This research was supported by the National Institutes of Health grant R01HL086939 (NCC).

In this protocol we demonstrate an isolated, ventilated, perfused mouse lung preparation that has previously been used to quantify the impact of pulmonary emboli and chronic hypoxia on pulsatile pulmonary vascular pressure-flow relationships (Tuchscherer, Webster, &amp; Chesler, 2006; Tuchscherer et al., 2007). In brief, the mouse lungs are surgically isolated from surrounding tissues, placed in a heated chamber (IL-1; Harvard Apparatus, Holliston, MA) and ventilated (Ventilatory Control Module (VCM)-R with timer counter module (TCM); Harvard Apparatus). The lung vasculature is perfused with heated RPMI 1640 cell culture medium with 3.5% Ficoll using a syringe pump (Cole-Parmer, Vernon Hills, IL) to generate the steady flow waveforms or a high-frequency oscillatory pump (Bose -Electro Force, Eden Prairie, MN) in parallel with the syringe pump to create pulsatile pulmonary vascular flow waveforms. Pressure transducers (P75, Harvard Apparatus) measure the instantaneous pulmonary artery pressure (PAP) and left atrial pressure (LAP). Instantaneous flow rate (Q) is measured with an in-line flow meter (Transonic Systems, Inc., Ithaca, NY). Pulsatile pressure-flow relationships are derived from these measurements, which provide insight into pulmonary vascular physiology and pathology and right ventricular afterload.
1. Equipment:
<ol>
	<li>Isolated lung set-up including mouse ventilator</li>
	<li>Data acquisition system and computer with LabView program</li>
	<li>Two pressure transducers and flowmeter for perfusate flow</li>
	<li>Pressure transducer and flowmeter (pnemotachometer) for airway flow</li>
	<li>High-frequency oscillatory pump and computer with Win Test program</li>
	<li>Boom/zoom Microscope, lamp</li>
	<li>Heating bath with high output pump for IL-1 system</li>
</ol>
2. Preparing the IL-1 system
<ol>
	<li>Distilled water heated to 37&deg;C by the heating bath is circulated into the interior of the IL-1 system.</li>
	<li>All pumps, transducers and the IL-1 cannula are connected via clean tubing and all tubing is flushed with distilled water heated to 37&deg;C. Air bubbles, which could travel into the lung and cause edema, must be removed. Tubing from the oscillatory flow pump to the flow sensor and from the flow sensor to the pulmonary artery cannula are flushed with 1% PBS.</li>
	<li>The P75 pressure transducers are zeroed by closing the valve to the cannula, opening the valves to the atmosphere and then pushing the automatic zero button on the PLUGSYS amplifier. Then, the valve to the atmosphere is closed and the valve to the cannula opened.</li>
	<li>The ceramic porous piece in the ventilation pathway of the IL-1 system is wetted to provide humidity.</li>
</ol>
3. Solutions
<ol>
	<li>Prepare 3.5% by volume Ficoll-RPMI solution and sterile filter the media. Filtering the media ensures there are no large particles that could unintentionally embolize the lung. Using sterile media also reduces the likelihood of sudden edema developing in the lung. Fill one 10 ml syringe with RPMI for surgery and one 60 ml syringe with RPMI for each experimental trial. Heat the perfusate in a 37 &deg;C water bath.</li>
	<li>Prepare 1ml of heparin 500IU/100g body weight of the mouse (approximate). The heparin salt is 158IU/mg. For a ~25 g mouse, mix 1.25 mg heparin salt with 1ml PBS solution in a small microcentrifuge tube.</li>
</ol>
4. Ventilating a Mouse
<ol>
	<li>After an intraperitoneal injection of 150 mg pentobarbital in solution per kilogram into the mouse, ensure deep anaesthesia by performing a hard pinch on a paw. If there is no reaction, prepare the mouse for surgery by pinning its front paws to corkboard for stability. Note: Throughout the experiment anesthetic depth is monitored by carefully observing the animal. The pins in the paws and frequent incisions act as noxious stimuli, the lack of response to which confirms that the animal remains in a surgical plane of anesthesia.</li>
	<li>Spray the chest with 95% alcohol to wet down the fur and use straight forceps to grab the skin at the neck. Cut a 1 cm opening in the skin using the straight scissors.</li>
	<li>Once the inside of the neck is exposed, remove all the white glandular tissue and superficial muscle, looking for the esophagus and trachea. Isolate the esophagus and trachea from tissue on both sides and posteriorly.</li>
	<li>Insert the small bent forceps under the trachea and grab a piece of suture on the other side. Pull the suture under the trachea and tie a loose surgeons knot. Do not tighten the suture or tie the knot.</li>
