This work was supported by funding from the Canadian Institutes of Health Research [MOP-1112666 to CVB &#38; ME].

Ethics statement: All experiments and procedures were conducted in agreement with the Laboratory Biosafety guidelines of the University of British Columbia, Vancouver, Canada.
1. Preparation of 35 mm Glass-bottom Culture Dishes for FRET-based Confocal Microscopy
<ol>
	<li>Prepare plates using the following procedure 36-48 hr prior to experiments.</li>
	<li>Retrieve chosen gelatinous protein mixture and 0.25% trypsin-EDTA from -20 &#176;C storage and keep them in mobile ice bath until use. 
	NOTE: The coating protein mixture should only be exposed to RT for a brief period of time to prevent the mixture from solidifying before use. Trypsin solution should also be keep chilled until needed; alternatively, trypsin can be retrieved from -20 &#176;C storage following completion of step 1.5.</li>
	<li>Prepare a 1:25 dilution of protein mixture: DMEM (no added serum, chilled). Transfer appropriate volume of protein mixture/DMEM to each plate being prepared. For 35 mm glass-bottomed plates, add approximately 200 &#181;l to fully cover inner glass circle on bottom of plate.</li>
	<li>Leave plates to sit for at least 30 min inside the safety cabinet and at RT. Warm DMEM with 10% NCS to 37 &#176;C in water bath during this time period.</li>
	<li>Warm trypsin in water bath to 37 &#176;C immediately prior to next steps.</li>
	<li>Remove DMEM that is currently in culture flask using a glass pipette and suction. Wash floor of the flask with warm sterile PBS to remove remaining DMEM residue.</li>
	<li>Dislodge previously cultured smooth muscle cells from flask bottom: 
	NOTE: Flask was established days to this step to allow cells to reach desired confluency (approximately 80%) before plate preparation. Cells were originally suspended in the flask in 20 ml of DMEM with 10% NCS and incubated at 37 &#176;C with 5% CO2 supply for 24 hr, after which DMEM supply was refreshed and cells were returned to incubator. DMEM was refreshed every 48 hr following these initial steps until cells reached confluency.
	<ol>
		<li>Wash flask quickly with 1 ml of trypsin and then remove 1 ml trypsin using suction.</li>
		<li>Add in another 1 ml of trypsin and leave for longer period (approximately 30-60 sec), swishing flask to ensure the enzyme makes contact with the entire bottom of the flask.</li>
		<li>Observe how much cells have detached under a microscope until satisfied with separation from flask. Alternate swishing trypsin solution and checking cell detachment until satisfied with proportion of cells separated from flask. 
		NOTE: The flask can be returned to the incubator for approximately 30-60 sec to speed up this process at 37 &#176;C.</li>
		<li>Once cells have been dislodged using trypsin, add fresh DMEM with 10% NCS to the flask to stop the trypsin reaction (sufficient volume so that trypsin forms 1/8th of total volume in flask). 
		NOTE: For example, when using a 250 ml culture flask, this would be 7 ml DMEM added to the 1 ml of trypsin to result in 8 ml of cell solution.</li>
	</ol>
	</li>
	<li>Transfer resulting cell solution from the flask to a clean (labeled) 50 ml conical tube.</li>
	<li>Add 1 ml fresh DMEM plus 10% NCS to each plate being prepared (sufficient medium to last 24 hr).</li>
	<li>Gently invert tube containing cell solution multiple times to break up any pellet that may have formed.</li>
	<li>Pipette desired volume of gently mixed cell solution into each plate. 
	NOTE: The recommended SMCs number plated for this protocol is 0.5-0.8 x 106 per 35 mm culture dishes. These numbers may vary depending on the cell type and should be tested and modified by each laboratory.</li>
	<li>Swirl each plate thoroughly clockwise/counterclockwise to ensure equal distribution of cells across glass bottom.</li>
</ol>
2. Transient Transfection with Adenoviral Vector Carrying D1SR Ca2+ Indicator
<ol>
	<li>Following addition of medium and cell solution to each plate, leave plates to incubate at 37 &#176;C with 5% CO2 supply for at least 1 hr to allow cells to attach properly to glass on bottom of dishes. Check plates under a microscope at 15 min intervals until cells are clearly attached.</li>
	<li>Retrieve adenovirus-D1SR aliquot from -80 &#176;C storage and keep in mobile ice bath to transport to biological safety cabinet.</li>
	<li>Pipette desired dose (multiplicity of infection of 100) of chilled D1SR to each plate. 
