Name: a large animal model for pulmonary hypertension and right ventricular failure: left pulmonary artery ligation and progressive main pulmonary artery banding in sheep
Uploaded: 2021-07-15T14:00:00
Description: Watch this Scientific Journal Video about A Large Animal Model for Pulmonary Hypertension and Right Ventricular Failure: Left Pulmonary Artery Ligation and Progressive Main Pulmonary Artery Banding in

This work was funded by the National Institutes of Health R01HL140231. We thank the Division of Animal Care for their animal husbandry and veterinary care. We thank the SR Light Laboratory and its staff, Jamie Adcock, Susan Fultz, Codi VanRooyen, and Jos&#233; Diaz, for their dedicated technical support with large animal surgeries.

The Institutional Animal Care and Use Committee at Vanderbilt University Medical Center approved the protocol. The described procedures were conducted in accordance with the US National Research Council&#39;s Guide for the Care and Use of Laboratory Animals, 8th edition. The overview and the timeline of the experimental procedure are provided in Figure 1.&#160;Supplementary Table 1&#160; describes the sheep&#8217;s sex, weight, breed, source of sheep, and other rele...

<ol>
	<li>Campo, A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Outcomes+of+hospitalization+for+right+heart+failure+in+pulmonary+arterial+hypertension.">Outcomes of hospitalization for right heart failure in pulmonary arterial hypertension.</a> European Respiratory Journal. 38 (2), 359-367 (2011).</li><li>Tonelli, A. R., 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=Causes+and+circumstances+of+death+in+pulmonary+arterial+hypertension.">Causes and circumstances of death in pulmonary arterial hypertension.</a> American Journal of Respiratory and Critical Care Medicine. 188 (3), 365-369 (2013).</li><li>Urashima, T., 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=Molecular+and+physiological+characterization+of+RV+remodeling+in+a+murine+model+of+pulmonary+stenosis.">Molecular and physiological characterization of RV remodeling in a murine model of pulmonary stenosis.</a> American Journal of Physiology- Heart and Circulatory Physiology. 295 (3), (2008).</li><li>Sato, 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=Large+animal+model+of+chronic+pulmonary+hypertension.">Large animal model of chronic pulmonary hypertension.</a> ASAIO Journal. 54 (4), 396-400 (2008).</li><li>Pohlmann, J. 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(105), e53133 (2015).</li><li>Pereda, D., 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=Swine+model+of+chronic+postcapillary+pulmonary+hypertension+with+right+ventricular+remodeling:+Long-term+characterization+by+cardiac+catheterization,+magnetic+resonance,+and+pathology.">Swine model of chronic postcapillary pulmonary hypertension with right ventricular remodeling: Long-term characterization by cardiac catheterization, magnetic resonance, and pathology.</a> Journal of Cardiovascular Translational Research. 7 (5), 494-506 (2014).</li><li>Silva, K. A. S., Emter, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Large+animal+models+of+heart+failure:+A+translational+bridge+to+clinical+success.">Large animal models of heart failure: A translational bridge to clinical success.</a> JACC: Basic to Translational Science. 5 (8), 840-856 (2020).</li><li>Ukita, R., 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=Left+pulmonary+artery+ligation+and+chronic+pulmonary+artery+banding+model+for+inducing+right+ventricular+-+pulmonary+hypertension+in+sheep.">Left pulmonary artery ligation and chronic pulmonary artery banding model for inducing right ventricular - pulmonary hypertension in sheep.</a> ASAIO Journal (American Society for Artificial Internal Organs: 1992. 67 (1), 44-48 (2020).</li><li>Ukita, R., 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=Progression+toward+decompensated+right+ventricular+failure+in+the+ovine+pulmonary+hypertension+model.">Progression toward decompensated right ventricular failure in the ovine pulmonary hypertension model.</a> ASAIO Journal (American Society for Artificial Internal Organs: 1992. , (2021).</li><li>Mercier, O., 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=Piglet+model+of+chronic+pulmonary+hypertension.">Piglet model of chronic pulmonary hypertension.</a> Pulmonary Circulation. 3 (4), 908-915 (2013).</li><li>Guihaire, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Right+ventricular+plasticity+in+a+porcine+model+of+chronic+pressure+overload.">Right ventricular plasticity in a porcine model of chronic pressure overload.</a> Journal of Heart and Lung Transplantation. 33 (2), 194-202 (2014).</li><li>Tang, K. J., Robbins, I. M., Light, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Incidence+of+pleural+effusions+in+idiopathic+and+familial+pulmonary+arterial+hypertension+patients.">Incidence of pleural effusions in idiopathic and familial pulmonary arterial hypertension patients.</a> Chest. 136 (3), 688-693 (2009).</li><li>Luo, Y. 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=Frequency+of+pleural+effusions+in+patients+with+pulmonary+arterial+hypertension+associated+with+connective+tissue+diseases.">Frequency of pleural effusions in patients with pulmonary arterial hypertension associated with connective tissue diseases.</a> Chest. 140 (1), 42-47 (2011).</li><li>Brixey, A. G., Light, R. 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The presented PH-RVF model can reliably induce varying levels of disease severity to match the goals of the investigation. Two different approaches are used in combination to induce this disease model. First, the LPA ligation serves to increase pulmonary vascular resistance and decrease PA capacitance11,12, thereby establishing the starting point of the chronic model at an already increased RV afterload state. Then, the implantation of the PA cuff and its progres...

