the calculated values for the cell count, the total area of RFP signal (reflecting the area of calcification), as well as the normalized ratio of total area of RFP signal per cell can now be selected individually and exported to a spreadsheet. Display the results as the ratio of total area of RFP signal per cell and visualize (e.g., bar graphs). Apply an unpaired Student&#39;s t-test to compare two groups or an unpaired ANOVA to compare different groups at the same time point. Comparing the same group at different time points may require paired statistical analysis.</li>
</ol>

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Leon Schurgers has received institutional grants from Bayer, Boehringer Ingelheim, NattoPharma, and IDS. Leon Schurgers owns shares in Coagulation Profile. Willi Jahnen-Dechent is a co-founder and shareholder of CALCISCON AG.

In this manuscript, we describe a semi-automated method for in vitro calcification determination. For this method, three critical steps of hVSMC calcification should be optimized. First, cellular density is critical for hVSMC calcification development. Low densities of hVSMCs will result in slow or no calcification and cell death due to the lack of cell-to-cell contact and the stress that is induced under calcifying conditions21. High cellular densities result in over-confluency, after wh...

Vascular calcification (VC) is an independent risk factor for cardiovascular morbidity and mortality1,2,3. Long considered a passive chemical process of ectopic mineral deposition, it now appears a modifiable tissue healing response involving the active contribution of various cells including activated vascular smooth muscle cells (hVSMC) as a driver of the disease4,5. In vivo VC can be measured by multislice CT scans as an assessment of atherosclerotic burden6,7,8. Currently, a paradigm shift is underway, wherein VC severity is becoming recognized as a risk factor in cardiovascular disease, type II diabetes, chronic kidney disease, and ageing9,10,11,12,13,14,15.
hVSMCs are the most abundant cell type in the cardiovascular system and a principal actor in the development of VC. In vitro hVSMC-induced calcification is a widely used disease model to study cardiovascular disease16,17. However, most protocols for the detection of in vitro calcification use end-point measurements that can limit data acquisition, require greater use of cellular material, and can slow research. Common methods for the detection of in vitro hVSMC calcification include the o-cresolphthalein assay, which measures solubilized calcium deposition against total protein and requires cell lysis18. Also, Alizarin Red staining is used, which binds directly to calcium deposits on fixed cells or tissue19. To study hVSMC calcification over time with either o-cresolphthalein or Alizarin Red requires batches of replicates per time point, increasing the demand on biological material, and in turn, increasing the chance of variability.
In this paper, we detail the method for the application of a novel assay that utilizes hVSMCs with a fluorescent imaging probe to determine in vitro VC progression as well as function as a singular end-stage calcification assay. We previously demonstrated that this assay is directly comparable to the o-cresolphthalein and Alizarin Red methods and can be used to distinguish between varying culture conditions20. In addition to real-time measurements, this assay may be used to determine the propensity of serum or plasma samples as a surrogate marker for clinical VC development20. This will aid in the application of biological strategies of cardiovascular sciences and disease modeling. A further application of the assay may be as a translational BioHybrid system to assess VC severity or progression from blood constituents such as serum or plasma.

<table><tbody><tr><td>Calcium chloride, 93%, anhydrous</td><td>Thermo Fisher Scientific</td><td>349615000</td><td></td></tr><tr><td>Costar 6-well Clear TC-treated well plates</td><td>Corning</td><td>3516</td><td></td></tr><tr><td>Cytation 3 System</td><td>BioTek, Abcoude, The Netherlands</td><td></td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Merck</td><td>F7524-100ML</td><td></td></tr><tr><td>Fetuin-A-Alexa Fluor-546</td><td>Prepared in-house</td><td></td><td></td></tr><tr><td>Gen5 Software v3.10</td><td>BioTek</td><td></td><td></td></tr><tr><td>Gibco Medium 199</td><td>Thermo Fisher Scientific</td><td>11150059</td><td></td></tr><tr><td>Hoechst 33342, Trihydrochloride</td><td>Thermo Fisher Scientific</td><td>H3570</td><td></td></tr><tr><td>PBS (10X), pH 7.4</td><td>Thermo Fisher Scientific</td><td>70011044</td><td></td></tr><tr><td>Penicillin-Streptomycin</td><td>Thermo Fisher Scientific</td><td>15140122</td><td></td></tr><tr><td>Trypsin-EDTA (0.05%), phenol red</td><td>Thermo Fisher Scientific</td><td>25300062</td><td></td></tr></tbody></table>

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.

The outcome includes original images of HOECHST-stained nuclei, RFP-labeled calcification, and brightfield images. Different stages of calcification ranging from low (Figure 2) to high (Figure 3) may be detected and analyzed. Calcification can usually be spotted as black speckles using light microscopy (Figure 2D and Figure&#160;3B, arrows indicate calcification), which are useful for primary assessment...

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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.

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 ...

