This work was supported in part by the Center for Biomedical Engineering and Technology (BioMET); NIH (1U01HL116321) and (1R01HL142290) and the American Heart Association 10SDG4030042 (GZ), 19POST34450156 (HCJ).

All animal care was in accordance with the guidelines of the University of Maryland Baltimore and the Institutional Animal Care and Use Committee approved protocols.
1. Preparation of the solutions
NOTE: Prepare solutions in advance. Two types of basic solutions are used in the experiments: (1) physiological saline solutions (PSS) for bath superfusate and (2) Tyrode&#8217;s solutions for lumen perfusate. Continuous bubbling with CO2 is needed to maintain the pH of PSS. HEPES-buffered Tyrode&#8217;s solution is used in the lumen instead of PSS to avoid bubbles going into the vessels, since bubbles would damage the endothelial cells7 and occlude the flow.
<ol>
	<li>PSS solution: See the formula and composition of PSS solution in the Table 1. Make 10 L of Ca2+-free and glucose-free PSS stock solution and store at room temperature. 1 h before any procedure, take out 1 L of the stock solution and bubble with 5% CO2 (74% N2 and 21% O2) to keep the pH of solution at ~7.4. Add 1.8 g glucose and 1.8 mL CaCl2 (1 M stock)8. 
	NOTE: Add Ca2+ only after bubbling when pH is ~7.4, otherwise, Ca2+ will be precipitated.</li>
	<li>Tyrode&#8217;s solution: See the formula and composition of the Tyrode&#8217;s solution in Table 1. Filter the solution (0.22 &#956;m) and stored at 4 &#176;C for future use9.</li>
	<li>Tyrode&#8217;s solution with BDM (2,3-Butanedione monoxime): Add BDM as a reversible myofilament inhibitor to prevent the contraction of the myocytes10. Weigh 600 mg BDM and add to 200 mL of Tyrode&#8217;s solution. Stir the solution with BDM until it is completely dissolved. No further filtration is required.</li>
	<li>Store the Tyrode&#8217;s solution containing BDM at 4 &#176;C for future use.</li>
</ol>
2. Chamber preparation
<ol>
	<li>Coat both dissecting chamber and experimental chamber in advance with polydimethylsiloxane (PDMS) following the instructions provided by the manufacturer.</li>
	<li>Fill the dissecting chamber with the Tyrode&#8217;s solution containing BDM.</li>
	<li>Turn on the chiller 1 h before the procedure. Set the temperature at 4 &#176;C.</li>
</ol>
3. Cannula preparation
<ol>
	<li>Turn on the micropipette puller. Select the settings as per the manufacturer&#8217;s instructions.</li>
	<li>Take a clean borosilicate glass tube (the outer and inner diameter of the glass tubes used was 1.2 mm and 0.69 mm respectively).</li>
	<li>Insert a glass tube into the grooved clamps of a Sutter pipette puller with a U-shaped platinum heating filament.</li>
	<li>Activate the puller to produce two cannulae, each with a long thin tip.</li>
	<li>Cut the tip of each cannula using a fine pair of scissors under a dissecting microscope to make the final diameter of the tip around 100 to 150 &#181;m.</li>
	<li>Fire polish the tip of the cut cannula to slightly round the sharp edges.</li>
	<li>Bend the cannula along the shaft using a platinum wire (heated with fine control) positioned on the side of the shaft approximately 2 mm from tip. The angle of the bend in the cannula should be around 45&#176;.</li>
	<li>Insert the open end of the cannula into the holder of the pressure myograph chamber and tighten the fitting. Connect a 5 mL syringe filled with Tyrode&#39;s solution to the other side of the fitting. Connect the cannula to a micromanipulator mounted on the chamber (see Figure 1 and Figure 2).</li>
	<li>Fill the cannula with Tyrode&#8217;s solution using the syringe (5 mL), flushing it, the tubing and connector of air bubbles.</li>
	<li>Prior to the cannulation procedure, measure the perfusion pressure inside the cannula over a given range of flow (50 to 300 &#181;L/min, Figure 3). Do so only when a new cannula is used. 
	NOTE: The perfusion pressure inside the cannula is proportional to the flow and is linear fitted (Figure 3).</li>
</ol>
4. Extraction of mouse heart
<ol>
	<li>Put a C57BL/6 mouse (either sex, 8-16 weeks old) in the anesthesia box that is connected to the tanks of isoflurane and oxygen.</li>
	<li>Turn on oxygen tank and start the flow of isoflurane. Wait ~5 min until the mouse is fully anesthetized.</li>
	<li>After anesthesia is established, take the mouse out of the anesthesia box and inject heparin IP (into the peritoneal cavity, 360 units, 0.5 mL/mouse) to avoid blood clot formation in the vasculature. Then put the mouse back into the anesthesia box.</li>
	<li>10 min after the heparin has been injected, move the mouse to the heated bed in a supine position and stabilize its paws using labeling tape. Keep the mouse under full anesthesia with isoflurane using a nose cone.</li>
	<li>Lift the abdominal skin of the mouse above the diaphragm with forceps and use surgical scissors to cut and expose the diaphragm.</li>
	<li>Cut the diaphragm and the sternum. Open the chest.</li>
	<li>Dissect the heart out of the thoracic cavity, cutting as close as possible to the dorsal thoracic wall.</li>
	<li>Place the heart into ice-cold Tyrode&#8217;s solution containing 30 mM BDM.</li>
	<li>Clean the heart to remove the connective tissues such as lung.</li>
	<li>Transfer the heart into the pre-chilled dissecting chamber filled with Tyrode&#8217;s solution containing 30 mM BDM.</li>
</ol>
5. Preparation and cannulation of septal artery
<ol>
	<li>Turn on the servo pump. Set the servo pump to &#8220;flow&#8221; mode. Let it run at high speed until the tubing is filled with the Tyrode&#39;s solution that will be used to perfuse the vascular lumen. Make sure there are no bubbles in the tubing. Then lower the speed.</li>
	<li>Pin the heart onto the PDMS, avoiding any damage to the middle area of the heart.</li>
	<li>Remove both the right and left atria. Cut open the right ventricle and remove the right ventricular free wall using a dissecting microscope.</li>
