We thank Charles Hewitt for excellent technical assistance while establishing equipment and supplies needed for the current protocols. We thank Drs. Sean M. Wilson and Christopher G. Wilson, from the LLU Center for Perinatal Biology, for providing us with an additional inverted microscope and electrometer, respectively. This research has been supported by National Institutes of Health grant R00-AG047198 (EJB) and Loma Linda University School of Medicine new faculty start-up funds. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Before conducting the following experiments, ensure that all animal care use and protocols are approved by the Institutional Animal Care and Use Committee (IACUC) and performed in accord with the National Research Council&#39;s &#34;Guide for the Care and Use of Laboratory Animals&#34; (8th Edition, 2011) and the ARRIVE guidelines. The IACUC of Loma Linda University has approved all protocols used for this manuscript for male and female C57BL/6 mice (age range: 3 to 30 mo).
1. Equipment and Materials
NOTE: Details of materials required for the protocol can be found in the Table of Materials, Reagents, and manuals or websites associated with the respective vendors.
<ol>
	<li>Flow Chamber
	<ol>
		<li>Fasten a superfusion chamber with a glass coverslip bottom into an anodized aluminum platform. Secure the platform with chamber into a compact aluminum stage.</li>
		<li>Set a micromanipulator at each end of the platform on the aluminum stage for holding the pinning pipette. 
		NOTE: As a practical option, use this transferable stage apparatus with a flow chamber unit secured to facilitate the transition from isolation of the endothelial tube to performance of the experiments.</li>
	</ol>
	</li>
	<li>Microscopes
	<ol>
		<li>Use stereomicroscopes (5x to 50x magnification range) equipped with fiber optic light sources for macro- and micro-dissection procedures.</li>
		<li>For preparation of endothelial tubes, use an inverted microscope equipped with phase contrast or differential interference contrast (DIC) compatible objectives (10x, 20x, and 40x) and an aluminum stage.</li>
		<li>To isolate endothelial tubes from partially digested blood vessels, place a microsyringe pump controller adjacent to the endothelial tube preparation apparatus.</li>
		<li>For the experimental apparatus, use an inverted microscope equipped with standard objectives (4x and 10x) in addition to fluorescent objectives (20x and 40x, numerical aperture 0.75) and a manual aluminum stage on a vibration isolation table.</li>
	</ol>
	</li>
	<li>Intracellular Recording Equipment
	<ol>
		<li>To record the Vm of endothelial cells in isolated endothelial tubes, connect the electrometer to the compatible headstage. If necessary, connect a function generator or stimulator to the electrometer for experiments that require current injection.</li>
		<li>Connect the amplifier outputs to a data acquisition system, audible baseline monitors, and an oscilloscope. Secure the reference electrode (Ag/AgCl pellet) near the flow chamber exit during experiments.</li>
		<li>Use a photometric system with integrated components of a fluorescence system interface, high intensity arc lamp and power supply, hyperswitch, photomultiplier tube (or PMT), and camera to measure [Ca2+]i in endothelial cells.</li>
		<li>Use a temperature controller equipped with an inline heater to raise and maintain a physiological temperature (37 &#176;C) throughout the experiment.</li>
		<li>Use a six-reservoir platform connected to a valve controller with an inline flow control valve to control the delivery of respective solutions to the endothelial tube secured in the chamber.</li>
	</ol>
	</li>
	<li>Micropipettes and Sharp Electrodes 
	NOTE: To manufacture pipettes for trituration and the mechanical stabilization of isolated endothelial tubes, use an electronic puller for pulling the pipettes and a micro-forge for breaking and fire polishing.
	<ol>
		<li>To separate adventitia, smooth muscle, and the endothelial tube from the partially digested arterial segment, prepare trituration pipettes (each with an internal tip diameter of 80&#8211;120 &#181;m) as appropriate, using borosilicate glass capillary tubes, and fire polish the tip.</li>
		<li>To secure the endothelial tube against the bottom of the superfusion chamber, use pinning pipettes with a blunted, spherical end (heat-polished, OD of 100&#8211;150 &#181;m; prepared from thin-wall borosilicate glass tubes).</li>
		<li>To record Vm of an endothelial cell, prepare sharp electrodes with a tip resistance of ~150 &#177; 30 M&#937; from glass capillary tubes using an electronic puller.</li>
	</ol>
	</li>
</ol>
2. Preparation of Solutions and Drugs
<ol>
	<li>Physiological Salt Solution (PSS)
	<ol>
		<li>For one experiment with drug preparations, prepare a minimum of 1 L of PSS using: 140 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 10 mM N-2-hydroxyethylpiperazine-N&#39;-2-ethanesulfonic acid (HEPES), and 10 mM glucose.</li>
		<li>To prepare the required solutions for dissection, isolation of cerebral artery, and preparation of endothelial tube, prepare PSS lacking CaCl2 (zero Ca2+ PSS).</li>
		<li>Prepare all solutions in ultrapure deionized H2O, adjust the pH to 7.4, sterilize them through a filter (0.22 &#181;m pore size), and ensure that the osmolality of the solutions is between 290 and 300 mOsm. 
		NOTE: Solutions can be stored at 4 &#176;C for approximately two weeks.</li>
	</ol>
	</li>
	<li>10% Bovine Serum Albumin (BSA)
	<ol>
		<li>To prepare 10% BSA, dissolve 1 g of the lyophilized powder into a beaker of 10 mL zero Ca2+ PSS.</li>
		<li>Cover the beaker with paraffin film and allow a period of several hours to pass, with slow stirring for the BSA to dissolve.</li>
		<li>Filter the final solution using a 10 mL syringe plus a 0.22 &#181;m filter and make 1 mL aliquots to be stored at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Dissection Solution
	<ol>
		<li>To prepare 50 mL of dissection solution, add 500 &#181;L of BSA (10%) to 49.5 mL of zero Ca2+ PSS.</li>
		<li>Immediately after preparation, transfer this dissection solution into a Petri dish for arterial dissection.</li>
	</ol>
	</li>
	<li>Dissociation Solution
	<ol>
		<li>To prepare 50 mL of dissociation solution, add 5 &#181;L of CaCl2 (1 M) and 500 &#181;L of BSA (10%) to 49.5 mL of zero Ca2+ PSS. Allow the solution to sit at room temperature (RT).</li>
	</ol>
	</li>
	<li>Enzymes
	<ol>
		<li>For partial digestion of the arterial segments, prepare 1 mL of digestion solution with 0.31 mg/mL papain, 0.5 mg/mL dithioerythritol, 0.75 mg/mL collagenase (Type H blend), and 0.13 mg/mL elastase. 
		NOTE: Enzyme activities can vary; however, for our successful protocols, commercially supplied enzyme activities have been used as follows: papain &#8805;10 units/mg; collagenase &#8805;1 units/mg, for N-[3-(2-Furyl)acryloyl]-Leu-Gly-Pro-Ala or FALGPA substrate turnover; and elastase &#8805; 5 units/mg.</li>
	</ol>
	</li>
	<li>Preparation of Fura-2 and Pharmacological Agents
	<ol>
		<li>Prepare stock concentration (1 mM) of Fura-2 AM dye in DMSO. Prepare a working concentration (10 &#181;M) by adding 5 &#181;L of stock to 495 &#181;L of PSS for loading.</li>
		<li>Prepare &#8805;50 mL of working concentrations of pharmacological agents (e.g., ATP) in PSS as appropriate.</li>
	</ol>
	</li>
	<li>Conducting Solution
	<ol>
		<li>Prepare a conducting solution, 2 M KCl, by dissolving KCl in deionized H2O (7.455 g KCl in 50 mL H2O).</li>
		<li>Filter the solution through a 0.22 &#181;m syringe filter prior to backfilling the sharp electrodes.</li>
	</ol>
	</li>
	<li>Propidium Iodide (0.1%)
	<ol>
		<li>Dissolve 10 mg of lyophilized propidium iodide in 10 mL of 2 M KCl.</li>
		<li>Mix well and store at RT.</li>
	</ol>
	</li>
</ol>
3. Dissection and Isolation of Cerebral Artery
NOTE: All dissection procedures require specimen magnification (up to 50x) via stereomicroscopes and illumination provided by fiber optic light sources. To perform dissection procedures for isolation of the brain and arteries, use sharpened dissection instruments. Microdissection tools to isolate and clean arteries include sharpened fine-tipped forceps and Vannas style dissection scissors (3 to 9.5 mm blades).
<ol>
	<li>Isolation of Mouse Brain
	<ol>
		<li>Anesthetize a C57BL/6 mouse (3-30 months old; male or female) via inhalation of isoflurane (3% for 2&#8211;3 min), then decapitate the animal immediately following complete induction of anesthesia. Place the head in a Petri dish (diameter 10 cm, depth 1.5 cm) containing cold (4 &#176;C) zero Ca2+ PSS under a stereomicroscope.</li>
		<li>Viewing through the microscope, remove the skin and hair over the skull and remove excessive blood with cold zero Ca2+ PSS. Make an incision using only the tips of standard dissection scissors (e.g., 24 mm blade), starting with the occipital bone and extending up through the nasal bone of the skull.</li>
		<li>Open the skull carefully along the incision using coarse-tipped forceps, and separate connective tissue to isolate the brain with an intact Circle of Willis. 