	NOTE: The calculation for the volume of adenoviral solution needed per plate depends on the virus titer in the original stock solution, which is presented as the plaque forming unit (PFU). Based on our experience, we recommend a multiplicity of infection (MOI) of 100, which is interpreted as the number or virus particles needed to infect one smooth muscle cell (SMC) in the culture plate.</li>
	<li>Incubate the infected cells at 37 &#176;C with 5% CO2 supply O/N.</li>
	<li>Next day, replenish cells with 1 ml of fresh DMEM plus 10% NCS.</li>
	<li>Incubate plates in a humidified incubator at 37 &#176;C in 5% CO2 O/N.</li>
</ol>
3. Measurement of SR Luminal Ca2+ Using FRET-based Confocal Imaging
<ol>
	<li>Following O/N&#160;incubation, remove the culture medium from the plate.</li>
	<li>Wash the culture plate with 1 ml warm physiological HEPES-PSS buffer containing (in mmol&#183;L&#8722;1) NaCl 140, glucose 10, KCl 5, HEPES 5, CaCl2 1.5 and MgCl2 1 (pH 7.4) three times.</li>
	<li>Add 1 ml fresh warm HEPES-PSS buffer immediately prior to initiation of recording.</li>
	<li>Perform initial imaging using the FRET SE (Sensitized Emission) application (or similar feature) on the confocal microscope&#39;s associated software.
	<ol>
		<li>Once application is open, click into the &#34;Setup&#34; tab to adjust settings to achieve desired experimental conditions.</li>
		<li>Manually change channel settings so that cells are excited in sequence at 440 nm (for the donor and FRET channels) and 513 nm (for the acceptor channel). Collect the emission wavelengths at 488 nm (for donor) and 535 nm (for FRET and acceptor).</li>
		<li>Set the intensity of light at 15% transmission, with 150 ms excitation exposure time of cells (recording time for three sequential CFP donor, FRET, and YFP acceptor channels) and 10 sec intervals between exposures.</li>
	</ol>
	</li>
	<li>Stabilize plate on the microscope stage.</li>
	<li>Use the &#34;Live&#34; button to check image resolution. Tweak settings further until desired resolution is reached.</li>
	<li>Still under the &#34;Setup&#34; tab, take an image of the sample using the &#34;Capture image&#34; button.</li>
	<li>Moving on to the &#34;Evaluation&#34; tab, choose the appropriate method to calculate the FRET efficiencies for the experiments being done. Method 3, using the Ratiometric Calculation [EA(i) = B/A], is recommended for experiments involving cameleons such as the D1SR indicator.</li>
	<li>Switch over to the &#34;Graph&#34; tab to be able to observe the intensity values being recorded in graph form during the experiment.</li>
	<li>Draw an ROI in the image viewer to observe the corresponding average intensity of this area of interest on the graph as the experiment proceeds. 
	NOTE: Investigator may also choose to draw multiple ROIs and have corresponding intensities be recorded and displayed simultaneously on the graph. Alternatively, investigators may outline a single ROI for initial recording, and proceed to outline multiple ROIs at a later time by opening the experiment file on a data analysis-oriented version of the software. 
	NOTE: Further detail on the steps required to set up FRET-based experiments can be found in FRET application wizard manuals. If the microscope is not equipped with the FRET Wizard, the following protocol can be used to set up the parameters for recording manually.</li>
	<li>Start recording and allow data collection for at least 3 min to ensure a steady basal recording of the signal prior to introducing the agent of interest.</li>
	<li>Introduce Ca2+-moving agent (e.g. 2 &#181;M thapsigargin; 100 nm endothelin-1) as a tool for depleting SR Ca2+, by manually pipetting the predetermined dose directly into the plate at a spot as near to the region of interest being recorded as possible. Continue recording for a desired time post treatment.</li>
	<li>Capture ratiometric FRET serial images (512 x 512 pixels) with a 63X oil-immersion objective of the confocal inverted microscope using the FRET SE application.</li>
	<li>Collect data resulting from imaging from a cluster of 5-10 SMCs in each region of interest (ROI) and subsequently format and analyze using statistics-based software.
	<ol>
		<li>To save data for later use, right-click over the recorded trace and choose the &#34;Export&#34; option to save the spreadsheet with recorded mean signal intensities on the investigator&#39;s chosen drive.</li>
		<li>Normalize data from each individual experiment by dividing each data point by the starting intensity value observed at time point 0 sec.</li>
		<li>Import normalized data from spreadsheets into a statistical program project file by manually copying and pasting relevant data rows from their original spreadsheet/sec.</li>
