This research has been supported by grants from the National Institutes of Health (R00AG047198 &#38; R56AG062169 to EJB; R00HL140106 to PWP) and the Alzheimer&#39;s Association (AZRGD-21-805835 to PWP). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or the Alzheimer&#39;s Association.

Experimenters should ensure that designated use of animals and associated protocols are approved by their Institutional Animal Care and Use Committee (IACUC) and performed in accordance 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 and the University of Arizona has approved all protocols used for this manuscript for C57BL/6N and 3xTg-AD mice (males and females; age range: 2-30 months). See Figure 1 as an overview of the isolation and examination of arteriolar endothelial tubes freshly isolated from mouse parenchymal arterioles of the brain.
1. Materials and equipment
NOTE: See the Table of Materials for all reagents and materials required for this protocol. In addition, manuals and websites associated with the respective vendors can also be consulted as needed. Illustrations of dissection stations and experimental apparatuses have been previously provided13.
<ol>
	<li>Flow chamber
	<ol>
		<li>Fasten a superfusion chamber with a glass coverslip onto a platform composed of anodized aluminum. Secure the platform with the chamber onto an aluminum microscope stage.</li>
		<li>Set a micromanipulator holding a pinning pipette at each end of the platform on the aluminum microscope stage. 
		NOTE: If necessary, use a transferable stage apparatus with a flow chamber unit to move a secured, isolated endothelial tube from one microscope apparatus to another for experimentation.</li>
	</ol>
	</li>
	<li>Microscopes
	<ol>
		<li>Use stereomicroscopes (5x to 50x magnification range) and fiber optic light sources for dissection procedures.</li>
		<li>For isolation of endothelial tubes, use an inverted microscope rig equipped with phase contrast- or differential interference contrast (DIC)-compatible objectives (10x, 20x, and 40x) and an aluminum stage.</li>
		<li>Have a microsyringe pump controller ready next to the apparatus dedicated to isolating endothelial tubes from partially digested blood vessels.</li>
		<li>Set up the experimental apparatus by arranging an inverted microscope (objectives: 4x, 10x, 20x, 40x, and 60x) and a manual aluminum stage on a vibration isolation table.</li>
	</ol>
	</li>
	<li>Intracellular Vm recording equipment
	<ol>
		<li>Connect the electrometer to a compatible headstage. Use accessories, such as a function generator and stimulator, for protocols requiring current injection.</li>
		<li>Connect amplifier outputs to a data digitizer system, oscilloscope and audible baseline monitors. Secure the reference bath electrode (Ag/AgCl pellet) near the flow chamber exit.</li>
		<li>Assemble 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>Assemble a temperature controller equipped with an inline heater to raise and maintain a physiological temperature (37 &#176;C) throughout the experiment.</li>
		<li>Assemble a multichannel platform connected to a valve controller with an inline flow control valve to control delivery of solutions to endothelial tubes secured in the chamber.</li>
	</ol>
	</li>
	<li>Micropipettes and sharp electrodes 
	&#8203;NOTE: The experimenter will need an electronic glass puller and a microforge for preparing trituration and pinning pipettes.
	<ol>
		<li>To separate the endothelial tube from the partially digested arteriolar segment, prepare heat-polished trituration pipettes (internal tip diameter: 30-50 &#181;m) from borosilicate glass capillaries.</li>
		<li>To secure the endothelial tube in the superfusion chamber, prepare heat-polished pinning pipettes with a blunted, spherical end (outer diameter: 50-70 &#181;m) prepared from thin-wall borosilicate glass capillaries.</li>
		<li>To record Vm of an endothelial cell, prepare sharp electrodes with a tip resistance of ~150 &#177; 30 M&#937; from glass capillaries using the glass puller only.</li>
	</ol>
	</li>
</ol>
2. Solutions and drugs
<ol>
	<li>Physiological salt solution (PSS)
	<ol>
		<li>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>Prepare necessary solutions lacking CaCl2 (zero Ca2+ PSS) for dissection of cerebral arterioles and isolation of endothelial tube. 
		NOTE: Prepare all solutions in ultrapure deionized H2O, followed by filtration (0.22 &#181;m). Ensure that the final product contains a pH 7.4 with osmolality between 290 and 300 mOsm.</li>
	</ol>
	</li>
	<li>Bovine serum albumin (BSA, 10%)
	<ol>
		<li>Dissolve 1 g of the lyophilized BSA powder in a beaker of 10 mL of zero Ca2+ PSS. Cover the beaker and allow at least 2 h with slow stirring for the BSA to dissolve.</li>
		<li>Filter solution using a 10 mL syringe plus a 0.22 &#181;m filter and prepare 1 mL aliquots to be stored at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Dissection solution
	<ol>
		<li>Add 500 &#181;L of BSA (10%) to 49.5 mL of zero Ca2+ PSS for a total volume of 50 mL.</li>
		<li>Transfer solution to a Petri dish for arteriolar dissections.</li>
	</ol>
	</li>
	<li>Dissociation solution
	<ol>
		<li>Add 5 &#181;L of CaCl2&#160;solution (1 M) and 500 &#181;L of BSA (10%) to 49.5 mL of zero Ca2+ PSS for a total volume of 50 mL. Incubate solution at room temperature for at least 1 h.</li>
	</ol>
	</li>
	<li>Enzymes
	<ol>
		<li>Prepare 1 mL of digestion solution with 100 &#181;g/mL papain, 170 &#181;g/mL dithioerythritol, 250 &#181;g/mL collagenase (Type H blend), and 40 &#181;g/mL elastase. 
		NOTE: Enzyme activities can vary, although successful protocols will likely require the following enzymatic activities: &#8805; 10 units/mg for papain, &#8805; 1 unit/mg for collagenase, and &#8805; 5 units/mg for elastase.</li>
	</ol>
	</li>
	<li>Fura-2 and pharmacological agents
	<ol>
