This work was supported by the National Institutes of Health, including the National Institute of General Medical Sciences, the Centers of Biomedical Research Excellence (COBRE), the Center for Studies of Host Response to Cancer Therapy [P20-GM109005], the National Cancer Institute [1R37CA282349-01], and the American Heart Association Predoctoral Fellowship [Award Number: 23PRE1020738; https://doi.org/10.58275/AHA.23PRE1020738.pc.gr.161089]. The content is solely the responsibility of the authors and does not necessarily represent the official views of the NIH or AHA.&#160;Figure 1 and Figure 3 were created with BioRender.com.

Nine to 13-week-old male Sprague-Dawley rats were purchased from a commercial vendor. After arrival, all rats were housed and maintained at the University of Arkansas for Medical Sciences (UAMS) Division of Lab Animal Medicine (DLAM) facility on a standard laboratory diet and exposed to 12 h of light:dark cycle at 25 oC. All procedures were carried out as per the approved animal use protocol #4127 by the Institutional Animal Care and Use Committee (IACUC) of UAMS.
1. Dissection and cannulation of mesenteric LVs
NOTE: It is important to set up the perfusion chamber prior to the isolation of the mesenteric LVs to make sure there is no interruption of flow or leak that would disrupt the experiment.
<ol>
	<li>Perfusion bath preparation
	<ol>
		<li>Purchase borosilicate glass micropipettes (1.2 mm outer diameter, 0.68 mm inner diameter, and pulled to an outer tip diameter of 75-100 &#181;m) from a commercial vendor. Cut and polish micropipettes (aka cannula) to approximately 1-2 cm in length for mounting into the isolated vessel perfusion chamber.</li>
		<li>Connect each mounted glass cannula in the vessel perfusion chamber to independent pressure transducers situated in line with independent gravity-fed pressure regulators using polyethylene tubing. 
		NOTE: This allows for independent manipulation of inflow and outflow pressures depending on the study design. Figure 1 shows this setup in detail.</li>
		<li>Backfill the perfusion chamber (5 mL), glass micropipettes, and independent gravity-fed pressure regulator along with polyethylene tubing with physiological salt solution (PSS; 119 mM NaCl; 24 mM NaHCO3; 1.17mM NaH2PO4; 4.7 mM KCl; 1.17 mM MgSO4; 5.5 mM C6H12O6 (glucose); 0.026 mM C10H16N2O8 (EDTA); and 1.6 mM CaCl2) devoid of any air bubbles. Then, clamp the pressure so that the cannulas are not pressurized. 
		NOTE: The pH of this solution is ~7.5. A pH of 7.4 is maintained in the perfusion bath utilizing CO2 bubbling to act on the bicarbonate buffering system within the PSS as described in step 1.3.7. EDTA is used here to chelate excess Ca2+ ions.</li>
	</ol>
	</li>
	<li>Preparation of knots
	<ol>
		<li>Tie double overhand knots under the microscope using braided silk suture thread (size 8-0). Key steps are illustrated in Figure 2.
		<ol>
			<li>Separate a single filament.</li>
			<li>Use two Dumont #5 forceps with microblunted tips and using one forceps make a double loop around the tip of the other forceps.</li>
			<li>With the looped forceps, grab the loose end of the suture and pull it through both loops making sure not to pull the knot completely closed and leaving a small opening.</li>
			<li>Use Vanna spring scissors to cut excess suture from either side. Use these knots later to secure the LV onto the cannulas. 
			NOTE: Do not use the same forceps you will dissect or cannulate with because there is more danger of damaging forceps tips during knot tying.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Isolation and cannulation of mesenteric LVs
	<ol>
		<li>Deeply anesthetize animals by administering a 5% isoflurane with 1.5 L/min O2 overdose and exsanguinate by decapitation.</li>
		<li>Isolate whole mesenteries for dissection of mesenteric LVs by first making a lengthwise cut along the midline of the abdominal wall, exteriorizing the mesentery, and then snipping the connection just below the pyloric sphincter and ~2-3 cm above the cecum as well as the connection to the rectum.</li>
		<li>Wash the dissected whole mesenteries in 200 mL of ice-cold PSS and then transfer to and pin down in a silicone-lined (8-10 mm) Petri-dish (100 mm) containing ice-cold PSS.</li>
		<li>Dissect second-order mesenteric LVs from surrounding fat and connective tissue using a stereomicroscope, dissection Dumont #5 Inox fine forceps, and Vanna spring scissors. To help identify the ends of the LVs once removed from the tissue, leave a small piece of fat on the proximal end of the LV.</li>
		<li>Transfer the dissected LVs to the vessel perfusion chamber for cannulation on glass cannulas.</li>
		<li>Use two presharpened Dumont #5 Inox Fine Forceps, straight tip (0.05 x 0.01 mm) to cannulate LVs with the distal end of the LV on the P1 cannula (inflow) and the proximal end of the vessel on the P2 cannula (outflow) to mimic the direction of lymph flow. 
		NOTE: P1 and P2 cannulas are identical and only differ by which end of the LV is connected. It is important to use forceps that meet perfectly at the tip and without damage to facilitate grabbing the thin vessel wall.
		<ol>
			<li>Slide a single pretied knot onto each glass cannula to later secure the vessel onto the cannulas.</li>
			<li>Using the small piece of fat to help orient LV direction, first cannulate the distal end onto P1.</li>
			<li>Slide the knot down the cannula and tighten to secure the LV. Make sure not to overtighten and break the cannula tip.</li>
			<li>Pressurize the LV to 4-5 mm Hg in the perfusion chamber. 
			NOTE: For our purposes, we set P1 and P2 pressure to be equal; however, depending on the experimental conditions, the pressure may be adjusted at each cannula to induce shear stress or backflow.</li>
			<li>Repeat steps 1.3.6.1-1.3.6.4 to cannulate the proximal end of the LV onto P2.</li>
		</ol>
		</li>
		<li>Place the bubbling stone with 7% carbon dioxide (CO2)/93% oxygen (O2) in the bath chamber to maintain physiological pH.</li>
		<li>Connect the chamber to the temperature regulator and set it at 37 oC for LVs to equilibrate and develop stable, spontaneous contractions (approximately 30 min). 
		NOTE: Figure 1 shows this setup in detail.</li>
	</ol>
	</li>
</ol>
2. Measurement of absolute concentrations of [Ca2+]i in LVs
<ol>
	<li>Fura-2AM staining of cannulated LVs
	<ol>
		<li>After spontaneous contractions develop (from step 1.3.8), incubate LVs with Fura-2-acetoxymethyl ester (Fura-2AM; 2 &#181;M or 10 &#181;L/5 mL) and pluronic acid (PA; 0.02% W/V or 5 &#181;L/5 mL of 20% PA) for 30 min in the dark. 
		NOTE: After the addition of Fura-2AM, all remaining steps must be performed in the dark.</li>
		<li>After 30 min, exchange the solution in the perfusion chamber 3x by emptying the complete bath volume with a negative pressure vacuum, replacing it with temperature-matched (37 oC) reagent-free PSS. 
		NOTE: This should be done quickly to minimize the time the LV is suspended in the air. Use both hands, for example, the right hand for vacuum and the left hand for replacing new reagent-free PSS.</li>
		<li>After washing, incubate LVs for 15 min in the dark to remove excess indicator and allow for de-esterification.</li>
	</ol>
	</li>
	<li>Capturing Ca2+ fluorescence and vessel diameter
	<ol>
		<li>Transfer the chamber onto the inverted fluorescent microscope stage equipped with an LED light source, a 20x S Fluor objective, a cell framing adaptor, and a fast CMOS video camera system, which enables frame-by-frame fluorescence capture at 15 Hz. 
		NOTE: A schematic of this workflow setup is presented in Figure 3. Ca2+ signal may be captured using a CCD camera with at least 15 fps.</li>
		<li>Connect the microscope to the computer equipped with imaging software to record fluorescence and edge detection. 
		NOTE: The referenced software also enables simultaneous pressure recording; however, that is not included here.</li>
		<li>Turn on the LED light source and Fluorescent System interface. 
		NOTE: The software instructions described here are for the referenced software, but other software can be used to obtain this data.</li>
		<li>Open software (IonWizard).</li>
		<li>Under File tab, select New.</li>
		<li>Under Collect tab, select Experiment.</li>
		<li>Load the desired experimental template and click OK. 
		NOTE: You will need to set up an experimental template with the parameters to be measured. Instructions for template setup can be found in the software manual29.</li>