		<li>Automatically generate graphs corresponding to the data sets pasted into the program. 
		NOTE: The program&#39;s default settings generate line graphs from imported data. Type of graph displayed can be modified by clicking the &#34;Choose a different type of graph&#34; button under the &#34;Change&#34; heading found within the main toolbar.</li>
		<li>Pool data representing multiple independent trials within a larger group of identical experiments into a single data sheet to produce an average trace for the specific test performed.</li>
	</ol>
	</li>
</ol>
4. Preparation of the Cytoplasmic Ca2+ Indicator
<ol>
	<li>Dissolve 0.0125 g Pluronic Acid (PA) in 1 ml of dimethyl sulfoxide (DMSO) in a black microcentrifuge tube (may have to warm the tube in mobile water bath to help dissolve PA).</li>
	<li>Retrieve a tube (50 &#181;g) of the cytoplasmic Ca2+ indicator dye (Fluo-4 AM) from -20 &#176;C storage.</li>
	<li>Wrap the tube of dye in foil to protect it as much as possible from contact with light.</li>
	<li>Pipette 45 &#181;l of the DMSO+PA solution into the tube containing the dye. Wrap DMSO+PA solution in foil and store at RT.</li>
	<li>Mix contents of the tube thoroughly by pipetting solution up and down multiple times. Alternatively, sonicate the dye tube for 10 min. 
	NOTE: Either method of thoroughly distributing the dye throughout the DMSO+PA solution will be sufficient; it is simply a matter of the investigator&#39;s preference. If investigator chooses to sonicate solution, it should be done in an isolated area/room where others will be unaffected by the noise. Investigator should also use heavy mufflers to protect ears.
	<ol>
		<li>If sonicating solution, insert foil-wrapped tube containing the dye inside a foam tube holder and place inside the 2/3 full (with dH2O) sonicator bath.</li>
		<li>Place sonicator bath in isolated area/room and power machine on (ensure protective headwear is in place prior to initiating sonication process). Leave room and leave machine running for 10 min before shutting it off and retrieving the tube (keep wrapped in foil to protect from light for as long as possible).</li>
	</ol>
	</li>
	<li>Aliquot the now thoroughly mixed solution into dark microcentrifuge tubes (5 &#181;l per tube, should be able to make 10 tubes total).</li>
	<li>Wrap each aliquot tube in foil. Label tubes and store in desiccant at -20 &#176;C conditions until use.</li>
</ol>
5. Measurement of Cytoplasmic Ca2+
<ol>
	<li>Follow steps in 1.1-1.12 to prepare culture plates of rat aortic SMCs.</li>
	<li>Remove the culture medium from the plate at 48 hr post culture.</li>
	<li>Remove the previously prepared aliquot of cytoplasmic Ca2+ indicator (5 &#181;l) from -20 &#176;C storage and maintain in mobile ice bath until use.</li>
	<li>Wash the culture plate with 1 ml warm physiological HEPES-PSS buffer containing (in mmol&#183;L&#8722;1) NaCl 140, glucose 10, KCl 5, HEPES 5, CaCl2 1.5 and MgCl2 1 (pH 7.4) three times.</li>
	<li>Mix the indicator with 1 ml of warm HEPES-PSS and load it onto the culture plate.</li>
	<li>Incubate SMCs with the indicator for 1 hr at RT (24 &#176;C).</li>
	<li>Remove indicator from culture plate and wash with 1 ml warm physiological HEPES-PSS buffer three times.</li>
	<li>Add 1 ml fresh warm HEPES-PSS buffer immediately prior to initiation of recording.</li>
	<li>Capture serial images (512 x 512 pixels) with a 63X oil-immersion objective of the confocal inverted microscope.
	<ol>
		<li>Manually change settings so that cells are excited at 488 nm, and the emission wavelengths are collected at 555 nm with a scanning speed of 700 Hz.</li>
		<li>Set the intensity of light at 15% transmission, with 150 msec excitation exposure time of cells (the amount of time that cells are exposed to light at designated time intervals during recording) and 5 sec intervals between exposures.</li>
	</ol>
	</li>
	<li>Stabilize plate on the microscope stage and start recording.</li>
	<li>Record for at least 3 min to ensure a steady basal recording of the signal prior to introducing the agent of interest.</li>
	<li>Introduce Ca2+-moving agent (e.g. 2&#181;M thapsigargin) as a tool for depleting SR Ca2+ (as described in step 3.16) and continue recording for desired amount of time post treatment.</li>
	<li>Record signal intensity using microscope-specific software by averaging fluorescence signals (F/F0) from a cluster of 5-10 SMCs in each region of interest (ROI). Collect, format, and analyze data resulting from imaging using data analysis software. See Steps 3.18.1-3.18.5 for description of data collection and analysis using this software.</li>
</ol>