	<li>Expose and identify the septal artery. Place a nylon thread (30 &#181;m diameter) under the septal artery and tie a loose knot for future use. 
	NOTE: The septal artery can be easily identified using a dissecting microscope. The septal artery is a major artery on the septum originating from the right coronary artery in most cases.</li>
	<li>Remove the left ventricular free wall using fine scissors.</li>
	<li>Transfer the papillary muscle preparation into the experimental chamber in which the cannula was positioned. The chamber is filled with Tyrode&#8217;s solution containing BDM. The bottom of this chamber is coated with a thin (~2 mm) layer of PDMS.</li>
	<li>Adjust the position of the cannula (using the micromanipulator) to enable cannulation of the septal artery. Tighten the knot.</li>
	<li>Secure the papillary muscle using small pins to the chamber floor so that the microscope objective has a clear view of the vasculature. Pay attention to the angle and the position of the cannula and ensure that the tip of the cannula is paralleled to the arterial wall.</li>
	<li>Test cannulation by gradually expelling the solution from the syringe through the cannula. If all elements function properly, residual blood in the papillary muscle will exit the tissue at the beginning of this process and only Tyrode&#39;s solution will be seen later.</li>
</ol>
6. Stabilization of the preparation
<ol>
	<li>Turn on the peristaltic pump that will provide the superfusion solution. Then constantly superfuse the preparation in the bath with pre-gassed PSS at the rate of 3-4 mL/min.</li>
	<li>Turn on the temperature controller for the superfusion solution. Adjust the temperature of the bath superfusate to ~35-37 &#176;C.</li>
	<li>Connect the cannula to the servo pump. Read the pressure displayed on the pressure servo controller.</li>
	<li>Monitor the flow and pressure. The flow (&#181;L/min) and arterial perfusion pressure (mm Hg) is digitized using digitizer and recorded using an associated software (Figure 2).</li>
	<li>Adjust the flow to set the pressure ~10 mmHg and let it run for ~10 min.</li>
	<li>Increase the flow of the luminal solution to make the pressure of artery ~60 mmHg. Obtain the final perfusion pressure of the arteriole itself by subtracting the pressure drop of the cannula (Figure 3). 
	NOTE: If the &#8220;pressure&#8221; mode on pressure servo controller is selected, the luminal pressure is set ~60 mmHg for these experiments. Then monitor the change of the flow.</li>
	<li>Let the sample stabilize for ~30-60 min by monitoring the flow and/or pressure levels. A gradual increase of arterial pressure is normally observed at the beginning of the perfusion after cannulation. A new steady state will get establish by itself in about 30 min (Figure 4).</li>
	<li>Initiate an experiment after the perfusion pressure is stabilized (at constant flow).</li>
</ol>
7. Loading the preparation with fluorescently tagged wheat-germ agglutinin (WGA)
<ol>
	<li>Prepare Tyrode&#39;s solution containing Alexa-Fluor-488 conjugated WGA by adding 100 &#956;g of conjugated WGA into 5 mL of Tyrode&#39;s solution.</li>
	<li>Carefully switch the perfusate from normal Tyrode&#39;s solution to one containing fluorescently tagged WGA. It is important to carefully switch the perfusate so that no bubbles are introduced.</li>
	<li>After ~30 min of WGA perfusion or when WGA solution is about to run out, change the perfusate back to normal Tyrode&#8217;s solution.</li>
</ol>
8. Confocal imaging of arterioles and capillaries
<ol>
	<li>Turn on the Nipkow disk confocal microscope system.</li>
	<li>Use the microscope to locate the septal artery using a low power objective (4x) in transmitted light mode. 
	NOTE: Septal artery can be located by following the position of the cannula.</li>
	<li>Start confocal imaging with the spinning disk confocal and choose 488 nm excitation laser, 40x objective magnification and adjust the laser intensity and the sampling rate (10 - 500 ms per image).</li>
	<li>Set up top and bottom imaging positions for the confocal microscope to define the imaging range and enter the parameters for the z-stack imaging.</li>
	<li>Set the step size as recommended for the system that is being used or set the step size as desired.</li>
	<li>Choose the z-stack setting and/or time series settings.</li>
	<li>Select or set a new folder to store the new image.</li>
	<li>Click &#8220;Run&#8221; to start recording imaging for future analysis (Figure 5).</li>
</ol>
9. Example vasodilator experiment: pinacidil-induced vasodilation (Video 1).
<ol>
	<li>Prepare pinacidil stock solution (100 mM in DMSO) and stored at 4 &#176;C.</li>
	<li>Prepare 10 mL of Tyrode&#8217;s solution. Add 10 &#181;L of pinacidil stock solution to make the final concentration of pinacidil 100 &#181;M.</li>
	<li>Image the papillary muscle at low magnification (4x objective) to include the septal artery and the branch arterioles.</li>
	<li>Switch luminal solution to the pinacidil containing solution.</li>
</ol>
10. Example blood flow control experiments: vasoconstrictor-induced arterial perfusion pressure increase at constant flow (Figure 6)
<ol>
	<li>Prepare endothelin-1 (ET-1) stock solution (10 &#181;M) and stored at -20 &#176;C.</li>
	<li>Prepare 10 mL of Tyrode&#8217;s solution. Add 10 &#181;L of ET-1 stock solution (10 &#181;M) to make the final concentration of ET-1 10 nM.</li>
	<li>Record the luminal pressure with constant flow and image the papillary muscle.</li>
	<li>Switch luminal solution to the ET-1 containing solution.</li>
</ol>
11. Example images of capillary with pericytes (Figure 7)
<ol>
	<li>Sacrifice an NG2DsRedBAC transgenic mouse as described in section 4 of this protocol.</li>
	<li>Extract the heart, cannulate the septal artery and load the sample with Alexa-Fluor-488 conjugated WGA following protocols 5-7.</li>
	<li>Image the papillary muscle at 488 nm and 560 nm excitation, and 510-525 nm, 575-625 nm emission using confocal. The arteriole pressure will be around 40 mmHg.</li>
</ol>