		<li>Adjust the on-screen traces to see Vessel Diameter, Numerator (340 signal), Denominator (380 signal), and Ratio in descending order.</li>
		<li>Adjust the y-axis scale as needed to help visualize the traces.
		<ol>
			<li>Under Traces, select Edit User Limits. Ensure that Automatic Limits is unchecked.</li>
			<li>Select the parameter you wish to adjust, enter the minimum and maximum values for the axis, and then select OK.</li>
		</ol>
		</li>
		<li>To begin the experiment click START (at the bottom of the screen).</li>
		<li>To measure LV diameter simultaneously, use the edge-detection software incorporated into the imaging system, which generates contractile traces of LVs at 3 Hz. Use these measurements to analyze contractile and rhythmicity parameters described in section 3 and shown in Figure 4.
		<ol>
			<li>Be sure to adjust the lighting so that the LV wall appears as dark lines.</li>
			<li>Select a region of interest (ROI) that is devoid of fat and debris. Once the experiment has started, do not move this ROI.</li>
			<li>Ensure the threshold is set so that the vessel wall edge is detected throughout the entire contraction cycle.</li>
		</ol>
		</li>
		<li>For fluorescence measurements, turn on the Photomultiplier Tube (PMT).</li>
		<li>Excite Fura-2, a ratiometric indicator, by alternating 340 and 380 nm wavelengths in 50 ms exposures using LED illuminators, and capture the emission spectra at 510 nm at 15 Hz across the whole imaging field. 
		NOTE: It is important to keep all optical parameters (excitation settings, emission filters, objective lens, and dichroic mirrors) the same for an entire series of experiments to obtain reproducible Ca2+ measurements.</li>
		<li>Measure the signal-to-background ratio by first obtaining the 340 and 380 fluorescence with the LV in the middle of the field of view (signal) and then, using the stage manipulators to move the field of view to the edge of the bath with no vessel (taking care to avoid the silicone- lined edge and/or by removing any debris from the bath) to capture the background. 
		NOTE: It is important to navigate back to the original section of the vessel each time for Ca2+ measurements.</li>
		<li>Exchange the bath solution with temperature-matched reagent-free PSS to remove the excess indicator in the bath. It may take several bath exchanges to remove excess Fura-2, so repeat this exchange until the signal to background ratio is approximately 10:1.</li>
		<li>Record baseline Fura-2 fluorescence signal and spontaneous contractions for approximately 30 min followed by a cumulative concentration response of nifedipine (NIF; 0.1-100 nM), a voltage-dependent Cav1.x antagonist. Obtain background measurements for each drug concentration.</li>
		<li>At the end of each experiment, wash LVs with temperature-matched Ca2+-free PSS to obtain the minimum Fura-2 fluorescence signal (Rmin) and maximum diameter of the LVs in the absence of Ca2+. Be sure to measure the background. 
		NOTE: The Ca2+-free PSS has the same composition as PSS but without CaCl2, and EDTA is replaced with 1 mM EGTA (C14H24N2O10) (pH ~7.5).</li>
		<li>Exchange the bath solution with temperature-matched PSS containing 10 mM Ca2+ and ionomycin (10 &#181;M IONO), a Ca2+ ionophore, to obtain the maximum Fura-2 fluorescence signal (Rmax) and minimum diameter of the LVs in conditions of saturating Ca2+. Be sure to measure the background.</li>
	</ol>
	</li>
	<li>The formula for measuring absolute [Ca2+]i&#8203;
	<ol>
		<li>Use Rmin and Rmax to calibrate the ratio of 340 and 380 nm wavelengths and calculate absolute cytosolic free Ca2+ concentration ([Ca2+]i).</li>
		<li>Calculate absolute cytosolic free Ca2+ concentration ([Ca2+]i) using equation (1)26: 
		<img alt="Equation 1" src="/files/ftp_upload/66535/66535eq01.jpg" style="margin: auto;" />&#160; &#160; &#160;(1) 
		Where Kd = 225 nM (dissociation constant for Fura-2)26, R = 340/380 ratio, Rmin = 340/380 ratio in the absence of Ca2+, Rmax = 340/380 ratio with saturating Ca2+ conditions, Sfree = 380 signal in the absence of Ca2+, Sbound = 380 signal with saturating Ca2+ conditions 
		NOTE: All fluorescent signals were corrected for background fluorescence.</li>
		<li>Figure 5 is an example Ca2+ trace detailing which parameters are recorded. Define baseline Ca2+ as the lowest resting Ca2+ prior to the Ca2+ spike and peak Ca2+ as the highest Ca2+ achieved during the Ca2+ spike. Amplitude is the difference between Peak and Baseline Ca2+. Export all parameters directly from the imaging software, except the frequency of Ca2+ spike which is to be calculated offline as the number of Ca2+ spikes/s. Below are key steps within the referenced software to obtain these parameters. 
		NOTE: Entire traces can be exported into a .txt file and all parameters can be analyzed or calculated in your software of choice.</li>
		<li>For Rmin, highlight the section of the Numerator trace that corresponds to the background during Ca2+-free PSS. A dialog box will open and provide the values for this portion of the trace.</li>
		<li>Under Operations, select Constants.</li>
		<li>Select Calcium-Numeric background and input the background numbers for the Numerator and Denominator from the previous step. Click OK.</li>
		<li>Highlight the section of the Ratio trace that corresponds to the lowest ratio during Ca2+-free PSS. This is Rmin. Also make note of the Denominator value for this section; this is Sfree.</li>
		<li>Repeat steps 2.3.4 through 2.3.7 for Rmax and Sbound with the section of the trace that corresponds to high Ca2+ free PSS and highest ratio.</li>
		<li>Under Operations, select Constants.</li>
		<li>Select Calcium-Calcium Calibration and input the values listed in Equation 1. Click OK.</li>
		<li>Adjust one of the on-screen traces as described in Step 2.2.8 so that you can see Calcium-Numeric subtracted Calcium.</li>
		<li>Perform a Monotonic Transient Analysis to acquire the remaining parameters. Instructions for this can be found in the software manual30.</li>
		<li>Alternatively, Under Export, select Current Trace.</li>
		<li>Select the location you would like to export the .txt file to and click OK. 
		NOTE: Make sure to click on the individual trace you want to export. You can export the entire trace or selected sections of the trace.</li>
	</ol>
	</li>
</ol>
3. Measurement of LV contractility and rhythmicity
<ol>
	<li>The edge-detection software incorporated into the imaging system generates contractile traces for LV diameter measurements as described above. Use these measurements to analyze contractile and rhythmicity parameters. Figure 4 is an example contractile trace detailing which contractile parameters are to be recorded. Export all parameters directly from the imaging software using the Monotonic Transient Analysis function on the Diameter trace30, except frequency of contractions, calculated flow, and interval, which are to be calculated offline. 
	NOTE: Entire traces can be exported into a .txt file and all parameters can be analyzed or calculated in your software of choice using the equations below.</li>
	<li>EDD, ESD, amplitude, frequency, and calculated flow measurements
	<ol>
		<li>Measure the maximum and minimum diameters (end diastolic diameter [EDD] and end systolic diameter [ESD], respectively) that the LVs can attain during their rhythmic and spontaneous contractions.</li>
		<li>Calculate the amplitude of contractions (AMP) as the difference between EDD and ESD.</li>
		<li>Calculate the frequency as the number of contractions per measurement period (in s).</li>
		<li>Calculate the calculated flow per &#181;m using equation (2): 
		Calculated flow = &#960;/4(EDD2- ESD2)F&#160; &#160; &#160;&#160;(2) 
		Where EDD2 = vessel cross-sectional area during relaxed state, ESD2 = measure of the vessel cross-sectional area during constriction, F = frequency of contractions/s</li>
	</ol>
	</li>
	<li>Rhythmicity: contraction and relaxation time:
	<ol>
		<li>Other measures of LV rhythmicity are contraction time and relaxation time.</li>
		<li>Define contraction time as the time taken by the LV to reach ESD with each contraction.</li>
		<li>Define relaxation time as the time taken by the LV to reach the EDD for each relaxation, which will give an overall indication of rhythmicity correlated with absolute [Ca2+]i within a specific time frame.</li>
		<li>Calculate the interval time (&#916;T) from equation (3). 
		&#916;t = t2 (ESD2)-t1 (ESD1)&#160;(3)</li>
	</ol>
	</li>
</ol>