		<li>Prepare Fura-2 AM stock in dimethyl sulfoxide (DMSO; 1 mM). Prepare 500 &#181;L of working concentration (10 &#181;M) by adding 5 &#181;L of the stock to 495 &#181;L of PSS for loading.</li>
		<li>Prepare at least 50 mL of working concentrations of pharmacological agents&#160;in PSS or DMSO as appropriate.</li>
	</ol>
	</li>
	<li>Conducting solution
	<ol>
		<li>Prepare 2 M KCl by dissolving KCl in deionized H2O (7.455 g of KCl in 50 mL of H2O). Pass the solution through a syringe with a 0.22 &#181;m filter prior to backfilling the sharp electrodes.</li>
	</ol>
	</li>
	<li>Fluorescent organelle trackers and antibodies
	<ol>
		<li>Prepare plasma membrane or organellar (e.g., nucleus, endoplasmic reticulum) trackers in the appropriate physiological saline solution according to the manufacturer&#39;s instructions.</li>
		<li>Prepare primary and secondary antibodies according to the manufacturer&#39;s instructions in the appropriate physiological salt solution.</li>
	</ol>
	</li>
</ol>
3. Dissection and isolation of mouse cerebral arterioles
NOTE: Stereomicroscopes and sharpened microdissection tools (e.g., fine-tipped forceps, Vannas-style dissection scissors) must be used for specimen magnification (up to 50x) in all these dissection procedures.
<ol>
	<li>Isolation of mouse brain
	<ol>
		<li>Anesthetize a standard laboratory mouse (e.g., C57BL/6, 3xTg-AD; 2-30 months old, male or female) using isoflurane inhalation (3% for 2-3 min), followed by immediate decapitation using sharp scissors or guillotine per IACUC approval. Place the mouse head in a Petri dish (diameter 10 cm, depth 1.5 cm) containing cold (4 &#176;C) dissection solution.</li>
		<li>While viewing under a stereomicroscope, remove the skin and hair over the skull and wash away excessive blood with cold dissection solution. Using the tips of standard dissection scissors (e.g., 24 mm blades), make an incision starting with the occipital bone and extending up through the nasal bone of the skull. Use coarse-tipped forceps to carefully open the skull along the incision, and separate connective tissue (meningeal membrane) to isolate the brain containing an intact Circle of Willis.</li>
		<li>Wash the isolated brain with cold dissection solution in a beaker or Petri dish to remove remaining blood from the surface of the brain. Place the brain ventral side facing up in a chamber containing cold dissection solution for isolation of parenchymal arterioles (Figure 2A).</li>
	</ol>
	</li>
	<li>Isolation of parenchymal arterioles 
	NOTE: Arterioles arising from the middle cerebral arteries (MCAs) and embedded in brain parenchyma are chosen for this protocol. The arterioles are isolated as previously described11, with modification.
	<ol>
		<li>Use steel pins (diameter: 0.2 mm, length: 11 to 12 mm) to secure the isolated brain in cold dissection solution in a Petri dish containing a charcoal-infused silicon polymer coating (depth &#8805; 50 cm).</li>
		<li>Using sharp and aligned dissection scissors, cut a rectangle of brain tissue (length: 5 mm, width: 3 mm) (Figure 2B) around the MCA while ensuring that the upper part of the tissue segment is past the branching point from the Circle of Willis. Cut another rectangle of brain tissue from the other hemisphere of the brain to access more arterioles if necessary.</li>
		<li>Secure the separated brain tissue segment into the dish with the MCA facing upwards (distal from the Circle of Willis) using steel pins (diameter: 0.1 mm, length: 13 to 14 mm). Carefully make a shallow incision near the pins to remove the pia with small forceps, gently peeling from one end towards the other (Figure 2C). Remove the pia from the other tissue segment to access more arterioles if needed. Carefully secure the isolated pia with parenchymal arterioles branched from MCA in the dish with the pins, and dissect the parenchymal arterioles (Figure 2D), ensuring that the arterioles are not damaged. 
		NOTE: If the arterioles are not easily pulled out with the pia from the parenchyma, use fine forceps and dig into the brain, starting at the MCA branch point from the Circle of Willis. Locate the arteriole, carefully loosen the tissue around arterioles (Figure 2E), and gently pull the arteriole out of the parenchyma, holding the upper end of the arteriole (Figure 2F). Multiple arterioles (length: ~1.5-2 mm) can be isolated from one rectangle of brain tissue.</li>
		<li>Ensure the arteriole is clean with no tissue attached to it, and cut off any remaining distal branches (Figure 3A). Use this clean, intact arteriole for enzymatic digestion. Alternatively, cut each arteriole into two pieces (length: 0.75-1 mm) for enzymatic digestion for the preparation of endothelial tubes if desired.</li>
	</ol>
	</li>
</ol>
4. Digestion of parenchymal arterioles and preparation of endothelial tubes
<ol>
	<li>Arrangement of equipment and pipettes 
	NOTE: Isolation of arterial endothelial tubes from the brain has been described previously13,18. The current protocol incorporates modifications for the isolation of endothelium from parenchymal (intracerebral) arterioles. Multiple arterioles or/and pieces of arterioles can be used together for enzymatic digestion.
	<ol>
		<li>Assemble the trituration apparatus13 for preparing endothelial tubes using an aluminum stage supporting a chamber and micromanipulators. Assemble a microscope equipped with objectives (5x-60x) and a camera connected to a computer monitor. Secure a microsyringe with a pump controller next to the stage and specimen.</li>
		<li>Secure a trituration pipette backfilled with mineral oil over the microsyringe piston. Use the microsyringe with the pump controller to withdraw the dissociation solution into the trituration pipette (~130 nL) on top of the mineral oil while taking care to avoid air bubbles in the pipette.</li>
	</ol>
	</li>
	<li>Partial digestion of arteriolar segments
	<ol>