<ol>
	<li>Zhao, G., Joca, H. C., Nelson, M. T., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=ATP-+and+voltage-dependent+electro-metabolic+signaling+regulates+blood+flow+in+heart.">ATP- and voltage-dependent electro-metabolic signaling regulates blood flow in heart.</a> Proceedings of the National Academy of Sciences of the United States of America. 117, 7461-7470 (2020).</li><li>Schouten, V. J., Allaart, C. P., Westerhof, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+perfusion+pressure+on+force+of+contraction+in+thin+papillary+muscles+and+trabeculae+from+rat+heart.">Effect of perfusion pressure on force of contraction in thin papillary muscles and trabeculae from rat heart.</a> Journal of Physiology. 451, 585-604 (1992).</li><li>Tillmanns, H., 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=Microcirculation+in+the+ventricle+of+the+dog+and+turtle.">Microcirculation in the ventricle of the dog and turtle.</a> Circulation Research. 34, 561-569 (1974).</li><li>Martini, J., Honig, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+measurement+of+intercapillary+distance+in+beating+rat+heart+in+situ+under+various+conditions+of+O2+supply.">Direct measurement of intercapillary distance in beating rat heart in situ under various conditions of O2 supply.</a> Microvascular Research. 1, 244-256 (1969).</li><li>Nellis, S. H., Liedtke, A. J., Whitesell, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Small+coronary+vessel+pressure+and+diameter+in+an+intact+beating+rabbit+heart+using+fixed-position+and+free-motion+techniques.">Small coronary vessel pressure and diameter in an intact beating rabbit heart using fixed-position and free-motion techniques.</a> Circulation Research. 49, 342-353 (1981).</li><li>Marcus, M. 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=Understanding+the+coronary+circulation+through+studies+at+the+microvascular+level.">Understanding the coronary circulation through studies at the microvascular level.</a> Circulation. 82, 1-7 (1990).</li><li>Ralevic, V., Kristek, F., Hudlicka, O., Burnstock, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+protocol+for+removal+of+the+endothelium+from+the+perfused+rat+hind-limb+preparation.">A new protocol for removal of the endothelium from the perfused rat hind-limb preparation.</a> Circulation Research. 64, 1190-1196 (1989).</li><li>Zhao, G., Adebiyi, A., Blaskova, E., Xi, Q., Jaggar, J. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Type+1+inositol+1,4,5-trisphosphate+receptors+mediate+UTP-induced+cation+currents,+Ca2++signals,+and+vasoconstriction+in+cerebral+arteries.">Type 1 inositol 1,4,5-trisphosphate receptors mediate UTP-induced cation currents, Ca2+ signals, and vasoconstriction in cerebral arteries.</a> Amercian Journal of Physiology-Cell Physiology. 295, 1376-1384 (2008).</li><li>Zhao, G., Li, T., Brochet, D. X., Rosenberg, P. B., Lederer, W. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=STIM1+enhances+SR+Ca2++content+through+binding+phospholamban+in+rat+ventricular+myocytes.">STIM1 enhances SR Ca2+ content through binding phospholamban in rat ventricular myocytes.</a> Proceedings of the National Academy of Sciences of the United States of America. 112, 4792-4801 (2015).</li><li>Stowe, D. F., Boban, M., Graf, B. M., Kampine, J. P., Bosnjak, Z. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Contraction+uncoupling+with+butanedione+monoxime+versus+low+calcium+or+high+potassium+solutions+on+flow+and+contractile+function+of+isolated+hearts+after+prolonged+hypothermic+perfusion.">Contraction uncoupling with butanedione monoxime versus low calcium or high potassium solutions on flow and contractile function of isolated hearts after prolonged hypothermic perfusion.</a> Circulation. 89, 2412-2420 (1994).</li><li>Lawton, P. F., 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=a+Low-Cost+and+Open+Source+Pressure+Myograph+System+for+Vascular+Physiology.">a Low-Cost and Open Source Pressure Myograph System for Vascular Physiology.</a> Frontiers in Physiology. 10, 99 (2019).</li><li>Kim, K. J., Filosa, J. 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In the present work, we have introduced a remarkably simple yet highly practical ex vivo method to study the coronary microcirculation in heart under physiological conditions. This method was modified from mechanical investigations using rats2. The challenging addition was the imaging technology with high speed and high optical resolution. We, therefore, were able to take advantage of the advanced optical imaging systems that are now commercially available. By careful dissection and placement of t...