Real-Time Analysis of Calcium and Contractility in Isolated Lymph Vessels

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M., Breslin, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+cytosolic+Ca2++in+isolated+contractile+lymphatics.">Measurement of cytosolic Ca2+ in isolated contractile lymphatics.</a> J Vis Exp. (58), e3438 (2011).</li><li>Grynkiewicz, G., Poenie, M., 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=A+new+generation+of+Ca2++indicators+with+greatly+improved+fluorescence+properties.">A new generation of Ca2+ indicators with greatly improved fluorescence properties.</a> J Biol Chem. 260 (6), 3440-3450 (1985).</li><li>Hill-Eubanks, D. C., Werner, M. E., Heppner, T. J., Nelson, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+signaling+in+smooth+muscle.">Calcium signaling in smooth muscle.</a> Cold Spring Harb Perspect Biol. 3 (9), 1-20 (2011).</li><li>Harmer, A. R., Abi-Gerges, N., Morton, M. J., Pullen, G. F., Valentin, J. P., Pollard, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Validation+of+an+in+vitro+contractility+assay+using+canine+ventricular+myocytes.">Validation of an in vitro contractility assay using canine ventricular myocytes.</a> Toxicol Appl Pharmacol. 260 (2), 162-172 (2012).</li><li>IonOptix LLC. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> IonWizard 7.2 Acquisition. , (2017).</li><li>IonOptix LLC. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=IonWizard+7.2+Core+&amp;+Analysis+Functions.">IonWizard 7.2 Core &amp; Analysis Functions.</a> IonOptix LLC. , (2017).</li><li>Roe, M. W., Lemasters, J. J., Herman, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+Fura-2+for+measurements+of+cytosolic+free+calcium.">Assessment of Fura-2 for measurements of cytosolic free calcium.</a> Cell Calcium. 11 (2-3), 63-73 (1990).</li><li>Zawieja, S. D., 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=Ano1+mediates+pressure-sensitive+contraction+frequency+changes+in+mouse+lymphatic+collecting+vessels.">Ano1 mediates pressure-sensitive contraction frequency changes in mouse lymphatic collecting vessels.</a> J Gen Physiol. 151 (4), 532-554 (2019).</li><li>Behringer, E. J., 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=Calcium+and+electrical+dynamics+in+lymphatic+endothelium.">Calcium and electrical dynamics in lymphatic endothelium.</a> J Physiol. 595 (24), 7347-7368 (2017).</li><li>Ferrusi, I., Zhao, J., van Helden, D., vonder Weid, P. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclopiazonic+acid+decreases+spontaneous+transient+depolarizations+in+guinea+pig+mesenteric+lymphatic+vessels+in+endothelium-dependent+and+-independent+manners.">Cyclopiazonic acid decreases spontaneous transient depolarizations in guinea pig mesenteric lymphatic vessels in endothelium-dependent and -independent manners.</a> Am J Physiol Heart Circ Physiol. 286 (6), H2287-H2295 (2004).</li></ol>

The authors have no conflicts of interest to disclose.

