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><tbody><tr><td>&lt;strong&gt;Cell Culture Reagents&lt;/strong&gt;</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&#039;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>&lt;strong&gt;Calcium Mobilization Reagents&lt;/strong&gt;</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&#039;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-NH&lt;sub&gt;2&lt;/sub&gt; (TFA salt)</td><td></td><td></td><td>Prepared by Trudy Holyst at the Versiti Blood Research Institute</td></tr><tr><td>&lt;strong&gt;Materials&lt;/strong&gt;</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&quot;</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 &amp;micro;L</td><td>Biotix</td><td>63300042</td><td>sterile, filtered tips</td></tr><tr><td>Pipette tips, 200 &amp;micro;L</td><td>Biotix</td><td>63300044</td><td>sterile, filtered tips</td></tr><tr><td>Pipette tips, 1250 &amp;micro;L</td><td>Biotix</td><td>63300047</td><td>sterile, filtered tips</td></tr><tr><td>Prism</td><td>GraphPad</td><td></td><td>volume 6 used</td></tr><tr><td>Serological pipette, 5 mL</td><td>Tradewinds Direct</td><td>&amp;nbsp;07-5005</td><td></td></tr><tr><td>Serological pipette, 10 mL</td><td>Tradewinds Direct</td><td>&amp;nbsp;07-5010</td><td></td></tr><tr><td>Serological pipette, 25 mL</td><td>Tradewinds Direct</td><td>&amp;nbsp;07-5025</td><td></td></tr></tbody></table>

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

403 Views.  Function Therapeutics. Ons laboratorium probeert te begrijpen hoe de modulatie van door protease geactiveerde receptoren ontstekingen kan remmen. We hebben een nieuwe klasse van kleine moleculen ontwikkeld, parmodulinen genaamd, die de activiteit van PAR1 moduleren. Het doel is om parmodulinen met verbeterde eigenschappen te ontdekken voor mogelijk gebruik als kandidaat-geneesmiddel, en ook om hun werkingsmechanisme te begrijpen.In preklinische studies met onze medewerkers h...

Video: Auteur in de schijnwerpers: Parmodulines ontwikkelen om protease-geactiveerde receptoren aan te pakken voor ontstekingscontrole

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 ...

Cell-based Calcium Assay for Medium to High Throughput Screening of TRP Channel Functions using FlexStation 3

The Molecular Devices' FlexStation 3 is a benchtop multi-mode microplate reader capable of automated fluorescence measurement in multi-well plates. It is ideal for medium- to high-throughput screens in academic settings. It has an integrated fluid transfer module equipped with a multi-channel pipetter and the machine reads one column at a time to monitor fluorescence changes of a variety of fluorescent reagents. For example, FlexStation 3 has been used to study the function of Ca2+-permeable ion channels and G-protein coupled receptors by measuring the changes of intracellular free Ca2+ levels. Transient receptor potential (TRP) channels are a large family of nonselective cation channels that play important roles in many physiological and pathophysiological functions. Most of the TRP channels are calcium permeable and induce calcium influx upon activation. In this video, we demonstrate the application of FlexStation 3 to study the pharmacological profile of the TRPA1 channel, a molecular sensor for numerous noxious stimuli. HEK293 cells transiently or stably expressing human TRPA1 channels, grown in 96-well plates, are loaded with a Ca2+-sensitive fluorescent dye, Fluo-4, and real-time fluorescence changes in these cells are measured before and during the application of a TRPA1 agonist using the FLEX mode of the FlexStation 3. The effect of a putative TRPA1 antagonist was also examined. Data are transferred from the SoftMax Pro software to construct concentration-response relationships of TRPA1 activators and inhibitors.

This video provides a detailed protocol for studying the pharmacological profile of human TRPA1 channels using FlexStation 3. The protocol covers details of cell preparation, dye loading and operation of the microplate reader, FlexStation 3.

The Molecular Devices' FlexStation 3 is a benchtop multi-mode microplate reader capable of automated fluorescence measurement in multi-well plates.  It is ...

Bioengineering

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 ...