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Research

JoVE Journal

Biochemistry

1.9K Views.  Maastricht University. Onze methode maakt het mogelijk om de verkalkingsneiging van patiëntmonsters te meten met behulp van menselijke vasculaire gladde spiercellen. Dit zal helpen bij het stimuleren van onderzoek naar vasculaire verkalkingsstoornissen, wat helpt bij de ontwikkeling van nieuwe therapieën. Deze techniek maakt gebruik van een fluorescerende detectie, die de gevoeligheid verhoogt en het mogelijk maakt om monsters in de loop van de tijd te meten, waardoor de behoefte aan replicaties wordt verminderd ...

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

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

Measuring Fast Calcium Fluxes in Cardiomyocytes

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to extracellular agents is predominantly regulated by the phospholipase C&beta;- G&alpha;q pathway localized on the plasma membrane which is stimulated by agents such as acetylcholine 3,4. We have recently found that plasma membrane protein domains called caveolae5,6 can entrap activated G&alpha;q 7. This entrapment has the effect of stabilizing the activated state of G&alpha;q and resulting in prolonged Ca2+ signals in cardiomyocytes and other cell types8. We uncovered this surprising result by measuring dynamic calcium responses on a fast scale in living cardiomyocytes. Briefly, cells are loaded with a fluorescent Ca2+ indicator. In our studies, we used Ca2+ Green (Invitrogen, Inc.) which exhibits an increase in fluorescence emission intensity upon binding of calcium ions. The fluorescence intensity is then recorded for using a line-scan mode of a laser scanning confocal microscope. This method allows rapid acquisition of the time course of fluorescence intensity in pixels along a selected line, producing several hundreds of time traces on the microsecond time scale. These very fast traces are transferred into excel and then into Sigmaplot for analysis, and are compared to traces obtained for electronic noise, free dye, and other controls. To dissect Ca2+ responses of different flux rates, we performed a histogram analysis that binned pixel intensities with time. Binning allows us to group over 500 traces of scans and visualize the compiled results spatially and temporally on a single plot. Thus, the slow Ca2+ waves that are difficult to discern when the scans are overlaid due to different peak placement and noise, can be readily seen in the binned histograms. Very fast fluxes in the time scale of the measurement show a narrow distribution of intensities in the very short time bins whereas longer Ca2+ waves show binned data with a broad distribution over longer time bins. These different time distributions allow us to dissect the timing of Ca2+fluxes in the cells, and to determine their impact on various cellular events.

We present a method to isolate rapid (microsecond) calcium events from slower fluxes in living cells using laser scanning confocal microscopy. The method measures fluorescence intensity fluctuations of calcium indicators by recording line scans of several hundred pixels in a cell. Histogram analysis allows us to isolate the time scales of different calcium fluxes.

Cardiomyocytes have multiple Ca2+ fluxes of varying duration that work together to optimize function 1,2. Changes in Ca2+ activity in response to ...

Biology

A Fluorescent Intravital Imaging Approach to Study Load-Induced Calcium Signaling Dynamics in Mouse Osteocytes

Bone tissue is exquisitely sensitive to differences in mechanical load magnitude. Osteocytes, dendritic cells that form a syncytium throughout the bone, are responsible for the mechanosensory function of bone tissue. Studies employing histology, mathematical modeling, cell culture, and ex vivo bone organ cultures have greatly advanced the understanding of osteocyte mechanobiology. However, the fundamental question of how osteocytes respond to and encode mechanical information at the molecular level in vivo is not well understood. Intracellular calcium concentration fluctuations in osteocytes offer a useful target for learning more about acute bone mechanotransduction mechanisms. Here, we report a method for studying osteocyte mechanobiology in vivo, combining a mouse strain with a fluorescently genetically encoded calcium indicator expressed in osteocytes with an in vivo loading and imaging system to directly detect osteocyte calcium levels during loading. This is achieved with a three-point bending device that can deliver well-defined mechanical loads to the third metatarsal of living mice while simultaneously monitoring fluorescently indicated calcium responses of osteocytes using two-photon microscopy. This technique allows for direct in vivo observation of osteocyte calcium signaling events in response to whole bone loading and is useful in the endeavor to reveal mechanisms in osteocyte mechanobiology.

The current article describes performing an intravital imaging approach to observe mechanically induced calcium signaling of embedded osteocytes in vivo in real-time in response to tissue-level mechanical loading of the mouse third metatarsal.

Bone tissue is exquisitely sensitive to differences in mechanical load magnitude. Osteocytes, dendritic cells that form a syncytium throughout the bone, ...

Bioengineering

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.

Calcium is involved in numerous physiological and pathophysiological signaling pathways. Live cell imaging requires specialized equipment and can be time consuming. A quick, simple method using a flow cytometer to determine relative changes in cytosolic calcium in adherent epithelial cells brought into suspension was optimized.

A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial ...

