We thank Irene Hernandez, Trudy Holyst, Dr. Hartmut Weiler (Versiti Blood Research Institute), and Dr. Leggy Arnold (University of Wisconsin-Milwaukee) for providing space and indirect support of this project, and Dr. John McCorvy (Medical College of Wisconsin) for pertinent advice. We thank the National Heart, Lung, and Blood Institute (R15HL127636), the U.S. Dept. of Defense (W81XWH22101), and the National Science Foundation (2223225) for grant support.

All media exchanges/additions made in steps 1 and 2 of the following protocol are performed in a sterile hood. Unless otherwise noted, all plasticware used in the sterile hood must be purchased sterilized and sealed or sterilized appropriately via autoclave.
1. Initiation of EA.hy926 cell line
<ol>
	<li>Acquire EA.hy926 cells.</li>
	<li>Store vial(s) of cells in the vapor phase of a liquid nitrogen tank.</li>
	<li>Prepare DMEM complete medium by first warming DMEM, FBS, and pen/strep (see Table of Materials for specific details) in a 37 &#176;C water bath for 15-30 min. Add 50 mL of FBS and 1 mL of pen/strep to 500 mL of DMEM. Gently mix the solution by inversion. 
	NOTE: Mixing too vigorously causes excessive amounts of bubbles.</li>
	<li>Remove one vial of cells from liquid nitrogen and warm in a 37 &#176;C water bath for 5 min.</li>
	<li>Sterilize the vial of cells and bottle of DMEM complete medium by spraying with alcohol, then move to the sterile hood.</li>
	<li>Use a 1,000 &#181;L pipette to carefully transfer the cells to a 15 mL sterile centrifuge tube. Add 1 mL of DMEM to the vial to rinse and add to the centrifuge tube.</li>
	<li>Make a pellet of cells using a centrifuge (150 &#215; g, 5 min, 22 &#176;C). Remove the medium from the centrifuge tube by aspiration; be careful not to disturb the cell pellet.</li>
	<li>Add 3 mL of DMEM complete medium to the centrifuge tube and gently break apart the pellet by mixing the cell suspension using a 1,000 &#181;L pipette. 
	NOTE: Causing bubbles by mixing too vigorously should be avoided as this could activate or damage the cells.</li>
	<li>Mix 10 &#181;L of the cell suspension with 10 &#181;L of trypan blue in a microcentrifuge tube using a pipette with either 20 &#181;L or 200 &#181;L tips. Add 10 &#181;L of the cell suspension/trypan blue mixture to a hemocytometer and count the cells. 
	NOTE: Using 0.1-10 &#181;L pipette tips can shear the cells. Tips of this size should not be used while handling the cell suspension.</li>
	<li>Add DMEM complete medium to the cell suspension to make a density of 66,000 cells/mL using either a 10 or 25 mL serological pipette. Pipette 15&#160;mL of the cell suspension into T-75 culture flask(s). Ensure equal dispersion of the cell suspension by moving the flask north to south and east to west.</li>
	<li>Incubate the cells until they reach confluency (typically after 4-6 days), as estimated by examination under a microscope. At this point, use the cells in the experiment (beginning with section 2) or seed them into new flasks for expansion to prepare frozen stocks (at a recommended seeding density of 1,000,000 cells in 15 mL of complete medium per T-75 flask). 
	NOTE: Cells can be grown in the culture flasks for as long as 3 weeks without a noticeable difference in shape, health, or performance in the assay. Complete medium should be exchanged every 3-4 days.</li>
</ol>
2. Addition of EA.hy926 cells to 96-well plates
NOTE: Cells from one confluent T-75 culture flask can be used to prepare two or three assay plates or optionally expanded into as many as 15 fresh T-75 flasks. At least five assay plates can be screened using the protocol in sections 4 and 5 in a normal workday. The following instructions describe preparing and testing one assay plate, but additional plates can be prepared by repeating steps 2.2, 2.7, 2.12, and 3.2 to prepare the desired number of assay plates. Most commonly, four assay plates are prepared per day to measure concentration-responses with up to 16 compounds, which requires two T-75 flasks with confluent cells.
<ol>
	<li>Warm DMEM complete medium in a 37 &#176;C water bath for 15-30 min. Sterilize the bottle of DMEM complete medium by spraying it with alcohol, then move it to the sterile hood.</li>
	<li>Add 100 &#181;L of a sterilized 0.4% gelatin solution to all wells of a black-walled, 96-well, clear-bottom plate with a lid. Incubate the plate with the gelatin solution in an incubator (37 &#176;C, 5% CO2) for 30 min.</li>
	<li>Confirm that cells in a T-75 culture flask are 80-100% confluent using an inverted microscope. Remove DMEM complete medium from the T-75 flask via aspiration.</li>
	<li>Wash the cells in the flask with 10 mL PBS and gently move the flask North to South and East to West. Remove the PBS from the flask via aspiration.</li>
	<li>Add 5 mL of cell dissociation solution to the flask and incubate the flask&#160;in an incubator (37 &#176;C, 5% CO2) for ~12 min. Tap the flask after 6 min to help facilitate dissociation. 