		NOTE: Alternatively, Littauer bone-cutting forceps can be used to open the skull and expose the brain.</li>
		<li>Gently wash the isolated brain with cold zero Ca2+ PSS in a beaker to remove blood. Place the brain ventral side facing up in a chamber containing cold dissection solution for isolation of cerebral arteries (Figure 1A).</li>
	</ol>
	</li>
	<li>Isolation of Cerebral Arteries 
	NOTE: The posterior cerebral artery has been selected as a representative cerebrovascular&#160;endothelial study model for this protocol.
	<ol>
		<li>Secure the isolated brain in cold dissection solution using stainless steel pins (diameter 0.2 mm, length ~11&#8211;12 mm) to be inserted into a charcoal-infused silicon polymer coating the bottom (depth &#8805;50 cm) of a glass Petri dish.</li>
		<li>To maintain a chilled temperature of PSS during dissection, use a dissection chamber cooled by a refrigerant system continuously cycling a 1:1 water-ethylene glycol mixture. Alternatively, dissection can be performed on fresh ice.</li>
		<li>Surgically isolate the posterior cerebral arteries (~0.3 to 0.5 cm segments, no axial tension) from the posterior communicating and basilar arteries using microdissection instruments Vannas style scissors and sharpened fine-tipped forceps (Figure 1B). Use stainless steel pins (diameter 0.1 mm, length ~13&#8211;14 mm) to secure both isolated posterior cerebral arteries in the dissection solution in the Petri dish (Figure 1C).</li>
		<li>Clean the isolated posterior cerebral arteries carefully by removing connective tissue using sharpened fine-tipped forceps (Figure 1D). Cut intact arteries into segments (length 1&#8211;2 mm) for enzymatic digestion (Figure 1D; inset).</li>
	</ol>
	</li>
</ol>
4. Preparation of Endothelial Tube and Superfusion
NOTE: The endothelial tube is prepared as described previously12, with modifications for the cerebral artery6.
<ol>
	<li>Prepare the trituration apparatus using a microscope equipped with objectives (10x, 20x, and 40x), a camera, and an aluminum stage holding a chamber and micromanipulators. Secure a microsyringe with a pump controller adjacent to the stage and specimen (Figure 2A).</li>
	<li>Completely backfill a trituration pipette with mineral oil and secure it over the micro-syringe piston. Then, using the micro-syringe with pump controller, withdraw dissociation solution into the pipette (~130 nl) on top of the mineral oil while ensuring the absence of air bubbles in the pipette.</li>
	<li>Place intact arterial segments into 1 mL of dissociation solution in a 10 mL glass tube (Figure 2B), containing 0.31 mg/mL papain, 0.5 mg/mL dithioerythritol, 0.75 mg/mL collagenase, and 0.13 mg/mL elastase. Incubate at 34 &#176;C for 10&#8211;12 min for partial digestion.</li>
	<li>Following the digestion, replace the enzyme solution with ~5 mL of fresh dissociation solution. Using a 1 mL pipette, transfer one segment into a chamber containing the dissociation solution at RT.</li>
	<li>Place the pipette into the dissociation solution in the chamber and position it close to one end of the digested vessel. Set a rate within the range of 2 to 5 nL/s on the pump controller for gentle trituration (Figure 2C; inset).</li>
	<li>While viewing through 100x to 200x magnification, withdraw and eject the arterial segment to dissociate smooth muscle cells while producing an endothelial tube. If necessary, carefully use fine-tipped forceps to separate dissociated adventitia and internal elastic lamina from the endothelial tube. Confirm that all smooth muscle cells are dissociated and that only endothelial cells remain as an intact &#34;tube&#34; (Figure 2C).</li>
	<li>Using micromanipulators, secure each end of the endothelial tube on the glass cover slip of the superfusion chamber using borosilicate glass pinning pipettes (Figure 2D).</li>
	<li>Wash-out dissociated adventitia and smooth muscle cells from the chamber and replace the dissociation solution with 2 mM CaCl2 PSS. Transfer the mobile platform with secured endothelial tube onto the microscope of superfusion and experimental rig.</li>
	<li>Use 6 clean 50 mL reservoirs (Figure 3A) for continuous delivery of PSS and respective drug solutions during the experiment, as appropriate. Use the inline flow control valve to manually set the flow rate throughout as consistent with laminar flow while matching flow feed to vacuum suction. Deliver PSS to the chamber (Figure 3B) for the superfusion of the endothelial tube for &#8805; 5 min before recording the background data and dye loading.</li>
</ol>
5. Dye Load, Wash-Out, and Temperature Settings
<ol>
	<li>Turn on all components of the photometry system for measuring [Ca2+]i. Use the microscope on the experimental rig (Figure 3C) to view the endothelial tube at 400x magnification, and focus on cells in the photometric window using the photometry system software suite.</li>
	<li>Measure the diameter of the endothelial tube and record background autofluorescence values in accord with 510 nm emission during alternate excitation at 340 and 380 nm (&#8805;10 Hz).</li>
	<li>To measure [Ca2+]i responses, load the endothelial tube with Fura-2 AM dye (10 &#181;M final concentration) at RT and allow ~30&#8211;40 min in the absence of light.</li>
	<li>Restart superfusion of the endothelial tube with fresh PSS for ~30&#8211;40 min to wash-out excess dye and allow intracellular Fura-2 AM to de-esterify. During the wash-out period, raise the temperature gradually from RT to 37 &#176;C using the temperature controller (Figure 3A) with an inline heater, then maintain at 37 &#176;C throughout the experiment. 
	NOTE: The recommended incremental transition between RT to 37 &#176;C entails three steps (e.g., 5 &#176;C) with a &#8805;5 min equilibration time at each step.</li>
</ol>
6. Simultaneous Measurement of [Ca2+]i and Vm
<ol>
	<li>Turn on all other equipment and the electrometer software suite for measuring Vm (see Figure 3, Figure 4). Adjust data acquisition rate accordingly (&#8805;10 Hz).</li>
	<li>Pull a sharp electrode and backfill with 2 M KCl. For studies of intercellular coupling via dye transfer, backfill the microelectrodes with 0.1% propidium iodide dissolved in 2 M KCl.</li>
	<li>Secure the electrode over a silver wire coated with chloride in the pipette holder attached to an electrometer head stage secured with a micromanipulator. Use the micromanipulator to briefly position the tip of the electrode into the flowing PSS in the chamber while viewing through the 4x objective.</li>
	<li>Set the resting Vm to 0 as consistent with grounded bath potential. Position the tip of electrode just over a cell of the endothelial tube. If desired, use audible baseline monitors linked to electrometers to associate sound pitch with potential recordings.</li>
	<li>Increase magnification to 400x using the 40x objective and re-position the tip of the electrode as needed. Adjust the photometric window using the photometry software to focus on ~50 to 80 endothelial cells.</li>
	<li>Gently place the electrode into one of the cells of the endothelial tube using the micromanipulator and wait &#8805;2 min for the resting Vm to stabilize. Similarly, insert a second electrode in another cell (distance &#8805;100 &#181;m) from the first cell impaled with first electrode.</li>
	<li>Once resting Vm is stable at its expected values (-30 to -40 mV)6, turn on the PMT on the fluorescence interface in the absence of light and begin acquisition of intracellular [Ca2+]i&#160;by exciting Fura-2 alternately (10 Hz) at 340 and 380 nm while collecting fluorescence emission at 510 nm. 
	NOTE: To test intercellular electrical coupling, current (&#177; 0.5-3 nA, ~20 s pulse duration) can be delivered via one electrometer as &#34;Site 1&#34;, and changes of Vm can be recorded with another electrometer as &#34;Site 2&#34; (distance &#8805;100 &#181;m).</li>
	<li>Once simultaneous measurements of Vm and [Ca2+]i&#160;are established, allow ~5 min for superfusion of endothelial tube with PSS at a constant laminar flow rate before application of drugs.</li>
	<li>Apply the drug (e.g., ATP) prepared in PSS to the superfusion chamber with the constant flow rate. After ~3 min (or a period of time appropriate for the kinetics of the drug), wash with PSS until Vm and the F340/F380 ratio return to their baseline conditions. Mark respective recordings as temporally synchronized across the photometer and electrometer software suites during each transition of solutions.</li>
	<li>Once the experiment is done, withdraw electrode from the cell using the micromanipulator and notice Vm ~0 mV as referenced by the bath electrode. Stop respective recordings of Vm and [Ca2+]i and save the records on file for data analysis.</li>
</ol>
7. Visualization of Cell-to-Cell Coupling
<ol>
	<li>To visualize cell-to-cell coupling via propidium iodide, use a 40x or 60x fluorescent objective with a relatively high numerical aperture (e.g., 0.95) and rhodamine filter set with solid-state illumination, a high-resolution camera (e.g., 16-megapixels), and an imaging software suite.</li>
</ol>

