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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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...

Watch this Scientific Journal Video about A Semi-Automated and Reproducible Biological-Based Method to Quantify Calcium Deposition In Vitro at JoVE.com

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. 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. 

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

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating these studies, reincorporation of detergent-solubilized membrane proteins into liposomes is a stochastic process where protein topology is impossible to enforce. This paper offers an alternative method to these challenging techniques that utilizes a liposome-based scaffold. Protein solubility is enhanced by deletion of the transmembrane domain, and these amino acids are replaced with a tethering moiety, such as a His-tag. This tether interacts with an anchoring group (Ni2+ coordinated by nitrilotriacetic acid (NTA(Ni2+)) for His-tagged proteins), which enforces a uniform protein topology at the surface of the liposome. An example is presented wherein the interaction between Dynamin-related protein 1 (Drp1) with an integral membrane protein, Mitochondrial Fission Factor (Mff), was investigated using this scaffold liposome method. In this work, we have demonstrated the ability of Mff to efficiently recruit soluble Drp1 to the surface of liposomes, which stimulated its GTPase activity. Moreover, Drp1 was able to tubulate the Mff-decorated lipid template in the presence of specific lipids. This example demonstrates the effectiveness of scaffold liposomes using structural and functional assays and highlights the role of Mff in regulating Drp1 activity.

Using Scaffold Liposomes to Reconstitute Lipid-proximal Protein-protein Interactions In Vitro

This paper describes a method for assessing the interactions and assemblies of integral membrane proteins in vitro with various partner factors in a lipid-proximal environment.

Studies of integral membrane proteins in vitro are frequently complicated by the presence of a hydrophobic transmembrane domain. Further complicating ...

Semi-automated Biopanning of Bacterial Display Libraries for Peptide Affinity Reagent Discovery and Analysis of Resulting Isolates

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. Peptide affinity reagents can be used for similar applications to antibodies, including sensing and therapeutics, but are more robust and able to perform in more extreme environments. Specific enrichment of peptide capture agents to a protein target of interest is enhanced using semi-automated sorting methods which improve binding and wash steps and therefore decrease the occurrence of false positive binders. A semi-automated sorting method is described herein for use with a commercial automated magnetic-activated cell sorting device with an unconstrained bacterial display sorting library expressing random 15-mer peptides. With slight modifications, these methods are extendable to other automated devices, other sorting libraries, and other organisms. A primary goal of this work is to provide a comprehensive methodology and expound the thought process applied in analyzing and minimizing the resulting pool of candidates. These techniques include analysis of on-cell binding using fluorescence-activated cell sorting (FACS), to assess affinity and specificity during sorting and in comparing individual candidates, and the analysis of peptide sequences to identify trends and consensus sequences for understanding and potentially improving the affinity to and specificity for the target of interest.

Biopanning bacterial display libraries is a proven technique for discovery of peptide affinity reagents, a robust alternative to antibodies. The semi-automated sorting method herein has streamlined biopanning to decrease the occurrence of false positives. Here we illustrate the thought process and techniques applied in evaluating candidates and minimizing downstream analysis.

Biopanning bacterial display libraries is a proven technique for peptide affinity reagent discovery for recognition of both biotic and abiotic targets. ...

Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ability to accurately measure the influx and efflux of Ca2+ from mitochondria is important for determining the role of mitochondrial Ca2+ handling in these processes. In this report, we present two methods for the measurement of mitochondrial Ca2+ handling in both isolated mitochondria and cultured cells. We first detail a plate reader-based platform for measuring mitochondrial Ca2+ uptake using the Ca2+ sensitive dye calcium green-5N. The plate reader-based format circumvents the need for specialized equipment, and the calcium green-5N dye is ideally suited for measuring Ca2+ from isolated tissue mitochondria. For our application, we describe the measurement of mitochondrial Ca2+ uptake in mitochondria isolated from mouse heart tissue; however, this procedure can be applied to measure mitochondrial Ca2+ uptake in mitochondria isolated from other tissues such as liver, skeletal muscle, and brain. Secondly, we describe a confocal microscopy-based assay for measurement of mitochondrial Ca2+ in permeabilized cells using the Ca2+ sensitive dye Rhod-2/AM and imaging using 2-dimensional laser-scanning microscopy. This permeabilization protocol eliminates cytosolic dye contamination, allowing for specific recording of changes in mitochondrial Ca2+. Moreover, laser-scanning microscopy allows for high frame rates to capture rapid changes in mitochondrial Ca2+ in response to various drugs or reagents applied in the external solution. This protocol can be applied to measure mitochondrial Ca2+ uptake in many cell types including primary cells such as cardiac myocytes and neurons, and immortalized cell lines.

Here, we present two protocols for the measurement of mitochondrial Ca2+ influx in isolated mitochondria and cultured cells. For isolated mitochondria, we detail a plate reader-based Ca2+ import assay using the Ca2+ sensitive dye calcium green-5N. For cultured cells, we describe a confocal microscopy method using the Ca2+ dye Rhod-2/AM.

