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/pubmed?term=((Calcium+regulation+of+growth+and+differentiation+of+mouse+epidermal+cells+in+culture%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">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/pubmed?term=((The+roles+of+calcium+and+cyclic+AMP+in+cell+proliferation%5BTitle%5D)+AND+%22Ann+N+Y+Acad+Sci%22%5BJournal%5D)">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/pubmed?term=((The+role+of+calcium+in+apoptosis%5BTitle%5D)+AND+%22Cell+Calcium%22%5BJournal%5D)">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/pubmed?term=((Calcium+flickers+steer+cell+migration%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">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/pubmed?term=((ER+calcium+and+the+functions+of+intracellular+organelles%5BTitle%5D)+AND+%22Sem+Cell+Dev+Biol%22%5BJournal%5D)">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/pubmed?term=((ER+calcium+and+Alzheimer%27s+disease%3A+in+a+state+of+flux%5BTitle%5D)+AND+%22Sci+signal%22%5BJournal%5D)">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/pubmed?term=((A+trip+to+the+ER%3A+coping+with+stress%5BTitle%5D)+AND+%22Trends+Cell+Biol%22%5BJournal%5D)">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/pubmed?term=((ER+stress+and+the+unfolded+protein+response%5BTitle%5D)+AND+%22Mutat+Res%22%5BJournal%5D)">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/pubmed?term=((Measuring+calcium+signaling+using+genetically+targetable+fluorescent+indicators%5BTitle%5D)+AND+%22Nat+Protoc%22%5BJournal%5D)">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/pubmed?term=((Mitochondrial+regulation+of+sarcoplasmic+reticulum+Ca2%2B+content+in+vascular+smooth+muscle+cells%5BTitle%5D)+AND+%22Circ+Res%22%5BJournal%5D)">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/pubmed?term=((Waves+of+calcium+depletion+in+the+sarcoplasmic+reticulum+of+vascular+smooth+muscle+cells%3A+an+inside+view+of+spatiotemporal+Ca2%2B+regulation%5BTitle%5D)+AND+%22PloS+one%22%5BJournal%5D)">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/pubmed?term=((Fluorescence+and+bioluminescence+measurement+of+cytoplasmic+free+calcium%5BTitle%5D)+AND+%22Biochem+J%22%5BJournal%5D)">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/pubmed?term=((Perinuclear%2C+perigranular+and+sub-plasmalemmal+mitochondria+have+distinct+functions+in+the+regulation+of+cellular+calcium+transport%5BTitle%5D)+AND+%22The+EMBO+journal%22%5BJournal%5D)">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/pubmed?term=((Intracellular+calcium+stores+in+Toxoplasma+gondii+govern+invasion+of+host+cells%5BTitle%5D)+AND+%22J+Cell+Sci%22%5BJournal%5D)">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/pubmed?term=((Matrigel%3A+basement+membrane+matrix+with+biological+activity%5BTitle%5D)+AND+%22Semin+Cancer+Biol%22%5BJournal%5D)">Matrigel: basement membrane matrix with biological activity.</a> Semin Cancer Biol. 15 (5), 378-386 (2005).</li>
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</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><tbody><tr><td>Dulbecco&#039;s Modified Eagle&#039;s Medium (high glucose, pyruvate)</td><td>Life technologies</td><td>11995-065</td><td>Warm in 37 &amp;deg;C water bath before use</td></tr><tr><td>Matrigel Basement Membrane Matrix Growth Factor Reduced, 5 ml</td><td>Corning</td><td>356230</td><td>Keep at -20 &amp;deg;C until right before use</td></tr><tr><td>Newborn Calf Serum</td><td>Sigma-Aldrich</td><td></td><td>Keep at -20 &amp;deg;C until right before use</td></tr><tr><td>35 mm glass-bottomed plates</td><td>MatTek Corporation</td><td>P35G-1.5-14-C</td><td></td></tr><tr><td>250 ml 0.2 &amp;mu;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 &amp;deg;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 &amp;deg;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></tbody></table>

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

Biology

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

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for muscle regeneration. Upon injury, which occurs frequently in skeletal muscles, SCs are activated. They proliferate as myoblasts and differentiate to repair muscle lesions. Among many events that take place during muscle differentiation, cytosolic Ca2+ signals are of great importance. These Ca2+ signals arise from Ca2+ release from internal Ca2+ stores, as well as from Ca2+ entry from the extracellular space, particularly the store-operated Ca2+ entry (SOCE). This paper describes a methodology used to obtain a pure population of human myoblasts from muscle samples collected after orthopedic surgery. The tissue is mechanically and enzymatically digested, and the cells are amplified and then sorted by flow cytometry according to the presence of specific membrane markers. Once obtained, human myoblasts are expanded and committed to differentiate by removing growth factors from the culture medium. The expression levels of specific transcription factors and in vitro immunofluorescence are used to assess the myogenic differentiation process in control conditions and after silencing proteins involved in Ca2+ signaling. Finally, we detail the use of Fura-2 as a ratiometric Ca2+ probe that provides reliable and reproducible measurements of SOCE.

Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for ...

Developmental Biology

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+ ...

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 ...