	NOTE: Incubating cells for longer than 15 min with the referenced cell dissociation agent causes cells to begin to clump together, which makes obtaining an accurate cell count more difficult and may interfere with the assay.</li>
	<li>Add 5 mL of the prewarmed DMEM complete medium to the flask and gently mix into the cell dissociation solution/cell suspension. Using a 10 mL serological pipette, add the cell suspension into a sterile 50 mL centrifuge tube. Make a pellet of cells using a centrifuge (150 &#215; g, 5 min, 22&#160;&#176;C).</li>
	<li>While the pellet is being formed, remove the gelatin solution from the 96-well plate via aspiration using a Pasteur pipette connected to a vacuum. Remove the plate lid from the time of gelatin removal until the solution is ready to be plated. 
	NOTE: A multichannel pipette or sterile manifold connected to a vacuum may optionally be used.</li>
	<li>Remove the medium from the centrifuge tube by aspiration using a Pasteur pipette connected to a vacuum or a serological pipette; be careful not to disturb the cell pellet.</li>
	<li>Add fresh DMEM complete medium to the centrifuge tube using a 5 or 10 mL serological pipette. Gently break up the cell pellet by mixing the DMEM complete medium with a 1,000 &#181;L pipette or a 5 mL serological pipette. 
	NOTE: Eight milliliters of DMEM are typically used, but higher volumes can be added to make cell counting easier.</li>
	<li>Mix 10 &#181;L of the cell suspension with 10 &#181;L of trypan blue in a microcentrifuge tube using a pipette with either 20 &#181;L or 200 &#181;L tips. Add 10 &#181;L of the cell suspension/trypan blue mixture to a hemocytometer and count the cells.</li>
	<li>Add DMEM complete media to the cell suspension to make a density of 600,000 cells/mL using either a 10 or 25 mL serological pipette. This will give a final density of 60,000 cells/well in a 96-well plate (determined to be the optimal density by experimentation). 
	NOTE: A normal successful cell culture will yield between 12,000,000 and 15,000,000 cells, which can seed two to three 96-well plates.</li>
	<li>Mix the cell suspension by gently inverting the centrifuge tube and add it to a 50 mL multi-channel pipette reservoir. Mixing the suspension in the reservoir every 30 s by gently rocking the reservoir side to side, add 100 &#181;L of the cell suspension to each well using a multi-channel pipette. Replace the transparent plate cover and gently shake the plate by sliding it on the surface of the sterile hood from North to South and East to West to ensure even cell distribution. Incubate the plate in the incubator (37 &#176;C, 5% CO2) for 16-24 h. 
	NOTE: Additional cell suspension can be used to prepare additional plates, or steps 1.10-1.11 can be repeated to continue the cell line.</li>
</ol>
3. Calcium mobilization assay preparation
<ol>
	<li>Prepare assay reagents.
	<ol>
		<li>Prepare probenecid solution (250 mM, aqueous) by adding 36 mg of probenecid to a microcentrifuge tube and dissolving it in 0.6 M NaOH (500 &#181;L). Vortex the solution. 
		NOTE: Probenecid improves the retention of dye in the cell by inhibiting organic ion transporters75,76,77.</li>
		<li>To make HBSS-HEPES assay buffer, add HEPES (1.19 g) to a 500 mL bottle of Ca/Mg/phenol red-free HBSS (making a 10 mM solution of HEPES). Supplement the buffer with MgCl2 (1 M aqueous solution, 500 &#181;L) and CaCl2 (1 M aqueous solution, 500 &#181;L). Mix the solution and store in a refrigerator at 5 &#176;C when not in use.
		<ol>
			<li>For each 96-well plate, add 50 mL of the HBSS-HEPES assay buffer to a 50 mL centrifuge tube, supplement with 500 &#181;L of the probenecid solution, and mix well. Warm the buffer to room temperature prior to use.</li>
		</ol>
		</li>
		<li>Make a 10% Pluronic F-127 solution in DMSO by adding Pluronic F-127 (20 mg) to a microcentrifuge tube or HPLC&#160;vial and dissolving it with 200 &#181;L of DMSO. 
		NOTE: This solution can be stored at room temperature and is sufficient for about 30 plates. Pluronic F-127 dissolves very slowly at room temperature and should be gently heated to facilitate dissolution.</li>
		<li>Prepare the dye loading buffer by first adding 24 &#181;L of DMSO to a vial of Fluo-4/AM (50 &#181;g) to dissolve the dye. In a foil-wrapped 15&#160;mL centrifuge tube, add 6 mL of HBSS-HEPES assay buffer and supplement with 6 &#181;L of 10% Pluronic F-127 and 6 &#181;L of Fluo-4/AM solution. Briefly vortex the solution and allow to sit at room temperature, out of light, for 10 min. 
		NOTE: To prevent photobleaching of Fluo-4, keep the dye out of light by wrapping aluminum foil around the centrifuge tube containing the dye solution and the lid of the assay plate.</li>
	</ol>
	</li>
	<li>Prepare assay plates.
	<ol>
		<li>Remove the assay plate from the incubator and confirm confluency using an inverted microscope. Manually remove the DMEM complete media by flicking it into a sink and blotting the top of the plate with a paper towel. 
		NOTE: Medium is sufficiently removed by holding the plate horizontally and flicking, followed by rotating the plate 180&#176;and repeating.</li>
