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). I...

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><tbody><tr><td>Glucose</td><td>Sigma-Aldrich (St. Louis, MO, USA)</td><td>G7021</td><td></td></tr><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>M2670</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma</td><td>223506</td><td></td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td></td></tr><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&amp;nbsp;</td><td>Warner Instruments, Camden, CT, USA</td><td>RC-27</td><td></td></tr><tr><td>Glass coverslip bottom (2.4&amp;nbsp;&amp;times;&amp;nbsp;5.0&amp;nbsp;cm)</td><td>ThermoFisher Scientific</td><td>12-548-5M</td><td></td></tr><tr><td>Anodized aluminum platform (diameter: 7.8&amp;nbsp;cm)&amp;nbsp;</td><td>Warner Instruments</td><td>PM6 or PH6</td><td></td></tr><tr><td>Compact aluminum stage&amp;nbsp;</td><td>Siskiyou, Grants Pass, OR, USA</td><td>8090P</td><td></td></tr><tr><td>Micromanipulator</td><td>Siskiyou</td><td>&amp;nbsp;MX10</td><td></td></tr><tr><td>Stereomicroscopes&amp;nbsp;</td><td>Zeiss, NY, USA</td><td>Stemi 2000 &amp;amp; 2000-C</td><td></td></tr><tr><td>Fiber optic light sources</td><td>&amp;nbsp;Schott, Mainz, Germany &amp;amp; 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&amp;nbsp;</td><td>Nikon Instruments Inc</td><td>&amp;nbsp;(Ph1 DL; 10X &amp;amp; 20X)</td><td></td></tr><tr><td>Fluorescent objectives&amp;nbsp;</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 )&amp;nbsp;</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>&amp;nbsp;Micro-g</td><td></td></tr><tr><td>Amplifiers</td><td>Molecular Devices, Sunnyvale, CA, USA</td><td>Axoclamp 2B &amp;amp; Axoclamp 900A</td><td></td></tr><tr><td>Headstages&amp;nbsp;</td><td>Molecular Devices</td><td>HS-2A &amp;amp; HS-9A</td><td></td></tr><tr><td>Function generator&amp;nbsp;</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>&amp;nbsp;BM-A-TM</td><td></td></tr><tr><td>Digital Storage Oscilloscope</td><td>Tektronix, Beaverton, Oregon, USA</td><td>&amp;nbsp;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&amp;nbsp;&amp;nbsp;</td><td>Warner Instruments</td><td>TC-344B or C</td><td></td></tr><tr><td>Inline Heater&amp;nbsp;</td><td>Warner Instruments</td><td>SH- 27B</td><td></td></tr><tr><td>Valve Controller&amp;nbsp;</td><td>Warner Instruments</td><td>VC-6</td><td></td></tr><tr><td>Inline Flow Control Valve</td><td>Warner Instruments</td><td>&amp;nbsp;FR-50</td><td></td></tr><tr><td>Electronic Puller&amp;nbsp;</td><td>Sutter Instruments, Novato, CA, USA</td><td>P-97 or P-1000&amp;nbsp;</td><td></td></tr><tr><td>Microforge</td><td>Narishige, East Meadow, NY, USA</td><td>&amp;nbsp;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 &amp;micro;m)&amp;nbsp;&amp;nbsp;</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 &amp;amp; 9.5&amp;nbsp;mm)</td><td>World Precision Instruments</td><td>555640S, 14364</td><td></td></tr><tr><td>Scissors 3 &amp;amp; 7 mm blades</td><td>Fine Science Tools (or FST), Foster City, CA, USA</td><td>Moria MC52 &amp;amp; 15000-00</td><td></td></tr><tr><td>Sharpened fine-tipped forceps&amp;nbsp;</td><td>FST</td><td>Dumont #5 &amp;amp; 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...