	<li>Cut a small angled "v" in the trachea with small scissors; do not cut all the way through the trachea. Move the cork board and mouse to the heated IL-1 system. Using the two blunt forceps, grab the tracheal cannula and the trachea below the "v". Then, slide the tracheal cannula into the trachea through the "v" opening. Tighten the suture around trachea and tracheal cannula to secure the cannula inside the trachea. Tie the knot.</li>
	<li>Begin ventilation (50% inspiration, 90 breaths/min, with deep inspiration) with room air.</li>
</ol>
5. Perfusing a Ventilated Mouse
5.1. Access the right ventricle to inject heparin
<ol>
	<li>Spray mouse chest with alcohol again to re-wet the fur. Remove all of the skin on the chest above the ribs using straight forceps and straight scissors. Cut upward along the sternum. Lift the skin on each side and then cut the skin following the line of the lower ribs.</li>
	<li>Grasp the xiphoid process at the bottom of the sternum with tweezers and cut a hole into the diaphragm using the straight scissors. Grab the diaphragm with the tweezers and cut it away from the ribs.</li>
	<li>Grasp the xiphoid process again with the tweezers (left hand) and use the ball-tip, angled scissors to cut up the sternum and through ribs, being careful not to cut the lungs, the heart or blood vessels (use the ball-tip on the scissors to guide you). There will be blood but as long as the cutting edge of the scissors is against the sternum, the heart and lungs will not be cut.</li>
	<li>Grasp the ribs on the left side and cut away as much of the ribs as necessary to expose the heart. Slowly inject the right ventricle with 0.1 ml heparin solution. This step is important to preventing blood clots in the lung, which damage endothelial cells and will impair perfusion. The heparin must be injected while the heart is still beating.</li>
</ol>
5.2. Cannulate the main pulmonary artery
<ol>
	<li>Cut away the rest of the ribs (left and right side) using the back (rounded) end of the forceps to gently push the lungs away from the rib wall. Grabbing the lungs themselves will damage the delicate tissue. Incidental contact between the scissor tips and the lung tissue will also cause damage.</li>
	<li>Move the microscope into place over lung. Cut away the glandular and fatty tissue on top of the heart. Use the tweezers to pull it away from the arteries and veins and then cut with the spring scissors while the tissue is in tension.</li>
	<li>Using another blunt set of tweezers, scoop from right to left from the top of heart under left atrium/ventricle to get the tip of the tweezers under the aorta and pulmonary artery (PA). Do this carefully- there should not be resistance to the tweezers. Using blunt tweezers reduces the likelihood of accidentally puncturing the pulmonary artery or aorta.</li>
	<li>Once the tweezer tip is under the aorta and PA, grab a piece of suture and pull through. Tie a loose surgeons knot. Do not tighten the suture or tie the knot.</li>
	<li>Optional: use the angled scissors to remove the lower half of the body. Cut down through the ribs and spine; cutting the aorta and vena cava will cause a significant amount of blood to flow -- use a Q-tip to staunch the flow. Place in a bag for disposal.</li>
	<li>Prime PA cannula with 4 ml from a 10 ml syringe of RPMI. Double check that all tubing is free of air bubbles. Perfusion of the lung with distilled
water or bubbles will result in edema.</li>
	<li>Cut a small notch in the right ventricular free wall and insert the PA cannula, aiming down and to the right. The tip of the cannula should be visible through the transparent wall of the PA. Infuse a small amount of RPMI to confirm your location. Tighten the suture around the cannula, aorta and PA and tie the knot. Note that at this point, the aorta is also tied off so actual perfusion should not begin until the left atrium is cannulated.</li>
</ol>
5.3. Cannulate the left atrium
<ol>
	<li>Cut a notch in lower part of left ventricle and inert the left atrial (LA) cannula, aiming upwards. Slight pressure may be needed to open the mitral valve in this direction. In the correct location, the cannula tip will slip through and be secure without suture.</li>
</ol>
5.4. Begin perfusion
<ol>
	<li>Manually infuse RPMI from the 10 ml syringe at 0.3 ml per minute until RMPI is evident in the outflow tubing (pink-ish color in contrast to the clear PBS). If there is no flow in the outflow tubing, re-position the LA cannula. If no outflow can be obtained with re-positioning, check for a leak in the pulmonary artery. A leak or tear in the pulmonary artery cannot be repaired; this is cause for aborting the experiment.</li>