Decompensated right ventricular (RV) failure is the predominant cause of morbidity and mortality for patients with pulmonary hypertension (PH). RV failure is responsible for over 50% of hospitalizations in patients with PH and is a common cause of death in this patient population1,2. Although current medical treatments for PH can provide temporizing measures, they do not reverse the progression of the disease. As such, the only long-term treatment is lung transplantation. To explore and test novel medical treatments and interventions for PH and RVF, a clinically relevant animal model is needed to recapitulate the disease&#39;s complex pathophysiology. In particular, there is a great clinical need to develop RV-targeted therapeutics for PH patients to improve RV function. To date, most published animal studies of PH and RV dysfunction have relied on small mammals such as mice and rats3. On the other hand, there have only been a handful of large animal models to study the disease and RV pathophysiology from abnormal afterload4,5,6,7. In addition, none of the previously published large animal models include descriptions of experimental procedures for controlled titration of disease severity that differentially leads to compensated versus decompensated RV failure phenotypes. An animal model of PH that can be titrated to induce acute and chronic RV failure with varying degrees of compensation is needed to study disease mechanisms and to develop, test, and translate novel diagnostics and therapeutics for PH and RVF into clinical practice. Such a model in a large animal is especially valuable for the development of mechanical circulatory support devices8.
Here, a chronic, large animal PH-RVF model using left pulmonary artery (PA) ligation and progressive main PA banding in adult sheep is presented9,10. The ligation of the left PA (LPA) increases the pulmonary vascular resistance and decreases PA capacitance11,12. The progressive PA banding approach allows for precise titration of disease severity and RV adaptation. This platform can also be readily utilized for longitudinal investigation of disease progression toward RV decompensation. The procedures and processes required to execute this model are presented as a resource for investigators interested in a large animal platform to develop novel treatments for PH and RVF.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#160;0.9% Sodium Chloride Irrigation Pour Bottle by Baxter Healthcare, 1000 mL</td>
			<td>Medline</td>
			<td>&#160;BHL2F7124</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>0.25% Bupivacaine</td>
			<td>Hospira Inc</td>
			<td>0409-1160-18</td>
			<td>Medication, Intra-Operative</td>
		</tr>
		<tr>
			<td>0.9% Normal Saline, 1000 mL</td>
			<td>Baxter Healthcare Corp</td>
			<td>0338-0049-04</td>
			<td>Medication, Intra-Operative</td>
		</tr>
		<tr>
			<td>0.9% Normal Saline, 500 mL</td>
			<td>Baxter Healthcare Corp.,</td>
			<td>&#160;0338-0049-03</td>
			<td>Medication, Chronic PH</td>
		</tr>
		<tr>
			<td>16 mm Heavy Duty Occluder with actuating tubing</td>
			<td>Access Technologies</td>
			<td>&#160;OC-16HD</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>3-mL Skin Prep Applicator</td>
			<td>Medline</td>
			<td>&#160;MDF260400</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>70% isopropyl alcohol prep pads</td>
			<td>Medline</td>
			<td>MDS090670</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Adhesive bandage tape</td>
			<td>Patterson Veterinary</td>
			<td></td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Adson forceps</td>
			<td>V. Mueller</td>
			<td>NL1400</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Allis tissue forceps</td>
			<td>V. Mueller</td>
			<td>CH1560</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Aortic clamp, straight (bainbridge forceps)</td>
			<td>V. Mueller</td>
			<td>SU6001</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Backhaus towel forceps</td>
			<td>V. Mueller</td>
			<td>SU2900</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Bags, Infusion: Nonsterile Novaplus Infusion Bag, 500 mL</td>
			<td>Medline</td>
			<td>TCV4005H</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Berry sternal needle holder</td>
			<td>V. Mueller</td>
			<td>CH2540</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Blades, Electrode: Electrode Blade, 6.5&#34;, with 0.24 cm Shaft</td>
			<td>Medline</td>
			<td>&#160;VALE15516</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Blades: Stainless-Steel Sterile Surgical Blade, Size #10</td>
			<td>Medline</td>
			<td>&#160;B-D371210</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Blades: Stainless-Steel Sterile Surgical Blade, Size #11</td>
			<td>Medline</td>
			<td>&#160;B-D371211</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Blades: Stainless-Steel Sterile Surgical Blade, Size #15</td>
			<td>Medline</td>
			<td>&#160;B-D371215</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>BNC Male to BNC Male Cable</td>
			<td>Digi-Key</td>
			<td>415-0198-036</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Castroviejo needle holder</td>
			<td>V. Mueller</td>
			<td>CH8589</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Cefazolin</td>
			<td>Apotex Corp</td>
			<td>60505-6142-0</td>
			<td>Medication, Intra-Operative</td>
		</tr>
		<tr>
			<td>Ceftiofur Crystalline Free Acid</td>
			<td>Zoetis Inc</td>
			<td>54771-5223-1</td>
			<td>Medication, Post-Operative</td>
		</tr>
		<tr>
			<td>Chest Drain, with Dry Suction, Adult-Pediatric</td>
			<td>Medline</td>
			<td>&#160;DEKA6000LFH</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Chest tube passer</td>
			<td>V. Mueller</td>
			<td>CH04189</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>COnfidence Flowprobes for Research (PAU-Series)</td>
			<td>Transonic</td>
			<td>24PAU</td>
			<td>Equipment, Perivascular Flow Probe</td>
		</tr>
		<tr>
			<td>Cooley tangential occlusion clamp</td>
			<td>V. Mueller</td>
			<td>CH6572</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Data Acquisition Hardware</td>
			<td>ADInstruments</td>
			<td>&#160;PowerLab 16/30</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>DeBakey Aorta clamp</td>
			<td>V. Mueller</td>
			<td>CH7247</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>DeBakey multi-purpose clamp</td>
			<td>V. Mueller</td>
			<td>CH7276</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Debakey tissue forceps, 12&#8217;&#8217;</td>
			<td>V. Mueller</td>
			<td>CH5906</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Debakey vascular tissue forceps 7 3/4&#8217;&#8217;</td>
			<td>V. Mueller</td>
			<td>CH5902</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Debakey vascular tissue forceps, 9&#8217;&#8217;</td>
			<td>V. Mueller</td>
			<td>CH5904</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Electrosurgical Generator</td>
			<td>Covidien</td>
			<td>&#160;Force FX-C</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Endotracheal Tube, 10mm</td>
			<td>Patterson Veterinary</td>
			<td>07-882-9008</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Enrofloxacin</td>
			<td>Norbrook Laboratories Limited</td>
			<td>55529-152-05</td>
			<td>Medication, Intra-Operative</td>
		</tr>
		<tr>
			<td>Fentanyl Transdermal Patch</td>
			<td>Apotex Corp</td>
			<td>60505-7007-2</td>
			<td>Medication, Pre-Operative</td>
		</tr>
		<tr>
			<td>Ferris smith tissue forceps</td>
			<td>V. Mueller</td>
			<td>SU2510</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Finochietto rib spreaders, large</td>
			<td>V. Mueller</td>
			<td>CH1220-1</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Finochietto rib spreaders, medium</td>
			<td>V. Mueller</td>
			<td>CH1215-1</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Flexsteel ribbon retractor, 1&#8221; x 13&#8221;</td>
			<td>V. Mueller</td>
			<td>SU3340</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Flexsteel ribbon retractor, 2&#8221; x 13&#8221;</td>
			<td>V. Mueller</td>
			<td>SU3346</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Foerster sponge forceps, curved</td>
			<td>V. Mueller</td>
			<td>GL660</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Gauze Sponges: Sterile X-ray Compatible Gauze Sponges, 16-Ply, 4&#34; x 4&#34;</td>
			<td>Medline</td>
			<td>&#160;PRM21430LFH</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Gerald-DeBakey forceps</td>
			<td>V. Mueller</td>
			<td>CH04242</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Glassman Allis</td>
			<td>V. Mueller</td>
			<td>SU6152</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Halsted mosquito forceps</td>
			<td>V. Mueller</td>
			<td>SU2702</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Harken clamp</td>
			<td>V. Mueller</td>
			<td>CH6462</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Heat Therapy Pump</td>
			<td>Gaymar/Stryker</td>
			<td>&#160;TP-400</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Fresenius Kabi,</td>
			<td>&#160;63323-540-31</td>
			<td>Medication, Chronic PH</td>
		</tr>
		<tr>
			<td>Hospira Primary IV Sets, 80&#34;</td>
			<td>Patterson Veterinary</td>
			<td>07-835-0123</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Hypertonic saline 3%</td>
			<td>Baxter Healthcare Corp.,</td>
			<td>&#160;0338-0054-03</td>
			<td>Medication, Chronic PH</td>
		</tr>
		<tr>
			<td>Hypodermic Needle with Bevel and Regular Wall, 20 G x 1&#34;</td>
			<td>Medline</td>
			<td>B-D305175Z</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Interface Cable, Edwards LifeScience Transducer to ADInstruments&#160; Bridge Amplifier</td>
			<td>Fogg System</td>
			<td>0395-2434</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Intravenous Infusion Pump</td>
			<td>Heska</td>
			<td>&#160;Vet/IV 2.2 Infusion Pump</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Patterson Veterinary</td>
			<td>14043-704-06</td>
			<td>Medication, Pre-Operative</td>
		</tr>
		<tr>
			<td>Kantrowitz thoracic clamp, 9-1/2&#8221;</td>
			<td>V. Mueller</td>
			<td>CH1722</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Kelly hemostats</td>
			<td>V. Mueller</td>
			<td>88-0314</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Lidocaine HCl, 2.46%</td>
			<td>PRN Pharmacal,</td>
			<td>&#160;49427-434-04</td>
			<td>Medication, Chronic PH</td>
		</tr>
		<tr>
			<td>Ligaclip Multiple-Clip Appliers by Ethicon</td>
			<td>Medline</td>
			<td>&#160;ETHMCS20</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Loop, Vessel, Mini, Red, 2/pk, Sterile</td>
			<td>Medline</td>
			<td>&#160;DYNJVL12</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Lorna non-perforating towel forceps</td>