Due to the fragile and diminutive nature of LVs, it is imperative to exercise the utmost care during both dissection and cannulation processes. Even minor damage to the vessel could lead to the development of a non-viable LV or give rise to abnormalities in [Ca2+]i transients. Consistency in excitation settings is equally crucial throughout the entire experimental series to ensure comparability in [Ca2+]i measurements between control and treated groups. Failing to maintain uniform settings poses a substantial risk of over- or underestimating [Ca2+]i across vessels within an experimental series. Similarly, it is equally important to accurately identify and monitor the same vessel region throughout each experiment.
The use of the ratiometric indicator Fura-2AM normalizes fluorescence variations caused by uneven tissue thickness, fluorophore distribution/leakage, or photobleaching, issues common with single wavelength dyes.31 This enables the continuous monitoring described in this protocol. However, since Fura-2 works by chelating Ca2+, it is possible to overload the LVs and reduce the [Ca2+]i available for contraction or drug response. In these cases, Ca2+ spikes may still be observed while rhythmic contractions are absent. Varying LV length also may contribute to this phenomenon. While these Ca2+ measurements likely may still be valid, it may be necessary to reduce the concentration of Fura-2AM in replicated setups to successfully achieve both Ca2+ and diameter measurements. Our results only include LVs for which both Ca2+ spikes and rhythmic contractions were present at baseline.
Measuring Rmin and Rmax are critical steps in calculating absolute [Ca2+]i. Because Rmin should be the Fura-2 ratio in the absence of Ca2+, a high concentration of EGTA has been added to the Ca2+-free PSS to ensure the chelation of any residual Ca2+. Initial studies were conducted with EDTA in the Ca2+-free PSS, and this resulted in sporadic vessel contractions with corresponding Ca2+ spikes. For Rmax, a high concentration of Ca2+ has been added to the PSS along with an ionophore, ionomycin, to maximize the [Ca2+]i signal. The high Ca2+ solution may precipitate, which may require the removal of the EDTA from the PSS. Importantly, these additional measurements of Rmin and Rmax provide the opportunity to evaluate physiologically relevant changes in [Ca2+]i, which can provide information on membrane excitability and contractility mechanisms27 as well as allow for baseline comparisons between experimental groups compared to protocols that only report 340/380 ratio for Fura-2. Failure to achieve adequate Rmin and Rmax values preclude the ability to calculate absolute [Ca2+]i.
Due to the contractile nature of the LVs, this method can only provide a measure of global Ca2+ levels rather than local Ca2+ release events that can be measured in paralyzed vessels32. However, this method is advantageous to correlate changes in absolute [Ca2+]i dynamics with contractility compared to methods using paralyzed vessels or individual cells28,32. For this approach, it is assumed that the majority of the Ca2+ measured originates from the lymph muscle cells. However, endothelial cells, which are also present in these isolated LVs, may contribute to the total Ca2+ signal observed33. This contribution could be estimated using LVs that have been denuded of endothelium34. LV contractions also may result in the vessel wall shifting slightly in and out of focus during the contraction cycle. Therefore, it is important to use short vessel segments that can be pulled taut but without stretching the vessel.
Beyond its application in LVs, this method could be used to study isolated vessels from other vascular beds, including arterioles and veins, and holds promise for potential utilization in neurobiology and other branches of vascular biology. Exploring the effects of various agonists or antagonists targeting different signal transduction pathways is another avenue for investigating underlying Ca2+ dynamics. Furthermore, this technique also can be used for comparative research involving control and treated samples from respective animals. Moreover, this approach is adaptable for implementation at the cellular level, such as in isolated lymphatic muscle cells, requiring minimal adjustments to the perfusion chamber and microscope objectives. In summary, this method provides physiologically relevant insight into global Ca2+ dynamics as it correlates to contractility and rhythmicity in LVs and provides a robust assessment of potential regulators of Ca2+ dynamics in collecting LVs.

The lymphatic vasculature is found in many organ systems including the brain, heart, lungs, kidney, and mesentery1,2,3,4,5,6, and operates by propelling fluid (lymph) from the interstitial spaces to the central venous ducts to maintain fluid homeostasis7,8,9,10. It starts with blind-ended lymphatic capillaries within the vascular capillary beds that drain into collecting lymph vessels (LVs). Collecting LVs are made of two layers of cells: a layer of endothelial cells encompassed by a layer of lymphatic muscle cells (LMCs)10,11. Lymph fluid transport is achieved through both extrinsic forces (e.g. new lymph formation, arterial pulsations, central venous pressure fluctuations) and intrinsic forces12.
The intrinsic force for lymph transport is the spontaneous rhythmic contraction of collecting LVs, which is the focus of the majority of studies investigating lymphatic function. This intrinsic lymphatic pump is principally regulated by the cyclic rise and fall of cytosolic, free Ca2+ ([Ca2+]i). Spontaneous depolarization of the plasma membrane in LMCs activates voltage-gated &#34;L-type&#34; Ca2+ (Cav1.x) channels triggering Ca2+ influx and subsequent LV rhythmic contraction8,9,10. This role was demonstrated by blocking Cav1.x with specific agents, like nifedipine, which inhibited LV contractions and caused vessel dilation13,14. The transient rise in [Ca2+]i or &#34;Ca2+ spike&#34; in the LMCs mediated by Cav1.x channels also may mobilize intracellular Ca2+ stores by activating inositol triphosphate (IP3) receptors and ryanodine receptors (RyRs) on the sarcoplasmic reticulum (SR)15,16,17,18. Current evidence suggests IP3 receptors contribute more Ca2+ required for normal LV contractions compared to RyRs15,16,19,20,21; however, RyRs may play a role during pathology or in response to pharmaceutical intervention17,18. Additionally, the activation of Ca2+-activated K+ channels22 and ATP-sensitive potassium (KATP) channels23,24 can hyperpolarize the LMC membrane and inhibit spontaneous contractile activity.
There are many other ion channels and proteins that may regulate Ca2+ dynamics in collecting LVs. Utilizing methods to study changes in Ca2+ and vessel contractility in response to pharmacological agents in real time is important to understand these potential regulators. An earlier method using Fura-2 to measure relative changes in LV [Ca2+]i has been described25. Because the dissociation constant for Fura-2 and Ca2+ is known26, it is possible to calculate actual concentrations of Ca2+, which broadens the application of this method and provides additional insight into Ca2+ signaling, membrane excitability, and contractility mechanisms27, as well as allowing for baseline comparisons between experimental groups. This latter approach has been used in cardiomyocytes28, and therefore, can be adapted to LVs. This paper presents an improved method that combines these two approaches to measure and calculate changes in absolute [Ca2+]i as well as vessel contractility/rhythmicity continuously in real time in isolated, pressurized LVs. We also provide representative results for LVs treated with nifedipine.