	<li>Connect a 60 ml syringe to the PA cannula through the IL-1 system and start 1ml/min infusion of perfusate, checking for leakage and making sure the lungs turn white, which shows that RPMI is replacing blood in the lungs. Perfuse with slow flow for two minutes.</li>
</ol>
6. Measuring Pulsatile and Steady Pulmonary Pressure-Flow Relationships
<ol>
	<li>For the pulsatile flow studies, first set the oscillatory pump piston displacements at the desired levels for each frequency in the WinTest program based on previous experiments. Due variability in lung structure and mechanics, the displacements have to be adjusted for each mouse. Set steady flow to the desired level. Open the valve to the oscillatory pump and start recording data right before running the oscillatory flow profile (Wintest program). Open the data file and plot the experimental flow (Q) in Excel. Adjust the oscillatory pump piston displacements at each frequency (Wintest program) so that Q max and Q min are as desired.</li>
	<li>For steady flow trials, close the valve to the oscillatory pump. If this valve is left open, the oscillatory pump acts as a capacitor, dampening the changes from one flow rate to another. Collect data for at least 10 seconds at each flow rate or until the PA pressure does not change by more than 5%.</li>
	<li>For either pulsatile or steady flow measurements, old ventilation at a constant pressure before data collection. Resume ventilation immediately after data collection.</li>
	<li>Watch for RPMI in the airway tubing; this is evidence of edema and is cause for aborting the experiment. Also, do not let RPMI reach the airway pressure or flow sensors because it will damage the transducers.</li>
</ol>
7. Representative Results:
Representative Steady Results:
In the isolated lung set-up the experimenter has the ability to independently control not only the pulmonary flow Q but also the airway pressure Pair and left atrial pressure LAP. This is advantageous because Q, Pair and LAP influence the pulmonary vasculature and the resulting pulmonary artery pressure PAP. Another advantage is that results obtained are independent of sympathetic nervous system tone 1, volume status, and anesthesia 2.
PAP changes induced by stair-step changes in Q for fixed Pair and either fixed or varying LAP are shown in Figure 1. Note that in the isolated lung preparation, the LAP cannula is typically connected to tubing that directs the perfusate into a waste container. With this tubing in place, LAP is linearly dependent on Q due to Poiseuille flow. However, the height of the outlet and waste container can be manually adjusted to provide a constant, non-zero LAP or the tubing can be removed to allow zero LAP that is independent of Q.
Representative Pulsatile Results:
While arbitrary pulsatile flow waveforms can be generated with this system, we typically generate flow of the form Q = 3 + 2 sin (2f&pi;t) ml/min at frequencies of f = 1, 2, 5, 10, 15 and 20 Hz to assess the linearized impedance of the pulmonary vasculature (Figure 2: top panel). From the resulting PAP, LAP and Q measurements, pulmonary vascular impedance magnitude (Z) and phase (&theta;) are calculated by first decomposing one full sinusoidal cycle of &Delta;P = PAP-LAP and Q at each imposed sinusoidal flow rate frequency into a series of sinusoidal harmonics using a Fourier transform. The ratio of the pressure transform to flow transform yields the pulmonary vascualar impedance, PVZ = FFT (&Delta;P)/FFT(Q), which has magnitude Z and phase &theta;. Input impedance Z0, characteristic impedance ZC, and index of wave reflection RW, are calculated from the impedance magnitude. In particular, Z0 is calculated from Z at the 0th harmonic (f = 0 Hz) averaged over all frequencies, ZC is calculated as the average of Z between the first minimum (5 Hz) and 20 Hz, and RW is calculated as (Z0-ZC)/(Z0+ZC) 3.
<img src="/files/ftp_upload/2690/2690fig1.jpg" alt="Figure 1" /> 
Figure 1. Steady flow waveforms (top row) and resulting pressures (second row: PAP, Pair, LAP) as a function of time with different combinations of LAP and Pair. The bottom row shows PAP vs. Q. In (A) and (B), LAP increases and decreases with Q because the outflow tubing was in place. This tubing was removed for (C) so that LAP is constant and independent of Q. In (D) and (E) the height of the outlet tubing was adjusted manually so that LAP is higher but independent of Q. Pair was either at end-inspiratory (A, C, D) or end-expiratory (B, E) pressure.
<img src="/files/ftp_upload/2690/2690fig2.jpg" alt="Figure 2" /> 
Figure 2. Pulsatile flow waveform Q (top panel) and resulting pressures (bottom panel: PAP, LAP and Pair) as a function of time. From these pulsatile pulmonary pressure-flow relationships, PVZ can be calculated, which reflects the total right ventricular afterload.