A Simple Pit Assay Protocol to Visualize and Quantify Osteoclastic Resorption In Vitro

Mature osteoclasts are multinucleated cells that can degrade bone through the secretion of acids and enzymes. They play a crucial role in various diseases (e.g., osteoporosis and bone cancer) and are therefore important objects of research. In vitro, their activity can be analyzed by the formation of resorption pits. In this protocol, we describe a simple pit assay method using calcium phosphate (CaP) coated cell culture plates, which can be easily visualized and quantified. Osteoclast precursors derived from human peripheral blood mononuclear cells (PBMCs) were cultured on the coated plates in the presence of osteoclastogenic stimuli. After 9 days of incubation, osteoclasts were fixed and stained for fluorescence imaging while the CaP coating was counterstained by calcein. To quantify the resorbed area, the CaP coating on plates was stained with 5% AgNO3 and visualized by brightfield imaging. The resorption pit area was quantified using ImageJ.

A Simple Pit Assay Protocol to Visualize and Quantify Osteoclastic Resorption In Vitro

Here, we present a simple and effective assay procedure for resorption pit assays using calcium phosphate coated cell culture plates.

Mature osteoclasts are multinucleated cells that can degrade bone through the secretion of acids and enzymes. They play a crucial role in various diseases ...

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) ...

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.

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.

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

Calcification Assay to Measure Calcium Concentration in Aortic Valve Interstitial Cells

Source: Bouchareb, R. et al., <a href="https://app.jove.com/v/62419">Isolation of Mouse Interstitial Valve Cells to Study the Calcification of the Aortic Valve In Vitro.</a>&#160; J. Vis. Exp. &#160;(2021).
This video demonstrates an assay technique for assessing the calcification of the aortic valve interstitial cells in vitro. The assay helps to quantify the calcium concentration using the arsenazo III reagent, a calcium-sensitive dye that reacts with calcium ions to form a purple-colored calcium-arsenazo complex.

Source: Bouchareb, R. et al., Isolation of Mouse Interstitial Valve Cells to Study the Calcification of the Aortic Valve In Vitro.&#160; J. Vis. Exp. ...

JoVE Encyclopedia of Experiments

Biological Techniques

Assay Techniques

Cell-based assays

In Vitro Mineralization Assay: A Colorimetric Method to Quantify Osteoblast Calcium Deposits on Bone Graft Substitute by Alizarin Red S Staining and Extraction

Transfer the stained beta-tricalcium phosphate disks to a new well, and scan the plates with a flatbed scanner to record mineralization.
After image capture, add 250 microliters of 10% cetylpyridinium chloride solution and shake for 15 minutes to extract the Alizarin red S dye. Next, transfer the solution from the wells to individual 1.5-milliliter tubes, and centrifuge at 17,000 times g for 5 minutes at room temperature.
Dilute the extract with 10% cetylpyridinium chloride solution at a dilution ratio of 1:10 to 1:20 in a 1.5-milliliter tube. Then, transfer 300 microliters of each diluted sample into the wells of the 96-well plate. Include two blank wells containing only 10% cetylpyridinium chloride.
Next, prepare seven Alizarin red S reference standards ranging from 4 to 400 micromolar by diluting 40 millimolar Alizarin red S staining solution with 10% cetylpyridinium chloride solution to generate a standard curve.
Load 300 microliters of each standard into the plate. Lastly, read the absorbance of the samples, blanks, and reference standards at 520 nanometers.

Biochemical assays

EoE Sub Articles

Resorption Pit Assay: An In Vitro Technique to Quantify Calcium Resorption Activity of Mature Osteoclasts using Calcium Phosphate-Coated Culture Plates

Resuspend PBMCs in a complete alpha-MEM containing 20 ng/mL macrophage colony-stimulating factor, and seed 2.5 x 105 cells/cm2 PBMCs in a flask. Incubate the calcium phosphate-coated plates with 50 microliters of fetal bovine serum at 37 degrees Celsius for one hour.
Next, wash the flask twice with PBS to detach the osteoclast precursors from dead or non-adherent cells. Then, add 4 milliliters of trypsin per 75 cm2 flask. After 30 minutes, stop the digestion by adding 4 milliliters of complete alpha-MEM. Then, carefully detach the cells using a cell scraper.
Transfer the cells into a 50-milliliter tube, and count the total number of cells using a counting chamber. Centrifuge the tube at 350 x g for seven minutes to pellet the cells, and resuspend the pellet in complete alpha-MEM containing M-CSF and RANKL to get 1 x 106 cells/mL.
Aspirate the fetal bovine serum from the calcium phosphate-coated plate, and pipette 200 microliters of cell suspension per well. After the incubation, wash the cells twice with PBS. Fix the cells with 4% paraformaldehyde for 10 minutes, and wash again with PBS.
For staining, add permeabilization buffer to the fixed cells, and incubate them for five minutes.
To stain the actin filaments, incubate the cells with 100 microliters of AlexaFluor 546-labeled phalloidin solution in PBS for 30 minutes. Then, aspirate the staining solution, and stain the nuclei using 100 microliters of Hoechst for 10 minutes.
Stain calcium phosphate coating with 100 microliters of 10 micromolar calcein in PBS for 10 minutes. After washing thrice with PBS, capture images.

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

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