Appropriate coronary pressure-flow regulation assures sufficient blood supply to the heart to meet its metabolic demands1. However, it has only recently become clear how coronary pressure-flow is dynamically regulated in heart, despite extensive studies that have been performed in vivo and in vitro for the past decades. One of the reasons is the difficulty in establishing a physiological working model for such studies due to the constant beating of the heart. Regardless, a variety of methods have been established for the observation of the coronary micro-vessels in living tissues or animals, but none of these methods were able to achieve constant/stable focus and the measurements of the pressure, flow and microvascular diameter at the same time2,3. The direct visualization of coronary arterial micro-vessels in beating heart was introduced decades ago4,3, but the diameter measurements in small vessels was challenging and the specific functions of the many specialized cell types associated with the microcirculation was equally vexing. Even the stroboscopic method and the floating objective system could not provide the above information simultaneously5. Nevertheless, a significant amount of valuable information has been obtained using the aforementioned technologies, which have helped us understanding more about the regulation of coronary blood flow6. The method we are describing in this paper will help one investigate and understand in detail how components of coronary arteries, the arterioles and the microvasculature respond differently to stimulations and metabolic demands.
The working model we established to pursue these studies was built on the previous work of Westerhof et al.2. Following cannulation of the septal artery of the mouse heart, physiological saline solution was used to perfuse that artery to keep the myocytes and other components of the heart tissue nourished. The arterial perfusion pressure, the flow and the vascular diameter was monitored among other physiological functions using appropriate fluorescent indicators. This method enables us to visualize the coronary microvascular bed under physiological pressure in living tissue and study the cellular mechanisms underlying microcirculation regulation for the first time.

<table>
	<tbody>
		<tr>
			<td>1 M CaCl2 solution</td>
			<td>MilliporeSigma, USA</td>
			<td>21115</td>
			<td></td>
		</tr>
		<tr>
			<td>1 M MgCl2 solution</td>
			<td>MilliporeSigma, USA</td>
			<td>M1028</td>
			<td></td>
		</tr>
		<tr>
			<td>AxoScope software</td>
			<td>Molecular Devices, San Jose, CA, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Chiller/water incubator</td>
			<td>FisherScientific, USA</td>
			<td>Isotemp 3016S</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal</td>
			<td>Nikon Instruments, USA</td>
			<td>A1R</td>
			<td></td>
		</tr>
		<tr>
			<td>Custom glass tubing</td>
			<td>Drummond Scientific Company</td>
			<td>9-000-3301</td>
			<td></td>
		</tr>
		<tr>
			<td>Digidata 1322A</td>
			<td>Molecular Devices, San Jose, CA, USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td>Olympus, Japan</td>
			<td>SZX12</td>
			<td></td>
		</tr>
		<tr>
			<td>Endothelin-1</td>
			<td>MilliporeSigma, USA</td>
			<td>E7764</td>
			<td></td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>Fine Scientific Tools</td>
			<td>11295-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin Sodium Salt</td>
			<td>Sigma-Aldrich, USA</td>
			<td>H3393</td>
			<td></td>
		</tr>
		<tr>
			<td>Inline solution Heater</td>
			<td>Warner Istruments, Hamden, CT, USA</td>
			<td>SH-27B</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>VETone, Idaho, USA</td>
			<td>502017</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter Instruments, Novato, CA, USA</td>
			<td>P-97</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette/cannula holder</td>
			<td>Warner Istruments, Hamden, CT, USA</td>
			<td>64-0981</td>
			<td></td>
		</tr>
		<tr>
			<td>NG2DsRedBAC transgenic mouse</td>
			<td>The Jackson Laboratory</td>
			<td>#008241</td>
			<td></td>
		</tr>
		<tr>
			<td>Nylon thread for tying blood vessels</td>
			<td>Living Systems Instrumentation, Burlington, Vt, USA</td>
			<td>THR-G</td>
			<td></td>
		</tr>
		<tr>
			<td>PDMS (polydimethylsiloxane)</td>
			<td>SYLGARD, Germantown, WI, USA</td>
			<td>184 SIL ELAST KIT</td>
			<td></td>
		</tr>
		<tr>
			<td>Peristaltic pump</td>
			<td>Gilson, Middleton, WI, USA</td>
			<td>minipuls 3</td>
			<td></td>
		</tr>
		<tr>
			<td>Pressure Servo Controller</td>
			<td>Living Systems Instrumentation, Burlington, Vt, USA</td>
			<td>PS-200-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Fine Scientific Tools, Foster City, CA, USA</td>
			<td>15000-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Servo Pump</td>
			<td>Living Systems Instrumentation, Burlington, Vt, USA</td>
			<td>PS-200-P</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature controller</td>
			<td>Warner Instruments, Hamden, CT, USA</td>
			<td>TC-324B</td>
			<td></td>
		</tr>
		<tr>
			<td>Wheat Germ Agglutinin, Alexa Fluor 488 Conjugate</td>
			<td>ThermoFisher Scientific, Waltham, MA USA</td>
			<td>W11261</td>
			<td></td>
		</tr>
	</tbody>
</table>

dynamic measurement and imaging of capillaries, arterioles, and pericytes in mouse heart