			<td>V. Mueller</td>
			<td>SU2937</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Mayo dissecting scissors, curved</td>
			<td>V. Mueller</td>
			<td>SU1826</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Mayo dissecting scissors, straight</td>
			<td>V. Mueller</td>
			<td>SU1821</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Medipore Dress-It Pre-Cut Dressing Covers by 3M</td>
			<td>Medline</td>
			<td>&#160;MMM2955Z</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Meloxicam</td>
			<td>Patterson Veterinary</td>
			<td>14043-909-10</td>
			<td>Medication, Post-Operative</td>
		</tr>
		<tr>
			<td>Mixter thoracic forceps, 9&#8221;</td>
			<td>V. Mueller</td>
			<td>CH1730-003</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Mosquito hemostats</td>
			<td>V. Mueller</td>
			<td>88-0301</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Multi-Channel Research Consoles</td>
			<td>Transonic</td>
			<td>T402/T403</td>
			<td>Equipment, Perivascular Flow Meter</td>
		</tr>
		<tr>
			<td>Multi-Lumen Central Venous Catheterization Kits</td>
			<td>Medline</td>
			<td>&#160;ARW45703XP1AH</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Multi-Parameter Vital Signs Monitor</td>
			<td>Smiths Medical</td>
			<td>&#160;SurgiVet Advisor 3</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Needles: Hypodermic Needle with Regular Bevel, Sterile, 18 G x 1.5&#34;</td>
			<td>Medline</td>
			<td>&#160;B-D305185Z</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>No. 3 knife handle</td>
			<td>V. Mueller</td>
			<td>SU1403-001</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>No. 7 knife handle</td>
			<td>V. Mueller</td>
			<td>SU1407</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Non-Vented Male Luer Cap</td>
			<td>Qosina</td>
			<td>13614</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Octal Bridge Amplifier</td>
			<td>ADInstruments</td>
			<td>&#160;FE228</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Ophthalmic Ointment</td>
			<td>Akorn Animal Health</td>
			<td>59399-162-35</td>
			<td>Medication, Pre-Operative</td>
		</tr>
		<tr>
			<td>Penrose Tubing, 6 mm x 46 cm, 11 mm Flat</td>
			<td>Medline</td>
			<td>&#160;SWD514604H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Perma-Hand Black Braided Silk:&#160; 2-0 SH Taperpoint Needle, Control Release, 30&#34;</td>
			<td>Medline</td>
			<td>&#160; ETHD8552</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Perma-Hand Suture, Black Braided, Size 0, 6 x 30&#8221;</td>
			<td>Medline</td>
			<td>&#160; ETHA306H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Perma-Hand Suture, Black Braided, Size 4-0, 12 x 30&#34;</td>
			<td>Medline</td>
			<td>&#160;ETHA303H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Phenylephrine</td>
			<td>West-Ward</td>
			<td>0641-6142-25</td>
			<td>Medication, Intra-Operative</td>
		</tr>
		<tr>
			<td>Polyhesive Cordless Patient Return Electrodes, Adult</td>
			<td>Medline</td>
			<td>&#160;SWDE7509</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Port-A-Cath Huber Needle, Straight, 22 G x 1-1/2&#34;</td>
			<td>Medline</td>
			<td>AAKM21200724</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>PROLENE Monofilament Suture, Blue, Size 4-0, 36&#34;, Double Arm, RB-1 Needle</td>
			<td>Medline</td>
			<td>&#160;ETHD7143</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>PROLENE Polypropylene Monofilament Suture, Blue, Double-Armed, RB-1 Needle, Size 5-0, 24&#34;</td>
			<td>Medline</td>
			<td>&#160;ETH8555H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Regional Block Needles, 22-gauge</td>
			<td>Medline</td>
			<td>&#160;B-D408348Z</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Schnidt tonsil artery forceps</td>
			<td>V. Mueller</td>
			<td>M01700</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Skin staple extractor</td>
			<td>Medline</td>
			<td>CND3031</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Skin stapler 35 wide, with counter</td>
			<td>Medline</td>
			<td>&#160;STAPLER35W</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Sphygmomanometer</td>
			<td>Medline</td>
			<td></td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Sponge bowl</td>
			<td>V. Mueller</td>
			<td>GE-75</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Sponge, Lap: X-Ray Detectable Sterile Lap Sponge, 18&#34; x 18&#34;, 5/Pack</td>
			<td>Medline</td>
			<td>&#160;MDS241518HH</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Sponge, Peanut: X-Ray Detectable Sterile Peanut Sponge, Small, 3/8&#34;</td>
			<td>Medline</td>
			<td>&#160;MDS72038</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Sterile Disposable Deluxe OR Towel, Blue, 17&#39;&#39; x 27&#39;&#39;, 2/Pack</td>
			<td>Medline</td>
			<td>&#160;MDT2168202</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Sterile Luer-Lock Syringe, 3 mL</td>
			<td>Medline</td>
			<td>SYR103010Z</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Sterile Luer-Lock Syringe, 5 mL</td>
			<td>Medline</td>
			<td>SYR105010Z</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Sterile Surgical Equipment Probe Covers</td>
			<td>Medline</td>
			<td>&#160;DYNJE5930</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Stopcock: 3-Way Stopcock with Handle in OFF Position, Rotating Adaptor Male Collar Fitting, 45 PSI</td>
			<td>Medline</td>
			<td>&#160;DYNJSC301</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Stopcock: 3-Way Stopcock with Handle in OFF Position, Rotating Adaptor Male Collar Fitting, 45 PSI</td>
			<td>Medline</td>
			<td>DYNJSC301</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Subcutaneous Port with 5-French Connector and Blue Boot</td>
			<td>Access Technologies</td>
			<td>CP2AC-5NC</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Super cut metzenbaum dissecting scissors</td>
			<td>V. Mueller</td>
			<td>CH2032-S</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Super cut nelson-metzenbaum dissecting scissors</td>
			<td>V. Mueller</td>
			<td>CH2025-S</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Syringes: Sterile Luer-Lock Syringe, 10 mL</td>
			<td>Medline</td>
			<td>&#160;SYR110010Z</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Thoracic Catheter, Straight, 28 Fr x 20&#34;</td>
			<td>Medline</td>
			<td>SWD570549H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Three-quarter surgical drape</td>
			<td>Medline</td>
			<td>&#160;DYNJP2414H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Tiletamine + Zolazepam</td>
			<td>Zoetis Inc</td>
			<td>54771-9050-1</td>
			<td>Medication, Pre-Operative</td>
		</tr>
		<tr>
			<td>TourniKwik Tourniquet Set with Four 7.5&#34; Bronze-Colored Tubes and 1 Snare, 12 French</td>
			<td>Medline</td>
			<td>&#160;CVR79013</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Transducer clip</td>
			<td>Edwards LifeScience</td>
			<td>TCLIP05</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Trigger Aneroid Gauge (Sphygmomanometer)</td>
			<td>Patterson Veterinary</td>
			<td>07-815-0464</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>TruWave Disposable Pressure Transducer Kits by Edwards Lifesciences</td>
			<td>Medline</td>
			<td>&#160;VSYPX260</td>
			<td>Surgical Disposable and Chronic PH</td>
		</tr>
		<tr>
			<td>TS420 Perivascular Flow Module</td>
			<td>Transonic</td>
			<td>TS420</td>
			<td>Equipment, Perivascular Flow Meter</td>
		</tr>
		<tr>
			<td>Tubing, Suction: Sterile Universal Suction Tubing with Straight Ribbed Connectors, 1/4&#34; x 12&#39;</td>
			<td>Medline</td>
			<td>&#160;OR612</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Tubing: Pressure Monitoring Tubing with Fixed Male Luer Lock and Female Fitting, Low Pressure, 72&#34; L</td>
			<td>Medline</td>
			<td>DYNJPMTBG72MF</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Tubing: Pressure Monitoring Tubing with Fixed Male Luer Lock and Female Fitting, Low Pressure, 72&#34; L</td>
			<td>Medline</td>
			<td>DYNJPMTBG72MF</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Tubular Elastic Dressing Retainer</td>
			<td>Medline</td>
			<td>DERGL711</td>
			<td>Disposable, Chronic PH</td>
		</tr>
		<tr>
			<td>Tuffier rib retractor</td>
			<td>V. Mueller</td>
			<td>CD1101</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Tygon E-3603 Flexible Tubings</td>
			<td>Fisher Scientific</td>
			<td>14-171-227</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>U.S.A retractor</td>
			<td>V. Mueller</td>
			<td>SU3660</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Umbilical Tape, Cotton, 3-Strand, 1/8 x 36&#34;</td>
			<td>Medline</td>
			<td>&#160;ETHU12TH</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Valleylab Button Switch Pencil</td>
			<td>Medline</td>
			<td>&#160;VALE2516H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Vanderbilt deep vessel forceps</td>
			<td>V. Mueller</td>
			<td>CH1687</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Veterinary Anesthesia Machine</td>
			<td>Midmark</td>
			<td>&#160;Matrx VMC</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Veterinary Anesthesia Ventilator</td>
			<td>Hallowell EMC</td>
			<td>&#160;Model 2000</td>
			<td>Equipment</td>
		</tr>
		<tr>
			<td>Vicryl: Undyed Coated Vicryl 0 CT-1 36&#34; Suture</td>
			<td>Medline</td>
			<td>&#160;ETHVCP946H</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Vicryl: Undyed Coated Vicryl 2 TP-1 Taper 54&#34; Suture</td>
			<td>Medline</td>
			<td>&#160;ETHVCP880T</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Vicryl: Undyed Coated Vicryl 2-0 CT-1 18&#34; Suture</td>
			<td>Medline</td>
			<td>&#160;ETHVCP739D</td>
			<td>Surgical Disposable</td>
		</tr>
		<tr>
			<td>Vital crile-wood needle holder, 10-3/8&#8221;</td>
			<td>V. Mueller</td>
			<td>CH2427</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Vital mayo-hegar needle holder, 7-1/4&#8221;</td>
			<td>V. Mueller</td>
			<td>CH2417</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Vital metzenbaum dissecting scissors, 14&#8217;&#8217;</td>
			<td>V. Mueller</td>
			<td>CH2009</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Vital metzenbaum dissecting scissors, 9&#8221;</td>
			<td>V. Mueller</td>
			<td>CH2006</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Vital ryder needle holder, 9&#8221;</td>
			<td>V. Mueller</td>
			<td>CH2510</td>
			<td>Surgical Instrument</td>
		</tr>
		<tr>
			<td>Yankauer, Bulb Tip: Sterile Rigid Yankauer with Bulb Tip, No Vent</td>
			<td>Medline</td>
			<td>&#160;DYND50130</td>
			<td>Surgical Disposable</td>
		</tr>
	</tbody>
</table>