<table><tbody><tr><td>20x S Fluor objective</td><td>Olympus Corporation of the Americas (Center Valley, PA, United States)</td><td>UPlanSApo</td><td></td></tr><tr><td>Borosilicate glass micropipettes</td><td>Living Systems Instrumentation (Burlington, VT, United States)</td><td>GCP-75-100</td><td></td></tr><tr><td>Calcium chloride (CaCl2)</td><td>Fisher Bioreagents (Waltham, MA, United States)</td><td>BP510-500</td><td></td></tr><tr><td>Carbon dioxide (CO2)</td><td>nexAir (Memphis, TN, United States)</td><td>UN3156</td><td></td></tr><tr><td>Dissection forceps</td><td>Fine Science Tools (Foster City, CA, United States)</td><td>11254-20</td><td></td></tr><tr><td>EDTA (C10H16N2O8)</td><td>Fisher Bioreagents (Waltham, MA, United States)</td><td>BP118-500</td><td></td></tr><tr><td>EGTA (C14H24N2O10)</td><td>Fisher Bioreagents (Waltham, MA, United States)</td><td>O2783-100</td><td></td></tr><tr><td>Fura-2AM</td><td>Invitrogen (Waltham, MA, United States)</td><td>F1221</td><td></td></tr><tr><td>Glucose (C6H12O6)</td><td>Fisher Bioreagents (Waltham, MA, United States)</td><td>D16-500</td><td></td></tr><tr><td>Gravity-Fed Pressure regulator</td><td>custom-made in the lab</td><td></td><td></td></tr><tr><td>Heating unit</td><td>Living Systems Instrumentation (Burlington, VT, United States)</td><td>TC-09S</td><td></td></tr><tr><td>Imaging software</td><td>IonOptix (Westwood, MA, United States)</td><td></td><td></td></tr><tr><td>Inverted fluorescent microscope</td><td>Olympus Corporation of the Americas (Center Valley, PA, United States)</td><td>IX73</td><td></td></tr><tr><td>Ionomycin</td><td>Invitrogen (Waltham, MA, United States)</td><td>I24222</td><td></td></tr><tr><td>IonOptix Cell Framing Adaptor</td><td>IonOptix (Westwood, MA, United States)</td><td>665 DXR</td><td></td></tr><tr><td>Isoflurane</td><td>Piramal Critical Care (Telangana, India)</td><td>NDC 66794-017-10</td><td></td></tr><tr><td>Isolated vessel perfusion chamber</td><td>Living Systems Instrumentation (Burlington, VT, United States)</td><td>CH-1</td><td></td></tr><tr><td>Knot preparation forceps</td><td>Fine Science Tools (Foster City, CA, United States)</td><td>11253-20</td><td></td></tr><tr><td>LED light source</td><td>Olympus Corporation of the Americas (Center Valley, PA, United States)</td><td>TL4</td><td></td></tr><tr><td>Magnesium sulfate (MgSO4)</td><td>Acros Organics (New Jersey, NJ, Unites States)</td><td>213115000</td><td></td></tr><tr><td>MyoCam-S3 Fast CMOS video system</td><td>IonOptix (Westwood, MA, United States)</td><td>MCS300</td><td></td></tr><tr><td>Nifedipine</td><td>Sigma (St. Louis, MO, United States)</td><td>N7634</td><td></td></tr><tr><td>Ophthalmic sutures</td><td></td><td></td><td></td></tr><tr><td>Oxygen (O2)</td><td>nexAir (Memphis, TN, United States)</td><td>UN1072</td><td></td></tr><tr><td>Pluronic acid</td><td>Sigma (St. Louis, MO, United States)</td><td>P2443</td><td></td></tr><tr><td>Potassium chloride (KCl)</td><td>Fisher Bioreagents (Waltham, MA, United States</td><td>BP366-500</td><td></td></tr><tr><td>Pressure monitor system</td><td>Living Systems Instrumentation (Burlington, VT, United States)</td><td>PM-4</td><td></td></tr><tr><td>Pressure Transducer</td><td>Living Systems Instrumentation (Burlington, VT, United States)</td><td>PT-F</td><td></td></tr><tr><td>Silicone-lined petri-dish</td><td>custom-made in the lab</td><td></td><td></td></tr><tr><td>Sodium bicarbonate (NaHCO3)</td><td>Fisher Bioreagents (Waltham, MA, United States</td><td>BP328-500</td><td></td></tr><tr><td>Sodium chloride (NaCl)</td><td>Fisher Bioreagents (Waltham, MA, United States</td><td>BP358-212</td><td></td></tr><tr><td>Sodium phosphate (NaH2PO4)</td><td>Fisher Bioreagents (Waltham, MA, United States</td><td>BP329-500</td><td></td></tr><tr><td>Sprague-Dawley rats</td><td>Envigo RMS (Indianapolis, IN, USA)</td><td>Male</td><td>9-13 weeks old</td></tr><tr><td>Stereomicroscope</td><td>Leica Microsystems (Wetzlar, Germany)</td><td>S9D</td><td></td></tr><tr><td>Vannas spring scissors</td><td>Fine Science Tools (Foster City, CA, United States)</td><td>15000-03</td><td></td></tr></tbody></table>

real-time evaluation of absolute, cytosolic, free ca2+ and corresponding contractility in isolated, pressurized lymph vessels

The lymphatic vasculature, now often referred to as &#34;the third circulation,&#34; is located in many vital organ systems. A principal mechanical function of the lymphatic vasculature is to return fluid from extracellular spaces back to the central venous ducts. Lymph transport is mediated by spontaneous rhythmic contractions of lymph vessels (LVs). LV contractions are largely regulated by the cyclic rise and fall of cytosolic, free calcium ([Ca2+]i).
This paper presents a method to concurrently calculate changes in absolute concentrations of [Ca2+]i and vessel contractility/rhythmicity in real time in isolated, pressurized LVs. Using isolated rat mesenteric LVs, we studied changes in [Ca2+]i and contractility/rhythmicity in response to drug addition. Isolated LVs were loaded with the ratiometric Ca2+-sensing indicator Fura-2AM, and video microscopy coupled with edge-detection software was used to capture [Ca2+]i&#160;and diameter measurements continuously in real time.
The Fura-2AM signal from each LV was calibrated to the minimum and maximum signal for each vessel and used to calculate absolute [Ca2+]i. Diameter measurements were used to calculate contractile parameters (amplitude, end diastolic diameter, end systolic diameter, calculated flow) and rhythmicity (frequency, contraction time, relaxation time) and correlated with absolute [Ca2+]i measurements.