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion pressure. However, it is difficult to monitor the dynamic changes of the coronary arterioles and the capillaries in the whole heart, primarily due to its motion and non-stop beating. Here we describe a method that enables monitoring of arterial perfusion rate, pressure and the diameter changes of the arterioles and capillaries in mouse right ventricular papillary muscles. The mouse septal artery is cannulated and perfused at a constant flow or pressure with the other dynamically measured. After perfusion with a fluorescently labeled lectin (e.g., Alexa Fluor-488 or -633 labeled Wheat-Germ Agglutinin, WGA), the arterioles and capillaries (and other vessels) in right ventricle papillary muscle and septum could be readily imaged. The vessel-diameter changes could then be measured in the presence or absence of heart contractions. When genetically encoded fluorescent proteins were expressed, specific features could be monitored. For examples, pericytes were visualized in mouse hearts that expressed NG2-DsRed. This method has provided a useful platform to study the physiological functions of capillary pericytes in heart. It is also suitable for studying the effect of reagents on the blood flow in heart by measuring the vascular/capillary diameter and the arterial luminal pressure simultaneously. This preparation, combined with a state-of-the-art optic imaging system, allows one to study the blood flow and its control at cellular and molecular level in the heart under near-physiological conditions.

When a fluorescence vascular marker is perfused in vascular lumen (here WGA conjugated with Alexa Fluor-488), it is possible to visualize whole vascular trees as shown in Figure 5 (Left panel) using high-speed confocal microscope. Further magnification enables the imaging of capillary in detail (Figure 5, Right Panel). Since the pressurized system supports a constant monitoring of luminal pressure, this preparation can be used for associate changes in arterial d...

Watch this Scientific Journal Video about Dynamic Measurement and Imaging of Capillaries, Arterioles, and Pericytes in Mouse Heart at JoVE.com

Dynamic Measurement and Imaging of Capillaries, Arterioles, and Pericytes in Mouse Heart

Presented here is a protocol to study the coronary microcirculation in living murine heart tissue by ex vivo monitoring of the arterial perfusion pressure and flow that maintains the pressure, as well as vascular tree components including the capillary beds and pericytes, as the septal artery is cannulated and pressurized.

Coronary arterial tone along with the opening or closing of the capillaries largely determine the blood flow to cardiomyocytes at constant perfusion ...

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5.2K Views.  University of Maryland School of Medicine. Presented here is a protocol to study the coronary microcirculation in living murine heart tissue by ex vivo monitoring of the arterial perfusion pressure and flow that maintains the pressure, as well as vascular tree components including the capillary beds and pericytes, as the septal artery is cannulated and pressurized.

Video: Dynamic Measurement and Imaging of Capillaries, Arterioles, and Pericytes in Mouse Heart

Quantitative Visualization and Detection of Skin Cancer Using Dynamic Thermal Imaging

In 2010 approximately 68,720 melanomas will be diagnosed in the US alone, with around 8,650 resulting in death 1. To date, the only effective treatment for melanoma remains surgical excision, therefore, the key to extended survival is early detection 2,3. Considering the large numbers of patients diagnosed every year and the limitations in accessing specialized care quickly, the development of objective in vivo diagnostic instruments to aid the diagnosis is essential. New techniques to detect skin cancer, especially non-invasive diagnostic tools, are being explored in numerous laboratories. Along with the surgical methods, techniques such as digital photography, dermoscopy, multispectral imaging systems (MelaFind), laser-based systems (confocal scanning laser microscopy, laser doppler perfusion imaging, optical coherence tomography), ultrasound, magnetic resonance imaging, are being tested. Each technique offers unique advantages and disadvantages, many of which pose a compromise between effectiveness and accuracy versus ease of use and cost considerations. Details about these techniques and comparisons are available in the literature 4.
Infrared (IR) imaging was shown to be a useful method to diagnose the signs of certain diseases by measuring the local skin temperature. There is a large body of evidence showing that disease or deviation from normal functioning are accompanied by changes of the temperature of the body, which again affect the temperature of the skin 5,6. Accurate data about the temperature of the human body and skin can provide a wealth of information on the processes responsible for heat generation and thermoregulation, in particular the deviation from normal conditions, often caused by disease. However, IR imaging has not been widely recognized in medicine due to the premature use of the technology 7,8 several decades ago, when temperature measurement accuracy and the spatial resolution were inadequate and sophisticated image processing tools were unavailable. This situation changed dramatically in the late 1990s-2000s. Advances in IR instrumentation, implementation of digital image processing algorithms and dynamic IR imaging, which enables scientists to analyze not only the spatial, but also the temporal thermal behavior of the skin 9, allowed breakthroughs in the field.
In our research, we explore the feasibility of IR imaging, combined with theoretical and experimental studies, as a cost effective, non-invasive, in vivo optical measurement technique for tumor detection, with emphasis on the screening and early detection of melanoma 10-13. In this study, we show data obtained in a patient study in which patients that possess a pigmented lesion with a clinical indication for biopsy are selected for imaging. We compared the difference in thermal responses between healthy and malignant tissue and compared our data with biopsy results. We concluded that the increased metabolic activity of the melanoma lesion can be detected by dynamic infrared imaging.

We demonstrated that malignant pigmented lesions with increased metabolic activity generate quantifiable amounts of heat and the measurement of the transient thermal response of the skin to a cooling excitation allows quantitative identification of melanoma and other skin cancers (vs. non-proliferative nevi) at an early stage of the disease.