A "Dual-Addition" Calcium Fluorescence Assay for the High-Throughput Screening of Recombinant G Protein-Coupled Receptors

G protein-coupled receptors (GPCRs) represent the largest superfamily of receptors and are the targets of numerous human drugs. High-throughput screening (HTS) of random small molecule libraries against GPCRs is used by the pharmaceutical industry for target-specific drug discovery. In this study, an HTS was employed to identify novel small-molecule ligands of invertebrate-specific neuropeptide GPCRs as probes for physiological studies of vectors of deadly human and veterinary pathogens.
The invertebrate-specific kinin receptor was chosen as a target because it regulates many important physiological processes in invertebrates, including diuresis, feeding, and digestion. Furthermore, the pharmacology of many invertebrate GPCRs is poorly characterized or not characterized at all; therefore, the differential pharmacology of these groups of receptors with respect to the related GPCRs in other metazoans, especially humans, adds knowledge to the structure-activity relationships of GPCRs as a superfamily. An HTS assay was developed for cells in 384-well plates for the discovery of ligands of the kinin receptor from the cattle fever tick, or southern cattle tick, Rhipicephalus microplus. The tick kinin receptor was stably expressed in CHO-K1 cells.
The kinin receptor, when activated by endogenous kinin neuropeptides or other small molecule agonists, triggers Ca2+ release from calcium stores into the cytoplasm. This calcium fluorescence assay combined with a &#34;dual-addition&#34; approach can detect functional agonist and antagonist &#34;hit&#34; molecules in the same assay plate. Each assay was conducted using drug plates carrying an array of 320 random small molecules. A reliable Z&#39; factor of 0.7 was obtained, and three agonist and two antagonist hit molecules were identified when the HTS was at a 2 &#181;M final concentration. The calcium fluorescence assay reported here can be adapted to screen other GPCRs that activate the Ca2+ signaling cascade.

In this work, a high-throughput, intracellular calcium fluorescence assay for 384-well plates to screen small molecule libraries on recombinant G protein-coupled receptors (GPCRs) is described. The target, the kinin receptor from the cattle fever tick, Rhipicephalus microplus, is expressed in CHO-K1 cells. This assay identifies agonists and antagonists using the same cells in one "dual-addition" assay.

G protein-coupled receptors (GPCRs) represent the largest superfamily of receptors and are the targets of numerous human drugs. High-throughput screening ...

A Kinetic Fluorescence-based Ca2+ Mobilization Assay to Identify G Protein-coupled Receptor Agonists, Antagonists, and Allosteric Modulators

G protein-coupled receptors (GPCRs) are of great importance to the pharmaceutical industry as they are involved in many human diseases and include well-validated targets for therapeutic intervention. Discovery of lead compounds, including small synthetic molecules, that specifically inhibit the receptor&#39;s function, is an important initial step in drug development and relies on sensitive, specific, and robust cell-based assays. Here, we describe a kinetic cellular assay with a fluorescent readout primarily designed to identify receptor-specific antagonists that inhibit the intracellular Ca2+ release evoked upon the activation of the CXC chemokine receptor 4 (CXCR4) by its endogenous ligand, the CXC chemokine ligand 12 (CXCL12). A key advantage of this method is that it also enables screening of compounds endowed with intrinsic agonistic properties (i.e., compounds eliciting an increase in intracellular Ca2+ concentration in the absence of CXCL12) or compounds modulating the receptor&#39;s function via interaction with allosteric binding sites (i.e., positive and negative allosteric modulators (PAMs and NAMs, respectively)). On the down side, autofluorescent compounds might interfere with the assay&#39;s readout, thereby hampering reliable data interpretation. Most likely this assay can be implemented, with minimal adaptations, as a generic drug discovery assay for many other GPCRs of which the activation leads to a release of intracellular Ca2+.

A Kinetic Fluorescence-based Ca2+ Mobilization Assay to Identify G Protein-coupled Receptor Agonists, Antagonists, and Allosteric Modulators

The described cellular assay is designed for the identification of CXC chemokine receptor 4 (CXCR4)-interacting agents that inhibit or stimulate, either competitively or allosterically, the intracellular Ca2+ release initiated by CXCR4 activation.

G protein-coupled receptors (GPCRs) are of great importance to the pharmaceutical industry as they are involved in many human diseases and include ...