<ol>
	<li>Pace, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sympathetic+control+of+pulmonary+vascular+impedance+in+anesthetized+dogs.">Sympathetic control of pulmonary vascular impedance in anesthetized dogs.</a> Circ Res. 29, 555-568 (1971).</li><li>Ewalenko, P., Stefanidis, C., Holoye, A., Brimioulle, S., Naeije, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulmonary+vascular+impedance+vs.+resistance+in+hypoxic+and+hyperoxic+dogs:+effects+of+propofol+and+isoflurane.">Pulmonary vascular impedance vs. resistance in hypoxic and hyperoxic dogs: effects of propofol and isoflurane.</a> J Appl Physiol. 74, 2188-2193 (1993).</li><li>Nichols, W. W., O'Rourke, M. F., Hartley, C., McDonald, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> McDonald's blood flow in arteries : theoretical, experimental, and clinical principles. , (2005).</li><li>Tuchscherer, H. A., Webster, E. B., Chesler, N. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulmonary+Vascular+Resistance+and+Impedance+in+Isolated+Mouse+Lungs:+Effects+of+Pulmonary+Emboli.">Pulmonary Vascular Resistance and Impedance in Isolated Mouse Lungs: Effects of Pulmonary Emboli.</a> Annals of Biomedical Engineering. 34, 660-668 (2006).</li><li>Tuchscherer, H. A., Vanderpool, R. R., Chesler, N. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulmonary+vascular+remodeling+in+isolated+mouse+lungs:+Effects+on+pulsatile+pressure-flow+relationships.">Pulmonary vascular remodeling in isolated mouse lungs: Effects on pulsatile pressure-flow relationships.</a> Journal of Biomechanics. 40, 993-1001 (2007).</li><li>Vanderpool, R., Naeije, R., Chesler, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Impedance+in+Isolated+Mouse+Lungs+for+the+Determination+of+Site+of+Action+of+Vasoactive+Agents+and+Disease.">Impedance in Isolated Mouse Lungs for the Determination of Site of Action of Vasoactive Agents and Disease.</a> Ann Biomed Eng. , (2010).</li><li>Vanderpool, R., Kim, A., Molthen, R., Chesler, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+acute+rho+kinase+inhibition+on+chronic+hypoxia-induced+changes+in+proximal+and+distal+pulmonary+arterial+structure+and+function.">Effects of acute rho kinase inhibition on chronic hypoxia-induced changes in proximal and distal pulmonary arterial structure and function.</a> Journal of Applied Physiology. , (2010).</li><li>El-Bizri, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Smooth+muscle+protein+22alpha-mediated+patchy+deletion+of+Bmpr1a+impairs+cardiac+contractility+but+protects+against+pulmonary+vascular+remodeling.">Smooth muscle protein 22alpha-mediated patchy deletion of Bmpr1a impairs cardiac contractility but protects against pulmonary vascular remodeling.</a> Circ Res. 102, 380-388 (2008).</li><li>Champion, H., Michelakis, E., Hassoun, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comprehensive+invasive+and+noninvasive+approach+to+the+right+ventricle-pulmonary+circulation+unit:+state+of+the+art+and+clinical+and+research+implications.">Comprehensive invasive and noninvasive approach to the right ventricle-pulmonary circulation unit: state of the art and clinical and research implications.</a> Circulation. 120, 992-1007 (2009).</li><li>Gan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Noninvasively+assessed+pulmonary+artery+stiffness+predicts+mortality+in+pulmonary+arterial+hypertension.">Noninvasively assessed pulmonary artery stiffness predicts mortality in pulmonary arterial hypertension.</a> Chest. 132, 1906-1912 (2007).</li><li>Mahapatra, S., Nishimura, R., Sorajja, P., Cha, S., McGoon, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relationship+of+pulmonary+arterial+capacitance+and+mortality+in+idiopathic+pulmonary+arterial+hypertension.">Relationship of pulmonary arterial capacitance and mortality in idiopathic pulmonary arterial hypertension.</a> J Am Coll Cardiol. 47, 799-803 (2006).</li><li>Bogaard, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chronic+pulmonary+artery+pressure+elevation+is+insufficient+to+explain+right+heart+failure.">Chronic pulmonary artery pressure elevation is insufficient to explain right heart failure.</a> Circulation. 120, 1951-1960 (2009).</li></ol>