a large animal model for pulmonary hypertension and right ventricular failure: left pulmonary artery ligation and progressive main pulmonary artery banding in sheep

Decompensated right ventricular failure (RVF) in pulmonary hypertension (PH) is fatal, with limited medical treatment options. Developing and testing novel therapeutics for PH requires a clinically relevant large animal model of increased pulmonary vascular resistance and RVF. This manuscript discusses the latest development of the previously published ovine PH-RVF model that utilizes left pulmonary artery (PA) ligation and main PA occlusion. This model of PH-RVF is a versatile platform to control not only the disease severity but also the RV&#39;s phenotypic response.
Adult sheep (60-80 kg) underwent left PA (LPA) ligation, placement of main PA cuff, and insertion of RV pressure monitor. PA cuff and RV pressure monitor were connected to subcutaneous ports. Subjects underwent progressive PA banding twice per week for 9 weeks with sequential measures of RV pressure, PA cuff pressures, and mixed venous blood gas (SvO2). At the initiation and endpoint of this model, ventricular function and dimensions were assessed using echocardiography. In a representative group of 12 animal subjects, RV mean and systolic pressure increased from 28 &#177; 5 and 57 &#177; 7 mmHg at week 1, respectively, to 44 &#177; 7 and 93 &#177; 18 mmHg (mean &#177; standard deviation) by week 9. Echocardiography demonstrated characteristic findings of PH-RVF, notably RV dilation, increased wall thickness, and septal bowing. The longitudinal trend of SvO2 and PA cuff pressure demonstrates that the rate of PA banding can be titrated to elicit varying RV phenotypes. A faster PA banding strategy led to a precipitous decline in SvO2 &#60; 65%, indicating RV decompensation, whereas a slower, more paced strategy led to the maintenance of physiologic SvO2 at 70%-80%. One animal that experienced the accelerated strategy developed several liters of pleural effusion and ascites by week 9. This chronic PH-RVF model provides a valuable tool for studying molecular mechanisms, developing diagnostic biomarkers, and enabling therapeutic innovation to manage RV adaptation and maladaptation from PH.