Medicine

Screening Peptides that Activate MRGPRX2 using Engineered HEK Cells

Identifying ligands specific to therapeutically significant cell receptors is crucial for many applications, including the design and development of new therapeutics. Mas related G-protein receptor-X2 (MRGPRX2) is an important receptor that regulates mast cell activation and, thus, directs the general immune response. Numerous ligands for MRGPRX2 have been identified and include endogenous peptides like PAMPs, defensins, LL-37 and other protein fragments (i.e., degraded albumin). Further identification of MRGPRX2 specific ligands requires the screening of a large number of peptides (i.e., peptide library); however, mast cells are difficult and expensive to maintain in vitro and, therefore, not economical to use for screening large numbers of molecules. The present paper demonstrates a method to design, develop, and screen a library of small peptide molecules using MRGPRX2 expressing HEK cells. This cell line is relatively easy and inexpensive to maintain and can be used for in vitro high-throughput analysis. A calcium sensitive Fura-2 fluorescent dye to mark intracellular calcium flux upon activation was used to monitor the activation. The ratio of fluorescence intensity of Fura-2 at 510 nm against excitation wavelengths of 340 and 380 nm was used to calculate calcium concentration. The peptide library used to verify this system was based on the endogenous proadrenomedullin N-terminal 12 (PAMP-12) secretagogue, which is known to bind MRGPRX2 with high specificity and affinity. Subsequent peptides were generated through amino acid truncation and alanine scanning techniques applied to PAMP-12. The method described here is simple and inexpensive yet robust for screening a large library of compounds to identify binding domains and other important parameters that play an important role in receptor activation.

Techniques for generating a library of short peptides that can activate mast cells via the MRGPRX2 receptor are described. Associated techniques are easy, inexpensive, and can be extended to other cell receptors.

Identifying ligands specific to therapeutically significant cell receptors is crucial for many applications, including the design and development of new ...

Immunology and Infection

Characterization of G Protein-coupled Receptors by a Fluorescence-based Calcium Mobilization Assay

For more than 20 years, reverse pharmacology has been the preeminent strategy to discover the activating ligands of orphan G protein-coupled receptors (GPCRs). The onset of a reverse pharmacology assay is the cloning and subsequent transfection of a GPCR of interest in a cellular expression system. The heterologous expressed receptor is then challenged with a compound library of candidate ligands to identify the receptor-activating ligand(s). Receptor activation can be assessed by measuring changes in concentration of second messenger reporter molecules, like calcium or cAMP. The fluorescence-based calcium mobilization assay described here is a frequently used medium-throughput reverse pharmacology assay. The orphan GPCR is transiently expressed in human embryonic kidney 293T (HEK293T) cells and a promiscuous G&#945;16 construct is co-transfected. Following ligand binding, activation of the G&#945;16 subunit induces the release of calcium from the endoplasmic reticulum. Prior to ligand screening, the receptor-expressing cells are loaded with a fluorescent calcium indicator, Fluo-4 acetoxymethyl. The fluorescent signal of Fluo-4 is negligible in cells under resting conditions, but can be amplified more than a 100-fold upon the interaction with calcium ions that are released after receptor activation. The described technique does not require the time-consuming establishment of stably transfected cell lines in which the transfected genetic material is integrated into the host cell genome. Instead, a transient transfection, generating temporary expression of the target gene, is sufficient to perform the screening assay. The setup allows medium-throughput screening of hundreds of compounds. Co-transfection of the promiscuous G&#945;16, which couples to most GPCRs, allows the intracellular signaling pathway to be redirected towards the release of calcium, regardless of the native signaling pathway in endogenous settings. The HEK293T cells are easy to handle and have proven their efficacy throughout the years in receptor deorphanization assays. However, optimization of the assay for specific receptors may remain necessary.

The here described fluorescence-based calcium mobilization assay is a medium-throughput reverse pharmacology screening system for the identification of functionally activating ligand(s) of orphan G protein-coupled receptors (GPCRs).

For more than 20 years, reverse pharmacology has been the preeminent strategy to discover the activating ligands of orphan G protein-coupled receptors ...

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

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