<ol>
	<li>Hennings, H., Michael, D., Cheng, C., Steinert, P., Holbrook, K., Yuspa, S. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+regulation+of+growth+and+differentiation+of+mouse+epidermal+cells+in+culture.">Calcium regulation of growth and differentiation of mouse epidermal cells in culture.</a> Cell. 19 (1), 245-254 (1980).</li><li>Whitfield, J. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+roles+of+calcium+and+cyclic+AMP+in+cell+proliferation.">The roles of calcium and cyclic AMP in cell proliferation.</a> Ann N Y Acad Sci. 339 (1), 216-240 (1980).</li><li>Nicotera, P., Orrenius, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+calcium+in+apoptosis.">The role of calcium in apoptosis.</a> Cell Calcium. 23 (2-3), 173-180 (1998).</li><li>Wei, C., Wang, X., Chen, M., Ouyang, K., Song, L. S., Cheng, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+flickers+steer+cell+migration.">Calcium flickers steer cell migration.</a> Nature. 457 (7231), 901-905 (2009).</li><li>Ashby, M. C., Tepikin, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ER+calcium+and+the+functions+of+intracellular+organelles.">ER calcium and the functions of intracellular organelles.</a> Sem Cell Dev Biol. 12 (1), 11-17 (2001).</li><li>Mattson, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ER+calcium+and+Alzheimer's+disease:+in+a+state+of+flux.">ER calcium and Alzheimer's disease: in a state of flux.</a> Sci signal. 3 (114), (2010).</li><li>Rutkowski, D. T., Kaufman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+trip+to+the+ER:+coping+with+stress.">A trip to the ER: coping with stress.</a> Trends Cell Biol. 14 (1), (2004).</li><li>Schroder, M., Kaufman, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ER+stress+and+the+unfolded+protein+response.">ER stress and the unfolded protein response.</a> Mutat Res. 569 (1-2), 29-63 (2005).</li><li>Palmer, A. E., Tsien, R. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+calcium+signaling+using+genetically+targetable+fluorescent+indicators.">Measuring calcium signaling using genetically targetable fluorescent indicators.</a> Nat Protoc. 1 (3), 1057-1065 (2006).</li><li>Poburko, D., Liao, C. H., van Breemen, C., Demaurex, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+regulation+of+sarcoplasmic+reticulum+Ca2++content+in+vascular+smooth+muscle+cells.">Mitochondrial regulation of sarcoplasmic reticulum Ca2+ content in vascular smooth muscle cells.</a> Circ Res. 104 (1), 104-112 (2009).</li><li>Esfandiarei, M., Fameli, N., Choi, Y. Y., Tehrani, A. Y., Hoskins, J. G., van Breemen, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Waves+of+calcium+depletion+in+the+sarcoplasmic+reticulum+of+vascular+smooth+muscle+cells:+an+inside+view+of+spatiotemporal+Ca2++regulation.">Waves of calcium depletion in the sarcoplasmic reticulum of vascular smooth muscle cells: an inside view of spatiotemporal Ca2+ regulation.</a> PloS one. 8 (2), e55333 (2013).</li><li>Cobbold, P. H., Rink, T. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+and+bioluminescence+measurement+of+cytoplasmic+free+calcium.">Fluorescence and bioluminescence measurement of cytoplasmic free calcium.</a> Biochem J. 248 (2), 313-328 (1987).</li><li>Park, M. K., Ashby, M. C., Erdemli, G., Petersen, O. H., Tepikin, A. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perinuclear,+perigranular+and+sub-plasmalemmal+mitochondria+have+distinct+functions+in+the+regulation+of+cellular+calcium+transport.">Perinuclear, perigranular and sub-plasmalemmal mitochondria have distinct functions in the regulation of cellular calcium transport.</a> The EMBO journal. 20 (8), 1863-1874 (2001).</li><li>Lovett, J. L., Sibley, L. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intracellular+calcium+stores+in+Toxoplasma+gondii+govern+invasion+of+host+cells.">Intracellular calcium stores in Toxoplasma gondii govern invasion of host cells.</a> J Cell Sci. 116 (14), 3009-3016 (2003).</li><li>Kleinman, H. K., Martin, G. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Matrigel:+basement+membrane+matrix+with+biological+activity.">Matrigel: basement membrane matrix with biological activity.</a> Semin Cancer Biol. 15 (5), 378-386 (2005).</li><li>Palmer, A. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ca2++indicators+based+on+computationally+redesigned+calmodulin-peptide+pairs.">Ca2+ indicators based on computationally redesigned calmodulin-peptide pairs.</a> Chem Biol. 13 (5), 521-530 (2006).</li></ol>