		<li>Add HBSS-HEPES assay buffer plus probenecid solution to a 50 mL multi-channel pipette solvent reservoir. 
		NOTE: This solution will be used for later washing steps and should be put aside and covered with aluminum foil until needed.</li>
		<li>Add 100 &#181;L of the HBSS-HEPES assay buffer plus probenecid solution to each well with a multi-channel pipette and allow the cells to sit in the presence of probenecid for 5 min. Remove the HBSS-HEPES assay buffer by flicking into the sink in the same manner as in step 3.2.1.</li>
		<li>Add dye loading buffer to a separate 50 mL multi-channel pipette solvent reservoir. Add 50 &#181;L of the dye loading buffer to each well of the assay plate using a multi-channel pipette. Incubate the plate for 45 min (37 &#176;C, 5% CO2).</li>
		<li>While the plate is in the incubator, prepare the agonist/antagonist solutions.
		<ol>
			<li>Antagonist solutions are stored as 31.6 mM solutions in DMSO. In a PCR tube, dilute 2 &#181;L of the stock solution with 38 &#181;L of 0.1% BSA/water to obtain a 1.58 mM solution. 2 &#181;L of this solution will give a final concentration of 31.6 &#181;M in the assay plate. Dilute 4 &#181;L of the 1.58 mM antagonist solution with 36 &#181;L of 5% DMSO/water (no BSA addition), and prepare the remaining concentrations through serial dilutions.</li>
			<li>Prepare a 108.3 &#181;M (16.7x) solution of TFLLRN-NH2 in HBSS-HEPES assay buffer, which will give a final concentration of 6.5 &#181;M when 6 &#181;L is added to each well of the assay plate. For example, dissolve a 2 mg aliquot of TFLLRN-NH2 with 5.244 mL of HBSS-HEPES assay buffer to give a 0.5 mM stock solution. Mix 1.083 mL of this solution with 3.917 mL of HBSS-HEPES assay buffer to give a 108.3 &#181;M solution. 
			NOTE: TFLLRN-NH2 solutions should be stored in a -20 &#176;C freezer when not in use.</li>
		</ol>
		</li>
		<li>Remove the plate from the incubator and ensure dye uptake by using a fluorescence microscope at a suitable setting (typically for green fluorescent protein, see Figure 1 for an example of Fluo-4 successfully loaded into cells). Remove the dye loading buffer by flicking the plate in the same manner as step 3.2.1.</li>
		<li>Wash the cells 2x with 50 &#181;L of the HBSS-HEPES assay buffer plus probenecid solution (from step 3.2.3) in each well (remove by flicking medium into the sink).</li>
		<li>Add 92 &#181;L of HBSS-HEPES assay buffer to each well and again view the cells with a fluorescence microscope to ensure excess dye has been fully removed, and that cells remain adherent.</li>
		<li>Use a multi-channel pipette to add 2 &#181;L of the antagonist solutions and vehicle to appropriate wells (see Table 1 for a representative plate map).</li>
	</ol>
	</li>
</ol>
4. Performing the calcium mobilization assay
<ol>
	<li>Set the plate reader as follows: temperature: 37 &#176;C; excitation wavelength: 485 nm; emission wavelength: 525 nm; measurement height: 7.8 mm (optimized); number of flashes: 100; number of repeats: 20.</li>
	<li>Incubate the plate for 15 min in the plate reader at 37 &#176;C. Measure the background fluorescence in columns 1 and 2 (five scans per well).</li>
	<li>Eject the plate from the plate reader and quickly add 6 &#181;L of agonist solution to each well in columns 1 and 2 from an 8-tube PCR strip using a multi-channel pipette. 
	NOTE: For this step, we use an electronic 8-channel pipette with 0.1-10 &#181;L pipette tips.</li>
	<li>Measure the change in fluorescence as calcium mobilization occurs in the cells. Perform 20 scans of each well according to the settings above. 
	NOTE: Scanning each well in two columns 20x each takes approximately 5 min (i.e., ~15 s between scans of each well).</li>
	<li>Repeat steps 4.2-4.4 for columns 3-12 in two column increments.</li>
</ol>
5. Data analysis
<ol>
	<li>Export data from the assay as separate spreadsheets for each two-column group.</li>
	<li>Find the average of the basal fluorescence readings by using the function AVG(first scan: last scan). Do this for every well in both columns.</li>
	<li>Find the maximum fluorescence after the addition of agonist by using the function MAX(first scan: last scan).</li>
	<li>Calculate the change in fluorescence by subtracting the basal fluorescence from the maximum fluorescence.</li>
	<li>Subtract the change in fluorescence calculated for the negative control (addition of vehicle without agonist) from each well in the same column.</li>
	<li>Find the relative (normalized) change in fluorescence by dividing the change in fluorescence in each well by the change in fluorescence in the vehicle (0.1% DMSO/water) + 6.5 &#181;M TFLLRN-NH2 well.</li>
	<li>Copy the non-control values, as percentages, into a statistics and graphing software program. If using the referenced software, choose the XY table option, with the Numbers chosen as the X-axis and replicate values in side-by-side subcolumns as the Y-axis.</li>
	<li>Use the program to plot concentration-response curves (CRCs) from the data. If using the referenced software, choose the log(inhibitor) vs. response - Variable slope (four parameters) option.</li>
</ol>