No conflicts of interest declared.

Critical steps in the surgery
It is critical that care is taken when cutting the rib cage away from the lungs. The lungs must be fully exposed and uninhibited by surrounding tissues during inflation but not damaged during the process of isolation. The use of a flat object like the back end of the forceps can be used to hold the lungs away from the chest wall so that there is a clear path for scissors to cut. Another critical step is the placement of the suture around the pulmonary artery and the...

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>1 ml syringe</td>
<td>&nbsp;</td>
<td>Fisher Sci</td>
<td>14-829-10F</td>
<td>&nbsp;</td>
<tr>
<td>10 ml syringe</td>
<td>&nbsp;</td>
<td>Fisher Sci</td>
<td>14-823-2A </td>
<td>&nbsp;</td>
<tr>
<td>60 ml syringe</td>
<td>&nbsp;</td>
<td>Fisher Sci</td>
<td>13-689-8 </td>
<td>&nbsp;</td>
<tr>
<td>RPMI with GLN 6/PK</td>
<td>&nbsp;</td>
<td>Fisher Sci</td>
<td>MT10040CV </td>
<td>&nbsp;</td>
<tr>
<td>Bottle Top Filters</td>
<td>&nbsp;</td>
<td>Fisher Sci</td>
<td>09-761-57</td>
<td>&nbsp;</td>
<tr>
<td>Ficoll PM 70</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>F2878-100g</td>
<td>&nbsp;</td>
<tr>
<td>Heparin</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Y27632</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>Y0503</td>
<td>&nbsp;</td>
<tr>
<td>Angled Ball Iris scissors</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>14109-09</td>
<td>&nbsp;</td>
<tr>
<td>Vannas Spring Scissors - 4mm Blades</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>15018-10</td>
<td>&nbsp;</td>
<tr>
<td>Fine Iris Scissors - straight</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>14106-09</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #5/45 Forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11251-35</td>
<td>&nbsp;</td>
<tr>
<td>Dumont Medical Biology Forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools</td>
<td>11254-20</td>
<td>&nbsp;</td>
<tr>
<td>Lauda E100 ECO-line 003 </td>
<td>&nbsp;</td>
<td>VWR</td>
<td>Comparable to Lauda-Brinkmann E-103, 62400-922</td>
<td>&nbsp;</td>
<tr>
<td>IL-1 Isolated perfused mouse lung system </td>
<td>&nbsp;</td>
<td>Harvard Apparatus </td>
<td>739904 </td>
<td>&nbsp;</td>
<tr>
<td>Blood Pressure Transducer P75 for PLUGSYS Module</td>
<td>&nbsp;</td>
<td>Harvard Apparatus</td>
<td>730020</td>
<td>&nbsp;</td>
<tr>
<td>TS410 Flow Modules</td>
<td>&nbsp;</td>
<td>Transonic </td>
<td>TS410</td>
<td>&nbsp;</td>
<tr>
<td>ME 4 PXN Precision PXN Inline Flowsensors</td>
<td>&nbsp;</td>
<td>Transonic</td>
<td>ME 4 PXN</td>
<td>&nbsp;</td>
<tr>
<td>Cole-Parmer Multi-Syringe Pumps</td>
<td>&nbsp;</td>
<td>Cole-Parmer </td>
<td>EW-74900-20</td>
<td>&nbsp;</td>
<tr>
<td>Nembutal 50MG/ML 20ML Vial</td>
<td>&nbsp;</td>
<td>Amatheon</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

characterization of the isolated, ventilated, and instrumented mouse lung perfused with pulsatile flow

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured with independent control over pulmonary arterial flow rate, flow rate waveform, airway pressure and left atrial pressure. Pulmonary vascular resistance is calculated based on multi-point, steady pressure-flow curves; pulmonary vascular impedance is calculated from pulsatile pressure-flow curves obtained at a range of frequencies. As now recognized clinically, impedance is a superior measure of right ventricular afterload than resistance because it includes the effects of vascular compliance, which are not negligible, especially in the pulmonary circulation. Three important metrics of impedance - the zero hertz impedance Z0, the characteristic impedance ZC, and the index of wave reflection RW - provide insight into distal arterial cross-sectional area available for flow, proximal arterial stiffness and the upstream-downstream impedance mismatch, respectively. All results obtained in isolated, ventilated and perfused lungs are independent of sympathetic nervous system tone, volume status and the effects of anesthesia. We have used this technique to quantify the impact of pulmonary emboli and chronic hypoxia on resistance and impedance, and to differentiate between sites of action (i.e., proximal vs. distal) of vasoactive agents and disease using the pressure dependency of ZC. Furthermore, when these techniques are used with the lungs of genetically engineered strains of mice, the effects of molecular-level defects on pulmonary vascular structure and function can be determined.

Watch this Scientific Journal Video about Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow at JoVE.com

Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

The following protocol outlines the process of isolating, ventilating and instrumenting mouse lungs to measure steady or pulsatile pulmonary vascular pressure-flow relationships in order to quantify the effects of blood flow, airflow, airway changes and vascular changes on right ventricular afterload.

