Large Animal Models for Left Ventricular Assist Device Research and Development
Monreal, Gretel*†; Sherwood, Leslie C.*‡; Sobieski, Michael A.*†; Giridharan, Guruprasad A.*†§; Slaughter, Mark S.*†§; Koenig, Steven C.*†§
From the *Cardiovascular Innovation Institute, University of Louisville, Louisville, Kentucky; †Division of Thoracic and Cardiovascular Surgery, University of Louisville, Louisville, Kentucky; ‡Research Resources Facilities, University of Louisville, Louisville, Kentucky; and §Department of Bioengineering, University of Louisville, Louisville, Kentucky.
Submitted for consideration May 2013; accepted for publication in revised form August 2013.
Disclosures: The authors have no conflicts of interest to report.
Reprint Requests: Steven C. Koenig, PhD, Departments of Bioengineering & Surgery, University of Louisville, Cardiovascular Innovation Institute, 302 East Muhammad Ali Blvd, Room 408, Louisville, KY 40202. Email: firstname.lastname@example.org.
In vivo preclinical testing of left ventricular assist devices (LVADs) warrants a large animal model that faithfully simulates human etiology. Although LVAD recipients are in end-stage heart failure (HF), healthy, young animals have served as the experimental platform for most LVAD research and development (R&D) to demonstrate device safety, reliability, and biocompatibility. The rapidly growing HF epidemic, donor heart shortage, and clinical acceptance of LVAD for bridge-to-transplant therapy (BTT) has led to the expanded role of LVAD for destination therapy and bridge-to-recovery therapy. New paradigms for the clinical care of these emerging patient populations are needed. Clinically relevant, robust, and reproducible large animal models of HF are required to demonstrate efficacy, investigate physiologic responses, elucidate genetic, molecular, and cellular mechanism(s), and develop LVAD control strategies. The animal model must be comparable in size, anatomical structure, and phenotype; the technique used to initiate HF must reflect the clinical portrait, should be technically and financially feasible, result in predictable, stable, and irreversible HF, and demonstrate bidirectionality of the remodeling cascade. In this review, large animal species commonly used in cardiac research, techniques used to create chronic HF, and the combined applicability to preclinical LVAD R&D studies are presented.
Heart failure (HF) is a serious health problem affecting 5.7 million Americans. Despite advances in clinical care, prevalence of HF is predicted to increase 25% by the year 2030.1 As a result of the paucity of available donor hearts for transplantation, bridge-to-transplant therapy (BTT) with a left ventricular assist device (LVAD) has been the primary alternative choice for end-stage HF patients. Achievement of comparable patient outcomes has expanded the role of LVAD to long-term support or destination therapy (DT).2 Clinical reports of spontaneous myocardial recovery in LVAD recipients3,4 have also generated great interest in the application of LVAD as platform technology for “rest and recovery” and “adjunctive combination” cellular-based treatment paradigms. The need for temporary support, replacement, and recovery therapies has driven advances in cardiac assist device research and development (R&D), necessitating an investigational evolution in test model platforms, including computer-based, in vitro mock circulatory flow loop, and acute and chronic in vivo models.
Preclinical evaluation of LVAD requires a large animal model that closely replicates the human device recipient. Device and cannula design, fit, surgical approach, performance, and the integration within the cardiovascular system must translate from the animal to the human patient. Human patients are almost always in end-stage HF or cardiogenic shock; nevertheless, healthy animals have been used for most device implant studies. The failing heart, a combination of energetic derangements and intra- and extracellular remodeling, has been shown to undergo reverse remodeling during LVAD support5,6; yet, the concomitant wall thinning, sphericalization, diminished septal elasticity, and augmented myocardial plasticity leaves the heart susceptible to LVAD-induced geometric alterations (septal shift, left ventricular [LV] decompression, right ventricular [RV] overload), which can result in RV failure.7–9 These sequelae are absent in a healthy heart; therefore, careful consideration of a robust and reproducible preclinical model that faithfully represents the human etiology of HF is warranted. In this review, a brief overview of the features of large animals commonly used in device research, techniques for establishing HF in these species, and the application of these models to device research are presented. The optimal combination of an appropriate animal species with a clinically relevant method of creating HF is necessary for elucidating insights into novel cardiac assist devices and their relationship with the failing heart to achieve the goals of improving patient outcomes and restoring their quality of life.