Studying Protease-Activated Receptor Ligands in Endothelial Cells

C.D. is inventor on patents involving parmodulins and is the founder and a shareholder of Function Therapeutics, Inc., which is developing parmodulins for clinical use.

While the previously reported protocol72 was generally reliable and allowed us to identify a new lead parmodulin, NRD-21,62 a more efficient protocol was desired. The assay was further compromised during the supply shortage caused by the COVID-19 pandemic. Acquiring tips for the automated liquid handler became difficult, and attempting to wash, sterilize, and reuse the tips produced CRCs with significant variance. This facilitated an urgent series of experiments designed to...

Protease-activated receptors (PARs)1,2,3 are a subfamily of class A G protein-coupled receptors (GPCRs) that are expressed in a variety of cell types, including platelets and endothelial cells4,5,6,7. Unlike the majority of GPCRs, PARs have a unique intramolecular mode of activation. Most GPCRs are activated by soluble ligands interacting with a distinct binding pocket, but PARs are activated by the proteolytic cleavage of the N-terminus, which results in a new tethered ligand that can interact with the extracellular loop 2 domain on the surface of a cell6,8,9. This interaction activates the receptor and can initiate several signaling pathways, promoting effects such as inflammation and platelet activation4,10,11,12. Different proteases can activate PARs through cleavage at unique sites on the N-terminus, revealing different tethered ligands (TL) that stabilize receptor conformations, which initiate different signaling pathways9,13,14,15. For example, in the most well-studied member of the subfamily, PAR1, cleavage by thrombin is used to support numerous biological processes, including platelet activation and leukocyte recruitment to the endothelium, but can lead to deleterious effects when the receptor is overexpressed or overactivated4,16,17,18,19,20,21. Conversely, cleavage by activated protein C (aPC) can promote anti-inflammatory effects and maintenance of endothelial barriers15,22,23,24,25,26,27,28,29. PARs can also be activated by peptide analogs of the TLs in an intermolecular fashion13,30,31. These peptides are routinely used to measure PAR inhibition (modulation) in place of PAR-targeting proteases, and they are used in this protocol.
Numerous disorders are associated with pathological PAR1 signaling, including sepsis22,32, cardiovascular disease33,34,35,36,37,38, kidney disease39,40,41,42, sickle cell disease43, fibrosis44, osteoporosis and osteoarthritis45,46, neurodegeneration47,48,49,50,51, and cancer52,53,54,55,56,57,58,59. Antagonists of PAR1 have been studied since the 1990s as antiplatelet agents for cardiovascular disease, and the growing list of diseases associated with the receptor necessitates the identification of novel ligands for use as biological probes (tool compounds) or as potential therapeutics. Currently, there is only one FDA-approved PAR1 antagonist, vorapaxar, which is used to treat coronary artery disease in high-risk patients34,36,37,60. An alternative PAR1 antagonist, the pepducin PZ-128, completed a successful phase II study to prevent thrombosis in cardiac catheterization patients38. The Dockendorff group has focused on the medicinal chemistry and pharmacology of a separate class of small molecules, PAR1 ligands known as parmodulins61,62. Unlike reported PAR1 antagonists such as vorapaxar, parmodulins are allosteric, biased modulators of PAR1 that selectively block the G&#945;q pathway while promoting cytoprotective effects similar to aPC. Unlike potent orthosteric PAR1 antagonists such as vorapaxar, published parmodulins are also reversible63,64,65.
Initially, parmodulins were identified by Flaumenhaft and coworkers for&#160;their ability to inhibit P-selectin expression or granule secretion in platelets61,66. However, an alternative method was required to study the effects of parmodulins on endothelial cells. One common method to monitor GPCR-related signaling is to measure intracellular Ca2+ mobilization, an important secondary messenger that can be measured using a suitable intracellular calcium-binding dye67,68. Substantial evidence has been provided showing that calcium mobilization induced by PAR1 is through the activation of G&#945;q69,70. Once activated by its tethered ligand (or a suitable exogenous ligand), PAR1 undergoes a conformational change which causes guanosine diphosphate (GDP) bound to the G&#945;q subunit to be replaced by guanosine triphosphate (GTP)68. The G&#945;q subunit then activates phospholipase C&#946; (PLC-&#946;), which catalyzes the hydrolysis of phosphatidylinositol 4,5 bisphosphate (PIP2), forming 1,4,5-inositol triphosphate (IP3) and diacylglycerol (DAG). Finally, IP3 binds to IP3-sensitive Ca2+ channels in the membrane of the endoplasmic reticulum, allowing Ca2+ to be released into the cytoplasm, where it can bind to Ca2+-dependent fluorescent dyes, such as Fluo-4, that are added to the cells71.&#160;This process occurs within seconds and can increase the concentration of Ca2+ 100-fold, leading to a drastic change in the amount of calcium-bound dye and a robust fluorescence signal.
In 2018, the Dockendorff group disclosed a medium-throughput Ca2+ mobilization assay that could be used to identify antagonists of the G&#945;q pathway of PAR172. The assay used&#160;EA.hy92673, a hybrid human endothelial cell line, which can be used for multiple passages without a noticeable change in PAR1 expression, and is established for in vitro measurements of cytoprotective effects.&#160;