Ca2+ handling by mitochondria is a critical function regulating both physiological and pathophysiological processes in a broad spectrum of cells. The ...

A Colorimetric Method for Measuring Iron Content in Plants

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content is rather low in all organisms, amounting in plants to about 0.009% of dry weight. To date, one of the most accurate methods for measuring iron concentration in plant tissues is flame absorption atomic spectroscopy. However, this approach is time-consuming and expensive and requires specific equipment not commonly found in plant laboratories. Therefore, a simpler, yet accurate method that can be routinely used is needed. The colorimetric Prussian Blue method is regularly used for qualitative iron staining in animal and plant histological sections. In this study, we adapted the Prussian Blue method for quantitative measurements of iron in tobacco leaves. We validated the accuracy of this method using both atomic spectroscopy and Prussian Blue staining to measure iron content in the same samples and found a linear regression (R2 = 0.988) between the two procedures. We conclude that the Prussian Blue method for quantitative iron measurement in plant tissues is precise, simple, and inexpensive. However, the linear regression presented here may not be appropriate for other plant species, due to potential interactions between the sample and the reagent. Establishment of a regression curve is thus needed for different plant species.

We present a simple and reliable protocol for measuring iron content in plant tissues using the colorimetric Prussian Blue method.

Iron, one of the most important micronutrients in living organisms, is involved in basic processes, such as respiration and photosynthesis. Iron content ...

An In Vitro Assay to Detect tRNA-Isopentenyl Transferase Activity

N6-isopentenyladenosine RNA modifications are functionally diverse and highly conserved among prokaryotes and eukaryotes. One of the most highly conserved N6-isopentenyladenosine modifications occurs at the A37 position in a subset of tRNAs. This modification improves translation efficiency and fidelity by increasing the affinity of the tRNA for the ribosome. Mutation of enzymes responsible for this modification in eukaryotes are associated with several disease states, including mitochondrial dysfunction and cancer. Therefore, understanding the substrate specificity and biochemical activities of these enzymes is important for understanding of normal and pathologic eukaryotic biology. A diverse array of methods has been employed to characterize i6A modifications. Herein is described a direct approach for the detection of isopentenylation by Mod5. This method utilizes incubation of RNAs with a recombinant isopentenyl transferase, followed by RNase T1 digestion, and 1-dimensional gel electrophoresis analysis to detect i6A modifications. In addition, the potential adaptability of this protocol to characterize other RNA-modifying enzymes is discussed.

An In Vitro Assay to Detect tRNA-Isopentenyl Transferase Activity

Here, we describe a protocol for the biochemical characterization of the yeast RNA-modifying enzyme, Mod5, and discuss how this protocol could be applied to other RNA-modifying enzymes.

N6-isopentenyladenosine RNA modifications are functionally diverse and highly conserved among prokaryotes and eukaryotes. One of the most highly conserved ...

How to Quantify the Fraction of Photoactivated Fluorescent Proteins in Bulk and in Live Cells

Photoactivatable and -convertible fluorescent proteins (PA-FPs) have been used in fluorescence live-cell microscopy for analyzing the dynamics of cells and protein ensembles. Thus far, no method has been available to quantify in bulk and in live cells how many of the PA-FPs expressed are photoactivated to fluoresce.
Here, we present a protocol involving internal rulers, i.e., genetically coupled spectrally distinct (photoactivatable) fluorescent proteins, to ratiometrically quantify the fraction of all PA-FPs expressed in a cell that are switched on to be fluorescent. Using this protocol, we show that different modes of photoactivation yielded different photoactivation efficiencies. Short high-power photoactivation with a confocal laser scanning microscope (CLSM) resulted in up to four times lower photoactivation efficiency than hundreds of low-level exposures applied by CLSM or a short pulse applied by widefield illumination. While the protocol has been exemplified here for (PA-)GFP and (PA-)Cherry, it can in principle be applied to any spectrally distinct photoactivatable or photoconvertible fluorescent protein pair and any experimental set-up.

Here, we present a protocol that involves genetically coupled spectrally distinct photoactivatable and fluorescent proteins. These fluorescent protein chimeras permit quantification of the PA-FP fraction that is photoactivated to be fluorescent, i.e., the photoactivation efficiency. The protocol reveals that different modes of photoactivation yield different photoactivation efficiencies.

Photoactivatable and -convertible fluorescent proteins (PA-FPs) have been used in fluorescence live-cell microscopy for analyzing the dynamics of cells ...

Using High Content Imaging to Quantify Target Engagement in Adherent Cells

Quantitating the interaction of small molecules with their intended protein target is critical for drug development, target validation and chemical probe validation. Methods that measure this phenomenon without modification of the protein target or small molecule are particularly valuable though technically challenging. The cellular thermal shift assay (CETSA) is one technique to monitor target engagement in living cells. Here, we describe an adaptation of the original CETSA protocol, which allows for high throughput measurements while retaining subcellular localization at the single cell level. We believe this protocol offers important advances to the application of CETSA for in-depth characterization of compound-target interaction, especially in heterogeneous populations of cells.