A representative group of 12 sheep is used to show the efficacy of this model for developing varying degrees of PH-RVF. Among these sheep, the mean PA cuff pressure increased from 32 &#177; 20 mmHg at week 1 to 1002 &#177; 429 mmHg at week 9. This resulted in increasing the RV mean and systolic pressures from 28 &#177; 5 and 57 &#177; 7 mmHg at week 1, respectively, to 44 &#177; 7 and 93 &#177; 18 mmHg by week 9. Furthermore, PA cuff pressure profile was superimposed onto mixed venous oxygen saturation (SvO2) ...

Watch this Scientific Journal Video about A Large Animal Model for Pulmonary Hypertension and Right Ventricular Failure: Left Pulmonary Artery Ligation and Progressive Main Pulmonary Artery Banding in

A Large Animal Model for Pulmonary Hypertension and Right Ventricular Failure: Left Pulmonary Artery Ligation and Progressive Main Pulmonary Artery Banding in Sheep

This manuscript describes the surgical technique and experimental approach to develop severe right ventricular pressure overload to model their adaptive and maladaptive phenotypes.

Decompensated right ventricular failure (RVF) in pulmonary hypertension (PH) is fatal, with limited medical treatment options. Developing and testing ...

This protocol is a large animal model for studying right ventricular adaptation and failure. This model is therefore vital for developing RV-targeted therapeutics to treat pulmonary hypertension. This model progressively increases the right RV after load which can be used as a strategy to titrate the RV adaptation.No previous large animal models have this control over the RV phenotype. Demonstrating that surgical procedure will be Dr.Matthew Bacchetta, the laboratory director and the surgical fellows doctors John Stokes, Kelly Wu and Yatrik Patel. I will be demonstrating the pulmonary artery banding procedure.After anesthetization, begin by shaving the surgical field from the sheep's neck to its upper abdomen. Clean the surgical field off dirt and other contaminants using soap or a scrub brush. Prep the neck in the chest with chlorhexidine or betaine solution and drape the surgical field in a sterile fashion.To obtain media style exposure perform a muscle sparing mini thoracotomy at the left fourth intercostal space with incision length less than eight centimeters. Use a small or medium Finochietto retractor to separate the ribs in a tuffier retractor to sit perpendicular to the Finochietto within the intercostal space which will retract the soft tissue within the intercostal space, improving exposure without injuring the phrenic nerve incise the pericardia interior to it and create a pericardial well with 2/0 silk sutures to expose the main pulmonary artery and right ventricle. Identify the left atrial appendage within the exposure as a landmark for the level of the pulmonary artery bifurcation dissect around the main pulmonary artery and isolate it with a umbilical tape.Perform intrapericardial dissection of the left pulmonary artery and encircle with a umbilical tape. Place a heavy duty silicone vascular occluder around the main pulmonary artery. Using an 0 silk suture on a Keith needle.Secure the end to the vascular occluder together with a U-stitch. Once secured around the main pulmonary artery slide the occluder distally along the main pulmonary artery. To establish a right ventricular pressure line, start by placing a 5/0 monofilament non-absorbable polypropylene purser suture with pledgets surrounding the selected location and see a vascular snare.Prepare the right ventricular pressure line by cutting off the male end of a sterile 36 inch pressure tubing at a 30 degree angle to facilitate insertion through the myocardium. Then use a 2/0 silk tie to mark the pressure line at an optimal depth for placement within the right ventricle. Using a biopsy punch, make a small cardiotomy in the right ventricular outflow tract or RVOT free wall within the previously placed purser string suture.Control the bleeding with manual pressure or by tightening the snare on the purser suture. Insert and secure the cut end of the pressure tubing into the RVOT. Tie down the purse string then secure the purse string to the pressure tubing to secure the pressure line.Extend the RVOT tubing by connecting an additional pressure tubing to the RVOT pressure line. Monitor for the measurement of the baseline, right ventricular pressure. Carefully dissect around the left pulmonary artery Encircle the left pulmonary artery with umbilical tape tying it down to ligate it.Bring the RVOT pressure line and pulmonary artery occluder tubing out of the chest one intercostal space below the thoracotomy incision. To mark sites for indwelling ports form two subdural pockets along the fascia layer on the left dorsum of the sheep as far posteriorly toward the spine as feasible within sterile field. Use a chest tube polar to tunnel the RVOT pressure line and occluder tubing from the chest incision out to the left dorsum port sites.Secure both the occluder tubing and right ventricular pressure line to the port's barb connections. Anchor the occluder and pressure tubing around the port connectors with additional ties. To prevent port migration anchor the ports in three locations around their rims to the underlying fascia with 3/0 polypropylene sutures.Reapproximate the subcutaneous tissue dermis and skin in layers with polyglactin 910 sutures. Flush the RVOT port with five milliliters of heparin sodium. Close the thoracotomy with figure of eight number two polyglactin 910 sutures Then close the pectorals muscle layer with a running number 0 polyglactin 910.Finally, close the subcutaneous tissue in layers of running number 2/0 polyglactin 910 sutures and staple the skin. Prepare two pressure transducers for monitoring, right ventricular and occluder cuff pressures. After the animal is mildly restrained insert the Huber needle from the right ventricular pressure transducer to the right ventricular port.Attach a five milliliter syringe to the three-way stop cock and attempt to draw blood back into the syringe from the right ventricular port. Once the right ventricular pressure line is established, connect the Huber needle from the pulmonary artery cuff transducer. Capture the starting values of rip ventricular and pulmonary artery cuff pressures, noting any drastic changes from previous readings.Slowly inject 3%hypertonic saline into the occluder port while paying attention to rip ventricular and cuff pressures. The once the PA cuff is inflated to the desired amount remove the Huber needle from the cuff port. Flush the right ventricular port with 10 milliliters of saline then flush the port with five milliliters of 1000 units per milliliter of heparin sodium.Among the 12 sheep. The mean pulmonary artery cuff pressure increased with an increase in time from the first to the ninth week. Faster pulmonary artery banding lead to a more rapid decline in venous oxygen saturation.In contrast, those that experienced a more gradual pulmonary artery banding strategy maintain a physiologic range of venous oxygen saturation between 70 and 80%By controlling the pulmonary artery cuff pressure. This model has ability to control and titrate ride ventricular adaptation and phenotype.

a-large-animal-model-for-pulmonary-hypertension-right-ventricular

Research

JoVE Journal

Bioengineering

Autologous Endothelial Progenitor Cell-Seeding Technology and Biocompatibility Testing For Cardiovascular Devices in Large Animal Model