Contractility of LVs and corresponding alterations in cytosolic, free Ca2+ ([Ca2+]i) were assessed in isolated rat mesenteric LVs upon exposure to varying concentrations of nifedipine (NIF; 0.1-100 nM) (Figure 6). The parameters, including Ca2+ spike amplitude, baseline Ca2+, and peak Ca2+, exhibited a concentration-dependent reduction with the incremental addition of NIF to the perfusion chamber (Figure 7A). Concurrently, contractile parameters such as contraction amplitude and calculated flow also demonstrated a stepwise decrease (Figure 7B). There was a small increase in EDD diameter with NIF (Figure 7B). Ca2+ spike frequency and contraction frequency appear to be an all-or-none response. However, this effect occurred at 10 nM for one LV, while all LVs had ceased contractions by 100 nM. Thus, the combined data generate graphs that resemble a graded concentration response. This effect is consistent with earlier publications that used NIF on LVs in other preparations (wire and pressure myography13,14). Manhattan plots show the individual LV responses for measures of rhythmicity, including interval, contraction time, and relaxation time (Figure 7C). This type of data representation allows the researcher to tease out these all-or-none responses or variability in contraction rhythms to provide additional insight into underlying mechanisms. Ultimately, the decreases in contraction amplitude and frequency resulted in a reduction in calculated flow through these isolated LVs, which serves as a surrogate indicator for in vivo function. Overall, the decline in LV contractility correlated with the reduction in [Ca2+]i. Our findings provide direct evidence that within the 100 nM range, NIF effectively halted contractions and [Ca2+]i oscillations in LVs by antagonizing Cav1.x channels present in lymph muscle cells (LMCs).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66535/66535fig01v2.jpg" /> 
Figure 1: Image of the isolated vessel chamber setup. Vessel perfusion studies used an isolated vessel chamber equipped with a thermoregulator. Gravity was used to control pressure via a PSS reservoir. Pressure was monitored by transducers connected to both inflow (P1) and outflow (P2) cannulas. Abbreviation: PSS = physiological salt solution. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig01largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66535/66535fig02.jpg" /> 
Figure 2: Preparation of knot at-a-glance. (A) Double loop preparation under the dissection microscope&#160;using a single filament of 3-ply silk suture thread, (B) grabbing the loose end and pulling it through both loops, (C) pulling the knot from both ends to keep a small opening, and (D) cutting the excess filament from either side and the blue box showing a ready to use complete double overhand knot. Scale bar = 1.5 mm. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66535/66535fig03.jpg" /> 
Figure 3: Schematic of experimental workflow for data acquisition. A healthy rat was anesthetized with 5% isoflurane induction and decapitation was performed to remove trunk blood. A midline incision was performed to expose and isolate the mesentery. The isolated mesentery was spread out in ice-cold PSS solution and a&#160;LV was dissected free from fat for cannulation in an isolated vessel perfusion chamber. The bath was placed on the stage of the inverted microscope using a 20x objective lens. The vessel was excited alternatively with 340 and 380 nm wavelength light and the emission fluorescent spectra were collected using a CCD camera at 510 nm. The computer connected to the microscope generated the contractile and Ca2+ traces using fluorescence capture and edge detection imaging software. Scale bar = 1 mm. Abbreviations: PSS = physiological salt solution; LV = lymph vessel; CCD = charge-coupled device. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66535/66535fig04.jpg" /> 
Figure 4: Representative LV contractile trace. (A) Example recording of changes in diameter of cannulated LVs loaded with Ca2+ imaging indicator Fura 2 AM in PSS and (B) a zoomed-in trace to show all the parameters related to vessel contractility: EDD, ESD, AMP, and frequency. These values were used to calculate rhythmicity and flow. Abbreviations: PSS = physiological salt solution; LV = lymph vessel; EDD = end-diastolic diameter; ESD = end-systolic diameter; AMP = amplitude. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66535/66535fig05.jpg" /> 
Figure 5: Representative LV Ca2+ imaging trace. (A) Example recording of changes in absolute&#160;[Ca2+]i in cannulated LVs loaded with Fura-2 in PSS and (B) a zoomed-in trace to show all the parameters (Peak, Amplitude, and Baseline) related to [Ca2+]i (not background-corrected). Abbreviation: PSS = physiological salt solution. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/66535/66535fig06.jpg" /> 
Figure 6: LV contractility and Ca2+ imaging at a glance. Representative traces corresponding to (A) diameter, (B) 340/380 ratio, and (C) absolute [Ca2+]i of PSS baseline, nifedipine, a Cav1.x (Ca2+) channel antagonist, concentration response, including Rmin and Rmax. Abbreviations: PSS = physiological salt solution; LV = lymph vessel; NIF = nifedipine; Rmin = minimum Fura-2 fluorescence signal; Rmax = maximum Fura-2 fluorescence signal. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/66535/66535fig07.jpg" /> 
Figure 7: Ca2+ oscillation and corresponding contractility blocked by nifedipine in LVs. (A) Ca2+ (n = 3) and (B) contractile (n = 3) parameters decreased in a concentration-dependent manner with the addition of nifedipine, a voltage-dependent Cav1.x (Ca2+) channel antagonist. (C) Representative Manhattan plots show mean time Interval (&#916;t) between contractions and contraction and relaxation times. Data presented as mean &#177; SEM. Abbreviations: PSS = physiological salt solution; LV = lymph vessel; NIF = nifedipine; EDD = end-diastolic diameter; AMP = amplitude. <a href="https://www.jove.com/files/ftp_upload/66535/66535fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Read this Scientific Article Protocol about Author Spotlight: Innovative Methods in Lymphedema and Hypertension Research at JoVE.com

Author Spotlight: Innovative Methods in Lymphedema and Hypertension Research

This protocol describes a method to simultaneously measure cytosolic, free calcium [Ca2+]i and vessel diameter in contracting lymph vessels in real time and then calculate absolute Ca2+ concentrations as well as contractility/rhythmicity parameters. This protocol can be used to study Ca2+ and contractile dynamics across a variety of experimental conditions.