		<li>Place intact arteriolar segments into 1 mL of dissociation solution in a 10 mL glass tube containing 100 &#181;g/mL papain, 170 &#181;g/mL dithioerythritol, 250 &#181;g/mL collagenase, and 40 &#181;g/mL elastase. Incubate arteriolar segments at 34 &#176;C for 10-12 min.</li>
		<li>After digestion, replace the enzyme solution with 3-4 mL of fresh dissociation solution at room temperature.</li>
	</ol>
	</li>
	<li>Isolation of arteriolar endothelial tube 
	NOTE: Following digestion and replacement of the enzyme solution, transfer one or multiple partially digested segments into the dissociation solution in a superfusion chamber attached to a mobile platform. While viewing at 100x to 200x magnification, select one partially digested but unbroken vessel segment and focus on it under the microscope.
	<ol>
		<li>Place the trituration pipette attached with the microsyringe injector in the dissociation solution in the chamber and position it close to one end of the digested vessel segment. Set a rate within the range of 1-3 nL/s on the pump controller for gentle trituration.</li>
		<li>While maintaining 100x to 200x magnification, withdraw the arteriolar segment into the pipette and then eject to dissociate the adventitia and smooth muscle cells (Figure 3B). Triturate the vessel segment until all smooth muscle cells are dissociated, and only endothelial cells remain as an intact &#34;tube.&#34; 
		NOTE: If necessary during trituration, carefully use fine-tipped forceps to remove the dissociated adventitia and internal elastic lamina from the endothelial tube. Typically, only a few cycles of trituration yield an intact endothelial tube.</li>
		<li>Use micromanipulators to secure each end of the endothelial tube on the glass coverslip in the chamber with borosilicate glass pinning pipettes (Figure 3C,D).</li>
	</ol>
	</li>
	<li>Securing the arteriolar endothelial tube
	<ol>
		<li>Replace the dissociation solution with superfusion solution (PSS containing 2 mM CaCl2) while ensuring that nonendothelial material is washed out of the chamber.</li>
		<li>Transfer secured endothelial tube in the mobile platform to the microscope rig designed for continuous superfusion and intracellular or fluorescent recordings/imaging.</li>
		<li>Apply continuous delivery of PSS during the experiment. Manually set the flow rate (5-7 mL/min) throughout the experiment using the inline flow control valve, consistent with the laminar flow, while matching the feed of solution flow to the extent of vacuum suction. Allow &#8805;5 min of PSS flow to the chamber for superfusion of the endothelial tube before acquiring background data and/or dye loading.</li>
	</ol>
	</li>
</ol>
5. Utilization of arteriolar endothelial tubes for the examination of cellular physiology
NOTE: Isolated and secured arteriolar endothelial tubes can be used for intracellular recordings of [Ca2+]i dynamics and Vm using photometry and sharp electrode electrophysiology, respectively, as previously illustrated13 (Figure 4). [Ca2+]i and Vm can be measured as separate or combined experimental variables as needed13 (Figure 4). However, arteriolar endothelial tubes are more delicate than arterial endothelium, and experimentation time should not exceed 1 h.
<ol>
	<li>Measurement of [Ca2+]i
	<ol>
		<li>Turn on the equipment and software for [Ca2+]i recordings while maintaining continuous superfusion at a flow rate of 5-7 mL/min.</li>
		<li>Load the endothelial tube with the Ca2+ dye Fura-2 AM for 30 min at room temperature. Wash the cells with superfusion solution for another 20-30 min while gradually raising the bath temperature to 37 &#176;C. Maintain the temperature at 37 &#176;C throughout the experiment.</li>
		<li>Manually adjust the imaging window using photometry software to focus on ~20 endothelial cells (Figure 4A). In the absence of light, turn the PMT on the fluorescence interface and begin acquisition of [Ca2+]i by exciting Fura-2 alternately (&#8805;10 Hz) at 340 nm and 380 nm while collecting fluorescence emission at 510 nm. Once a stable baseline recording of [Ca2+]i is established, apply pharmacological agents (e.g., purinergic receptor agonists) per experimental objective (Figure 4B).</li>
	</ol>
	</li>
	<li>Measurement of Vm&#8203;
	<ol>
		<li>Turn on the equipment and software for Vm recordings and set the data acquisition rate (&#8805;10 Hz) while maintaining continuous superfusion at a flow rate of 5-7 mL/min. Gradually raise the bath temperature to 37 &#176;C and maintain it until the end of the experiment.</li>
		<li>Pull a sharp electrode using a borosilicate glass capillary, backfill with 2 M KCl, and secure it over a silver wire coated with chloride in the pipette holder attached to an electrometer, that in turn, is held by a micromanipulator.</li>
		<li>While viewing through the 4x objective, use a micromanipulator to carefully position the sharp electrode tip just over a cell of the arteriolar endothelial tube into the flowing PSS in the chamber.</li>
		<li>Gradually increase magnification to 400x and reposition the electrode tip as needed.</li>
		<li>Using the micromanipulator, gently insert the tip of a sharp electrode into one of the cells of the endothelial tube and start recording Vm using an electrometer (Figure 4A).</li>
		<li>Once the endothelial resting Vm is stable (&#8722;30 to &#8722;40 mV), apply the desired pharmacological agents per experimental objective (Figure 4C).</li>
	</ol>
	</li>
</ol>
6. Cellular imaging
NOTE: Endothelial tubes secured in the chamber of the mobile platform can also be used for microscopic imaging in both live and fixed conditions19 using standard fluorescence or confocal microscopy. Immunohistochemistry with different antibodies for receptors and ion channels can also be applied as previously described20.
<ol>
	<li>Load the endothelial tube with the fluorescence tracker for the plasma membrane or desired organelle (e.g., nucleus, endoplasmic reticulum) at 37 &#176;C for 15-30 min.</li>