The authors declare that they have no competing financial interests.

This protocol describes the transfection of VSMCs with the D1SR adenovirus to study Ca2+ concentrations within the lumen of the SR. This method allows for the direct measurement of Ca2+ within this organelle as well as its movement into/out of the SR as a result of the application of different calcium-moving agents. This method has many advantages for investigators working towards a comprehensive understanding of how changes to SR and cytoplasmic Ca2+ levels affect overall cellular health...

Ca2+ is a very important ion found in abundance within every single cell in the body. It has significant roles in many different cellular functions, including growth, proliferation, migration, and apoptosis1-4. It is well established that Ca2+ can affect these processes through both direct and indirect actions, and therefore, changes to the normal intracellular concentrations of this ion can easily result in negative outcomes for affected cells. The SR is considered as the largest intracellular Ca2+ store within the cell5. Steady state levels of Ca2+ in both the SR and cytoplasm are normally maintained through constant flux into and out of the Ca2+-transporting channels of this organelle. The stressful conditions imposed upon cells due to any form of injury or disease are commonly associated with significant changes in SR Ca2+ that go on to have long-lasting effects on the health of the cell. In the most severe of cases, the inability of a stressed cell to recover and maintain steady state SR Ca2+ levels may even culminate in death by apoptosis6-8.
Current research into cellular Ca2+ dynamics is limited by the fact that few studies test organelle Ca2+ store content directly9-11. The most common practice instead involves measurement of cytoplasmic Ca2+ levels as indirect measurements of changes in SR Ca2+ content 12-14. In these experiments, Ca2+ is commonly induced to be released from the SR through the use of pharmacological agents causing the organelle&#39;s depletion (e.g. thapsigargin). Conclusions are then drawn with regard to changes to SR Ca2+ based on fluctuations in the cytoplasmic Ca2+ concentrations. Despite the ability remaining for investigators to draw such conclusions in a roundabout manner, this method of SR Ca2+ measurement is clearly an indirect way to glean such information, with many limitations concerning the interpretation of collected data. In order to bypass this clear restriction, it is necessary to measure the amount of Ca2+ found directly within the SR luminal network.
Vital to the final outcome of being able to directly record intraluminal SR Ca2+ levels are the cell culture tools and the Ca 2+ indicator used. For the data referenced in the current manuscript, it is important to note that VSMCs used came from a frozen cell line. Cells were cultured in Dulbecco&#8217;s Modified Eagle&#8217;s medium (DMEM) + 10% newborn calf serum (NCS)&#160;over passages 22-26 for the outlined experiments, incubated at a constant 37 &#176;C with 5% CO2 supply. Using the current method, for example, cells have been very successfully grown on a protein mixture that simulates the extracellular environment of many different types of tissue15. Important for success along the vein of SR Ca2+ research is the type of protein mixture used; in this case, a low growth factor variety was necessary to avoid components of this tool from affecting the regular Ca2+ signaling and movements constantly occurring within the tested cells. Following successful growth of test cells, the Ca2+ indicator must also be effectively introduced to these cultured cells. This has become possible by using an adenoviral vector that carries the SR-residing Ca2+ indicator D1SR. To achieve a high transfection efficiency, cells must be incubated with viral vectors for at least 36-48 hr prior to imaging. This preparation provides a reliable tool to measure Ca2+ transients within the SR lumen with high accuracy and reproducibility.
D1SR indicator used in this protocol is a modified variant of the D1ER Ca2+ indicator that was originally created by Dr. Roger Y. Tsien&#39;s laboratory at the University of California, San Diego, USA9,16. The original D1ER belongs to a second generation of genetically encoded Ca2+ indicators called cameleons that display Ca2+ sensitivities over a much broader range (0.5-160 &#181;M) as compared to previous Ca2+ indicators9,16. The new variant D1SR, a kind gift from Dr. Wayne Chen (University of Alberta, Canada), however, carries a mutant calsequestrin sequence (instead of a calreticulin sequence as in the original D1ER). The mutant calsequestrin has a reduced binding to Ca2+ that eliminates the issue of competing with endogenous calsequestrin in binding to Ca2+ within the SR lumen11. The D1SR indicator carries a truncated enhanced cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP) that bind by a linker protein containing modified calmodulin (CaM) and M13 (the 26-residue peptide of myosin light-chain kinase that binds to CaM) sequences. The CaM-M13 sequence has been modified to prevent M13 from binding to endogenous calmodulin. Also, to ensure SR retention, a calsequestrin sequence has been added on the 5&#180; end of CFP11. When bound to Ca2+, the CaM-M13 domain goes through conformational changes that result in an increase in the energy transfer between the flanking CFP and YFP, which is recorded as an increase in the FRET signal intensity. On the other hand, when the concentration of SR luminal Ca2+ drops, the CaM-M13 domain goes through reverse conformational changes, resulting in a decrease in the energy transfer between the flanking CFP and YFP, and a significant drop in FRET signal intensity.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle&#39;s Medium (high glucose, pyruvate)</td>
			<td>Life technologies</td>
			<td>11995-065</td>
			<td>Warm in 37&#176;C water bath before use</td>
		</tr>
		<tr>
			<td>Matrigel Basement Membrane Matrix Growth Factor Reduced, 5mL</td>
			<td>Corning</td>
			<td>356230</td>
			<td>Keep at -20&#176;C until right before use</td>
		</tr>
		<tr>
			<td>Newborn Calf Serum</td>
			<td>Sigma-Aldrich</td>
			<td></td>
			<td>Keep at -20&#176;C until right before use</td>
		</tr>
		<tr>
			<td>35mm glass-bottomed plates</td>
			<td>MatTek Corporation</td>
			<td>P35G-1.5-14-C</td>
			<td></td>
		</tr>
		<tr>
			<td>250 mL 0.2 &#956;M vented blue plug seal cap flask</td>
			<td>BD Falcon</td>
			<td>353136</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin/EDTA Solution</td>
			<td>Life technologies</td>
			<td>25200056</td>
			<td>Keep at -20&#176;C until right before use</td>
		</tr>
		<tr>
			<td>50 mL Polypropylene Conical Tube</td>
			<td>BD Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>D1SR</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>Gift</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO)</td>
			<td>Sigma-Aldrich</td>
			<td>D8779</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F-127</td>
			<td>Sigma-Aldrich</td>
			<td>P2443</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluo 4-AM</td>
			<td>Life technologies</td>
			<td>F-23917</td>
			<td>Keep at -20&#176;C until right before use</td>
		</tr>
		<tr>
			<td>1.5 mL Black Microcentrifuge Tubes</td>
			<td>Argos Technologies</td>
			<td>T7100BK</td>
			<td></td>
		</tr>
		<tr>
			<td>Leica TCS SP5 confocal microscope</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Leica Application Suite 2.6.3 (with FRET SE Wizard application)</td>
			<td>Leica Microsystems</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>GraphPad Prism 5.0</td>
			<td>GraphPad Software, Inc.</td>
			<td></td>
			<td></td>
		</tr>
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measurement of calcium fluctuations within the sarcoplasmic reticulum of cultured smooth muscle cells using fret-based confocal imaging