The isolated, ventilated and instrumented mouse lung preparation allows steady and pulsatile pulmonary vascular pressure-flow relationships to be measured ...

characterization-isolated-ventilated-instrumented-mouse-lung-perfused

Research

JoVE Journal

Biology

16.2K vues.  University of Wisconsin – Madison. Le protocole suivant décrit le processus d'isolement, de ventilation et d'instrumenter les poumons de souris pour mesurer stable ou pulsatile vasculaire pulmonaire pression-débit des relations afin de quantifier les effets de la circulation sanguine, circulation de l'air, les changements des voies aériennes et sur les modifications vasculaires postcharge ventriculaire droite.

Vidéo: Characterization of the Isolated, Ventilated, and Instrumented Mouse Lung Perfused with Pulsatile Flow

Optical Mapping of Langendorff-perfused Rat Hearts

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in experimental models that range in scale from cell cultures to whole-organs[1, 2]. Using state-of-the-art optical imaging systems, generation and propagation of action potentials during normal cardiac rhythm or throughout initiation and maintenance of arrhythmias can be visualized almost instantly[1]. The latest commercially-available systems can provide information at exceedingly high spatiotemporal resolutions and were based on custom-built equipment initially developed to overcome the obstacles imposed by more conventional electrophysiological methods[1]. Advancements in high-resolution and high-speed complementary metal-oxide-semiconductor (CMOS) cameras and intensely-bright, light-emitting diodes (LEDs) as well as voltage-sensitive dyes, optics, and filters have begun to make electrical signal acquisition practical for cardiovascular cell biologists who are more accustomed to working with microscopes. Although the newest generation of CMOS cameras can acquire 10,000 frames per second on a 16,384 pixel array, depending on the type of sample preparation, long-established fluorescence acquisition technologies such as photodiode arrays, laser scanning systems, and cooled charged-coupled device (CCD) cameras still have some distinct advantages with respect to dynamic range, signal-to-noise ratio, and quantum efficiency[1, 3]. In the present study, Lewis rat hearts were perfused ex vivo with a crystalloid perfusate (Krebs-Henseleit solution) at 37&deg;C on a modified Langendorff apparatus. After a 20 minute stabilization period, the hearts were intermittently perfused with 11 mMol/L 2,3-butanedione monoxime to eliminate contraction-associated motion during image acquisition. For optical mapping, we loaded hearts with the fast-response potentiometric probe di-8-ANEPPS[4] (5 &mu;Mol/L) and briefly illuminated the preparation with 475&plusmn;15 nm excitation light. During a typical 2 second period of illumination, &gt;605 nm light emitted from the cardiac preparation was imaged with a high-speed CMOS camera connected to a horizontal macroscope. For this demonstration, hearts were paced at 300 beats per minute with a coaxial electrode connected to an isolated electrical stimulation unit. Simultaneous bipolar electrographic recordings were acquired and analyzed along with the voltage signals using readily-available software. In this manner, action potentials on the surface of Langendorff-perfused rat hearts can be visualized and registered with electrographic signals.

This article describes a high temporal and spatial resolution technique to optically image action potential movement on the surface of Langendorff-perfused rat hearts using a potentiometric dye (di-8-ANEPPS).

Optical mapping of the cardiac surface with voltage-sensitive fluorescent dyes has become an important tool to investigate electrical excitation in ...

Flow Cytometry Purification of Mouse Meiotic Cells

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the unique differentiation programs that are manifest during meiosis. Two principal methods have been developed to purify, to varying degrees, various meiotic fractions from both adult and immature animals: elutriation or Staput (sedimentation) using BSA and/or percoll gradients. Both of these methods rely on cell size and density to separate meiotic cells1-5. Overall, except for few cell populations6, these protocols fail to yield sufficient purity of the numerous meiotic cell populations that are necessary for detailed molecular analyses. Moreover, with such methods usually one type of meiotic cells can be purified at a given time, which adds an extra level of complexity regarding the reproducibility and homogeneity when comparing meiotic cell samples.
Here, we describe a refined method that allows one to easily visualize, identify, and purify meiotic cells, from germ cells to round spermatids, using FACS combined with Hoechst 33342 staining7,8. This method provides an overall snapshot of the entire meiotic process and allows one to highly purify viable cells from most stage of meiosis. These purified cells can then be analyzed in detail for molecular changes that accompany progression through meiosis, for example changes in gene expression9,10and the dynamics of nucleosome occupancy at hotspots of meiotic recombination11.

An efficient method to obtain highly purified viable meiotic fractions from mouse testis is described, which combines a refined cell dissociation protocol with fluorescent activated cell sorting (FACS). This method takes advantage of differences in the DNA content and nuclear density of discrete meiotic fractions.

The heterogeneous nature of cell types in the testis and the absence of meiotic cell culture models have been significant hurdles to the study of the ...