	<li>Wash the cells with fresh superfusion PSS and image live cells under the microscope at the excitation wavelength of respective dyes (Figure 4D,E). 
	NOTE: Immunohistochemistry can be performed on this tube model using antibodies of interest.</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aging+impairs+electrical+conduction+along+endothelium+of+resistance+arteries+through+enhanced+Ca2+-activated+K++channel+activation.">Aging impairs electrical conduction along endothelium of resistance arteries through enhanced Ca2+-activated K+ channel activation.</a> Arteriosclerosis Thrombosis and Vascular Biology. 33 (8), 1892-1901 (2013).</li><li>Attems, J., Jellinger, K. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+overlap+between+vascular+disease+and+Alzheimer's+disease--lessons+from+pathology.">The overlap between vascular disease and Alzheimer's disease--lessons from pathology.</a> BMC Medicine. 12, 206 (2014).</li><li>Fisher, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+arterial+lesions+underlying+lacunes.">The arterial lesions underlying lacunes.</a> Acta Neuropathologica. 12 (1), 1-15 (1968).</li><li>Behringer, E. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+and+electrical+signaling+in+arterial+endothelial+tubes:+New+insights+into+cellular+physiology+and+cardiovascular+function.">Calcium and electrical signaling in arterial endothelial tubes: New insights into cellular physiology and cardiovascular function.</a> Microcirculation. 24 (3), (2017).</li><li>Dunn, K. M., Nelson, M. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurovascular+signaling+in+the+brain+and+the+pathological+consequences+of+hypertension.">Neurovascular signaling in the brain and the pathological consequences of hypertension.</a> American Journal of Physiology-Heart and Circulatory Physiology. 306 (1), 1-14 (2014).</li><li>Cipolla, M. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Increased+pressure-induced+tone+in+rat+parenchymal+arterioles+vs.+middle+cerebral+arteries:+role+of+ion+channels+and+calcium+sensitivity.">Increased pressure-induced tone in rat parenchymal arterioles vs. middle cerebral arteries: role of ion channels and calcium sensitivity.</a> Journal of Applied Physiology. 117 (1), 53-59 (2014).</li><li>Cipolla, M. J., Smith, J., Kohlmeyer, M. M., Godfrey, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=SKCa+and+IKCa+Channels,+myogenic+tone,+and+vasodilator+responses+in+middle+cerebral+arteries+and+parenchymal+arterioles:+effect+of+ischemia+and+reperfusion.">SKCa and IKCa Channels, myogenic tone, and vasodilator responses in middle cerebral arteries and parenchymal arterioles: effect of ischemia and reperfusion.</a> Stroke. 40 (4), 1451-1457 (2009).</li><li>Chen, Y. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium+signal+profiles+in+vascular+endothelium+from+Cdh5-GCaMP8+and+Cx40-GCaMP2+mice.">Calcium signal profiles in vascular endothelium from Cdh5-GCaMP8 and Cx40-GCaMP2 mice.</a> Journal of Vascular Research. 58 (3), 159-171 (2021).</li><li>Bando, Y., Sakamoto, M., Kim, S., Ayzenshtat, I., Yuste, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+evaluation+of+genetically+encoded+voltage+indicators.">Comparative evaluation of genetically encoded voltage indicators.</a> Cell Reports. 26 (3), 802-813 (2019).</li><li>Pires, P. W., Sullivan, M. N., Pritchard, H. A., Robinson, J. J., Earley, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Unitary+TRPV3+channel+Ca2++influx+events+elicit+endothelium-dependent+dilation+of+cerebral+parenchymal+arterioles.">Unitary TRPV3 channel Ca2+ influx events elicit endothelium-dependent dilation of cerebral parenchymal arterioles.</a> American Journal of Physiology-Heart and Circulatory Physiology. 309 (12), 2031-2041 (2015).</li><li>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=Tuning+electrical+conduction+along+endothelial+tubes+of+resistance+arteries+through+Ca2+-activated+K++channels.">Tuning electrical conduction along endothelial tubes of resistance arteries through Ca2+-activated K+ channels.</a> Circulation Research. 110 (10), 1311-1321 (2012).</li><li>Behringer, E. J., Socha, M. J., Polo-Parada, L., 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=Electrical+conduction+along+endothelial+cell+tubes+from+mouse+feed+arteries:+confounding+actions+of+glycyrrhetinic+acid+derivatives.">Electrical conduction along endothelial cell tubes from mouse feed arteries: confounding actions of glycyrrhetinic acid derivatives.</a> British Journal of Pharmacology. 166 (2), 774-787 (2012).</li><li>Thomsen, M. S., Routhe, L. J., Moos, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+vascular+basement+membrane+in+the+healthy+and+pathological+brain.">The vascular basement membrane in the healthy and pathological brain.</a> Journal of Cerebral of Blood Flow and Metabolism. 37 (10), 3300-3317 (2017).</li><li>Jambusaria, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Endothelial+heterogeneity+across+distinct+vascular+beds+during+homeostasis+and+inflammation.">Endothelial heterogeneity across distinct vascular beds during homeostasis and inflammation.</a> elife. 9, 51413 (2020).</li><li>Diaz-Otero, J. M., Garver, H., Fink, G. D., Jackson, W. F., Dorrance, A. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aging+is+associated+with+changes+to+the+biomechanical+properties+of+the+posterior+cerebral+artery+and+parenchymal+arterioles.">Aging is associated with changes to the biomechanical properties of the posterior cerebral artery and parenchymal arterioles.</a> American Journal of Physiology-Heart and Circulatory Physiology. 310 (3), 365-375 (2016).</li><li>Chen, M. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brain+endothelial+cells+are+exquisite+sensors+of+age-related+circulatory+cues.">Brain endothelial cells are exquisite sensors of age-related circulatory cues.</a> Cell Reports. 30 (13), 4418-4432 (2020).</li></ol>