Implantable cardiovascular devices are manufactured from artificial materials (e.g. titanium (Ti), expanded polytetrafluoroethylene), which pose the risk of thromboemboli formation1,2,3. We have developed a method to line the inside surface of Ti tubes with autologous blood-derived human or porcine endothelial progenitor cells (EPCs)4. By implanting Ti tubes containing a confluent layer of porcine EPCs in the inferior vena cava (IVC) of pigs, we tested the improved biocompatibility of the cell-seeded surface in the prothrombotic environment of a large animal model and compared it to unmodified bare metal surfaces5,6,7 (Figure 1). This method can be used to endothelialize devices within minutes of implantation and test their antithrombotic function in vivo. 
Peripheral blood was obtained from 50 kg Yorkshire swine and its mononuclear cell fraction cultured to isolate EPCs4,8. Ti tubes (9.4 mm ID) were pre-cut into three 4.5 cm longitudinal sections and reassembled with heat-shrink tubing. A seeding device was built, which allows for slow rotation of the Ti tubes. 
We performed a laparotomy on the pigs and externalized the intestine and urinary bladder. Sharp and blunt dissection was used to skeletonize the IVC from its bifurcation distal to the right renal artery proximal. The Ti tubes were then filled with fluorescently-labeled autologous EPC suspension and rotated at 10 RPH x 30 min to achieve uniform cell-coating9. After administration of 100 USP/ kg heparin, both ends of the IVC and a lumbar vein were clamped. A 4 cm veinotomy was performed and the device inserted and filled with phosphate-buffered saline. As the veinotomy was closed with a 4-0 Prolene running suture, one clamp was removed to de-air the IVC. At the end of the procedure, the fascia was approximated with 0-PDS (polydioxanone suture), the subcutaneous space closed with 2-0 Vicryl and the skin stapled closed. 
After 3 - 21 days, pigs were euthanized, the device explanted en-block and fixed. The Ti tubes were disassembled and the inner surfaces imaged with a fluorescent microscope.
We found that the bare metal Ti tubes fully occluded whereas the EPC-seeded tubes remained patent. Further, we were able to demonstrate a confluent layer of EPCs on the inside blood-contacting surface.
Concluding, our technology can be used to endothelialize Ti tubes within minutes of implantation with autologous EPCs to prevent thrombosis of the device. Our surgical method allows for testing the improved biocompatibility of such modified devices with minimal blood loss and EPC-seeded surface disruption.

A method for seeding titanium blood-contacting biomaterials with autologous cells and testing biocompatibility is described. This method uses endothelial progenitor cells and titanium tubes, seeded within minutes of surgical implantation into porcine venae cavae. This technique is adaptable to many other implantable biomedical devices.

Implantable cardiovascular devices are manufactured from artificial materials (e.g. titanium (Ti), expanded polytetrafluoroethylene), which pose the risk ...

Measuring Left Ventricular Pressure in Late Embryonic and Neonatal Mice

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable around embryonic day (E) 8.51. Systolic left ventricular (LV) pressure is 2 mmHg at E9.5 and 11 mmHg at E14.52. At these mid-embryonic stages, the LV is clearly visible through the chest wall for invasive pressure measurements because the ribs and skin are not fully developed. Between E14.5 and birth (approximately E21) imaging methods must be used to view the LV. After birth, mean arterial pressure increases from 30 - 70 mmHg from postnatal day (P) 2 - 353. Beyond P20, arterial pressure can be measured with solid-state catheters (i.e. Millar or Scisense). Before P20, these catheters are too big for developing mouse arteries and arterial pressure must be measured with custom pulled plastic catheters attached to fluid-filled pressure transducers3 or glass micropipettes attached to servo null pressure transducers4.
Our recent work has shown that the greatest increase in blood pressure occurs during the late embryonic to early postnatal period in mice5-7. This large increase in blood pressure may influence smooth muscle cell (SMC) phenotype in developing arteries and trigger important mechanotransduction events. In human disease, where the mechanical properties of developing arteries are compromised by defects in extracellular matrix proteins (i.e. Marfan's Syndrome8 and Supravalvular Aortic Stenosis9) the rapid changes in blood pressure during this period may contribute to disease phenotype and severity through alterations in mechanotransduction signals. Therefore, it is important to be able to measure blood pressure changes during late embryonic and neonatal periods in mouse models of human disease.
We describe a method for measuring LV pressure in late embryonic (E18) and early postnatal (P1 - 20) mice. A needle attached to a fluid-filled pressure transducer is inserted into the LV under ultrasound guidance. Care is taken to maintain normal cardiac function during the experimental protocol, especially for the embryonic mice. Representative data are presented and limitations of the protocol are discussed.

Measuring left ventricular pressure (LV) in embryonic and neonatal mice is described. Pressure is measured by inserting a needle connected to a fluid-filled transducer into the LV under ultrasound guidance. Care must be taken to maintain normal cardiac function during the experimental protocol.

Blood pressure increases significantly during embryonic and postnatal development in vertebrate animals. In the mouse, blood flow is first detectable ...

Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation

Lung cancer is the leading cause of cancer-related deaths1. Squamous cell and small cell cancers typically arise in association with the conducting airways, whereas adenocarcinomas are typically more peripheral in location. Lung malignancy detection early in the disease process may be difficult due to several limitations: radiological resolution, bronchoscopic limitations in evaluating tissue underlying the airway mucosa and identifying early pathologic changes, and small sample size and/or incomplete sampling in histology biopsies. High resolution imaging modalities, such as optical frequency domain imaging (OFDI), provide non-destructive, large area 3-dimensional views of tissue microstructure to depths approaching 2 mm in real time (Figure 1)2-6. OFDI has been utilized in a variety of applications, including evaluation of coronary artery atherosclerosis6,7 and esophageal intestinal metaplasia and dysplasia6,8-10.
Bronchoscopic OCT/OFDI has been demonstrated as a safe in vivo imaging tool for evaluating the pulmonary airways11-23 (Animation). OCT has been assessed in pulmonary airways16,23 and parenchyma17,22 of animal models and in vivo human airway14,15. OCT imaging of normal airway has demonstrated visualization of airway layering and alveolar attachments, and evaluation of dysplastic lesions has been found useful in distinguishing grades of dysplasia in the bronchial mucosa11,12,20,21. OFDI imaging of bronchial mucosa has been demonstrated in a short bronchial segment (0.8 cm)18. Additionally, volumetric OFDI spanning multiple airway generations in swine and human pulmonary airways in vivo has been described19. Endobronchial OCT/OFDI is typically performed using thin, flexible catheters, which are compatible with standard bronchoscopic access ports. Additionally, OCT and OFDI needle-based probes have recently been developed, which may be used to image regions of the lung beyond the airway wall or pleural surface17.
While OCT/OFDI has been utilized and demonstrated as feasible for in vivo pulmonary imaging, no studies with precisely matched one-to-one OFDI:histology have been performed. Therefore, specific imaging criteria for various pulmonary pathologies have yet to be developed. Histopathological counterparts obtained in vivo consist of only small biopsy fragments, which are difficult to correlate with large OFDI datasets. Additionally, they do not provide the comprehensive histology needed for registration with large volume OFDI. As a result, specific imaging features of pulmonary pathology cannot be developed in the in vivo setting. Precisely matched, one-to-one OFDI and histology correlation is vital to accurately evaluate features seen in OFDI against histology as a gold standard in order to derive specific image interpretation criteria for pulmonary neoplasms and other pulmonary pathologies. Once specific imaging criteria have been developed and validated ex vivo with matched one-to-one histology, the criteria may then be applied to in vivo imaging studies. Here, we present a method for precise, one to one correlation between high resolution optical imaging and histology in ex vivo lung resection specimens. Throughout this manuscript, we describe the techniques used to match OFDI images to histology. However, this method is not specific to OFDI and can be used to obtain histology-registered images for any optical imaging technique. We performed airway centered OFDI with a specialized custom built bronchoscopic 2.4 French (0.8 mm diameter) catheter. Tissue samples were marked with tissue dye, visible in both OFDI and histology. Careful orientation procedures were used to precisely correlate imaging and histological sampling locations. The techniques outlined in this manuscript were used to conduct the first demonstration of volumetric OFDI with precise correlation to tissue-based diagnosis for evaluating pulmonary pathology24. This straightforward, effective technique may be extended to other tissue types to provide precise imaging to histology correlation needed to determine fine imaging features of both normal and diseased tissues.