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P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Activation+of+endothelial+transient+receptor+potential+C3+channel+is+required+for+small+conductance+calcium-activated+potassium+channel+activation+and+sustained+endothelial+hyperpolarization+and+vasodilation+of+cerebral+artery.">Activation of endothelial transient receptor potential C3 channel is required for small conductance calcium-activated potassium channel activation and sustained endothelial hyperpolarization and vasodilation of cerebral artery.</a> Journal of the American Heart Association. 3 (4), (2014).</li><li>Zhang, L., Papadopoulos, P., Hamel, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endothelial+TRPV4+channels+mediate+dilation+of+cerebral+arteries:+impairment+and+recovery+in+cerebrovascular+pathologies+related+to+Alzheimer's+disease.">Endothelial TRPV4 channels mediate dilation of cerebral arteries: impairment and recovery in cerebrovascular pathologies related to Alzheimer's disease.</a> British Journal of Pharmacology. 170 (3), 661-670 (2013).</li><li>Socha, M. J., Domeier, T. L., Behringer, E. J., Segal, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coordination+of+intercellular+Ca2++signaling+in+endothelial+cell+tubes+of+mouse+resistance+arteries.">Coordination of intercellular Ca2+ signaling in endothelial cell tubes of mouse resistance arteries.</a> Microcirculation. 19 (8), 757-770 (2012).</li></ol>