The authors declare no conflicts of interest.

Growing evidence suggests that cerebrovascular disease (CVD), aging, and Alzheimer&#39;s disease are strongly correlated and are a current topic of dementia research4,8,14,21. Thus, it is obvious that studies of the cerebrovascular network would have a broad impact on health while requiring continued extensive investigation during conditions of&#160;disease. As a significant point of vascular r...

Parenchymal arterioles directly deliver essential oxygen and nutrients throughout the brain1. While interfacing with capillaries, highly vasoactive arterioles respond to retrograde signaling initiated by capillary ion channels that sense metabolic signals from specific neuronal regions2. With brain parenchyma having historically received the bulk of investigation, a role for endothelial dysfunction has now emerged for clarifying pathological mechanisms associated with various cerebrovascular disorders that underlie dementia (e.g., ischemic stroke, Alzheimer&#39;s disease)3,4,5,6. The endothelium is integral to perfusion of the brain in accord with the heterogeneity of genetics, structure, and function throughout vascular segments7. Pial arteries have been extensively studied due to their relatively large size, high segmental vascular resistance, and role in blood flow distribution to the underlying cerebrum8,9. Thus, a better understanding of arteriolar endothelial mechanisms will likely enhance the understanding of brain blood flow regulation in health and disease towards the development of novel therapeutic regimens.
Emerging evidence highlights the importance of studying parenchymal arterioles in relation to different signaling pathways and diseases8,10. However, this approach has been limited to using intact pressurized arteriole11 and/or capillary-parenchymal arteriole (CaPA) preparations12. Freshly isolated, native cerebral arteriolar endothelial cells devoid of other cell types and confounding factors have not been examined, likely due to technical difficulties in their isolation. This paper advances a previous technique highlighting the isolation of pial arterial endothelium13 to now reliably and reproducibly isolate the endothelium of brain parenchymal arterioles (width: ~25 &#181;m, length: ~250 &#181;m). This technique helps achieve optimal resolution of electrically and chemically coupled cells in their individual orientation and cellular networks.
Key pathways of interest have included the interaction of intracellular Ca2+ ([Ca2+]i) signaling and hyperpolarization of membrane potential (Vm)14,15&#8212;integral to vasodilation16&#8212;to allow blood to enter the capillaries and deliver oxygen and nutrients to active parenchyma17. These preparations allow for real-time electrophysiological recordings of ion channels, including Ca2+-permeant, transient receptor potential (TRP) and K+ channels and/or fluorescent imaging of intracellular organelles within endothelial cell tubes in near-physiological conditions. This is a suitable technique for researchers interested in physiological cellular mechanisms that govern endothelial cell control of cerebral blood flow delivery to the brain parenchyma. Altogether, this technique will help researchers better understand fundamental endothelial signaling pathways and network communication of arterioles embedded in brain parenchyma while&#160;addressing questions related to cerebrovascular physiology and pathology.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Amplifiers</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>Axoclamp 2B &#38; Axoclamp 900A</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>Bath Chiller (Isotemp 500LCU)</td>
			<td>ThermoFisher Scientific</td>
			<td>13874647</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries (Pinning)</td>
			<td>Warner Instruments</td>
			<td>G150T-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries (Sharp Electrodes)</td>
			<td>Warner Instruments</td>
			<td>GC100F-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Borosilicate glass capillaries (Trituration)</td>
			<td>World Precision Instruments (WPI), Sarasota, FL, USA</td>
			<td>1B100-4</td>
			<td></td>
		</tr>
		<tr>
			<td>BSA: Bovine Serum Albumin</td>
			<td>Sigma</td>
			<td>A7906</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2: Calcium Chloride</td>
			<td>Sigma</td>
			<td>223506</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase (Type H Blend)</td>
			<td>Sigma</td>
			<td>C8051</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Glass (2.4&#160;&#215;&#160;5.0&#160;cm)</td>
			<td>ThermoFisher Scientific</td>
			<td>12-548-5M</td>
			<td></td>
		</tr>
		<tr>
			<td>Data Acquision Digitizer</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>Digidata 1550A</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection Dish (Glass Petri with Charcoal Sylgard bottom)</td>
			<td>Living Systems Instrumentation, St. Albans City, VT, USA</td>
			<td>DD-90-S-BLK</td>
			<td></td>
		</tr>
		<tr>
			<td>Dithioerythritol</td>
			<td>Sigma</td>
			<td>D8255</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO: Dimethyl Sulfoxide</td>
			<td>Sigma</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr>
			<td>Elastase (porcine pancreas)</td>
			<td>Sigma</td>
			<td>E7885</td>
			<td></td>
		</tr>
		<tr>
			<td>Endoplasmic Reticulum Tracker (ER-Tracker Red, BODIPY TR Glibenclamide)</td>
			<td>ThermoFisher Scientific</td>
			<td>E34250</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>Flow Control Valve</td>
			<td>Warner Instruments</td>
			<td>&#160;FR-50</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescence system interface, ARC lamp &#38; power supply, hyperswitch and PMT</td>
			<td>Molecular Devices, Sunnyvale, CA, USA</td>
			<td>IonOptix Systems</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps (Fine-tipped, sharpened)</td>
			<td>FST</td>
			<td>Dumont #5 &#38; Dumont #55</td>
			<td></td>
		</tr>
		<tr>
			<td>Function Generator</td>
			<td>EZ Digital, Seoul, South Korea</td>
			<td>FG-8002</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>Glucose</td>
			<td>Sigma-Aldrich (St. Louis, MO, USA)</td>
			<td>G7021</td>
			<td></td>
		</tr>
		<tr>
			<td>HCl: Hydrochloric Acid</td>
			<td>ThermoFisher Scientific (Pittsburgh, PA, USA)</td>
			<td>A466250</td>
			<td></td>
		</tr>
		<tr>
			<td>Headstages</td>
			<td>Molecular Devices</td>
			<td>HS-2A &#38; HS-9A</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES: (4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid)</td>
			<td>Sigma</td>
			<td>H4034</td>
			<td></td>
		</tr>
		<tr>
			<td>Inline Solution Heater</td>
			<td>Warner Instruments</td>
			<td>SH-27B</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl: Potassium Chloride</td>
			<td>Sigma</td>
			<td>P9541</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2: Magnesium Chloride</td>
			<td>Sigma</td>
			<td>M2670</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>Micromanipulator</td>
			<td>Siskiyou</td>
			<td>&#160;MX10</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller (digital)</td>
			<td>Sutter Instruments, Novato, CA, USA</td>
			<td>P-97 or P-1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope (Nikon-inverted)</td>
			<td>Nikon Instruments Inc, Melville, NY, USA</td>
			<td>Ts2</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope (Nikon-inverted)</td>
			<td>Nikon Instruments Inc</td>
			<td>Eclipse TS100</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope objectives</td>
			<td>Nikon Instruments Inc</td>
			<td>20X (S-Fluor) and 40X (Plan Fluor)</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope platform (anodized aluminum; diameter, 7.8&#160;cm)</td>
			<td>Warner Instruments</td>
			<td>PM6 or PH6</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Stage (Aluminum)</td>
			<td>Siskiyou, Grants Pass, OR, USA</td>
			<td>8090P</td>
			<td></td>
		</tr>
		<tr>
			<td>Microsyringe Pump Controller</td>
			<td>World Precision Instruments (WPI), Sarasota, FL, USA</td>
			<td>SYS-MICRO4</td>
			<td></td>
		</tr>
		<tr>
			<td>MTA: 2-Methylthioadenosine diphosphate trisodium salt</td>
			<td>Tocris</td>
			<td>1624</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl: Sodium Chloride</td>
			<td>Sigma</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>NaOH: Sodium Hydroxide</td>
			<td>Sigma</td>
			<td>S8045</td>
			<td></td>
		</tr>
		<tr>
			<td>Nuclear Stain (NucBlue Live ReadyProbes Reagent; Hoechst 33342)</td>
			<td>ThermoFisher Scientific</td>
			<td>R37605</td>
			<td></td>
		</tr>
		<tr>
			<td>Oscilloscope</td>
			<td>Tektronix, Beaverton, Oregon, USA</td>
			<td>&#160;TDS 2024B</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Sigma</td>
			<td>P4762</td>
			<td></td>
		</tr>
		<tr>
			<td>Phase contrast objectives</td>
			<td>Nikon Instruments Inc</td>
			<td>&#160;(Ph1 DL; 10X &#38; 20X)</td>
			<td></td>
		</tr>
		<tr>
			<td>Plasma Membrane Stain (CellMask Deep Red)</td>
			<td>ThermoFisher Scientific</td>
			<td>C10046</td>
			<td></td>
		</tr>
		<tr>
			<td>Plexiglas superfusion chamber</td>
			<td>Warner Instruments, Camden, CT, USA</td>
			<td>RC-27</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors (3 mm &#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>Scissors (Vannas style; 9.5&#160;mm &#38; 3&#160;mm blades)</td>
			<td>World Precision Instruments</td>
			<td>555640S, 14364</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscopes</td>
			<td>Zeiss, NY, USA</td>
			<td>Stemi 2000 &#38; 2000-C</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe filter (0.22 &#181;m)</td>
			<td>ThermoFisher Scientific</td>
			<td>722-2520</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller (Dual Channel)</td>
			<td>Warner Instruments</td>
			<td>TC-344B or C</td>
			<td></td>
		</tr>
		<tr>
			<td>Valve Control System</td>
			<td>Warner Instruments</td>
			<td>VC-6</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>
	</tbody>
</table>