The original protocol used EA.hy926 cells in 96-well plates and loaded with Fluo-4/AM dye, which was chosen due to its intense fluorescence at 488 nm and high cell permeability. Once the dye was loaded into the cells, lengthy washing steps were performed with an automated 8-channel liquid handler (faster methods of liquid handling, such as a 96-channel washer, were inaccessible). The reproducibility of this assay was superior to that without the careful, automated robotic media changes. Antagonists were then incubated with the cells, PAR1 was activated through the sequential addition of a selective agonist (16 wells at a time), and changes in fluorescence resulting from calcium mobilization and dye binding were measured to determine activity.
While this protocol allows for the measurement of PAR1-mediated calcium mobilization, it is limited by the time required to assay each 96-well plate. Long experiment times are problematic not only because the number of compounds that can be screened each day is limited, but also because dye efflux occurs over time, narrowing the assay window by increasing the basal fluorescence. One contributor to the long experiment time is the use of an 8-channel liquid handler for plate washing, which adds over 30 min to each experiment. The required tips also became difficult to obtain due to supply chain problems. Here an updated protocol for the PAR-mediated calcium mobilization assay that does not require a liquid handler, and therefore can be run in higher throughput, is reported. This protocol should also be suitable for measuring signaling with other GPCRs that lead to intracellular calcium mobilization.&#160;This updated plate reader protocol&#160;is ideal for academic and small industrial labs that do not have the resources for expensive cell imaging instruments but have a need to rapidly screen numerous compounds. For an example of a calcium mobilization assay using a plate imager, see Caers et al.74.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Cell Culture Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Adherent EA.hy926 cells</td>
			<td>ATCC</td>
			<td>CRL-2922</td>
			<td></td>
		</tr>
		<tr>
			<td>CellStripper cell dissociation reagent</td>
			<td>Corning</td>
			<td>25-056-CI</td>
			<td>Trypsin can optionally be used, but should definitely be avoided with PAR2 assays.</td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium (DMEM) w/phenol red</td>
			<td>Corning</td>
			<td>10-013-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS)</td>
			<td>Avantor</td>
			<td>97068-091</td>
			<td></td>
		</tr>
		<tr>
			<td>Gelatin from porcine skin</td>
			<td>MilliporeSigma</td>
			<td>G2500</td>
			<td>Use to make an aqueous 0.4% (w/v) solution with deionized water. Autoclave before use to sterilize.</td>
		</tr>
		<tr>
			<td>Pen/Strep (100X)</td>
			<td>Corning</td>
			<td>30-002-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS)</td>
			<td>Corning</td>
			<td>21-040-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypan Blue (0.4% w/v)</td>
			<td>Corning</td>
			<td>25-900-CI</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Mobilization Reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES)</td>
			<td>Thermo</td>
			<td>172571000</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine serum albumin (BSA)</td>
			<td>Avantor</td>
			<td>97061-420</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Thermo</td>
			<td>42352-0250</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide</td>
			<td>Thermo</td>
			<td>J66650-AD</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluo-4/AM</td>
			<td>Invitrogen</td>
			<td>F14201</td>
			<td></td>
		</tr>
		<tr>
			<td>Hank&#39;s balanced salt solution (Ca/Mg/phenol-red free)</td>
			<td>Corning</td>
			<td>21-022-CV</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium chloride hexahydrate</td>
			<td>MilliporeSigma</td>
			<td>M2393</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F-127 (Poloxamer 407)</td>
			<td>Spectrum Chemical</td>
			<td>P1166</td>
			<td></td>
		</tr>
		<tr>
			<td>Probenecid</td>
			<td>TCI America</td>
			<td>P1975</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>VWR International</td>
			<td>BDH9292</td>
			<td></td>
		</tr>
		<tr>
			<td>TFLLRN-NH2 (TFA salt)</td>
			<td></td>
			<td></td>
			<td>Prepared by Trudy Holyst at the Versiti Blood Research Institute</td>
		</tr>
		<tr>
			<td>Materials</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>96-well culture-treated, black-walled, clear bottom assay plate</td>
			<td>Corning</td>
			<td>3603</td>
			<td>with transparent lids</td>
		</tr>
		<tr>
			<td>Centrifuge tube, 15 mL</td>
			<td>Avantor</td>
			<td>89039-664</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge tube, 50 mL</td>
			<td>Avantor</td>
			<td>89039-656</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture flask, T-75</td>
			<td>Corning</td>
			<td>353136</td>
			<td>tissue culture treated</td>
		</tr>
		<tr>
			<td>Disposable reagent reservoir, 50 mL</td>
			<td>Corning</td>
			<td>RES-V-50-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Enspire plate reader</td>
			<td>Perkin Elmer</td>
			<td></td>
			<td>Discontinued</td>
		</tr>
		<tr>
			<td>Microcentrifuge tube, 1.5 mL</td>
			<td>Avantor</td>
			<td>20170-038</td>
			<td></td>
		</tr>
		<tr>
			<td>Pasteur pipette, 9&#34;</td>
			<td>Fisher</td>
			<td>13-678-6B</td>
			<td>must be sterilized</td>
		</tr>
		<tr>
			<td>PCR tube strip with separate flat cap strips</td>
			<td>Avantor</td>
			<td>76318-802</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipette tips, 20 &#181;L</td>
			<td>Biotix</td>
			<td>63300042</td>
			<td>sterile, filtered tips</td>
		</tr>
		<tr>
			<td>Pipette tips, 200 &#181;L</td>
			<td>Biotix</td>
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			<td>Serological pipette, 10 mL</td>
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			<td>Serological pipette, 25 mL</td>
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characterizing modulators of protease-activated receptors with a calcium mobilization assay using a plate reader