The authors declare no conflicts of interest.

In light of recent developments6,15,16,17, we now demonstrate the method to isolate mouse cerebral arterial endothelium in preparation for simultaneous measurement of [Ca2+]i and Vm underlying EDH consistently for ~2 h at 37 &#176;C. Although technically difficult, we can measure cell-to-cell coupling as well (see reference6, Figure 1). In this manner, we can measure discrete and conducted signaling events simultaneously. For an account of the general challenges faced while isolating mouse arterial endothelium consult references11,12. We emphasize critical steps for isolating mouse cerebral arteries, our particular measurements of endothelial [Ca2+]i and Vm, suggestions for troubleshooting, and considerations for alternative methods in light of experimental strengths/weaknesses below.
The posterior cerebral artery requires careful isolation from the Circle of Willis and connective tissue without damage to smooth muscle and endothelial cells. Surgical tools (forceps and scissors) should be suitable for mouse microcirculation dissection procedures while maintained with clean, sharp edges. Relative to skeletal muscle12, we reduced the time period for enzymatic incubation by two-thirds and the concentrations of papain, collagenase, and dithioerythritol by half while adding use of an empirically optimized concentration of elastase. For consistent success in isolating cerebral arterial endothelial tubes, it is important to order enzymes in batches (2 to 4 vials) from a highly reputable vendor with consistent manufacturing practices in accord with frequency of use and shelf-life. Even with the potential for variation in enzyme activity from one lot of enzyme to the next, it is suggested that enzyme concentrations be maintained while optimizing the time of digestion within 10 to 12 min. It is worth noting that neither the gender nor age of the animal (3 to 30 months) has been a significant barrier to the preparation of isolated endothelium from the cerebral arteries in our experience. Thus, it is recommended that the composition of the enzyme cocktail and time of enzyme digestion remain the same in studies on age and gender.
As emphasized, caution is required during gentle trituration and isolation of the endothelium from adventitia and spindle-shaped smooth muscle cells in order to avoid damage to endothelial cells12. For the trituration step, the viscous mineral oil serves as a hydraulic medium between the piston of the micro-syringe and the aqueous PSS in order to withdraw or eject vessel segments bathed in PSS. It should be noted that if the vessel segment contacts the mineral oil, this step must be repeated using a clean trituration pipette with sequential backfilling of mineral oil and PSS. Before an experiment, it is recommended that an approximately linear in vivo length be restored to the endothelial tube to the extent possible, whereby individual cells assume a &#34;flat&#34;, longitudinal morphology (e.g., diameter ~5 &#181;m; length ~100 &#181;m; thickness ~0.5 &#181;m)8. However, axial tension should not be applied to the extent where cells at the edges of the specimen are pulling away from underneath respective pinning pipettes and thereby compromising mechanical stability for reliable cellular measurements during the experiment.
The photometric [Ca2+]i measuring system used in this technique is suitable for continuous measurements of endothelial [Ca2+]i using the&#160;ratiometric Fura-2 dye. Individual protocols can typically last &#8805;10 min without significant photobleaching. Further, we commonly use this approach for measuring [Ca2+]i because it is amenable to timely pauses in data acquisition and frequent movement of microscope objectives as needed to secure sharp electrodes for Vm measurements. Note that this is a photometric approach that only captures global, averaged [Ca2+]i among multiple cells. In general, for [Ca2+]i measurements, we recommend photometry for the initial characterization of endothelial cell responses to pharmacological agents (e.g., time course of peak and plateau phases) while securing plans to proceed to quantitate microdomain signaling events using standard confocal or multiphoton microscopy12,23.
The sharp electrode electrophysiology approach is amenable for integrated Vm recordings in physiological multicellular preparations. The measurement of intracellular Vm is by far the most technically challenging with numerous critical steps that can either facilitate or obliterate success of measurements. First, the experimenter needs to determine the optimal program conditions on a glass puller for consistently manufacturing sharp electrodes with a tip resistance of ~150 &#177; 30 M&#937;. Using a filament heat ramp test reading as a reference, it is recommended that the experimenter determine parameters empirically, particularly with respect to variation in the &#34;heat&#34; setting while maximizing the time of pull. Next, mechanical stability of the micromanipulators, headstages, pipette holders, and specimen is absolutely critical. Aside from the obvious necessity for a vibration isolation table, the experimenter will need to ensure that all fittings are secure and that the stage area around and underneath the manipulators is clean. Ideally, extraneous noise should be reduced to the extent that the resolution of sharp electrode recordings is within 1 mV. In our experience, the most common source of noise is silver wire secured in the pipette holder that is in need of a sufficient chloride coating or replacement altogether. If common 50/60 Hz noise is present, &#34;hum silencing&#34; technology can be used and comes standard with a modern electrometer. If noise appears completely unmanageable, check to see whether there is a leak in the bath or overflow where PSS will come into contact with resistors for temperature control. If complications remain with acquiring a stable Vm signal, check the integrity of electrical cords and t-connectors.
Altogether, if establishing a sharp electrode recording is unsuccessful, it is commonly due to an inappropriate electrode, mechanical instability, or poor cellular health. Alternatively, the experimenter may also want to consider a configuration of the patch-clamp technique on individual, electrically uncoupled cells where information on whole cell currents and single ion channel recordings can be obtained. Although faced with the general challenges of using a fluorescent dye (e.g., heterogeneous intracellular loading, bleaching, calibration using ionophores, modest detection resolution), voltage-sensitive dyes such as Di-8-ANEPPS are also commercially available.
Our understanding of the endothelium in blood flow regulation to and throughout the brain is crucial with a rapidly aging human population and the rising incidence of chronic cardiovascular and neurodegenerative diseases. Thus, we provide a method for isolating intact endothelium from a vital cerebral artery, which may be adapted to other vascular structures in the brain. In addition, we show a useful approach for the simultaneous measurements of underlying [Ca2+]i and Vm. Finally, using dual intracellular electrodes, cell-to-cell coupling through gap junctions can also be assessed. With carefully planned pharmacological and/or genetic interventions, the current protocol reasonably and quantitatively encompasses study of cerebral endothelial function in its native form. Our expectation is that experimenters will build from and adapt this information to advance vascular biology and neuroscience in general. In particular, it would be satisfactory to see common experimental struggles involved with electrical measurements minimized altogether. Although this protocol is not a stand-alone set of techniques by any means, it can be used as a complementary approach to determine precisely how endothelial biophysical determinants translate into changes in vascular resistance and how they fine-tune regulation of blood flow relative to conditions representative of health vs. disease.