Large Animal Models
Selecting the appropriate animal species for device-related investigations is critical to successful LVAD R&D and testing innovative and clinically relevant hypotheses. The most obvious difference between humans and large animals is their posture and resultant anatomical arrangement: in quadrupeds, the great vessels run along the length of the body parallel to the ground, with the heart as a pendant. Interspecies factors to comparatively consider include the overall size of the animal, anatomical variations (thorax dimensions, cardiac anatomy, coronary anatomy, and perfusion),10–16 and specific structures to be encountered during device implantation (e.g., pericardium,17 great vessels,18 valves,19 papillary muscles,20 blood).21–23 In addition to anatomical characteristics, social aspects to consider include whether the animal can be housed with kin to preserve innate herding and group behaviors, as well as the presence of enrichment items and activities for the study animals (e.g., toys, exercise, treats), which have been shown to reduce the stress of laboratory animals.24 Acclimation of the animals to blood collection, examinations, restraint, and equipment reduces stress to the animals and facilitates execution of the research objectives. This last point is critical in study design and data analysis because physiologic and hemodynamic parameters differ between conscious and sedated/anesthetized animals25,26; therefore, it is important to determine in advance whether the animal will tolerate these interactions or whether conditioning is needed. In this section, a brief overview of the large animals most commonly used in cardiac research is presented.
Cows, most commonly calves, are a useful model for investigations into cardiovascular studies and device research.27–35 Their large size and thoracic cavity make them excellent models for device-related studies, and they possess large peripheral vessels that facilitate vascular access for device cannulation, graft anastomosis, instrumentation, and blood collection. Calves (4–6 months) are more suitable than adult cattle for approximating the human body size (70–100 kg) and are physically easier to handle by study personnel. However, they have the potential for significant growth during long-term investigations as they have not yet reached full maturity. Because of this, their cardiac output can quickly exceed human and LVAD ranges and potentially limit LVAD study duration.36 In contrast to the human aortic arch, cattle possess a common brachiocephalic trunk that divides into the subclavian, vertebral, and carotid arteries18: this dissimilar anatomy may influence device cannulation/graft sites.30 The bovine pericardium is thicker than other large animal species but relatively compliant.17 Calves are ruminants and can bloat during prolonged periods of recumbency, despite preoperative fasting. Rumen bloat can compress the vena cava and hinder venous return; therefore, placement of an orogastric (OG) tube during surgical procedures may be necessary to decompress the rumen. The cost of acquisition and maintenance, as well as adequate vivarium and surgical space required for this model, is considerable, and specific study supplies (large animal laryngoscope blades and endotracheal tubes) may be required.
Dogs are one of the most extensively used and widely reported species for animal model development for cardiovascular investigations. They have historically been easy to procure; they have been bred into a wide range of sizes, proportions, and temperaments; and there is a substantial amount of reference literature on their physiology and pathophysiology. The canine myocardium is rich with collateral perfusion, and collateralization is further stimulated by ischemia37; thus, dog models of ischemia-induced injury can be challenging to establish as they often do not demonstrate substantial dysfunction.38 Dogs possess a comparatively thin but stiff pericardium rich in type III collagen.17 In contrast to humans, where 72–85% of hearts are right-dominant,39,40 dogs are left-dominant41 and additionally have only two vessels that branch from the aortic arch (the innominate and left subclavian arteries).42 Dogs possess a more narrow and deep chest cavity, as opposed to the more barrel-shaped chest of humans, which may influence surgical approach or cardiac assist device placement.
Goats have been used for pediatric device research and circulatory support devices designed for small adults. Goats are social and relatively pleasant large animals that are easy to care for and transport. As with other large animals, differing goat breeds provide a spectrum of varying body sizes. Several studies have characterized features of goat coronary anatomy,14,43 thoracic anatomy,44 the aorta,45 and echocardiography parameters,46 contributing useful references for investigators interested in using goats for cardiovascular research. Similar to dogs and in contrast to humans, goats are left-dominant.43 Like cattle, goats are ruminants and may require an OG tube for rumen decompression during surgical procedures. Goats have the potential to transmit zoonotics such as Orf (contagious ecthyma) and Q fever (Coxiella burnetii); therefore, quarantine of the animal may be necessary before study entrance.