Optical Frequency Domain Imaging of Ex vivo Pulmonary Resection Specimens: Obtaining One to One Image to Histopathology Correlation

A method to image ex vivo pulmonary resection specimens with optical frequency domain imaging (OFDI) and obtain precise correlation to histology is described, which is essential to developing specific OFDI interpretation criteria for pulmonary pathology. This method is applicable to other tissue types and imaging techniques to obtain precise imaging to histology correlation for accurate image interpretation and assessment. Imaging criteria established with this technique would then be applicable to image assessment in future in vivo studies.

Lung cancer is the leading cause of cancer-related deaths1. Squamous cell and small cell cancers typically arise in association with the conducting ...

Experimental Investigation of Secondary Flow Structures Downstream of a Model Type IV Stent Failure in a 180&#176; Curved Artery Test Section

The arterial network in the human vasculature comprises of ubiquitously present blood vessels with complex geometries (branches, curvatures and tortuosity). Secondary flow structures are vortical flow patterns that occur in curved arteries due to the combined action of centrifugal forces, adverse pressure gradients and inflow characteristics. Such flow morphologies are greatly affected by pulsatility and multiple harmonics of physiological inflow conditions and vary greatly in size-strength-shape characteristics compared to non-physiological (steady and oscillatory) flows 1 - 7.
Secondary flow structures may ultimately influence the wall shear stress and exposure time of blood-borne particles toward progression of atherosclerosis, restenosis, sensitization of platelets and thrombosis 4 - 6, 8 - 13. Therefore, the ability to detect and characterize these structures under laboratory-controlled conditions is precursor to further clinical investigations.
A common surgical treatment to atherosclerosis is stent implantation, to open up stenosed arteries for unobstructed blood flow. But the concomitant flow perturbations due to stent installations result in multi-scale secondary flow morphologies 4 - 6. Progressively higher order complexities such as asymmetry and loss in coherence can be induced by ensuing stent failures vis-&#224;-vis those under unperturbed flows 5. These stent failures have been classified as &#34;Types I-to-IV&#34; based on failure considerations and clinical severity 14.
This study presents a protocol for the experimental investigation of the complex secondary flow structures due to complete transverse stent fracture and linear displacement of fractured parts (&#34;Type IV&#34;) in a curved artery model. The experimental method involves the implementation of particle image velocimetry (2C-2D PIV) techniques with an archetypal carotid artery inflow waveform, a refractive index matched blood-analog working fluid for phase-averaged measurements 15 - 18. Quantitative identification of secondary flow structures was achieved using concepts of flow physics, critical point theory and a novel wavelet transform algorithm applied to experimental PIV data 5, 6, 19 - 26.

Stent implants in stenosed arterial curvatures are prone to "Type IV" failures involving the complete transverse fracture of stents and linear displacement of the fractured parts. We present a protocol for detection of secondary flow (vortical) structures in a curved artery model, downstream of clinically relevant "Type IV" stent failures.

The arterial network in the human vasculature comprises of ubiquitously present blood vessels with complex geometries (branches, curvatures and ...

Electric Cell-substrate Impedance Sensing for the Quantification of Endothelial Proliferation, Barrier Function, and Motility

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. To this end, cells are grown in special culture chambers on top of opposing, circular gold electrodes. A constant small alternating current is applied between the electrodes and the potential across is measured. The insulating properties of the cell membrane create a resistance towards the electrical current flow resulting in an increased electrical potential between the electrodes. Measuring cellular impedance in this manner allows the automated study of cell attachment, growth, morphology, function, and motility. Although the ECIS measurement itself is straightforward and easy to learn, the underlying theory is complex and selection of the right settings and correct analysis and interpretation of the data is not self-evident. Yet, a clear protocol describing the individual steps from the experimental design to preparation, realization, and analysis of the experiment is not available. In this article the basic measurement principle as well as possible applications, experimental considerations, advantages and limitations of the ECIS system are discussed. A guide is provided for the study of cell attachment, spreading and proliferation; quantification of cell behavior in a confluent layer, with regard to barrier function, cell motility, quality of cell-cell and cell-substrate adhesions; and quantification of wound healing and cellular responses to vasoactive stimuli. Representative results are discussed based on human microvascular (MVEC) and human umbilical vein endothelial cells (HUVEC), but are applicable to all adherent growing cells.

This protocol reviews Electric Cell-substrate Impedance Sensing, a method to record and analyze the impedance spectrum of adherent cells for the quantification of cell attachment, proliferation, motility, and cellular responses to pharmacological and toxic stimuli. Detection of endothelial permeability and assessment of cell-cell and cell-substrate contacts are emphasized.

Electric Cell-substrate Impedance Sensing (ECIS) is an in vitro impedance measuring system to quantify the behavior of cells within adherent cell layers. ...

A Microfluidic Model of Biomimetically Breathing Pulmonary Acinar Airways

Quantifying respiratory flow characteristics in the pulmonary acinar depths and how they influence inhaled aerosol transport is critical towards optimizing drug inhalation techniques as well as predicting deposition patterns of potentially toxic airborne particles in the pulmonary alveoli. Here, soft-lithography techniques are used to fabricate complex acinar-like airway structures at the truthful anatomical length-scales that reproduce physiological acinar flow phenomena in an optically accessible system. The microfluidic device features 5 generations of bifurcating alveolated ducts with periodically expanding and contracting walls. Wall actuation is achieved by altering the pressure inside water-filled chambers surrounding the thin PDMS acinar channel walls both from the sides and the top of the device. In contrast to common multilayer microfluidic devices, where the stacking of several PDMS molds is required, a simple method is presented to fabricate the top chamber by embedding the barrel section of a syringe into the PDMS mold. This novel microfluidic setup delivers physiological breathing motions which in turn give rise to characteristic acinar air-flows. In the current study, micro particle image velocimetry (&#181;PIV) with liquid suspended particles was used to quantify such air flows based on hydrodynamic similarity matching. The good agreement between &#181;PIV results and expected acinar flow phenomena suggest that the microfluidic platform may serve in the near future as an attractive in vitro tool to investigate directly airborne representative particle transport and deposition in the acinar regions of the lungs.