Changes in calcium concentration in cells are rapidly monitored in a high-throughput fashion with the use of intracellular, fluorescent, calcium-binding dyes and imaging instruments that can measure fluorescent emissions from up to 1,536 wells simultaneously. However, these instruments are much more expensive and can be challenging to maintain relative to widely available plate readers that scan wells individually. Described here is an optimized plate reader assay for use with an endothelial cell line (EA.hy926) to measure the protease-activated receptor (PAR)-driven activation of G&#945;q signaling and subsequent calcium mobilization using the calcium-binding dye Fluo-4. This assay has been used to characterize a range of PAR ligands, including the allosteric PAR1-targeting anti-inflammatory "parmodulin" ligands identified in the Dockendorff lab. This protocol obviates the need for an automated liquid handler and permits the medium-throughput screening of PAR ligands in 96-well plates and should be applicable to the study of other receptors that initiate calcium mobilization.

The purpose of this assay is generally to produce concentration-response curves (CRCs) for three to four new parmodulins. On each assay plate, an additional CRC for a known compound, such as NRD-21, is often generated that acts as a quality check for the experiment due to its known IC50. To generate CRCs, a plate map such as the one depicted in Table 1 should be planned. If single-point concentration-responses are desired instead, compounds at 10 &#181;M final concentrations (or other preferre...

Author Spotlight: Developing Parmodulins to Target Protease-Activated Receptors for Inflammation Control

An improved protocol for a calcium mobilization assay with endothelial cells, used to identify ligands of protease-activated receptors (PARs), has been developed. The new protocol reduces total assay time by 90-120 min and yields reproducible concentration-response curves.

Characterizing Modulators of Protease-Activated Receptors with a Calcium Mobilization Assay Using a Plate Reader

Changes in calcium concentration in cells are rapidly monitored in a high-throughput fashion with the use of intracellular, fluorescent, calcium-binding ...

characterizing-modulators-protease-activated-receptors-with-calcium

Research

JoVE Journal

Biochemistry

363 Views.  Function Therapeutics. An improved protocol for a calcium mobilization assay with endothelial cells, used to identify ligands of protease-activated receptors (PARs), has been developed. The new protocol reduces total assay time by 90-120 min and yields reproducible concentration-response curves.