Micro-Mechanical Characterization of Lung Tissue Using Atomic Force Microscopy

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3. On the macro scale the stiffness of tissues and organs within the human body span several orders of magnitude4. Much less is known about how stiffness varies spatially within tissues, and what the scope and spatial scale of stiffness changes are in disease processes that result in tissue remodeling. To better understand how changes in matrix stiffness contribute to cellular physiology in health and disease, measurements of tissue stiffness obtained at a spatial scale relevant to resident cells are needed. This is particularly true for the lung, a highly compliant and elastic tissue in which matrix remodeling is a prominent feature in diseases such as asthma, emphysema, hypertension and fibrosis. To characterize the local mechanical environment of lung parenchyma at a spatial scale relevant to resident cells, we have developed methods to directly measure the local elastic properties of fresh murine lung tissue using atomic force microscopy (AFM) microindentation. With appropriate choice of AFM indentor, cantilever, and indentation depth, these methods allow measurements of local tissue shear modulus in parallel with phase contrast and fluorescence imaging of the region of interest. Systematic sampling of tissue strips provides maps of tissue mechanical properties that reveal local spatial variations in shear modulus. Correlations between mechanical properties and underlying anatomical and pathological features illustrate how stiffness varies with matrix deposition in fibrosis. These methods can be extended to other soft tissues and disease processes to reveal how local tissue mechanical properties vary across space and disease progression.

The stiffness of the extracellular matrix strongly influences multiple behaviors of adherent cells. Matrix stiffness varies spatially throughout a tissue, and undergoes modification in various disease conditions. Here we develop methods to characterize spatial variations in stiffness in normal and fibrotic mouse lung tissue using atomic force microscopy microindentation.

Matrix stiffness strongly influences growth, differentiation and function of adherent cells1-3.  On the macro scale the stiffness of tissues and organs ...

Isolation &#38; Characterization of Hoechstlow CD45negative Mouse Lung Mesenchymal Stem Cells

Tissue resident mesenchymal stem cells (MSC) are important regulators of tissue repair or regeneration, fibrosis, inflammation, angiogenesis and tumor formation. Taken together these studies suggest that resident lung MSC play a role during pulmonary tissue homeostasis, injury and repair during diseases such as pulmonary fibrosis (PF) and arterial hypertension (PAH). Here we describe a technology to define a population of resident lung MSC. The definition of this population in vivo pulmonary tissue using a define set of markers facilitates the repeated isolation of a well-characterized stem cell population by flow cytometry and the study of a specific cell type and function.

Isolation &amp; Characterization of Hoechstlow CD45negative Mouse Lung Mesenchymal Stem Cells

In this article we demonstrate the isolation of murine resident lung mesenchymal stem cells (lung MSC), their expansion, characterization and analysis of immunomodulatory properties.

Tissue resident mesenchymal stem cells (MSC) are important regulators of tissue repair or regeneration, fibrosis, inflammation, angiogenesis and tumor ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

Quantifying Single Microvessel Permeability in Isolated Blood-perfused Rat Lung Preparation

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with real time imaging, it becomes feasible to determine permeability changes in individual pulmonary microvessels. Herein we describe steps to isolate rat lungs and perfuse them with autologous blood. Then, we outline steps to infuse fluorophores or agents via a microcatheter into a small lung region. Using these procedures described, we determined permeability increases in rat lung microvessels in response to infusions of bacterial lipopolysaccharide. The data revealed that lipopolysaccharide increased fluid leak across both venular and capillary microvessel segments. Thus, this method makes it possible to compare permeability responses among vascular segments and thus, define any heterogeneity in the response. While commonly used methods to define lung permeability require postprocessing of lung tissue samples, the use of real time imaging obviates this requirement as evident from the present method. Thus, the isolated lung preparation combined with real time imaging offers several advantages over traditional methods to determine lung microvascular permeability, yet is a straightforward method to develop and implement.

The isolated blood-perfused lung preparation makes it feasible to visualize microvessel networks on the lung surface. Here we describe an approach to quantify permeability of single microvessels in isolated lungs using real time fluorescence imaging.

The isolated blood-perfused lung preparation is widely used to visualize and define signaling in single microvessels. By coupling this preparation with ...

An Immunofluorescent Method for Characterization of Barrett&#8217;s Esophagus Cells

Esophageal adenocarcinoma (EAC) has an overall survival rate of less than 17% and incidence of EAC has risen dramatically over the past two decades. One of the primary risk factors of EAC is Barrett&#8217;s esophagus (BE), a metaplastic change of the normal squamous esophagus in response to chronic heartburn. Despite the well-established connection between EAC and BE, interrogation of the molecular events, particularly altered signaling pathways involving progression of BE to EAC, are poorly understood. Much of this is due to the lack of suitable in vitro models available to study these diseases. Recently, immortalized BE cell lines have become commercially available allowing for in vitro studies of BE. Here, we present a method for immunofluorescent staining of immortalized BE cell lines, allowing in vitro characterization of cell signaling and structure after exposure to therapeutic compounds. Application of these techniques will help develop insight into the mechanisms involved in BE to EAC progression and provide potential avenues for treatment and prevention of EAC.