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall health. A significant portion of intracellular Ca2+ content is found within the SR stores in VSMCs. As the only intracellular organelle with a close association to the surrounding extracellular space through plasma membrane-SR junctions, the SR can be considered to constitute the first line of response to any irregularity in Ca2+ transients, or stress experienced by the cell. Among its many functions, one of the most important is its role in the transmission of Ca2+-regulated signals throughout the cell to induce further cell-wide reactions downstream. The more common use of cytoplasmic Ca2+ indicators in this regard is overall insufficient for research into the highly dynamic changes to the intraluminal SR Ca2+ store that have yet to be fully characterized. Here, we provide a detailed protocol for the direct and clear measurement of luminal SR Ca2+. This tool is useful for investigation into the nuanced changes in SR Ca2+ that have significant subsequent effects on the normal function and health of the cell. Fluctuations in SR Ca2+ content specifically can provide us with a significant amount of information pertaining to cellular responses to disease or stress conditions experienced by the cell. In this method, a modified version of a SR-targeted Ca2+ indicator, D1SR, is used to detect Ca2+ fluctuations in response to the introduction of agents to cultured rat aortic smooth muscle cells (SMCs). Following incubation with the D1SR indicator, confocal fluorescence microscopy and fluorescence resonance energy transfer (FRET)-based imaging are used to directly observe changes to intraluminal SR Ca2+ levels under control conditions and with the addition of agonist agents that function to induce intracellular Ca2+ movement.

This section provides examples of the results that can be obtained using the described method to capture SR luminal Ca2+ concentration data, compared to the commonly employed manner of indirectly measuring changes to organelle Ca2+ store concentrations by observing fluctuations in cytoplasmic Ca2+.
Figure 1A shows a snapshot of a region of interest (ROI) of the SMC culture transf...

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Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall ...

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Research

JoVE Journal

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7.6K Views.  University of British Columbia. Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Video: Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Focal Ca2+ Transient Detection in Smooth Muscle

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ from internal stores, and allows multiple cells to be monitored simultaneously to assess, for example, coupling in syncytial tissue. Subcellular Ca2+ transients are common in smooth muscle, yet are difficult to measure accurately because of the problems caused by their stochastic occurrence, over an often wide field of view, in an organ that it prone to contract. To overcome this problem, we've developed a series of imaging protocols and analysis routines to acquire and then analyse, in an automated fashion, the frequency, location and amplitude of such events. While this approach may be applied in other contexts, our own work involves the detection of local purinergic Ca2+ transients for locating transmitter release with submicron resolution. 
ATP is released as a cotransmitter from autonomic nerves, where it binds to P2X1 receptors on the smooth muscle of the detrusor and vas deferens. Ca2+ enters the smooth muscle, resulting in purinergic neuroeffector Ca2+ transients (NCTs). The focal Ca2+ transients allow the optical monitoring of neurotransmitter release in a manner that has many advantages over electrophysiology. Apart from the greatly improved spatial resolution, optical recording has the additional advantage of allowing the recording of transmitter release from many distinguishable sites simultaneously. Furthermore, the optical plane of focus is easier to maintain or correct during long recording series than is the repositioning of an intracellular sharp microelectrode. 
In summary, a method for imaging of Ca2+ fluorescence is outlined which details the preparation of tissue, and the acquisition and analysis of data. We outline the use of several scripts for the analysis of such Ca2+ transients.

Focal Ca2+ Transient Detection in Smooth Muscle

Details methods for high-resolution Ca2+ imaging of smooth muscle within isolated organs, including: preparation of the tissue, image acquisition and data analysis.

Ca2+ imaging of smooth muscle provides insight into cellular mechanisms that may not result in changes of membrane potential, such as the release of Ca2+ ...