In 2010 approximately 68,720 melanomas will be diagnosed in the US alone, with around 8,650 resulting in death 1. To date, the only effective treatment ...

Segmentation and Measurement of Fat Volumes in Murine Obesity Models Using X-ray Computed Tomography

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are associated with obesity, though the precise underlying mechanisms that contribute to the disease are currently being delineated.2,3 Several small animal models of obesity have been developed and are employed in a variety of studies.4 A critical component to these experiments involves the collection of regional and/or total animal fat content data under varied conditions.
Traditional experimental methods available for measuring fat content in small animal models of obesity include invasive (e.g. ex vivo measurement of fat deposits) and non-invasive (e.g. Dual Energy X-ray Absorptiometry (DEXA), or Magnetic Resonance (MR)) protocols, each of which presents relative trade-offs. Current invasive methods for measuring fat content may provide details for organ and region specific fat distribution, but sacrificing the subjects will preclude longitudinal assessments. Conversely, current non-invasive strategies provide limited details for organ and region specific fat distribution, but enable valuable longitudinal assessment. With the advent of dedicated small animal X-ray computed tomography (CT) systems and customized analytical procedures, both organ and region specific analysis of fat distribution and longitudinal profiling may be possible. Recent reports have validated the use of CT for in vivo longitudinal imaging of adiposity in living mice.5,6 Here we provide a modified method that allows for fat/total volume measurement, analysis and visualization utilizing the Carestream Molecular Imaging Albira CT system in conjunction with PMOD and Volview software packages.

Fat content analysis is routinely conducted in studies utilizing murine obesity models. Emerging methods in small animal CT imaging and analysis are providing for longitudinal detail rich fat content analysis. Here we detail step by step procedures for performing small animal CT imaging, analysis, and visualization.

Obesity is associated with increased morbidity and mortality as well as reduced metrics in quality of life.1 Both environmental and genetic factors are ...

Non-invasive Optical Measurement of Cerebral Metabolism and Hemodynamics in Infants

Perinatal brain injury remains a significant cause of infant mortality and morbidity, but there is not yet an effective bedside tool that can accurately screen for brain injury, monitor injury evolution, or assess response to therapy. The energy used by neurons is derived largely from tissue oxidative metabolism, and neural hyperactivity and cell death are reflected by corresponding changes in cerebral oxygen metabolism (CMRO2). Thus, measures of CMRO2 are reflective of neuronal viability and provide critical diagnostic information, making CMRO2 an ideal target for bedside measurement of brain health.
Brain-imaging techniques such as positron emission tomography (PET) and single-photon emission computed tomography (SPECT) yield measures of cerebral glucose and oxygen metabolism, but these techniques require the administration of radionucleotides, so they are used in only the most acute cases.
Continuous-wave near-infrared spectroscopy (CWNIRS) provides non-invasive and non-ionizing radiation measures of hemoglobin oxygen saturation (SO2) as a surrogate for cerebral oxygen consumption. However, SO2 is less than ideal as a surrogate for cerebral oxygen metabolism as it is influenced by both oxygen delivery and consumption. Furthermore, measurements of SO2 are not sensitive enough to detect brain injury hours after the insult 1,2, because oxygen consumption and delivery reach equilibrium after acute transients 3. We investigated the possibility of using more sophisticated NIRS optical methods to quantify cerebral oxygen metabolism at the bedside in healthy and brain-injured newborns. More specifically, we combined the frequency-domain NIRS (FDNIRS) measure of SO2 with the diffuse correlation spectroscopy (DCS) measure of blood flow index (CBFi) to yield an index of CMRO2 (CMRO2i) 4,5.
With the combined FDNIRS/DCS system we are able to quantify cerebral metabolism and hemodynamics. This represents an improvement over CWNIRS for detecting brain health, brain development, and response to therapy in neonates. Moreover, this method adheres to all neonatal intensive care unit (NICU) policies on infection control and institutional policies on laser safety. Future work will seek to integrate the two instruments to reduce acquisition time at the bedside and to implement real-time feedback on data quality to reduce the rate of data rejection.

We combined frequency-domain near-infrared spectroscopy measures of cerebral hemoglobin oxygenation with diffuse correlation spectroscopy measures of cerebral blood flow index to estimate an index of oxygen metabolism. We tested the utility of this measure as a bedside screening tool to evaluate the health and development of the newborn brain.

Perinatal brain injury remains a significant cause of infant mortality  and morbidity, but there is not yet an effective bedside tool that can  accurately ...

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease and injury regional estimates, World Health Organization, 2004). Surgeries that often compromise the quality of life are required to correct heart defects, reminding us of the importance of finding the causes of CHD. Mutant mouse models and live imaging technology have become essential tools to study the etiology of this disease. Although advanced methods allow live imaging of abnormal hearts in embryos, the physiological and hemodynamic states of the latter are often compromised due to surgical and/or lengthy procedures. Noninvasive ultrasound imaging, however, can be used without surgically exposing the embryos, thereby maintaining their physiology. Herein, we use simple M-mode ultrasound to assess heart rates of embryos at E18.5 in utero. The detection of abnormal heart rates is indeed a good indicator of dysfunction of the heart and thus constitutes a first step in the identification of developmental defects that may lead to heart failure.