Measuring Fast Calcium Fluxes in Cardiomyocytes

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

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

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

Assessment of Sarcoplasmic Reticulum Calcium Reserve and Intracellular Diastolic Calcium Removal in Isolated Ventricular Cardiomyocytes

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Cardiac sarcoplasmic reticulum (SR) serves as a Ca2+ reservoir for contraction, which reuptakes intracellular Ca2+ during relaxation. The SR Ca2+ reserve available for beats is determinate for cardiac contractibility, and the removal of intracellular Ca2+ is critical for cardiac diastolic function. Under some pathophysiological conditions, such as diabetes and heart failure, impaired calcium clearance and SR Ca2+ store in cardiomyocytes may be involved in the progress of cardiac dysfunction. Here, we describe a protocol to evaluate SRCa2+ reserve and diastolic Ca2+ removal. Briefly, a single cardiomyocyte was enzymatically isolated, and the intracellular Ca2+ fluorescence indicated by Fura-2 was recorded by a calcium imaging system. To employ caffeine for inducing total SR Ca2+ release, we preset an automatic perfusion switch program by interlinking the stimulation system and the perfusion system. Then, the mono-exponential curve fitting was used for analyzing decay time constants of calcium transients and caffeine-induced calcium pulses. Accordingly, the contribution of the SR Ca2+-ATPase (SERCA) and Na+-Ca2+ exchanger (NCX) to diastolic calcium removal was evaluated.

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Here, we describe a protocol to evaluate sarcoplasmic reticulum Ca2+ reserve and diastolic calcium removal function in cardiomyocytes by a calcium imaging system.

Intracellular calcium recycling plays a critical role in regulation of systolic and diastolic function in cardiomyocytes. Cardiac sarcoplasmic reticulum ...

Medicine

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 ...

Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry

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

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

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

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of intracellular calcium release are the major intracellular calcium storage organelles, the endoplasmic reticulum (ER) and the more specialized sarcoplasmic reticulum (SR) in muscle cells. The dynamic release of calcium from these organelles is mediated by the ryanodine receptor (RyR) and the inositol 1,4,5-triphosphate receptor (IP3R) with refilling occurring through the sarco/endoplasmic reticulum calcium ATPase (SERCA) pump. A genetically encoded calcium sensor (GECI) called CatchER was created to monitor the rapid calcium release from the ER/SR. Here, the detailed protocols for the transfection and expression of the improved, ER/SR-targeted GECI CatchER+ in HEK293 and C2C12 cells and its application in monitoring IP3R, RyR, and SERCA pump-mediated calcium transients in HEK293 cells using fluorescence microscopy is outlined. The receptor agonist or inhibitor of choice is dispersed in the chamber solution and the intensity changes are recorded in real time. With this method, a decrease in ER calcium is seen with RyR activation with 4-chloro-m-cresol (4-cmc), the indirect activation of IP3R with adenosine triphosphate (ATP), and inhibition of the SERCA pump with cyclopiazonic acid (CPA). We also discuss protocols for determining the in situ Kd and quantifying basal [Ca2+] in C2C12 cells. In summary, these protocols, used in conjunction with CatchER+, can elicit receptor mediated calcium release from the ER with future application in studying ER/SR calcium related pathologies.

Monitoring ER/SR Calcium Release with the Targeted Ca2+ Sensor CatchER+

We present protocols for the application of our targeted genetically-encoded calcium indicator (GECI) CatchER+ for monitoring rapid calcium transients in the endoplasmic/sarcoplasmic reticulum (ER/SR) of HEK293 and skeletal muscle C2C12 cells using real-time fluorescence microscopy. A protocol for the in situ Kd measurement and calibration is also discussed.

Intracellular calcium (Ca2+) transients evoked by extracellular stimuli initiate a multitude of biological processes in living organisms. At the center of ...

Utilizing the Precision-Cut Lung Slice to Study the Contractile Regulation of Airway and Intrapulmonary Arterial Smooth Muscle

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, respectively, hence playing a critical role in the homeostasis of the pulmonary system. Deregulation of SMC contractility contributes to several pulmonary diseases, including asthma and pulmonary hypertension. However, due to limited tissue access and a lack of culture systems to maintain in vivo SMC phenotypes, molecular mechanisms underlying the deregulated SMC contractility in these diseases remain fully identified. The precision-cut lung slice (PCLS) offers an ex vivo model that circumvents these technical difficulties. As a live, thin lung tissue section, the PCLS retains SMC in natural surroundings and allows in situ tracking of SMC contraction and intracellular Ca2+ signaling that regulates SMC contractility. Here, a detailed mouse PCLS preparation protocol is provided, which preserves intact airways and intrapulmonary arteries. This protocol involves two essential steps before subjecting the lung lobe to slicing: inflating the airway with low-melting-point agarose through the trachea and infilling pulmonary vessels with gelatin through the right ventricle. The PCLS prepared using this protocol can be used for bioassays to evaluate Ca2+-mediated contractile regulation of SMC in both the airway and the intrapulmonary arterial compartments. When applied to mouse models of respiratory diseases, this protocol enables the functional investigation of SMC, thereby providing insight into the underlying mechanism of SMC contractility deregulation in diseases.

The present protocol describes preparing and utilizing mouse precision-cut lung slices to assess the airway and intrapulmonary arterial smooth muscle contractility in a nearly in vivo milieu.

Smooth muscle cells (SMC) mediate the contraction of the airway and the intrapulmonary artery to modify airflow resistance and pulmonary circulation, ...

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

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