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.

A demonstration of the protocol is shown in Figure 1 with arteriolar dissection and endothelial tube isolation steps as Figure 2 and Figure 3, respectively. Here, endothelial function was assessed by measuring [Ca2+]i and Vm using Fura-2 photometry and sharp electrode electrophysiology (Figure 4A) in response to a pharmacological agent [2-methylthioadenosine diphosphate ...

Watch this Scientific Journal Video about Isolation and Functional Analysis of Arteriolar Endothelium of Mouse Brain Parenchyma at JoVE.com

Isolation and Functional Analysis of Arteriolar Endothelium of Mouse Brain Parenchyma

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 ...

isolation-functional-analysis-arteriolar-endothelium-mouse-brain

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2.7K Views.  Loma Linda University. Intensive preparation of intact mouse cerebral endothelial "tubes" 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.

Video: Isolation and Functional Analysis of Arteriolar Endothelium of Mouse Brain Parenchyma

Analysis of Physiologic E-Selectin-Mediated Leukocyte Rolling on Microvascular Endothelium

E-selectin is a type-1 membrane protein on microvascular endothelial cells that helps initiate recruitment of circulating leukocytes to cutaneous, bone and inflamed tissues.  E-selectin expression is constitutive on dermal and bone microvessels and is inducible by pro-inflammatory cytokines, such as IL-1&#945;/  and TNF-&#945;, on microvessels in inflamed tissues.  This lectin receptor mediates weak binding interactions with carbohydrate counter-receptor ligands on circulating leukocytes, which results in a characteristic  rolling  behavior.  Because these interactions precede more stable adhesive events and diapedesis activity, characterization of leukocyte rolling activity and identification of leukocyte E-selectin ligands have been major goals in studies of leukocyte trafficking and inflammation and in the development of anti-inflammatory therapeutics (1-5).  The intent of this report is to provide a visual, comprehensive description of the most widely-used technology for studying E-selectin   E-selectin ligand interactions under physiologic blood flow conditions.  Our laboratory in conjunction with the Harvard Skin Disease Research Center uses a state-of-the-art parallel-plate flow chamber apparatus accompanied by digital visualization and new recording software, NIS-Elements.  This technology allows us to analyze adhesion events in real time for onscreen visualization as well as record rolling activity in a video format.  Cell adhesion parameters, such as rolling frequency, shear resistance and binding/tethering efficiency, are calculated with NIS-Elements software, exported to an Excel spreadsheet and subjected to statistical analysis.  In the demonstration presented here, we employed the parallel-plate flow chamber to investigate E-selectin-dependent leukocyte rolling activity on live human bone marrow endothelial cells (hBMEC).  Human hematopoietic progenitor KG1a cells, which express a high level of E-selectin ligand, were used as our leukocyte model, while an immortalized hBMEC cell line, HBMEC-60 cells, was used as our endothelial cell model (6).  To induce and simulate native E-selectin expression in the flow chamber, HBMEC-60 cells were first activated with IL-1 .  Our video presentation showed that parallel-plate flow analysis is a suitable method for studying physiologic E-selectin-mediated leukocyte rolling activities and that functional characterization of leukocyte E-selectin ligand(s) in the flow chamber can be ascertained by implementing protease or glycosidase digestions.