In utero Measurement of Heart Rate in Mouse by Noninvasive M-mode Echocardiography

Ultra-high frequency ultrasound is a powerful live imaging tool to examine cardiac abnormalities in small animals. Its noninvasiveness allows maintaining the physiologic state of embryos. Herein, we show the use of M-mode ultrasound to measure the heart rates of embryos at E18.5 in utero.

Congenital heart disease (CHD) is the most frequent noninfectious cause of death at birth. The incidence of CHD ranges from 4 to 50/1,000 births (Disease ...

Diagnostic Ultrasound Imaging of Mouse Diaphragm Function

Function analysis of rodent respiratory skeletal muscles, particularly the diaphragm, is commonly performed by isolating muscle strips using invasive surgical procedures. Although this is an effective method of assessing in vitro diaphragm activity, it involves non-survival surgery. The application of non-invasive ultrasound imaging as an in vivo procedure is beneficial since it not only reduces the number of animals sacrificed, but is also suitable for monitoring disease progression in live mice. Thus, our ultrasound imaging method may likely assist in the development of novel therapies that alleviate muscle injury induced by various respiratory diseases. Particularly, in clinical diagnoses of obstructive lung diseases, ultrasound imaging has the potential to be used in conjunction with other standard tests to detect the early onset of diaphragm muscle fatigue. In the current protocol, we describe how to accurately evaluate diaphragm contractility in a mouse model using a diagnostic ultrasound imaging technique.

Diagnostic ultrasound imaging has proven to be effective in diagnosing various respiratory diseases in human and animal subjects. We demonstrate a comprehensive ultrasound protocol utilized by Dr. Zuo's lab to analyze diaphragm kinetics specifically in mouse models. This is also a non-invasive research technique which can provide quantitative information on mouse respiratory muscle function.

Function analysis of rodent respiratory skeletal muscles, particularly the diaphragm, is commonly performed by isolating muscle strips using invasive ...

Dynamic Contrast Enhanced Magnetic Resonance Imaging of an Orthotopic Pancreatic Cancer Mouse Model

Dynamic contrast enhanced magnetic resonance imaging (DCE-MRI) has been limitedly used for orthotopic pancreatic tumor xenografts due to severe respiratory motion artifact in the abdominal area. Orthotopic tumor models offer advantages over subcutaneous ones, because those can reflect the primary tumor microenvironment affecting blood supply, neovascularization, and tumor cell invasion. We have recently established a protocol of DCE-MRI of orthotopic pancreatic tumor xenografts in mouse models by securing tumors with an orthogonally bent plastic board to prevent motion transfer from the chest region during imaging. The pressure by this board was localized on the abdominal area, and has not resulted in respiratory difficulty of the animals. This article demonstrates the detailed procedure of orthotopic pancreatic tumor modeling using small animals and DCE-MRI of the tumor xenografts. Quantification method of pharmacokinetic parameters in DCE-MRI is also introduced. The procedure described in this article will assist investigators to apply DCE-MRI for orthotopic gastrointestinal cancer mouse models.

The goal of this protocol is to apply dynamic contrast enhanced magnetic resonance imaging (DCE-MRI) for orthotopic pancreatic tumor xenografts in mice. DCE-MRI is a non-invasive method to analyze microvasculature in a target tissue, and useful to assess vascular response in a tumor following a novel therapy.

Dynamic contrast enhanced magnetic resonance imaging (DCE-MRI) has been limitedly used for orthotopic pancreatic tumor xenografts due to severe ...

Preparation Steps for Measurement of Reactivity in Mouse Retinal Arterioles Ex Vivo

Vascular insufficiency and alterations in normal retinal perfusion are among the major factors for the pathogenesis of various sight-threatening ocular diseases, such as diabetic retinopathy, hypertensive retinopathy, and possibly glaucoma. Therefore, retinal microvascular preparations are pivotal tools for physiological and pharmacological studies to delineate the underlying pathophysiological mechanisms and to design therapies for the diseases. Despite the wide use of mouse models in ophthalmic research, studies on retinal vascular reactivity are scarce in this species. A major reason for this discrepancy is the challenging isolation procedures owing to the small size of these retinal blood vessels, which is ~ &#8804; 30 &#181;m in luminal diameter. To circumvent the problem of direct isolation of these retinal microvessels for functional studies, we established an isolation and preparation technique that enables ex vivo studies of mouse retinal vasoactivity under near-physiological conditions. Although the present experimental preparations will specifically refer to the mouse retinal arterioles, this methodology can readily be employed to microvessels from rats.

Preparation Steps for Measurement of Reactivity in Mouse Retinal Arterioles Ex Vivo

Many vision-threatening ocular diseases are associated with dysfunctional retinal microvessels. Therefore, the measurement of retinal arteriole responses is important to investigate the underlying pathophysiological mechanisms. This article describes a detailed protocol for mouse retinal arteriole isolation and preparation to assess the effects of vasoactive substances on vascular diameter.

Vascular insufficiency and alterations in normal retinal perfusion are among the major factors for the pathogenesis of various sight-threatening ocular ...