Real-time Evaluation of Absolute, Cytosolic, Free Ca2+ and Corresponding Contractility in Isolated, Pressurized Lymph Vessels

The lymphatic vasculature, now often referred to as &#34;the third circulation,&#34; is located in many vital organ systems. A principal mechanical ...

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1.2K Views.  University of Arkansas for Medical Sciences. We bestuderen mechanismen van samentrekkingen van lymfevaten en lymfestroom om de onderliggende pathologie van kankergerelateerd lymfoedeem beter te begrijpen, evenals het verband tussen het lymfestelsel en hypertensie. Ons doel is om therapeutische doelen binnen het lymfevastraat te identificeren, zodat we nieuwe behandelingen voor lymfeziekten kunnen ontwikkelen. Recente ontdekkingen van verschillende ang-kanalen in lymfespier- en endotheelcellen, die bijdragen aan de onderliggende calciumd...

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Measurement of Cytosolic Ca2+ in Isolated Contractile Lymphatics

Lymphatic vessels comprise a multifunctional transport system that maintains fluid homeostasis, delivers lipids to the central circulation, and acts as a surveillance system for potentially harmful antigens, optimizing mucosal immunity and adaptive immune responses1. Lymph is formed from interstitial fluid that enters blind-ended initial lymphatics, and then is transported against a pressure gradient in larger collecting lymphatics. Each collecting lymphatic is made up of a series of segments called lymphangions, separated by bicuspid valves that prevent backflow. Each lymphangion possesses a contractile cycle that propels lymph against a pressure gradient toward the central circulation2. This phasic contractile pattern is analogous to the cardiac cycle, with systolic and diastolic phases, and with a lower contraction frequency4. In addition, lymphatic smooth muscle generates tone and displays myogenic constriction and dilation in response to increases and decreases in luminal pressure, respectively5. A hybrid of molecular mechanisms that support both the phasic and tonic contractility of lymphatics are thus proposed.
Contraction of smooth muscle is generally regulated by the cytosolic Ca2+ concentration ([Ca2+]i) plus sensitivity to Ca2+, of the contractile elements in response to changes in the environment surrounding the cell6. [Ca2+]i is determined by the combination of the movement of Ca2+ through plasma membrane ligand or voltage gated Ca2+ channels and the release and uptake of Ca2+ from internal stores. Cytosolic Ca2+ binds to calmodulin and activates enzymes such as myosin light chain (MLC) kinase (MLCK), which in turn phosphorylates MLC leading to actin-myosin-mediated contraction8. However, the sensitivity of this pathway to Ca2+ can be regulated by the MLC phosphatase (MLCP)9. MLCP activity is regulated by Rho kinase (ROCK) and the myosin phosphatase inhibitor protein CPI-17.
Here, we present a method to evaluate changes in [Ca2+]i over time in isolated, perfused lymphatics in order to study Ca2+-dependent and Ca2+-sensitizing mechanisms of lymphatic smooth muscle contraction. Using isolated rat mesenteric collecting lymphatics we studied stretch-induced changes in [Ca2+]i and contractile activity. The isolated lymphatic model offers the advantage that pressure, flow, and the chemical composition of the bath solution can be tightly controlled. [Ca2+]i was determined by loading lymphatics with the ratiometric, Ca2+-binding dye Fura-2. These studies will provide a new approach to the broader problem of studying the different molecular mechanisms that regulate phasic contractions versus tonic constriction in lymphatic smooth muscle.

Measurement of Cytosolic Ca2+ in Isolated Contractile Lymphatics

We introduce an approach to evaluate the cytosolic Ca2+ concentration in isolated lymphatics to study Ca2+-dependent and Ca2+-sensitizing mechanisms of lymphatic smooth muscle contraction.

Lymphatic vessels comprise a multifunctional transport system that maintains fluid homeostasis, delivers lipids to the central circulation, and acts as a ...

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 ...

Biology

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 ...

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 ...

Assessment of Myofilament Ca2+ Sensitivity Underlying Cardiac Excitation-contraction Coupling

Heart failure and cardiac arrhythmias are the leading causes of mortality and morbidity worldwide. However, the mechanism of pathogenesis and myocardial malfunction in the diseased heart remains to be fully clarified. Recent compelling evidence demonstrates that changes in the myofilament Ca2+ sensitivity affect intracellular Ca2+ homeostasis and ion channel activities in cardiac myocytes, the essential mechanisms responsible for the cardiac action potential and contraction in healthy and diseased hearts. Indeed, activities of ion channels and transporters underlying cardiac action potentials (e.g., Na+, Ca2+ and K+ channels and the Na+-Ca2+ exchanger) and intracellular Ca2+ handling proteins (e.g., ryanodine receptors and Ca2+-ATPase in sarcoplasmic reticulum (SERCA2a) or phospholamban and its phosphorylation) are conventionally measured to evaluate the fundamental mechanisms of cardiac excitation-contraction (E-C) coupling. Both electrical activities in the membrane and intracellular Ca2+ changes are the trigger signals of E-C coupling, whereas myofilament is the functional unit of contraction and relaxation, and myofilament Ca2+ sensitivity is imperative in the implementation of myofibril performance. Nevertheless, few studies incorporate myofilament Ca2+ sensitivity into the functional analysis of the myocardium unless it is the focus of the study. Here, we describe a protocol that measures sarcomere shortening/re-lengthening and the intracellular Ca2+ level using Fura-2 AM (ratiometric detection) and evaluate the changes of myofilament Ca2+ sensitivity in cardiac myocytes from rat hearts. The main aim is to emphasize that myofilament Ca2+ sensitivity should be taken into consideration in E-C coupling for mechanistic analysis. Comprehensive investigation of ion channels, ion transporters, intracellular Ca2+ handling, and myofilament Ca2+ sensitivity that underlie myocyte contractility in healthy and diseased hearts will provide valuable information for designing more effective strategies of translational and therapeutic value.

Assessment of Myofilament Ca2+ Sensitivity Underlying Cardiac Excitation-contraction Coupling

This paper describes a protocol that assesses the changes of myofilament Ca2+ sensitivity during contraction in isolated cardiac myocytes from rat heart. Together with cardiac electrophysiology, systolic/diastolic cytosol Ca2+ levels and contraction/relaxation, this measurement is imperative in underpinning the mechanisms mediating cardiac excitation-contraction coupling in healthy and diseased hearts.

Heart failure and cardiac arrhythmias are the leading causes of mortality and morbidity worldwide. However, the mechanism of pathogenesis and myocardial ...