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

Research

JoVE Journal

Biology

8.1K Views.  Loma Linda University. Deze methode kan helpen bij het beantwoorden van belangrijke vragen op het gebied van cerebrale vasculaire fysiologie als het gaat om cerebrale bloedstroom verordening tijdens veroudering en ontwikkeling van chronische ziekten. Het belangrijkste voordeel van deze techniek is dat het gelijktijdige meting van intracellulair calcium- en membraanpotentieel van cerebrale arteriële endotheel in zijn oorspronkelijke vorm onder fysiologische omstandigheden mogelijk maakt. Om de muishersenen te isole...

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

Optical Mapping of Action Potentials and Calcium Transients in the Mouse Heart

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. Cardiovascular physiological phenotyping of the mouse heart can be easily done using fluorescence imaging employing various probes for transmembrane potential (Vm), calcium transients (CaT), and other parameters. Excitation-contraction coupling is characterized by action potential and intracellular calcium dynamics; therefore, it is critically important to map both Vm and CaT simultaneously from the same location on the heart1-4. Simultaneous optical mapping from Langendorff perfused mouse hearts has the potential to elucidate mechanisms underlying heart failure, arrhythmias, metabolic disease, and other heart diseases. Visualization of activation, conduction velocity, action potential duration, and other parameters at a myriad of sites cannot be achieved from cellular level investigation but is well solved by optical mapping1,5,6. In this paper we present the instrumentation setup and experimental conditions for simultaneous optical mapping of Vm and CaT in mouse hearts with high spatio-temporal resolution using state-of-the-art CMOS imaging technology. Consistent optical recordings obtained with this method illustrate that simultaneous optical mapping of Langendorff perfused mouse hearts is both feasible and reliable.

This paper details the dissection procedure, instrumental setup, and experimental conditions during optical mapping of transmembrane potential (Vm) and intracellular calcium transient (CaT) in intact isolated Langendorff perfused mouse hearts.

The mouse heart is a popular model for cardiovascular studies due to the existence of low cost technology for genetic engineering in this species. ...

Bioengineering

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, ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Neuroscience

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of individual cells. Since an increase of intracellular calcium (Ca2+) in the cytosol often occurs because of the electrical activity, or in response to a myriad of other stimuli, this process can be monitored by the imaging of cells loaded with fluorescent calcium-sensitive dyes.&#160; However, it is difficult to image this response in an individual cell type within whole tissue because these dyes are taken up by all cell types within the tissue. In contrast, genetically encoded calcium indicators (GECIs) can be expressed by an individual cell type and fluoresce in response to an increase of intracellular Ca2+, thus permitting the imaging of Ca2+ signaling in entire populations of individual cell types. Here, we apply the use of the GECIs GCaMP3/6 to the mouse neuromuscular junction, a tripartite synapse between motor neurons, skeletal muscle, and terminal/perisynaptic Schwann cells. We demonstrate the utility of this technique in classic ex vivo tissue preparations. Using an optical splitter, we perform dual-wavelength imaging of dynamic Ca2+ signals and a static label of the neuromuscular junction (NMJ) in an approach that could be easily adapted to monitor two cell-specific GECI or genetically encoded voltage indicators (GEVI) simultaneously. Finally, we discuss the routines used to capture spatial maps of fluorescence intensity. Together, these optical, transgenic, and analytic techniques can be employed to study the biological activity of distinct cell subpopulations at the NMJ in a wide variety of contexts.

Ex Vivo Imaging of Cell-specific Calcium Signaling at the Tripartite Synapse of the Mouse Diaphragm

Here we present a protocol to image calcium signaling in populations of individual cell types at the murine neuromuscular junction.

The electrical activity of cells in tissues can be monitored by electrophysiological techniques, but these are usually limited to the analysis of ...