Quantitative [18F]-Naf-PET-MRI Analysis for the Evaluation of Dynamic Bone Turnover in a Patient with Facetogenic Low Back Pain

Imaging techniques that reflect dynamic bone turnover may aid in characterizing a wide range of bone pathologies. Bone is a dynamic tissue undergoing continuous remodeling with the competing activity of osteoblasts, which produce the new bone matrix, and osteoclasts, whose function is to eliminate mineralized bone. [18F]-NaF is a positron emission tomography (PET) radiotracer that enables visualization of bone metabolism. [18F]-NaF is chemically absorbed into hydroxyapatite in the bone matrix by osteoblasts and can thus noninvasively detect osteoblastic activity, which is occult to conventional imaging techniques. Kinetic modeling of dynamic [18F]-NaF-PET data provides detailed quantitative measures of bone metabolism. Conventional semi-quantitative PET data, which utilizes standardized uptake values (SUVs) as a measure of radiotracer activity, is referred to as a static technique due to its snapshot of tracer uptake in time.&#160; Kinetic modeling, however, utilizes dynamic image data where tracer levels are continuously acquired providing tracer uptake temporal resolution. From the kinetic modeling of dynamic data, quantitative values like blood flow and metabolic rate (i.e., potentially informative metrics of tracer dynamics) can be extracted, all with respect to the measured activity in the image data. When combined with dual modality PET-MRI, region-specific kinetic data can be correlated with anatomically registered high-resolution structural and pathologic information afforded by MRI. The goal of this methodological manuscript is to outline detailed techniques for performing and analyzing dynamic [18F]-NaF-PET-MRI data. The lumbar facet joint is a common site of degenerative arthritis disease and a common cause for axial low back pain.&#160; Recent studies suggest [18F]-NaF-PET may serve as a useful biomarker of painful facetogenic disease.&#160; The human lumbar facet joint will, therefore, be used as a prototypical region of interest for dynamic [18F]-NaF-PET-MRI analysis in this manuscript.

Quantitative [18F]-Naf-PET-MRI Analysis for the Evaluation of Dynamic Bone Turnover in a Patient with Facetogenic Low Back Pain

Imaging techniques that reflect dynamic bone turnover may aid in characterizing a wide range of bone pathologies. We present detailed methodologies for performing and analyzing dynamic [18F]-NaF-PET-MRI data in a patient with facetogenic low back pain using the lumbar facet joints as a prototypical region of interest.

Imaging techniques that reflect dynamic bone turnover may aid in characterizing a wide range of bone pathologies. Bone is a dynamic tissue undergoing ...

Isolation and Purification of Murine Cardiac Pericytes

Pericytes, perivascular cells of microvessels and capillaries, are known to play a part in angiogenesis, vessel stabilization, and endothelial barrier integrity. However, their tissue-specific functions in the heart are not well understood. Moreover, there is currently no protocol utilizing readily accessible materials to isolate and purify pericytes of cardiac origin. Our protocol focuses on using the widely used mammalian model, the mouse, as our source of cells. Using the enzymatic digestion and mechanical dissociation of heart tissue, we obtained a crude cell mixture that was further purified by fluorescence activating cell sorting (FACS) by a plethora of markers. Because there is no single unequivocal marker for pericytes, we gated for cells that were CD31-CD34-CD45-CD140b+NG2+CD146+. Following purification, these primary cells were cultured and passaged multiple times without any changes in morphology and marker expression. With the ability to regularly obtain primary murine cardiac pericytes using our protocol, we hope to further understand the role of pericytes in cardiovascular physiology and their therapeutic potential.

We have optimized a protocol to isolate and purify murine cardiac pericytes for basic research and investigation of their biology and therapeutic potential.

Pericytes, perivascular cells of microvessels and capillaries, are known to play a part in angiogenesis, vessel stabilization, and endothelial barrier ...

Whole-Mount Immunofluorescence Staining, Confocal Imaging and 3D Reconstruction of the Sinoatrial and Atrioventricular Node in the Mouse

The electrical signal physiologically generated by pacemaker cells in the sinoatrial node (SAN) is conducted through the conduction system, which includes the atrioventricular node (AVN), to allow excitation and contraction of the whole heart. Any dysfunction of either SAN or AVN results in arrhythmias, indicating their fundamental role in electrophysiology and arrhythmogenesis. Mouse models are widely used in arrhythmia research, but the specific investigation of SAN and AVN remains challenging.
The SAN is located at the junction of the crista terminalis with the superior vena cava and AVN is located at the apex of the triangle of Koch, formed by the orifice of the coronary sinus, the tricuspid annulus, and the tendon of Todaro. However, due to the small size, visualization by conventional histology remains challenging and it does not allow the study of SAN and AVN within their 3D environment.
Here we describe a whole-mount immunofluorescence approach that allows the local visualization of labelled mouse SAN and AVN. Whole-mount immunofluorescence staining is intended for smaller sections of tissue without the need for manual sectioning. To this purpose, the mouse heart is dissected, with unwanted tissue removed, followed by fixation, permeabilization and blocking. Cells of the conduction system within SAN and AVN are then stained with an anti-HCN4 antibody. Confocal laser scanning microscopy and image processing allow differentiation between nodal cells and working cardiomyocytes, and to clearly localize SAN and AVN. Furthermore, additional antibodies can be combined to label other cell types as well, such as nerve fibers.
Compared to conventional immunohistology, whole-mount immunofluorescence staining preserves the anatomical integrity of the cardiac conduction system, thus allowing the investigation of AVN; especially so into their anatomy and interactions with the surrounding working myocardium and non-myocyte cells.

We provide a step-by-step protocol for whole-mount immunofluorescence staining of the sinoatrial node (SAN) and atrioventricular node (AVN) in murine hearts.

The electrical signal physiologically generated by pacemaker cells in the sinoatrial node (SAN) is conducted through the conduction system, which includes ...