eoe sub articles

EoE Sub Articles

All procedures involving sample collection have been performed in accordance with the institute&#39;s IRB guidelines.
1. Dissection and Isolation of Cerebral Artery
NOTE:&#160;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)&#160;via&#160;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+&#160;Physiological salt solution (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+&#160;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:&#160;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+&#160;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 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>
2. Preparation of Endothelial Tube and Superfusion
<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 room temperature (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&#160;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>
3. 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 intracellular Ca2+ ([Ca2+]i) responses, load the endothelial tube with Fura-2 AM dye (Fura-2-acetoxymethyl ester, 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>
4. Simultaneous Measurement of [Ca2+]i&#160;and Vm
<ol>
	<li>Turn on all other equipment and the electrometer software suite for measuring membrane potential (Vm, see&#160;Figure 3,&#160;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&#160;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&#160;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&#160;is stable at its expected values (-30 to -40 mV), 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:&#160;To test intercellular electrical coupling, current (&#177; 0.5-3 nA, ~20 s pulse duration) can be delivered&#160;via&#160;one electrometer as &#34;Site 1&#34;, and changes of Vm&#160;can be recorded with another electrometer as &#34;Site 2&#34; (distance &#8805;100 &#181;m).</li>
	<li>Once simultaneous measurements of Vm&#160;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., adenosine triphosphate [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&#160;and the F340/F380&#160;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&#160;~0 mV as referenced by the bath electrode. Stop respective recordings of Vm&#160;and [Ca2+]i&#160;and save the records on file for data analysis.</li>
</ol>

<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>
		</tr>
		<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>
		</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&#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>
		</tr>
		<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 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.

<img alt="Figure 1" src="/files/ftp_upload/22848/22848_Figure1.jpg" /> 
Figure 1: Isolation of cerebral artery from brain.&#160;(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.&#160;Panel A). (C) Isolated posterior cerebral arteries secured with stainless steel pins in sp...

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

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

Take segments of a mouse posterior cerebral artery.
Incubate with an enzyme solution to degrade the tissue matrix, loosening the cells.
Replace the solution with a buffer, then transfer a segment to a recording chamber.
Mechanically dissociate the connective tissue and smooth muscle cell layers, isolating the underlying endothelial tube.
Secure the tube, remove the dissociated layers, and add a physiological buffer.
Place the chamber within a recording system with continuous buffer flow.
Introduce a fluorescent calcium indicator that enters the endothelial cells, then insert a recording electrode into a cell.
Administer a drug that triggers calcium ion influx and induces downstream signaling for calcium release from the endoplasmic reticulum.
The indicator binds to the calcium and, upon excitation, emits fluorescence.
Calcium increase activates calcium-dependent potassium channels, causing potassium outflow and altering the membrane potential.
Simultaneously record changes in intracellular calcium via the fluorescent signal and membrane potential using the electrode.

simultaneous-measurement-intracellular-calcium-membrane-potential

Universidad San Sebastian

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Imaging and Optical Techniques

44 Views. Źródło: Hakim, M. A. et al. Jednoczesne pomiary wewnątrzkomórkowego potencjału wapnia i błony w świeżo wyizolowanym i nienaruszonym śródbłonku mózgu myszy. J. Vis. Exp. (2019 rok) Wideo pokazuje jednoczesny pomiar wewnątrzkomórkowego poziomu wapnia i zmian potencjału błonowego w komórkach śródbłonka mysich odcinków tętnic mózgowych. Izolowany śródbłonek jest ładowany fluorescencyjnym wskaźnikiem wapnia i traktowany lekiem, który indukuje wewnątrzkomórkowe uwalnia...

Video: Jednoczesny pomiar wewnątrzkomórkowego potencjału wapnia i błony w śródbłonku mózgu myszy - Concept

Microelectrode Impalement Method to Record Membrane Potential from a Cannulated Middle Cerebral Artery

Membrane potential (Vm) of vascular smooth muscle cells determines vessel tone and thus blood flow to an organ. Changes in the expression and function of ion channels and electrogenic pumps that regulate Vm in disease conditions could potentially alter Vm, vascular tone, and blood flow. Thus, a basic understanding of electrophysiology and the methods necessary to accurately record Vm in healthy and diseased states are essential. This method will allow modulating Vm using different pharmacological agents to restore Vm. Although there are several methods, each with its advantages and disadvantages, this article provides protocols to record Vm from cannulated resistance vessels such as the middle cerebral artery using the microelectrode impalement method. Middle cerebral arteries are allowed to gain myogenic tone in a myograph chamber, and the vessel wall is impaled using high resistance microelectrodes. The Vm signal is collected through an electrometer, digitized, and analyzed. This method provides an accurate reading of the Vm of a vessel wall without damaging the cells and without changing the membrane resistance.

The primary goal of this article is to provide details of how to record membrane potential (Vm) from the middle cerebral artery using the microelectrode impalement method. The cannulated middle cerebral artery is equilibrated to gain myogenic tone, and the vessel wall is impaled using high resistance microelectrodes.

Membrane potential (Vm) of vascular smooth muscle cells determines vessel tone and thus blood flow to an organ. Changes in the expression and function of ...

JoVE Journal

Medicine

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+ ...

Biology

Whole Cell Electrophysiology of Primary Cultured Murine Enterochromaffin Cells

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory cells, EC cells sense luminal stimuli and convert them into serotonin (5-hyroxytryptamine, 5-HT) release events. However, the electrophysiology of these cells is poorly understood because they are difficult to culture and to identify. The method presented in this paper outlines primary EC cell cultures optimized for single cell electrophysiology. This protocol utilizes a transgenic cyan fluorescent protein (CFP) reporter to identify mouse EC cells in mixed primary cultures, advancing the approach to obtaining high-quality recordings of whole cell electrophysiology in voltage- and current-clamp modes.

Enterochromaffin (EC) cells comprise a small subset of gastrointestinal epithelial cells. EC cells are electrically excitable and release serotonin, yet difficulties in culturing and identifying EC cells have limited physiological studies. The method presented here establishes a primary culture model amenable to examination of single EC cells by electrophysiology.

Enterochromaffin (EC) cells in the gastrointestinal (GI) epithelium constitute the largest subpopulation of enteroendocrine cells. As specialized sensory ...

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

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