Blood flow throughout the brain is regulated by the coordination of vasodilation among cerebral arteries and arterioles in vascular networks1. Endothelial cells lining cerebral resistance arteries command changes in vascular diameter as needed to meet the metabolic demand of neurons1,2,3. In particular, during endothelium-derived hyperpolarization (commonly known as EDH), intracellular Ca2+ ([Ca2+]i) and electrical signaling in endothelial cells coordinate vasodilation among endothelial cells and their surrounding smooth muscle cells through gap junctions for arterial relaxation4. Physiological initiation of EDH sequentially entails stimulation of Gq-coupled receptors (GPCRs), an increase in [Ca2+]i, and activation of endothelial small- and intermediate-Ca2+-activated K+ (SKCa/IKCa) channels to hyperpolarize cerebral endothelial membrane potential (Vm)5,6,7. Thus, the intimate relationship of endothelial [Ca2+]i and Vm is integral to blood flow regulation and indispensable to cardio- and cerebrovascular function6,8. Throughout the broader literature, numerous studies have reported the association of vascular endothelial dysfunction with the development of chronic diseases (e.g., hypertension, diabetes, heart failure, coronary artery disease, chronic renal failure, peripheral artery disease)9,10, indicating the significance of studying endothelial function in physiological as well as pathological conditions.
Vascular endothelium is integral to the production of hyperpolarization, vasodilation, and tissue perfusion and thus, examination of its native cellular properties is crucial. As a general study model, preparation of the mouse arterial endothelial tube model has been published before for skeletal muscle11,12, gut13, lung14, and recently for the brain6. Studies of simultaneous [Ca2+]i and Vm measurements in particular have been published for skeletal muscle arterial endothelium15,16 as well as lymphatic vessel endothelium17. In addition to primary studies utilizing the endothelial tube approach, a comprehensive review of its advantages and disadvantages8 can be consulted to determine if this experimental tool is appropriate for a specific study. In brief, an advantage is that the key physiological components of endothelial cell function are retained (e.g., Ca2+ influx and intracellular release, hyperpolarization of Vm up to the Nernst potential for K+&#160;via SKCa/IKCa activation, and endothelial intercellular coupling via gap junctions) without confounding factors such as perivascular nerve input, smooth muscle voltage-gated channel function and contractility, circulation of blood, and hormonal influences8. In contrast, commonly used cell culture approaches introduce significant alterations in morphology18 and ion channel expression19 in a manner that can greatly obfuscate comparisons to physiological observations determined ex vivo or in vivo. Limitations include a lack of integration with other essential components for regulating blood flow, such as smooth muscle and restricted flexibility in an experimental schedule, as this model is optimally tested within 4 h of intact vascular segment isolation from the animal.
Building from a previous video protocol authored by Socha and Segal12 and recent experimental developments in the interim6,15,16, we hereby demonstrate the isolation of fresh endothelium from posterior cerebral arteries and simultaneous measurements of endothelial [Ca2+]i and Vm using Fura-2 photometry and intracellular sharp electrodes, respectively. Additionally, this experiment entails continuous superfusion of salt solutions and pharmacological agents during physiological conditions (pH 7.4, 37 &#176;C). We chose the posterior cerebral artery, as it yields isolated endothelium with the structural integrity (cells coupled through gap junctions) and sufficient dimensions (width &#8805;50 &#181;m, length &#8805;300 &#181;m) amenable for intra- and intercellular signaling along and among endothelial cells. In addition, studies of the rodent posterior cerebral artery are substantially represented in the literature and encompass examination of fundamental endothelial signaling mechanisms, vascular development/aging, and pathology20,21,22. This experimental application is expected to yield a high-throughput analysis of cerebral endothelial function (and dysfunction) and will thereby allow for significant advancements in the understanding of blood flow regulation throughout aging and the development of neurodegenerative disease.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Glucose</td>
			<td>Sigma-Aldrich (St. Louis, MO, USA)</td>
			<td>G7021</td>
			<td></td>
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		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Sigma</td>
			<td>M2670</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma</td>
			<td>223506</td>
			<td></td>
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		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H4034</td>
			<td></td>
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		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P9541</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>Sigma</td>
			<td>S8045</td>
			<td></td>
		</tr>
		<tr>
			<td>ATP</td>
			<td>Sigma</td>
			<td>A2383</td>
			<td></td>
		</tr>
		<tr>
			<td>HCl</td>
			<td>ThermoFisher Scientific (Pittsburgh, PA, USA)</td>
			<td>A466250</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase (Type H Blend)</td>
			<td>Sigma</td>
			<td>C8051</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithioerythritol</td>
			<td>Sigma</td>
			<td>D8255</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Sigma</td>
			<td>P4762</td>
			<td></td>
		</tr>
		<tr>
			<td>Elastase</td>
			<td>Sigma</td>
			<td>E7885</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td></td>
		</tr>
		<tr>
			<td>Propidium iodide</td>
			<td>Sigma</td>
			<td>P4170</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr>
			<td>Fura-2 AM dye</td>
			<td>Invitrogen, Carlsbad, CA, USA</td>
			<td>F14185</td>
			<td></td>
		</tr>
		<tr>
			<td>Recirculating chiller (Isotemp 500LCU)</td>
			<td>ThermoFisher Scientific</td>
			<td>13874647</td>
			<td></td>
		</tr>
		<tr>
			<td>Plexiglas superfusion chamber&#160;</td>
			<td>Warner Instruments, Camden, CT, USA</td>
			<td>RC-27</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip bottom (2.4&#160;&#215;&#160;5.0&#160;cm)</td>
			<td>ThermoFisher Scientific</td>
			<td>12-548-5M</td>
			<td></td>
		</tr>
		<tr>
			<td>Anodized aluminum platform (diameter: 7.8&#160;cm)&#160;</td>
			<td>Warner Instruments</td>
			<td>PM6 or PH6</td>
			<td></td>
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		<tr>
			<td>Compact aluminum stage&#160;</td>
			<td>Siskiyou, Grants Pass, OR, USA</td>
			<td>8090P</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator</td>
			<td>Siskiyou</td>
			<td>&#160;MX10</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscopes&#160;</td>
			<td>Zeiss, NY, USA</td>
			<td>Stemi 2000 &#38; 2000-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Fiber optic light sources</td>
			<td>&#160;Schott, Mainz, Germany &#38; KL200, Zeiss</td>
			<td>Fostec 8375</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon inverted microscope</td>
			<td>Nikon Instruments Inc, Melville, NY, USA</td>
			<td>Ts2</td>
			<td></td>
		</tr>
		<tr>
			<td>Phase contrast objectives&#160;</td>
			<td>Nikon Instruments Inc</td>
			<td>&#160;(Ph1 DL; 10X &#38; 20X)</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescent objectives&#160;</td>
			<td>Nikon Instruments Inc</td>
			<td>20X (S-Fluor), and 40X (Plan Fluor)</td>
			<td></td>
		</tr>
		<tr>
			<td>Nikon inverted microscope</td>
			<td>Nikon Instruments Inc</td>
			<td>Eclipse TS100</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsyringe pump controller (Micro4 )&#160;</td>
			<td>World Precision Instruments (WPI), Sarasota, FL, USA</td>
			<td>SYS-MICRO4</td>
			<td></td>
		</tr>
		<tr>
			<td>Vibration isolation table</td>
			<td>Technical Manufacturing, Peabody, MA, USA</td>
			<td>&#160;Micro-g</td>
			<td></td>
		</tr>
		<tr>
			<td>Amplifiers</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>Axoclamp 2B &#38; Axoclamp 900A</td>
			<td></td>
		</tr>
		<tr>
			<td>Headstages&#160;</td>
			<td>Molecular Devices</td>
			<td>HS-2A &#38; HS-9A</td>
			<td></td>
		</tr>
		<tr>
			<td>Function generator&#160;</td>
			<td>EZ Digital, Seoul, South Korea</td>
			<td>FG-8002</td>
			<td></td>
		</tr>
		<tr>
			<td>Data Acquision System</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>Digidata 1550A</td>
			<td></td>
		</tr>
		<tr>
			<td>Audible Baseline Monitors</td>
			<td>Ampol US LLC, Sarasota, FL, USA</td>
			<td>&#160;BM-A-TM</td>
			<td></td>
		</tr>
		<tr>
			<td>Digital Storage Oscilloscope</td>
			<td>Tektronix, Beaverton, Oregon, USA</td>
			<td>&#160;TDS 2024B</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence System Interface, ARC Lamp + Power Supply, Hyperswitch, PMT</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>IonOptix Systems</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller&#160;&#160;</td>
			<td>Warner Instruments</td>
			<td>TC-344B or C</td>
			<td></td>
		</tr>
		<tr>
			<td>Inline Heater&#160;</td>
			<td>Warner Instruments</td>
			<td>SH- 27B</td>
			<td></td>
		</tr>
		<tr>
			<td>Valve Controller&#160;</td>
			<td>Warner Instruments</td>
			<td>VC-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Inline Flow Control Valve</td>
			<td>Warner Instruments</td>
			<td>&#160;FR-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Electronic Puller&#160;</td>
			<td>Sutter Instruments, Novato, CA, USA</td>
			<td>P-97 or P-1000&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Microforge</td>
			<td>Narishige, East Meadow, NY, USA</td>
			<td>&#160;MF-900</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass Tubes (Trituration)</td>
			<td>World Precision Instruments (WPI), Sarasota, FL, USA</td>
			<td>1B100-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass Tubes (Pinning)</td>
			<td>Warner Instruments</td>
			<td>G150T-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate Glass Tubes (Sharp Electrodes)</td>
			<td>Warner Instruments</td>
			<td>GC100F-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe Filter (0.22 &#181;m)&#160;&#160;</td>
			<td>ThermoFisher Scientific</td>
			<td>722-2520</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Petri Dish + Charcoal Sylgard</td>
			<td>Living Systems Instrumentation, St. Albans City, VT, USA</td>
			<td>DD-90-S-BLK</td>
			<td></td>
		</tr>
		<tr>
			<td>Vannas Style Scissors (3 mm &#38; 9.5&#160;mm)</td>
			<td>World Precision Instruments</td>
			<td>555640S, 14364</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors 3 &#38; 7 mm blades</td>
			<td>Fine Science Tools (or FST), Foster City, CA, USA</td>
			<td>Moria MC52 &#38; 15000-00</td>
			<td></td>
		</tr>
		<tr>
			<td>Sharpened fine-tipped forceps&#160;</td>
			<td>FST</td>
			<td>Dumont #5 &#38; Dumont #55</td>
			<td></td>
		</tr>
	</tbody>
</table>