An Immunofluorescent Method for Characterization of Barrett&amp;#8217;s Esophagus Cells

There is a discernible need for improved information on molecular drivers of Barrett&#8217;s Esophagus. Immunofluorescent staining is a useful technique for understanding the effects of cell signaling on cell morphology. We present a simple, effective protocol for the use of immunofluorescent staining to assess therapeutic treatment in Barrett&#8217;s Esophagus cells.

Esophageal adenocarcinoma (EAC) has an overall survival rate of less than 17% and incidence of EAC has risen dramatically over the past two decades. One ...

Phenotyping Mouse Pulmonary Function In Vivo with the Lung Diffusing Capacity

The mouse is now the primary animal used to model a variety of lung diseases. To study the mechanisms that underlie such pathologies, phenotypic methods are needed that can quantify the pathologic changes. Furthermore, to provide translational relevance to the mouse models, such measurements should be tests that can easily be done in both humans and mice. Unfortunately, in the present literature few phenotypic measurements of lung function have direct application to humans. One exception is the diffusing capacity for carbon monoxide, which is a measurement that is routinely done in humans. In the present report, we describe a means to quickly and simply measure this diffusing capacity in mice. The procedure involves brief lung inflation with tracer gases in an anesthetized mouse, followed by a 1 min gas analysis time. We have tested the ability of this method to detect several lung pathologies, including emphysema, fibrosis, acute lung injury, and influenza and fungal lung infections, as well as monitoring lung maturation in young pups. Results show significant decreases in all the lung pathologies, as well as an increase in the diffusing capacity with lung maturation. This measurement of lung diffusing capacity thus provides a pulmonary function test that has broad application with its ability to detect phenotypic structural changes with most of the existing pathologic lung models.

Phenotyping Mouse Pulmonary Function In Vivo with the Lung Diffusing Capacity

We describe a means to quickly and simply measure the lung diffusing capacity in mice and show that it is sufficiently sensitive to phenotype changes in multiple common lung pathologies. This metric thus brings direct translational relevance to the mouse models, since diffusing capacity is also easily measured in humans.

The mouse is now the primary animal used to model a variety of lung diseases. To study the mechanisms that underlie such pathologies, phenotypic methods ...

The Mouse Isolated Perfused Kidney Technique

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

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

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

Analysis of Cell Suspensions Isolated from Solid Tissues by Spectral Flow Cytometry

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to the analysis of a few fluorochromes, currently there are dozens of different fluorescent dyes, and up to 14-18 different dyes can be combined at a time. However, several limitations still impair the analytical capabilities. Because of the multiplicity of fluorescent probes, data analysis has become increasingly complex due to the need of large, multi-parametric compensation matrices. Moreover, mutant mouse models carrying fluorescent proteins to detect and trace specific cell types in different tissues have become available, so the analysis (by flow cytometry) of auto-fluorescent cell suspensions obtained from solid organs is required. Spectral flow cytometry, which distinguishes the shapes of emission spectra along a wide range of continuous wavelengths, addresses some of these problems. The data is analyzed with an algorithm that replaces compensation matrices and treats auto-fluorescence as an independent parameter. Thus, spectral flow cytometry should be capable of discriminating fluorochromes with similar emission peaks and can provide a multi-parametric analysis without compensation requirements.
This protocol describes the spectral flow cytometry analysis, allowing for a 21-parameter (19 fluorescent probes) characterization and the management of an auto-fluorescent signal, providing high resolution in minor population detection. The results presented here show that spectral flow cytometry presents advantages in the analysis of cell populations from tissues difficult to characterize in conventional flow cytometry, such as the heart and the intestine. Spectral flow cytometry thus demonstrates the multi-parametric analytical capacity of high-performing conventional flow cytometry without the requirement for compensation and enables auto-fluorescence management.

This article describes spectral cytometry, a new approach in flow cytometry that uses the shapes of emission spectra to distinguish fluorochromes. An algorithm replaces compensations and can treat auto-fluorescence as an independent parameter. This new approach allows for the proper analysis of cells isolated from solid organs.

Flow cytometry has been used for the past 40 years to define and analyze the phenotype of lymphoid and other hematopoietic cells. Initially restricted to ...