Isolation of Human Umbilical Arterial Smooth Muscle Cells (HUASMC)

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded and their collection does not imply any added risk to the newborn or mother s health. Moreover no ethical concerns are raised. The UC is composed by one vein and two arteries from which both endothelial cells (ECs) 1 and smooth muscle cells (SMCs) 2, two of the main cellular components of blood vessels, can be isolated. In this project the SMCs were obtained after enzymatic treatment of the UC arteries accordingly the experimental procedure previously described by Jaffe et al 3. After cell isolation they were kept in t-flash with DMEM-F12 supplemented with 5% of fetal bovine serum and were cultured for several passages. Cells maintained their morphological and other phenotypic characteristics in the different generations. The aim of this study was to isolate smooth muscle cells in order to use them as models for future assays with constrictor drugs, isolate and structurally characterize L-type calcium channels, to study cellular and molecular aspects of the vascular function 4 and to use them in tissue engineering.

Isolation of Human Umbilical Arterial Smooth Muscle Cells HUASMC

The umbilical cords are used to isolate smooth muscle cells by different ways. In this work we used the enzymatic treatment to isolated smooth muscle cells.

The human umbilical cord (UC) is a biological sample that can be easily obtained just after birth. This biological sample is, most of the time, discarded ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Isolation of Pulmonary Artery Smooth Muscle Cells from Neonatal Mice

Pulmonary hypertension is a significant cause of morbidity and mortality in infants. Historically, there has been significant study of the signaling pathways involved in vascular smooth muscle contraction in PASMC from fetal sheep. While sheep make an excellent model of term pulmonary hypertension, they are very expensive and lack the advantage of genetic manipulation found in mice. Conversely, the inability to isolate PASMC from mice was a significant limitation of that system. Here we described the isolation of primary cultures of mouse PASMC from P7, P14, and P21 mice using a variation of the previously described technique of Marshall et al.26 that was previously used to isolate rat PASMC. These murine PASMC represent a novel tool for the study of signaling pathways in the neonatal period. Briefly, a slurry of 0.5% (w/v) agarose + 0.5% iron particles in M199 media is infused into the pulmonary vascular bed via the right ventricle (RV). The iron particles are 0.2 &#956;M in diameter and cannot pass through the pulmonary capillary bed. Thus, the iron lodges in the small pulmonary arteries (PA). The lungs are inflated with agarose, removed and dissociated. The iron-containing vessels are pulled down with a magnet. After collagenase (80 U/ml) treatment and further dissociation, the vessels are put into a tissue culture dish in M199 media containing 20% fetal bovine serum (FBS), and antibiotics (M199 complete media) to allow cell migration onto the culture dish. This initial plate of cells is a 50-50 mixture of fibroblasts and PASMC. Thus, the pull down procedure is repeated multiple times to achieve a more pure PASMC population and remove any residual iron. Smooth muscle cell identity is confirmed by immunostaining for smooth muscle myosin and desmin.

We have developed a novel and reproducible technique to isolate primary cultures of pulmonary artery smooth muscle cells (PASMC) from mice as young as P7, thereby allowing better study of the signaling pathways involved in neonatal smooth muscle cell contraction and relaxation.

Pulmonary hypertension is a significant cause of morbidity and mortality in infants. Historically, there has been significant study of the signaling ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Simultaneous pH Measurement in Endocytic and Cytosolic Compartments in Living Cells using Confocal Microscopy

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial for homeostatic control of physiological processes that include: protein, DNA and RNA synthesis, vesicular trafficking, cell growth and cell division. Alterations in cellular pH homeostasis can lead to detrimental functional changes and promote progression of various diseases2. Various methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of pH in the cytoplasm and in organelles. Here, we describe in detail a rapid and accurate method for the simultaneous measurement of cytoplasmic and organellar pH by using confocal microscopy on living cells3. This goal is achieved with the use of two pH-sensing ratiometric dyes that possess selective cellular compartment partitioning. For instance, SNARF-1 is compartmentalized inside the cytoplasm whereas HPTS is compartmentalized inside endosomal/lysosomal organelles. Although HPTS is commonly used as a cytoplasmic pH indicator, this dye can specifically label vesicles along the endosomal-lysosomal pathway after being taken up by pinocytosis3,4. Using these pH-sensing probes, it is possible to simultaneously measure pH within the endocytic and cytoplasmic compartments. The optimal excitation wavelength of HPTS varies depending on the pH while for SNARF-1, it is the optimal emission wavelength that varies. Following loading with SNARF-1 and HPTS, cells are cultured in different pH-calibrated solutions to construct a pH standard curve for each probe. Cell imaging by confocal microscopy allows elimination of artifacts and background noise. Because of the spectral properties of HPTS, this probe is better suited for measurement of the mildly acidic endosomal compartment or to demonstrate alkalinization of the endosomal/lysosomal organelles. This method simplifies data analysis, improves accuracy of pH measurements and can be used to address fundamental questions related to pH modulation during cell responses to external challenges.

Several methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of cytoplasmic and organellar pH. Here, we describe in detail a rapid and accurate methodology to simultaneously measure cytoplasmic and vesicular pH by ratiometric imaging of living cells.

Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1. Regulation of cellular pH is crucial ...