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to correlate this with local hemodynamics. This protocol uses transgenic mice that have been prepared with a chronic cranial window and express the genetically encoded calcium indicator, RCaMP1.07, under the &#945;-smooth muscle actin promoter to specifically label mural cells, such as vascular smooth muscle cells and ensheathing pericytes. Steps are outlined on how to prepare a tail vein catheter for intravenous injection of fluorescent dyes to trace blood flow, as well as how to measure brain pericyte calcium and local blood vessel hemodynamics (diameter, red blood cell velocity, etc.) by two photon microscopy in vivo through the cranial window in ketamine/xylazine anesthetized mice. Finally, details are provided for the analysis of calcium fluctuations and blood flow movies via the image processing algorithms developed by Barrett et al. 2018, with an emphasis on how these processes can be adapted to other cellular imaging data.

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

This protocol presents steps to acquire and analyze fluorescent calcium images from brain ensheathing pericytes and blood flow data from nearby blood vessels in anesthetized mice. These techniques are useful for studies of mural cell physiology and can be adapted to investigate calcium transients in any cell type.

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to ...

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 ...

Isolation and Functional Analysis of Arteriolar Endothelium of Mouse Brain Parenchyma

Cerebral blood flow is conveyed by vascular resistance arteries and downstream parenchymal arterioles. Steady-state vascular resistance to blood flow increases with decreasing diameter from arteries to arterioles that ultimately feed into capillaries. Due to their smaller size and location in the parenchyma, arterioles have been relatively understudied and with less reproducibility in findings than surface pial arteries. Regardless, arteriolar endothelial cell structure and function&#8212;integral to the physiology and etiology of chronic degenerative diseases&#8212;requires extensive investigation. In particular, emerging evidence demonstrates that compromised endothelial function precedes and exacerbates cognitive impairment and dementia.
In the parenchymal microcirculation, endothelial K+ channel function is the most robust stimulus to finely control the spread of vasodilation to promote increases in blood flow to areas of neuronal activity. This paper illustrates a refined method for freshly isolating intact and electrically coupled endothelial &#34;tubes&#34; (diameter, ~25 &#181;m) from mouse brain parenchymal arterioles. Arteriolar endothelial tubes are secured during physiological conditions (37 &#176;C, pH 7.4) to resolve experimental variables that encompass K+ channel function and their regulation, including intracellular Ca2+ dynamics, changes in membrane potential, and membrane lipid regulation. A distinct technical advantage versus arterial endothelium is the enhanced morphological resolution of cell and organelle (e.g., mitochondria) dimensions, which expands the usefulness of this technique. Healthy cerebral perfusion throughout life entails robust endothelial function in parenchymal arterioles, directly linking blood flow to the fueling of neuronal and glial activity throughout precise anatomical regions of the brain. Thus, it is expected that this method will significantly advance the general knowledge of vascular physiology and neuroscience concerning the healthy and diseased brain.

Intensive preparation of intact mouse cerebral endothelial &#34;tubes&#34; from cerebral parenchymal arterioles is illustrated for studying cerebral blood flow regulation. Further, we demonstrate the experimental strengths of this endothelial study model for fluorescence imaging and electrophysiology measurement of key cellular signaling pathways, including changes in intracellular [Ca2+] and membrane potential.

Cerebral blood flow is conveyed by vascular resistance arteries and downstream parenchymal arterioles. Steady-state vascular resistance to blood flow ...

Simultaneous Measurement of Intracellular Calcium and Membrane Potential in a Mouse Cerebral Endothelium

Source: Hakim, M. A. et al. <a href="https://app.jove.com/v/58832">Simultaneous Measurements of Intracellular Calcium and Membrane Potential in Freshly Isolated and Intact Mouse Cerebral Endothelium.</a> J. Vis. Exp. (2019)
The video demonstrates the simultaneous measurement of intracellular calcium levels and changes in membrane potential in endothelial cells of mouse cerebral artery segments. The isolated endothelial tube is loaded with a fluorescent calcium indicator and treated with a drug that induces intracellular calcium release, resulting in a change in membrane potential. The indicator&#39;s fluorescence enables the measurement of calcium levels, and a recording electrode is inserted into a cell to measure changes in membrane potential.

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Dissection and Isolation of ...

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

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