Measurements of drug target engagement are central to effective drug development and chemical probe validation. Here, we detail a protocol for measuring drug-target engagement using high content imaging in a microplate-compatible adaption of the cellular thermal shift assay (CETSA).

Quantitating the interaction of small molecules with their intended protein target is critical for drug development, target validation and chemical probe ...

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for the targeted expression of genetically encoded calcium sensors. This protocol outlines the use of these sensors for the in vivo imaging of calcium dynamics in targeted cells, specifically the body wall muscles of the worms. By utilizing the co-expression of presynaptic channelrhodopsin, stimulation of acetylcholine release from excitatory motor neurons can be induced using blue light pulses, resulting in muscle depolarization and reproducible changes in cytoplasmic calcium levels. Two worm immobilization techniques are discussed with varying levels of difficulty. Comparison of these techniques demonstrates that both approaches preserve the physiology of the neuromuscular junction and allow for the reproducible quantification of calcium transients. By pairing optogenetics and functional calcium imaging, changes in postsynaptic calcium handling and homeostasis can be evaluated in a variety of mutant backgrounds. Data presented validates both immobilization techniques and specifically examines the roles of the C. elegans sarco(endo)plasmic reticular calcium ATPase and the calcium-activated BK potassium channel in the body wall muscle calcium regulation.

In Vivo Calcium Imaging in C. elegans Body Wall Muscles

This method provides a way to couple optogenetics and genetically encoded calcium sensors to image baseline cytosolic calcium levels and changes in evoked calcium transients in the body wall muscles of the model organism C. elegans.

The model organism C. elegans provides an excellent system to perform in vivo calcium imaging. Its transparent body and genetic manipulability allow for ...

Real-Time, Semi-Automated Fluorescent Measurement of the Airway Surface Liquid pH of Primary Human Airway Epithelial Cells

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) hydration, mucus viscosity and activity of antimicrobial peptides, key parameters involved in innate defense of the lungs. This is of primary relevance in the field of chronic respiratory diseases such as cystic fibrosis (CF) where these parameters are dysregulated. While different groups have studied ASL pH both in vivo and in vitro, their methods report a relatively wide range of ASL pH values and even contradictory findings regarding any pH differences between non-CF and CF cells. Furthermore, their protocols do not always provide enough details in order to ensure reproducibility, most are low throughput and require expensive equipment or specialized knowledge to implement, making them difficult to establish in most labs. Here we describe a semi-automated fluorescent plate reader assay that enables the real-time measurement of ASL pH under thin film conditions that more closely resemble the in vivo situation. This technique allows for stable measurements for many hours from multiple airway cultures simultaneously and, importantly, dynamic changes in ASL pH in response to agonists and inhibitors can be monitored. To achieve this, the ASL of fully differentiated primary human airway epithelial cells (hAECs) are stained overnight with a pH-sensitive dye in order to allow for the reabsorption of the excess fluid to ensure thin film conditions. After fluorescence is monitored in the presence or absence of agonists, pH calibration is performed in situ to correct for volume and dye concentration. The method described provides the required controls to make stable and reproducible ASL pH measurements, which ultimately could be used as a drug discovery platform for personalized medicine, as well as adapted to other epithelial tissues and experimental conditions, such as inflammatory and/or host-pathogen models.

We present a protocol to make dynamic measurements of the airway surface liquid pH under thin film conditions using a plate-reader.

In recent years, the importance of mucosal surface pH in the airways has been highlighted by its ability to regulate airway surface liquid (ASL) ...

Induction of Eryptosis in Red Blood Cells Using a Calcium Ionophore

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic cells is the loss of compositional asymmetry of the cell membrane, leading to the translocation of phosphatidylserine to the membrane outer leaflet. This process is triggered by increased intracellular concentration of Ca2+, which activates scramblase, an enzyme that facilitates bidirectional movement of phospholipids between membrane leaflets. Given the importance of eryptosis in various diseased conditions, there have been efforts to induce eryptosis in vitro. Such efforts have generally relied on the calcium ionophore, ionomycin, to enhance intracellular Ca2+ concentration and induce eryptosis. However, many discrepancies have been reported in the literature regarding the procedure for inducing eryptosis using ionomycin. Herein, we report a step-by-step protocol for ionomycin-induced eryptosis in human erythrocytes. We focus on important steps in the procedure including the ionophore concentration, incubation time, and glucose depletion, and provide representative result. This protocol can be used to reproducibly induce eryptosis in the laboratory.

A protocol for the induction of eryptosis, programmed cell death in erythrocytes, using the calcium ionophore, ionomycin, is provided. Successful eryptosis is evaluated by monitoring the localization phosphatidylserine in the membrane outer leaflet. Factors affecting the success of the protocol have been examined and optimal conditions provided.

Eryptosis, erythrocyte programmed cell death, occurs in a number of hematological diseases and during injury to erythrocytes. A hallmark of eryptotic ...