Video: Author Spotlight: Developing Parmodulins to Target Protease-Activated Receptors for Inflammation Control

Quantification of Bacterial Histidine Kinase Autophosphorylation Using a Nitrocellulose Binding Assay

We demonstrate a useful method for quantifying autophosphorylation of purified bacterial histidine kinases. Histidine kinases are known for their involvement in two-component signal transduction, a ubiquitous system through which bacteria sense and respond to environmental stimuli. Two-component signaling features autophosphorylation of a histidine kinase, followed by phosphotransfer to the receiver domain of a response regulator protein, which ultimately leads to an output response. Autophosphorylation of the histidine kinase is responsive to the presence of a cognate environmental stimulus, thereby giving bacteria a means to detect and respond to changes in the environment. Despite their importance in bacterial biology, histidine kinases remain poorly understood due to the inherent lability of phosphohistidine. Conventional methods for studying these proteins, such as SDS-PAGE autoradiography, have significant shortcomings. We have developed a nitrocellulose binding assay that can be used to characterize histidine kinases. The protocol for this assay is simple and easy to execute. Our method is higher throughput, less time-consuming, and offers a greater dynamic range than SDS-PAGE autoradiography.

We report and demonstrate an optimized nitrocellulose binding assay that can be used to quantify autophosphorylation of purified bacterial histidine kinases. Our method has several advantages over traditional SDS-PAGE based techniques, providing a valuable alternative for characterizing these important proteins.

We demonstrate a useful method for quantifying autophosphorylation of purified bacterial histidine kinases. Histidine kinases are known for their ...

Hydrolysis of a Ni-Schiff-Base Complex Using Conditions Suitable for Retention of Acid-labile Protecting Groups

Unnatural amino acids, amino acids containing side-chain functionalities not commonly seen in nature, are increasingly found in synthetic peptide sequences. Synthesis of some unnatural amino acids often includes the use of a precursor consisting of a Schiff-base stabilized by a nickel cation. Unnatural side-chains can be installed on an amino acid backbone found in this Schiff-base complex. The resulting unnatural amino acid can then be isolated from this complex using hydrolysis of the Schiff-base, typically by employing reflux in strongly acidic solution. These highly acidic conditions may remove acid-labile side-chain protecting groups necessary for the unnatural amino acids to be used in microwave-assisted solid-phase peptide synthesis. In this work, we present an efficient hydrolysis and subsequent Fmoc protection of an amino acid isolated from a Ni-Schiff base complex. Hydrolysis conditions presented in this work are suitable for retention of acid-labile side-chain protecting groups and may be adaptable to a variety of unnatural amino acid substrates.

Here, we present an efficient hydrolysis and subsequent Fmoc protection of an amino acid isolated from a Ni-Schiff-base complex. Hydrolysis conditions presented here are suitable for use when retention of acid-labile side-chain protecting groups is required. This technique may be adaptable to a variety of unnatural amino acid substrates.

Unnatural amino acids, amino acids containing side-chain functionalities not commonly seen in nature, are increasingly found in synthetic peptide ...

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 ...

Detection of Small GTPase Prenylation and GTP Binding Using Membrane Fractionation and  GTPase-linked Immunosorbent Assay

The Rho GTPase family belongs to the Ras superfamily and includes approximately 20 members in humans. Rho GTPases are important in the regulation of diverse cellular functions, including cytoskeletal dynamics, cell motility, cell polarity, axonal guidance, vesicular trafficking, and cell cycle control. Changes in Rho GTPase signaling play an essential regulatory role in many pathological conditions, such as&#160;cancer, central nervous system diseases, and immune system-dependent diseases. The posttranslational modification of Rho GTPases (i.e., prenylation by mevalonate pathway intermediates) and GTP binding are key factors which affect the activation of this protein. In this paper, two essential and simple methods are provided to detect a broad range of Rho GTPase prenylation and GTP binding activities. Details of the technical procedures that have been used are explained step by step in this manuscript.

Here we describe a protocol to investigate the prenylation and guanosine-5'-triphosphate (GTP)-loading of Rho GTPase. This protocol consists of two detailed methods, namely membrane fractionation and a GTPase-linked immunosorbent assay. The protocol can be used for measuring the prenylation and GTP loading of different other small GTPases.

The Rho GTPase family belongs to the Ras superfamily and includes approximately 20 members in humans. Rho GTPases are important in the regulation of ...

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 ...

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 ...

Visualization of Estrogen Receptors in Colons of Mice with TNBS-Induced Crohn's Disease using Immunofluorescence

Crohn&#39;s disease is the most diagnosed type of inflammatory bowel disease. Chronic inflammation developing in the intestine leads to peristalsis disorder and damage of intestinal mucosa and seems to be associated with an increased risk of colon neoplastic transformation. Accumulating evidence indicates that estrogens and estrogen receptors affect not only hormone-sensitive tissues, but also other tissues not directly related to estrogens, such as the lungs or colon. Here, we describe the protocol for the successful immunofluorescence staining of estrogen receptors in colon obtained from a murine model of TNBS-induced Crohn&#39;s disease. A detailed protocol for the induction of Crohn&#39;s disease in mice and intestine preparation is provided as well as a step-by-step immunohistochemical procedure using formalin-fixed paraffin-embedded intestine sections. The described methods are not only useful for estrogen receptor detection and estrogen signaling investigation in vivo but can also be applied to for other proteins which may be involved in the development of colitis.