Soft-lithography was utilized to produce a representative true-scale model of pulmonary alveolated airways that expand and contract periodically, mimicking physiological breathing motion. This platform recreates respiratory acinar flows on a chip, and is anticipated to facilitate experimental investigation of inhaled aerosol dynamics and deposition in the pulmonary acinus.

Quantifying respiratory flow characteristics in the pulmonary acinar depths and how they influence inhaled aerosol transport is critical towards ...

Tunable Hydrogels from Pulmonary Extracellular Matrix for 3D Cell Culture

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung tissue from porcine, rat, or mouse, the tissue is perfused and submerged in subsequent chemical detergents to remove the cellular debris. Histological comparison of the tissue before and after processing confirms removal of over 95% of double stranded DNA and alpha galactosidase staining suggests the majority of cellular debris is removed. After decellularization, the tissue is lyophilized and then cryomilled into a powder. The matrix powder is digested for 48 hr in an acidic pepsin digestion solution and then neutralized to form the pregel solution. Gelation of the pregel solution can be induced by incubation at 37 &#176;C and can be used immediately following neutralization or stored at 4 &#176;C for up to two weeks. Coatings can be formed using the pregel solution on a non-treated plate for cell attachment. Cells can be suspended in the pregel prior to self-assembly to achieve a 3D culture, plated on the surface of a formed gel from which the cells can migrate through the scaffold, or plated on the coatings. Alterations to the strategy presented can impact gelation temperature, strength, or protein fragment sizes. Beyond hydrogel formation, the hydrogel stiffness may be increased using genipin.

This is a method to create a 3-dimensional cell culture scaffold from pulmonary extracellular matrix. Intact lung is processed into hydrogels that can support the growth of cells in three-dimensions.

Here we present a method for establishing multiple component cell culture hydrogels for in vitro lung cell culture. Beginning with healthy en bloc lung ...

Evaluation of Left Ventricular Structure and Function using 3D Echocardiography

Three-dimensional (3D) quantification of the left ventricle (LV) provides significant added value in terms of diagnostic accuracy and precise risk stratification in various cardiac disorders. Recently, 3D echocardiography became available in routine cardiology practice; however, high-quality image acquisition and subsequent analysis have a steep learning curve. The present article aims to guide the reader through a detailed 3D protocol by presenting tips and tricks and also by highlighting the potential pitfalls to facilitate the widespread but technically sound use of this important technique concerning the LV. First and foremost, we show the acquisition of a high-quality 3D dataset with optimal spatial and temporal resolution. Then, we present the analytical steps toward a detailed quantification of the LV by using one of the most widely applied built-in software. We will quantify LV volumes, sphericity, mass and also systolic function by measuring ejection fraction and myocardial deformation (longitudinal and circumferential strain). We will discuss and provide clinical examples about the essential scenarios where the transition from a conventional echocardiographic approach to a 3D-based quantification is highly recommended.

In this article, we provide a step-by-step acquisition and analysis protocol for the volumetric assessment and speckle-tracking analysis of the left ventricle by 3D echocardiography, particularly focusing on practical aspects that maximize the feasibility of this technique.

Three-dimensional (3D) quantification of the left ventricle (LV) provides significant added value in terms of diagnostic accuracy and precise risk ...

Three-Dimensional Echocardiographic Method for the Visualization and Assessment of Specific Parameters of the Pulmonary Veins

The dimensions of the pulmonary veins are important parameters when planning pulmonary vein isolation (PVI), especially with the cryoballoon ablation technique. Acknowledging the dimensions and anatomical variations of the pulmonary veins (PVs) may improve the outcome of the intervention. Conventional 2D transoesophageal echocardiography can only provide limited data about the dimensions of the PVs; however, 3D echocardiography can further evaluate relevant diameters and areas of the PVs, as well as their spatial relationship to surrounding structures. In previous literature data, parameters influencing the success rate of PVI have already been identified. These are the left lateral ridge, the intervenous ridge, the ostial area of the PVs and the ovality index of the ostium. Proper imaging of the PVs by 3D echocardiography is a technically challenging method. One crucial step is the collection of images. Three individual transducer positions are necessary to visualize the important structures; these are the left lateral ridge, the ostium of the PVs and the intervenous ridge of the left and right PVs. Next, 3D images are acquired and saved as digital loops. These datasets are cropped, which result in the en face views displaying spatial relationships. This step can also be employed to determine the anatomical variations of the PVs. Finally, multiplanar reconstructions are created to measure each individual parameter of the PVs.
Optimal quality and orientation of the acquired images are paramount for the appropriate assessment of PV anatomy. In the present work, we examined the 3D visibility of the PVs and the suitability of the above method in 80 patients. The aim was to provide a detailed outline of the essential steps and potential pitfalls of PV visualization and assessment with 3D echocardiography.

The dimensions of the pulmonary veins (PV) are important parameters when planning pulmonary vein isolation. 2D transoesophageal echocardiography can only provide limited data about the PVs; however, 3D echocardiography can evaluate relevant diameters and areas of the PVs, as well as their spatial relationship to surrounding structures.

The dimensions of the pulmonary veins are important parameters when planning pulmonary vein isolation (PVI), especially with the cryoballoon ablation ...

Evaluating Regional Pulmonary Deposition using Patient-Specific 3D Printed Lung Models

Development of targeted therapies for pulmonary diseases is limited by the availability of preclinical testing methods with the ability to predict regional aerosol delivery. Leveraging 3D printing to generate patient-specific lung models, we outline the design of a high-throughput, in vitro experimental setup for quantifying lobular pulmonary deposition. This system is made with a combination of commercially available and 3D printed components and allows the flow rate through each lobe of the lung to be independently controlled. Delivery of fluorescent aerosols to each lobe is measured using fluorescence microscopy. This protocol has the potential to promote the growth of personalized medicine for respiratory diseases through its ability to model a wide range of patient demographics and disease states. Both the geometry of the 3D printed lung model and the air flow profile setting can be easily modulated to reflect clinical data for patients with varying age, race, and gender. Clinically relevant drug delivery devices, such as the endotracheal tube shown here, can be incorporated into the testing setup to more accurately predict a device&#8217;s capacity to target therapeutic delivery to a diseased region of the lung. The versatility of this experimental setup allows it to be customized to reflect a multitude of inhalation conditions, enhancing the rigor of preclinical therapeutic testing.

We present a high-throughput, in vitro method for quantifying regional pulmonary deposition at the lobe level using CT scan-derived, 3D printed lung models with tunable air flow profiles.