This report provides a visual depiction of parallel-plate flow chamber analysis for studying leukocyte endothelial interactions under physiologic shear stress. This method is particularly useful for investigating the role of endothelial (E)-selectin and leukocyte E-selectin ligands that trigger leukocyte rolling on endothelial cell surfaces.

E-selectin is a type-1 membrane protein on microvascular endothelial cells that helps initiate recruitment of circulating leukocytes to cutaneous, bone ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Mapping the After-effects of Theta Burst Stimulation on the Human Auditory Cortex with Functional Imaging

Auditory cortex pertains to the processing of sound, which is at the basis of speech or music-related processing1. However, despite considerable recent progress, the functional properties and lateralization of the human auditory cortex are far from being fully understood. Transcranial Magnetic Stimulation (TMS) is a non-invasive technique that can transiently or lastingly modulate cortical excitability via the application of localized magnetic field pulses, and represents a unique method of exploring plasticity and connectivity. It has only recently begun to be applied to understand auditory cortical function 2. 
An important issue in using TMS is that the physiological consequences of the stimulation are difficult to establish. Although many TMS studies make the implicit assumption that the area targeted by the coil is the area affected, this need not be the case, particularly for complex cognitive functions which depend on interactions across many brain regions 3. One solution to this problem is to combine TMS with functional Magnetic resonance imaging (fMRI). The idea here is that fMRI will provide an index of changes in brain activity associated with TMS. Thus, fMRI would give an independent means of assessing which areas are affected by TMS and how they are modulated 4. In addition, fMRI allows the assessment of functional connectivity, which represents a measure of the temporal coupling between distant regions. It can thus be useful not only to measure the net activity modulation induced by TMS in given locations, but also the degree to which the network properties are affected by TMS, via any observed changes in functional connectivity.
Different approaches exist to combine TMS and functional imaging according to the temporal order of the methods. Functional MRI can be applied before, during, after, or both before and after TMS. Recently, some studies interleaved TMS and fMRI in order to provide online mapping of the functional changes induced by TMS 5-7. However, this online combination has many technical problems, including the static artifacts resulting from the presence of the TMS coil in the scanner room, or the effects of TMS pulses on the process of MR image formation. But more importantly, the loud acoustic noise induced by TMS (increased compared with standard use because of the resonance of the scanner bore) and the increased TMS coil vibrations (caused by the strong mechanical forces due to the static magnetic field of the MR scanner) constitute a crucial problem when studying auditory processing. 
This is one reason why fMRI was carried out before and after TMS in the present study. Similar approaches have been used to target the motor cortex 8,9, premotor cortex 10, primary somatosensory cortex 11,12 and language-related areas 13, but so far no combined TMS-fMRI study has investigated the auditory cortex. The purpose of this article is to provide details concerning the protocol and considerations necessary to successfully combine these two neuroscientific tools to investigate auditory processing. 
Previously we showed that repetitive TMS (rTMS) at high and low frequencies (resp. 10 Hz and 1 Hz) applied over the auditory cortex modulated response time (RT) in a melody discrimination task 2. We also showed that RT modulation was correlated with functional connectivity in the auditory network assessed using fMRI: the higher the functional connectivity between left and right auditory cortices during task performance, the higher the facilitatory effect (i.e. decreased RT) observed with rTMS. However those findings were mainly correlational, as fMRI was performed before rTMS. Here, fMRI was carried out before and immediately after TMS to provide direct measures of the functional organization of the auditory cortex, and more specifically of the plastic reorganization of the auditory neural network occurring after the neural intervention provided by TMS. 
Combined fMRI and TMS applied over the auditory cortex should enable a better understanding of brain mechanisms of auditory processing, providing physiological information about functional effects of TMS. This knowledge could be useful for many cognitive neuroscience applications, as well as for optimizing therapeutic applications of TMS, particularly in auditory-related disorders.

Auditory processing is the basis of speech and music-related processing. Transcranial Magnetic Stimulation (TMS) has been used successfully to study cognitive, sensory and motor systems but has rarely been applied to audition. Here we investigated TMS combined with functional Magnetic Resonance Imaging to understand the functional organization of auditory cortex.

Auditory cortex pertains to the processing of sound, which is at the basis of speech or music-related processing1. However, despite considerable recent ...

Isolation and Culture of Neonatal Mouse Cardiomyocytes

Cultured neonatal cardiomyocytes have long been used to study myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow for easy investigation and manipulation of biochemical pathways, and their effect on the biomechanical properties of spontaneously beating cardiomyocytes.	
The following 2-day protocol describes the isolation and culture of neonatal mouse cardiomyocytes. We show how to easily dissect hearts from neonates, dissociate the cardiac tissue and enrich cardiomyocytes from the cardiac cell-population. We discuss the usage of different enzyme mixes for cell-dissociation, and their effects on cell-viability. The isolated cardiomyocytes can be subsequently used for a variety of morphological, electrophysiological, biochemical, cell-biological or biomechanical assays. We optimized the protocol for robustness and reproducibility, by using only commercially available solutions and enzyme mixes that show little lot-to-lot variability. We also address common problems associated with the isolation and culture of cardiomyocytes, and offer a variety of options for the optimization of isolation and culture conditions.

Primary mouse cardiomyocyte cultures are one of the pivotal tools for the investigation of myofibrillar organization and function. The following protocol describes the isolation and culture of primary cardiomyocytes from neonatal mouse hearts. The resulting cardiomyocyte cultures may be subsequently used for a variety of biomechanical, biochemical and cell-biological assays.

Cultured neonatal cardiomyocytes have long been used to study  myofibrillogenesis and myofibrillar functions. Cultured cardiomyocytes allow  for easy ...

Isolation, Culture, and Functional Characterization of Adult Mouse Cardiomyoctyes

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of heart disease. In particular, the ability to study ion homeostasis, ion channel function, cellular excitability and excitation-contraction coupling and their alterations in diseased conditions and by disease-causing mutations have led to significant insights into cardiac diseases. Furthermore, the lack of an adequate immortalized cell line to mimic adult CMs, and the limitations of neonatal CMs (which lack many of the structural and functional biomechanics characteristic of adult CMs) in culture have hampered our understanding of the complex interplay between signaling pathways, ion channels and contractile properties in the adult heart strengthening the importance of studying adult isolated cardiomyocytes. Here, we present methods for the isolation, culture, manipulation of gene expression by adenoviral-expressed proteins, and subsequent functional analysis of cardiomyocytes from the adult mouse. The use of these techniques will help to develop mechanistic insight into signaling pathways that regulate cellular excitability, Ca2+ dynamics and contractility and provide a much more physiologically relevant characterization of cardiovascular disease.