simultaneous measurements of intracellular calcium and membrane potential in freshly isolated and intact mouse cerebral endothelium

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the lumen of blood vessels and command changes in vascular diameter as needed to meet the metabolic demand of neurons. Primary endothelial-dependent signaling pathways of hyperpolarization of membrane potential (Vm) and nitric oxide typically operate in parallel to mediate vasodilation and thereby increase blood flow. Although integral to coordinating vasodilation over several millimeters of vascular length, components of endothelium-derived hyperpolarization (EDH) have been historically difficult to measure. These components of EDH entail intracellular Ca2+ [Ca2+]i increases and subsequent activation of small- and intermediate conductance Ca2+-activated K+ (SKCa/IKCa) channels.
Here, we present a simplified illustration of the isolation of fresh endothelium from mouse cerebral arteries; simultaneous measurements of endothelial [Ca2+]i and Vm using Fura-2 photometry and intracellular sharp electrodes, respectively; and a continuous superfusion of salt solutions and pharmacological agents under physiological conditions (pH 7.4, 37 &#176;C). Posterior cerebral arteries from the Circle of Willis are removed free of the posterior communicating and the basilar arteries. Enzymatic digestion of cleaned posterior cerebral arterial segments and subsequent trituration facilitates removal of adventitia, perivascular nerves, and smooth muscle cells. Resulting posterior cerebral arterial endothelial &#34;tubes&#34; are then secured under a microscope and examined using a camera, photomultiplier tube, and one to two electrometers while under continuous superfusion. Collectively, this method can simultaneously measure changes in endothelial [Ca2+]i and Vm in discrete cellular locations, in addition to the spreading of EDH through gap junctions up to millimeter distances along the intact endothelium. This method is expected to yield a high-throughput analysis of the cerebral endothelial functions underlying mechanisms of blood flow regulation in the normal and diseased brain.