Measurement of Smooth Muscle Function in the Isolated Tissue Bath-applications to Pharmacology Research

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues. While this technique has been implemented for over 100 years, the versatility, simplicity and reproducibility of this assay helps it to remain an indispensable tool for pharmacologists and physiologists alike. Tissue bath systems are available in a wide array of shapes and sizes, allowing a scientist to evaluate samples as small as murine mesenteric arteries and as large as porcine ileum &#8211; if not larger. Central to the isolated tissue bath assay is the ability to measure concentration-dependent changes to isometric contraction, and how the efficacy and potency of contractile agonists can be manipulated by increasing concentrations of antagonists or inhibitors. Even though the general principles remain relatively similar, recent technological advances allow even more versatility to the tissue bath assay by incorporating computer-based data recording and analysis software. This video will demonstrate the function of the isolated tissue bath to measure the isometric contraction of an isolated smooth muscle (in this case rat thoracic aorta rings), and share the types of knowledge that can be created with this technique. Included are detailed descriptions of aortic tissue dissection and preparation, placement of aortic rings in the tissue bath and proper tissue equilibration prior to experimentation, tests of tissue viability, experimental design and implementation, and data quantitation. Aorta will be connected to isometric force transducers, the data from which will be captured using a commercially available analog-to-digital converter and bridge amplifier specifically designed for use in these experiments. The accompanying software to this system will be used to visualize the experiment and analyze captured data.

This protocol describes the measurement of isometric contraction in an isolated smooth muscle preparation, using an isolated tissue bath system and computer-based data acquisition.

Isolated tissue bath assays are a classical pharmacological tool for evaluating concentration-response relationships in a myriad of contractile tissues.  ...

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a significant role in certain types of arrhythmia. Because arrhythmias are spatially distinct, emergent phenomena, they must be investigated at the tissue level. However, methods for directly probing SR Ca2+ in the intact heart remain limited. This article describes the protocol for dual optical mapping of transmembrane potential (Vm) and free intra-SR [Ca2+] ([Ca2+]SR) in the Langendorff-perfused rabbit heart. This approach takes advantage of the low-affinity Ca2+ indicator Fluo-5N, which has minimal fluorescence in the cytosol where intracellular [Ca2+] ([Ca2+]i) is relatively low but exhibits significant fluorescence in the SR lumen where [Ca2+]SR is in the millimolar range. In addition to revealing SR Ca2+ characteristics spatially across the epicardial surface of the heart, this approach has the distinct advantage of simultaneous monitoring of Vm, allowing for investigations into the bidirectional relationship between Vm and SR Ca2+ and the role of SR Ca2+ in arrhythmogenic phenomena.

Optical Mapping of Intra-Sarcoplasmic Reticulum Ca2+ and Transmembrane Potential in the Langendorff-perfused Rabbit Heart

This article describes the detailed protocol and equipment necessary for dual optical mapping of transmembrane potential (Vm) and free intra-sarcoplasmic reticulum (SR) Ca2+ in the Langendorff-perfused rabbit heart. This method allows for direct observation and quantification of Vm and SR Ca2+ dynamics in the intact heart.

Sarcoplasmic reticulum (SR) Ca2+ handling plays a key role in normal excitation-contraction coupling and aberrant SR Ca2+ handling is known to play a ...

Monitoring Endoplasmic Reticulum Calcium Homeostasis Using a Gaussia Luciferase SERCaMP

The endoplasmic reticulum (ER) contains the highest level of intracellular calcium, with concentrations approximately 5,000-fold greater than cytoplasmic levels. Tight control over ER calcium is imperative for protein folding, modification and trafficking. Perturbations to ER calcium can result in the activation of the unfolded protein response, a three-prong ER stress response mechanism, and contribute to pathogenesis in a variety of diseases. The ability to monitor ER calcium alterations during disease onset and progression is important in principle, yet challenging in practice. Currently available methods for monitoring ER calcium, such as calcium-dependent fluorescent dyes and proteins, have provided insight into ER calcium dynamics in cells, however these tools are not well suited for in vivo studies. Our lab has demonstrated that a modification to the carboxy-terminus of Gaussia luciferase confers secretion of the reporter in response to ER calcium depletion. The methods for using a luciferase based, secreted ER calcium monitoring protein (SERCaMP) for in vitro and in vivo applications are described herein. This video highlights hepatic injections, pharmacological manipulation of GLuc-SERCaMP, blood collection and processing, and assay parameters for longitudinal monitoring of ER calcium.

Monitoring Endoplasmic Reticulum Calcium Homeostasis Using a Gaussia Luciferase SERCaMP

Endoplasmic reticulum calcium homeostasis is disrupted in diverse pathologies. A secreted ER calcium monitoring protein (SERCaMP) reporter can be used to detect disruptions in the ER calcium store. This protocol describes the use of a Gaussia luciferase SERCaMP to examine ER calcium homeostasis in vitro and in vivo.

The endoplasmic reticulum (ER) contains the highest level of intracellular calcium, with concentrations approximately 5,000-fold greater than cytoplasmic ...