Here we describe the isolation of adult mouse cardiomyoctyes using a Langendorff perfusion system. The resulting cells are Ca2+-tolerant, electrically quiescent and can be cultured and transfected with adeno- or lentiviruses to manipulate gene expression. Their functionality can also be analyzed using the MMSYS system and patch clamp techniques.

The use of primary cardiomyocytes (CMs) in culture has provided a powerful complement to murine models of heart disease in advancing our understanding of ...

Isolation and Physiological Analysis of Mouse Cardiomyocytes

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force used to pump blood. Individual cardiomyocytes were first isolated over 40 years ago in order to better study the physiology and structure of heart muscle. Techniques have rapidly improved to include enzymatic digestion via coronary perfusion. More recently, analyzing the contractility and calcium flux of isolated myocytes has provided a vital tool in the cellular and sub-cellular analysis of heart failure. Echocardiography and EKGs provide information about the heart at an organ level only. Cardiomyocyte cell culture systems exist, but cells lack physiologically essential structures such as organized sarcomeres and t-tubules required for myocyte function within the heart. In the protocol presented here, cardiomyocytes are isolated via Langendorff perfusion. The heart is removed from the mouse, mounted via the aorta to a cannula, perfused with digestion enzymes, and cells are introduced to increasing calcium concentrations. Edge and sarcomere detection software is used to analyze contractility, and a calcium binding fluorescent dye is used to visualize calcium transients of electrically paced cardiomyocytes; increasing understanding of the role cellular changes play in heart dysfunction. Traditionally used to test drug effects on cardiomyocytes, we employ this system to compare myocytes from WT mice and mice with a mutation that causes dilated cardiomyopathy. This protocol is unique in its comparison of live cells from mice with known heart function and known genetics. Many experimental conditions are reliably compared, including genetic or environmental manipulation, infection, drug treatment, and more. Beyond physiologic data, isolated cardiomyocytes are easily fixed and stained for cytoskeletal elements. Isolating cardiomyocytes via perfusion is an extremely versatile method, useful in studying cellular changes that accompany or lead to heart failure in a variety of experimental conditions.

Individual cardiomyocytes from wild type and mutant mice can be isolated from the heart in order to study their contractility and calcium transients. This allows characterization of the contribution of cellular dysfunction to heart dysfunction from any cause.

Cardiomyocytes, the workhorse cell of the heart, contain exquisitely organized cytoskeletal and contractile elements that generate the contractile force ...

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

A Rapid, Scalable Method for the Isolation, Functional Study, and Analysis of Cell-derived Extracellular Matrix

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and pathological processes. However, the isolation of ECM, from tissues or cell culture, is complicated by the insoluble and cross-linked nature of the assembled ECM and by the potential contamination of ECM extracts with cell surface and intracellular proteins. Here, we describe a method for use with cultured cells that is rapid and reliably removes cells to isolate a cell-derived ECM for downstream experimentation.
Through use of this method, the isolated ECM and its components can be visualized by in situ immunofluorescence microscopy. The dynamics of specific ECM proteins can be tracked by tracing the deposition of a tagged protein using fluorescence microscopy, both before and after the removal of cells. Alternatively, the isolated ECM can be extracted for biochemical analysis, such as sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and immunoblotting. At larger scales, a full proteomics analysis of the isolated ECM by mass spectrometry can be conducted. By conducting ECM isolation under sterile conditions, sterile ECM layers can be obtained for functional or phenotypic studies with any cell of interest. The method can be applied to any adherent cell type, is relatively easy to perform, and can be linked to a wide repertoire of experimental designs.

The extracellular matrix plays a major role in defining the microenvironment of cells and in modulating cell behavior and phenotype. We describe a rapid method for the isolation of cell-derived extracellular matrix, which can be adapted to different scales for microscopic, biochemical, proteomic, or functional studies.

The extracellular matrix (ECM) is recognized as a diverse, dynamic, and complex environment that is involved in multiple cell-physiological and ...

Isolation of Intact, Whole Mouse Mammary Glands for Analysis of Extracellular Matrix Expression and Gland Morphology

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal architecture. Here, a small pocket below the skin was created using Mayo scissors, allowing separation of the glands within the subcutaneous tissue from the underlying peritoneum. Visualization of the glands was aided by the use of 3.5x-R surgical micro loupes. The pelt was inverted and pinned back allowing identification of the intact mammary fat pads. Each of the #4 abdominal glands was bluntly dissected by sliding the scalpel blade laterally between the subcutaneous layer and the glands. Immediately post-harvest, glands were placed in 10% neutral buffered formalin for subsequent tissue processing. Excision of the entire gland is advantageous because it primarily eliminates the risk of excluding important tissue-wide interactions between ductal epithelial cells and other microenvironmental cellular populations that could be missed in a partial biopsy. One drawback of the methodology is the use of serial sections from fixed tissues which limits analyses of ductal morphogenesis and protein expression to discrete locations within the gland. As such, changes in ductal architecture and protein expression in 3 dimensions (3D) is not readily obtainable. Overall, the technique is applicable to studies requiring whole intact murine mammary glands for downstream investigations such as developmental ductal morphogenesis or breast cancer.

Here, we present a protocol for the isolation of whole, intact mouse mammary glands to investigate extracellular matrix (ECM) expression and ductal morphology. Mouse #4 abdominal glands were extracted from 8-10 week old female nulliparous mice, fixed in neutral buffered formalin, sectioned and stained using immunohistochemistry for ECM proteins.

The goal of this procedure was to harvest the #4 abdominal mammary glands from female nulliparous mice in order to assess ECM expression and ductal ...