The schematic demonstration of the protocol described above is shown in the attached figures. A brain isolated from a young adult male C57BL/6N mouse (5 months) is shown in Figure 1A. Posterior cerebral arteries are carefully isolated from the Circle of Willis, removed without connective tissue, and cut into segments (Figure 1B-D). From partially digested arterial segments, the intact endothelial tube is produced and secured on the glass cover slip using pinning pipettes. Gentle mechanical stretch is applied using two pinning pipettes to approximate linear in vivo length without vessel tortuosity. This has been further clarified in the manuscript (Figure 2A-D). Endothelial tubes isolated from posterior cerebral arteries are ~90&#8211;100 &#181;m in width and &#8805;300 &#181;m in length. The tube is loaded with Fura-2 AM followed by wash-out with PSS. Simultaneous measurement of [Ca2+]i and Vm in the endothelial tube is performed by Ca2+ photometry as well as sharp electrode electrophysiology (experimental rig shown in Figure 3 and Figure 4).
Functional assessment of the endothelial tube is performed by applying a classical endothelial GPCR agonist such as ATP into the flow chamber under physiological conditions. As shown in Figure 5, [Ca2+]i and Vm are recorded simultaneously in response to ATP (100 &#181;M). A significant hyperpolarization of&#160;Vm (&#8805;5 mV) occurs concomitantly with an intracellular [Ca2+]i increase, indicating that a rise in [Ca2+]i activates SKCa/IKCa channels and thereby allows K+ efflux across the plasma membrane to produce EDH. Evidence of intercellular coupling using current injection (&#177; 0.5 to 3 nA, ~20 s pulses) can be found in our recently published work (reference6, Figure 1). Note that propidium iodide has a mass of ~668 Da in reference to second messengers known for passing through gap junctions such as inositol trisphosphate (IP3, ~420 Da), cyclic adenosine monophosphate (cAMP, ~329 Da).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58832/58832fig1.jpg" /> 
Figure 1: Isolation of cerebral artery from brain. (A) Brain isolated with intact arteries (white arrow indicates the posterior cerebral artery) in dessection solution. (B) Magnified view of posterior cerebral artery (white arrow; ~3x vs. Panel A). (C) Isolated posterior cerebral arteries secured with stainless steel pins in specimen dish. (D) Posterior cerebral arteries following removal of connective tissues. Inset shows cut segments of arteries; Scale bar = 500 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58832/58832fig1large.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/58832/58832fig2.jpg" /> 
Figure 2: Preparation of cerebral endothelial tube. (A) Equipment used for triturating the partially digested arterial segments; a = micromanipulator, b = chamber for preparing tube, c = aluminum stage, d = microsyringe, e = microscope, f = microsyringe pump controller. (B) Arterial segments (white arrows) in solution for enzymatic digestion. (C) Endothelial tubes prepared by trituration; a = trituration pipette, b = intact endothelial tube. Inset shows the trituration process to remove adventitia and smooth muscle cells; Scale bar = 100 &#181;m. (D) An intact endothelial tube secured on the glass cover slip with pinning pipette; a = pinning pipette, b = intact endothelial tube with diameter of ~100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58832/58832fig2large.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/58832/58832fig3.jpg" /> 
Figure 3: Equipment for superfusion and simultaneous [Ca2+]i&#160;and Vm measurements. (A) Equipment for superfusion of the endothelial tube and temperature control, a = high intensity ARC lamp power supply, b = fluorescence system interface, c = temperature controller, d = six-reservoirs of superfusion system, e = fiber optic light sources, f = valve controller, g = hyperswitch of [Ca2+]i photometry system. (B) Superfusion chamber apparatus; a and b = headstages, c = ground electrode, d = inline heater, e = superfusion tubing, f = vacuum suction, g = superfusion chamber, h = aluminum stage holding the chamber. (C) Experimental platform on a vibration isolation table; a = cell framing adaptor with camera, b = microscope light, c = microscope, d = aluminum stage, e = micromanipulator for headstage control, f = vibration isolation table. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58832/58832fig3large.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/58832/58832fig4.jpg" /> 
Figure 4: Equipment for operating&#160;electrophysiology and data recording. a = oscilloscope, b = function generator, c = audible baseline monitor, d = 50/60 Hz noise filter, e and f = electrometers, g = data acquisition system. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58832/58832fig4large.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/58832/58832fig5.jpg" /> 
Figure 5: Simultaneous [Ca2+]i&#160;and Vm recordings. (A) Differential Interference Contrast image (400x) of a mouse cerebral arterial endothelial tube adjusted for experiment in photometric window using a 40x objective. A sharp electrode is placed into a cell as shown at the top of the image (white arrow). (B) Raw traces for simultaneous measurement of F340/F380 ratio (top) and Vm (bottom) in response to ATP (100 &#181;M).&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58832/58832fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium at JoVE.com

Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium

Demonstrated here are protocols for (1) freshly isolating intact cerebral endothelial "tubes" and (2) simultaneous measurements of endothelial calcium and membrane potential during endothelium-derived hyperpolarization. Further, these methods allow for pharmacological tuning of endothelial cell calcium and electrical signaling as individual or interactive experimental variables.

Cerebral arteries and their respective microcirculation deliver oxygen and nutrients to the brain via blood flow regulation. Endothelial cells line the ...

simultaneous-measurements-intracellular-calcium-membrane-potential

Nanyang Technological University

Research

JoVE Journal

Biology

8.0K Views.  Loma Linda University. Demonstrated here are protocols for (1) freshly isolating intact cerebral endothelial "tubes" and (2) simultaneous measurements of endothelial calcium and membrane potential during endothelium-derived hyperpolarization. Further, these methods allow for pharmacological tuning of endothelial cell calcium and electrical signaling as individual or interactive experimental variables.

Video: Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium

Measuring Near Plasma Membrane and Global Intracellular Calcium Dynamics in Astrocytes

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, increasing evidence suggests that astrocytes may also actively participate in brain function through functional interactions with neurons.  However, many fundamental aspects of astrocyte biology remain controversial, unclear and/or experimentally unexplored. One important issue is the dynamics of intracellular calcium transients in astrocytes. This is relevant because calcium is well established as an important second messenger and because it has been proposed that astrocyte calcium elevations can trigger the release of transmitters from astrocytes. However, there has not been any detailed or satisfying description of near plasma membrane calcium signaling in astrocytes. Total internal reflection fluorescence (TIRF) microscopy is a powerful tool to analyze physiologically relevant signaling events within about 100 nm of the plasma membrane of live cells. Here, we use TIRF microscopy and describe how to monitor near plasma membrane and global intracellular calcium dynamics almost simultaneously. The further refinement and systematic application of this approach has the potential to inform about the precise details of astrocyte calcium signaling. A detailed understanding of astrocyte calcium dynamics may provide a basis to understand if, how, when and why astrocytes and neurons undergo calcium-dependent functional interactions.