The protocol presents a complete validated TNBS-induced murine model of Crohn&#39;s disease and methods for visualization of estrogen receptors by immunohistochemistry using immunofluorescence of formalin-fixed colon sections embedded in paraffin.

Crohn&#39;s disease is the most diagnosed type of inflammatory bowel disease. Chronic inflammation developing in the intestine leads to peristalsis ...

Characterizing Cellular Proteins with In-cell Fast Photochemical Oxidation of Proteins

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. FPOP uses a 248 nm excimer laser to photolyze hydrogen peroxide producing hydroxyl radicals. These radicals oxidatively modify solvent exposed side chains of 19 of the 20 amino acids. Recently, this method has been used in live cells (IC-FPOP) to study protein interactions in their native environment. The study of proteins in cells accounts for intermolecular crowding and various protein interactions that are disrupted for in vitro studies. A custom single cell flow system was designed to reduce cell aggregation and clogging during IC-FPOP. This flow system focuses the cells past the excimer laser individually, thus ensuring consistent irradiation. By comparing the extent of oxidation produced from FPOP to the protein&#39;s solvent accessibility calculated from a crystal structure, IC-FPOP can accurately probe the solvent accessible side chains of proteins.

Here, we characterize protein structure and interaction sites in living cells using a protein footprinting technique termed in-cell fast photochemical oxidation of proteins (IC-FPOP).

Fast photochemical oxidation of proteins (FPOP) is a hydroxyl radical protein footprinting method used to characterize protein structure and interactions. ...

An Economical and Versatile High-Throughput Protein Purification System Using a Multi-Column Plate Adapter

Protein purification is imperative to the study of protein structure and function and is usually used in combination with biophysical techniques. It is also a key component in the development of new therapeutics. The evolving era of functional proteomics is fueling the demand for high-throughput protein purification and improved techniques to facilitate this. It was hypothesized that a multi column plate adaptor (MCPA) can interface multiple chromatography columns of different resins with multi-well plates for parallel purification. This method offers an economical and versatile method of protein purification that can be used under gravity or vacuum, rivaling the speed of an automated system. The MCPA can be used to recover milligram yields of protein by an affordable and time efficient method for subsequent characterization and analysis. The MCPA has been used for high-throughput affinity purification of SH3 domains. Ion exchange has also been demonstrated via the MCPA to purify protein post Ni-NTA affinity chromatography, indicating how this system can be adapted to other purification types. Due to its setup with multiple columns, individual customization of parameters can be made in the same purification, unachievable by the current plate-based methods.

A multi-column plate adapter allows chromatography columns to be interfaced with multi-well collection plates for parallel affinity or ion exchange purification providing an economical high throughput protein purification method. It can be used under gravity or vacuum yielding milligram quantities of protein via affordable instrumentation.

Protein purification is imperative to the study of protein structure and function and is usually used in combination with biophysical techniques. It is ...

Using Phage Display to Develop Ubiquitin Variant Modulators for E3 Ligases

Ubiquitin is a small 8.6 kDa protein that is a core component of the ubiquitin-proteasome system. Consequently, it can bind to a diverse array of proteins with high specificity but low affinity. Through phage display, ubiquitin variants (UbVs) can be engineered such that they exhibit improved affinity over wildtype ubiquitin and maintain binding specificity to target proteins. Phage display utilizes a phagemid library, whereby the pIII coat protein of a filamentous M13 bacteriophage (chosen because it is displayed externally on the phage surface) is fused with UbVs. Specific residues of human wildtype ubiquitin are soft and randomized (i.e., there is a bias towards to native wildtype sequence) to generate UbVs so that deleterious changes in protein conformation are avoided while introducing the diversity necessary for promoting novel interactions with the target protein. During the phage display process, these UbVs are expressed and displayed on phage coat proteins and panned against a protein of interest. UbVs that exhibit favorable binding interactions with the target protein are retained, whereas poor binders are washed away and removed from the library pool. The retained UbVs, which are attached to the phage particle containing the UbV's corresponding phagemid, are eluted, amplified, and concentrated so that they can be panned against the same target protein in another round of phage display. Typically, up to five rounds of phage display are performed, during which a strong selection pressure is imposed against UbVs that bind weakly and/or promiscuously so that those with higher affinities are concentrated and enriched. Ultimately, UbVs that demonstrate higher specificity and/or affinity for the target protein than their wildtype counterparts are isolated and can be characterized through further experiments.

Ubiquitination is a critical protein post-translational modification, dysregulation of which has been implicated in numerous human diseases. This protocol details how phage display can be utilized to isolate novel ubiquitin variants that can bind and modulate the activity of E3 ligases that control the specificity, efficiency, and patterns of ubiquitination.

Ubiquitin is a small 8.6 kDa protein that is a core component of the ubiquitin-proteasome system. Consequently, it can bind to a diverse array of proteins ...