Isolating Myofibrils from Skeletal Muscle Biopsies and Determining Contractile Function with a Nano-Newton Resolution Force Transducer

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of serially linked sarcomeres, the smallest contractile units in muscle. Sarcomeric dysfunction contributes to muscle weakness in patients with mutations in genes encoding for sarcomeric proteins. The study of myofibril mechanics allows for the assessment of actin-myosin interactions without potential confounding effects of damaged, adjacent myofibrils when measuring the contractility of single muscle fibers. Ultrastructural damage and misalignment of myofibrils might contribute to impaired contractility. If structural damage is present in the myofibrils, they likely break during the isolation procedure or during the experiment. Furthermore, studies in myofibrils provide the assessment of actin-myosin interactions in the presence of the geometrical constraints of the sarcomeres. For instance, measurements in myofibrils can elucidate whether myofibrillar dysfunction is the primary effect of a mutation in a sarcomeric protein. In addition, perfusion with calcium solutions or compounds is almost instant due to the small diameter of the myofibril. This makes myofibrils eminently suitable to measure the rates of activation and relaxation during force production. The protocol described in this paper employs an optical force probe based on the principle of a Fabry-P&#233;rot interferometer capable of measuring forces in the nano-Newton range, coupled to a piezo length motor and a fast-step perfusion system. This setup enables the study of myofibril mechanics with high resolution force measurements.

Presented here is a protocol to assess the contractile properties of striated muscle myofibrils with nano-Newton resolution. The protocol employs a setup with an interferometry-based, optical force probe. This setup generates data with a high signal-to-noise ratio and enables the assessment of the contractile kinetics of myofibrils.

Striated muscle cells are indispensable for the activity of humans and animals. Single muscle fibers are comprised of myofibrils, which consist of ...

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and calcium handling. For this purpose, atrial or ventricular cardiomyocytes of high quality are necessary to perform patch-clamp measurements or to explore calcium handling abnormalities. However, the limited yield of high-quality cardiomyocytes obtained by current isolation protocols does not allow both measurements in the same mouse. This article describes a method to isolate high-quality murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, for subsequent simultaneous measurements of calcium transients and L-type calcium current from one animal. Mouse hearts are obtained, and the aorta is rapidly cannulated to remove blood. Hearts are then initially perfused with a calcium-free solution (37 &#176;C) to dissociate the tissue at the level of intercalated discs and afterwards with an enzyme solution containing little calcium to disrupt extracellular matrix (37 &#176;C). The digested heart is subsequently dissected into atria and ventricles. Tissue samples are chopped into small pieces and dissolved by carefully pipetting up and down. The enzymatic digestion is stopped, and cells are stepwise reintroduced to physiologic calcium concentrations. After loading with a fluorescent Ca2+-indicator, isolated cardiomyocytes are prepared for simultaneous measurement of calcium currents and transients. Additionally, isolation pitfalls are discussed and patch-clamp protocols and representative traces of L-type calcium currents with simultaneous calcium transient measurements in atrial and ventricular murine myocytes isolated as described above are provided.

Isolation of High Quality Murine Atrial and Ventricular Myocytes for Simultaneous Measurements of Ca2+ Transients and L-Type Calcium Current

Mouse models allow studying key mechanisms of arrhythmogenesis. For this purpose, high quality cardiomyocytes are necessary to perform patch-clamp measurements. Here, a method to isolate murine atrial and ventricular myocytes via retrograde enzyme-based Langendorff perfusion, which allows simultaneous measurements of calcium-transients and L-type calcium current, is described.

Mouse models play a crucial role in arrhythmia research and allow studying key mechanisms of arrhythmogenesis including altered ion channel function and ...

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, ...

Real-Time Measurements of Calcium and Contractility Parameters in hiPSC-CMs

Real-Time Measurements of Calcium and Contractility Parameters in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) represent a powerful tool for studying mutation-mediated changes in cardiomyocyte function and defining the effects of stressors and drug interventions. In this study, it is demonstrated that this optics-based system is a powerful tool to assess the functional parameters of hiPSC-CMs in 2D. By using this platform, it is possible to perform paired measurements in a well-preserved temperature environment on different plate layouts. Moreover, this system provides researchers with instant data analysis.
This paper describes a method for measuring the contractility of unmodified hiPSC-CMs. Contraction kinetics are measured at 37 &#176;C based on pixel correlation changes relative to a reference frame taken at relaxation at a 250 Hz sampling frequency. Additionally, simultaneous measurements of intracellular calcium transients can be acquired by loading the cell with a calcium-sensitive fluorophore, such as Fura-2. Using a hyperswitch, ratiometric calcium measurements can be performed on a 50 &#181;m diameter illumination spot, corresponding to the area of the contractility measurements.

Author Spotlight: Real-Time Measurements of Calcium and Contractility Parameters in Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes  

Here, an established method is described to perform contractility and calcium measurements in human induced pluripotent stem cell-derived cardiomyocytes using an optics-based platform. This platform enables researchers to study the effect of mutations and the response to various stimuli in a rapid and reproducible manner.

Human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) represent a powerful tool for studying mutation-mediated changes in cardiomyocyte ...

Imaging Ca2+ Signals in Small Pulmonary Veins at Physiological Intraluminal Pressures

Pulmonary veins (PVs) carry oxygen-rich blood from the lungs back to the left heart, thus serving an important function in the delivery of oxygen-rich blood to vital organs. However, most studies of pulmonary vasculature have focused on pulmonary arteries and capillaries under normal and disease conditions. Ca2+ signals are critical regulators of vascular function. Despite the critical physiological roles of PVs, Ca2+ signals in small intrapulmonary PVs have not been recorded under physiological conditions. Here, we describe a technique to record Ca2+ signal activity in mouse PVs isolated, cannulated and pressurized at 5 mmHg. By incorporating a Ca2+ indicator, we can study Ca2+ signals in the myocyte layer of small PVs using high-speed, spinning disk confocal imaging under physiological conditions. Our representative data indicates that the Ca2+ signals in small PV myocytes are mediated by openings of ryanodine receptor ion channels. This method will be of considerable interest to researchers in the field of pulmonary vascular physiology and disorders.

In this protocol, we present a novel technique for recording and analyzing Ca2+ signals in intrapulmonary veins (small pulmonary veins or PVs) at physiological intraluminal pressures. The technique involves isolating small PVs, incubating them with a Ca2+ indicator, cannulating and pressurizing them, confocal imaging of Ca2+ signals, and data analysis.

Pulmonary veins (PVs) carry oxygen-rich blood from the lungs back to the left heart, thus serving an important function in the delivery of oxygen-rich ...

Real-time Evaluation of Absolute, Cytosolic, Free Ca²⁺ and Corresponding Contractility in Isolated, Pressurized Lymph Vessels

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