We describe how to measure near membrane and global intracellular calcium dynamics in cultured astrocytes using total internal reflection and epifluorescence microscopy.

The brain contains glial cells.  Astrocytes, a type of glial cell, have long been known to provide a passive supportive role to neurons. However, ...

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

The study of electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cardiac cellular Ca2+ handling abnormalities requires isolated cardiomyocytes. In addition, the possibility to investigate myocytes from patients using these techniques is an invaluable requirement to elucidate the molecular basis of cardiac diseases such as atrial fibrillation (AF).1 Here we describe a method for isolation of human atrial myocytes which are suitable for both patch-clamp studies and simultaneous measurements of intracellular Ca2+ concentrations. First, right atrial appendages obtained from patients undergoing open heart surgery are chopped into small tissue chunks ("chunk method") and washed in Ca2+-free solution. Then the tissue chunks are digested in collagenase and protease containing solutions with 20 &mu;M Ca2+. Thereafter, the isolated myocytes are harvested by filtration and centrifugation of the tissue suspension. Finally, the Ca2+ concentration in the cell storage solution is adjusted stepwise to 0.2 mM. We briefly discuss the meaning of Ca2+ and Ca2+ buffering during the isolation process and also provide representative recordings of action potentials and membrane currents, both together with simultaneous Ca2+ transient measurements, performed in these isolated myocytes.

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

We describe the isolation of human atrial myocytes which can be used for intracellular Ca2+ measurements in combination with electrophysiological patch-clamp studies.

The study of  electrophysiological properties of cardiac ion channels with the patch-clamp  technique and the exploration of cardiac cellular Ca2+ ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

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 ...

Single-channel Analysis and Calcium Imaging in the Podocytes of the Freshly Isolated Glomeruli

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells in the pathogenesis of various renal diseases has been established since podocyte injury leads to proteinuria and foot process effacement. It was previously reported that various endogenous agents may cause a dramatic overload in intracellular Ca2+ concentration in podocytes, presumably leading to albuminuria, and this likely occurs via calcium-conducting ion channels. Therefore, it appeared important to study calcium handling in the podocytes both under normal conditions and in various pathological states. However, available experimental approaches have remained somewhat limited to cultured and transfected cells. Although they represent a good basic model for such studies, they are essentially extracted from the native environment of the glomerulus. Here we describe the methodology of studying podocytes as a part of the freshly isolated whole glomerulus. This preparation retains the functional potential of the podocytes, which are still attached to the capillaries; therefore, podocytes remain in the environment that conserves the major parts of the glomeruli filtration apparatus. The present manuscript elaborates on two experimental approaches that allow 1) real-time detection of calcium concentration changes with the help of ratiometric confocal fluorescence microscopy, and 2) the recording of the single ion channels activity in the podocytes of the freshly isolated glomeruli. These methodologies utilize the advantages of the native environment of the glomerulus that enable researchers to resolve acute changes in the intracellular calcium handling in response to applications of various agents, measure basal concentration of calcium within the cells (for instance, to evaluate disease progression), and assess and manipulate calcium conductance at the level of single ion channels.

Changes in the intracellular calcium levels in the podocytes are one of the most important means to control the filtration function of glomeruli. Here we explain a high-throughput approach that allows detection of real-time calcium handling and single ion channels activity in the podocytes of the freshly isolated glomeruli.

Podocytes (renal glomerular epithelial cells) are known to regulate glomerular permeability and maintain glomerular structure; a key role for these cells ...

Implementing Patch Clamp and Live Fluorescence Microscopy to Monitor Functional Properties of Freshly Isolated PKD Epithelium

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal fluid accumulation and extracellular matrix formation. Activity of ion channels and intracellular calcium signaling are key physiologic parameters which determine functions of tubular epithelium. We developed a method suitable for real-time observation of ion channels activity with patch-clamp technique and registration of intracellular Ca2+ level in epithelial monolayers freshly isolated from renal cysts. PCK rats, a genetic model of autosomal recessive polycystic kidney disease (ARPKD), were used here for ex vivo analysis of ion channels and calcium flux. Described here is a detailed step-by-step procedure designed to isolate cystic monolayers and non-dilated tubules from PCK or normal Sprague Dawley (SD) rats, and monitor single channel activity and intracellular Ca2+ dynamics. This method does not require enzymatic processing and allows analysis in a native setting of freshly isolated epithelial monolayer. Moreover, this technique is very sensitive to intracellular calcium changes and generates high resolution images for precise measurements. Finally, isolated cystic epithelium can be further used for staining with antibodies or dyes, preparation of primary cultures and purification for various biochemical assays.

Ion channels expressed in renal tubular epithelium play a significant role in the pathology of polycystic kidney disease. Here we describe experimental protocols used to perform patch-clamp analysis and intracellular calcium level measurements in cystic epithelium freshly isolated from rodent kidneys.

Cyst initiation and expansion during polycystic kidney disease is a complex process characterized by abnormalities in tubular cell proliferation, luminal ...

The Mouse Isolated Perfused Kidney Technique

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a prerequisite for studying the physiology of the isolated organ and for many innovative applications that may be possible in the future, including perfusion decellularization for kidney bioengineering or the administration of anti-rejection or genome-editing drugs in high doses to prime the kidney for transplantation. During the time of the perfusion, the kidney can be manipulated, renal function can be assessed, and various pharmaceuticals administered. After the procedure, the kidney can be transplanted or processed for molecular biology, biochemical analysis, or microscopy.
This paper describes the perfusate and the surgical technique needed for the ex vivo perfusion of mouse kidneys. Details of the perfusion apparatus are given and data are presented showing the viability of the kidney&#39;s preparation: renal blood flow, vascular resistance, and urine data as functional, transmission electron micrographs of different nephron segments as morphological readouts, and western blots of transport proteins of different nephron segments as molecular readout.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. The buffers and surgical technique are described in detail.

The mouse isolated perfused kidney (MIPK) is a technique for keeping a mouse kidney under ex vivo conditions perfused and functional for 1 hr. This is a ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

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 ...