Pigs are a popular species for animal model development in cardiovascular research because of their ease of procurement. The porcine aortic valve is most similar to humans19 and has been a critical component of transplantation research. Porcine coronary artery distribution and perfusion is right-dominant as in humans10,11; however, there are some cardiac anatomical differences (including the presence of only two pulmonary vein ostia in the left atrium and sharper angles that the vena cava enter the right atrium) that have been documented in detail by Crick et al.47 Literature on intrinsic coronary collaterals and angiogenic formation in pigs is mixed, with some studies reporting the presence48 and absence49 of collaterals. The peripheral vessels of pigs are small and deep, presenting some challenges in vascular access particular in studies where repeated procedures or sampling is involved. Conscious pigs can be challenging to interact with during examinations (e.g., physical examinations, body temperature acquisition, echocardiography) than other large animal species because of their stature and tendency to flee; however, they can be trained with positive reinforcement techniques.
Sheep are commonly used in cardiac research as their heart size and hemodynamic responses are comparable with humans. Similar to humans, sheep do not have a prevalent network of coronary collaterals; thus, infarction-based studies result in distinct injuries with sharp border zone regions of ischemic versus perfused territory. The coronary sinus of sheep drains the LV (and none of the RV)50; therefore, investigations into LV-exclusive energetics and myocardial substrate utilization can be performed.51 In contrast to humans, sheep are left-dominant, and the left and right coronaries perfuse the LV and RV, respectively, with only minor overlap.52 As with calves and goats, sheep are ruminants and may require an OG tube for rumen decompression before surgical studies. Sheep often have substantial amounts of wool and need to be shorn regularly, particularly in long-term studies or where body weight or conditioning plays a role in the data. Similar to goats, sheep can also transmit zoonotic diseases and may require a period of prestudy quarantine.
Large Animal Models of Heart Failure
Determining the appropriate technique to create a clinically relevant large animal HF model for LVAD R&D is critical to the success of the study. Most LVAD implantation studies have been performed in healthy animal models.33–35,52–55 Only a few studies have used chronically failing models.8,51,56–58 Healthy animals can adjust to LVAD unloading by reducing flow through their heart8,59; thus, LVAD implantation in HF models may yield different results than those in healthy animals. There are a myriad of well-established techniques for creating HF in large animals, including pacing, which is nonischemic and has been shown to reverse upon cessation,60 direct current shock, which triggers myocyte necrosis coupled with heart block,61 and cardiotoxins.56,62–64 Cardiotoxin models have been used in LVAD research56; however, the agents affect the whole animal, are difficult to restrict to the heart (let alone specific targeted regions of the heart), and instigate dysfunction via oxidative damage and transmembrane ion alterations,65,66 which is not reflective of human HF.
Coronary embolization is achieved by injecting material into the coronary circulation to produce numerous focal microinfarcts resulting in a diffuse area of ischemia. In humans, coronary embolization, primarily platelet and cholesterol debris from shedding plaques, is also a complication from percutaneous coronary interventions (PCI)67 and suspected to be an instigator of subclinical myocardial infarction (MI) and non-ST-segment MI.68,69 In large animals, coronary embolization can be performed minimally invasively via catheters introduced from the groin or neck. This is advantageous to LVAD studies, as the resultant small incision minimizes pain and complications and preserves the native chest environment for future device implants. In addition, strategic targeting of specific myocardial regions can be performed based on the vessel(s) embolized: investigators have targeted the right coronary artery,70 the circumflex coronary artery,71 the left main coronary artery,72 and multiples coronaries.28,73 This technique has been performed using a variety of agents, including microspheres,28,71–73 sponge,74 plugs,75 platelet aggregates,76 mercury,70 and coils.77 Embolization results in permanent occlusion of the coronary microvessels, with the magnitude of injury influenced by the size or volume of agent delivered. During emboli administration, there is a risk that emboli will enter the systemic circulation instead of the coronary artery; therefore, end-organ evaluation at necropsy for the presence of damage may be warranted. In addition, multiple embolization procedures are sometimes required to achieve a predetermined degree of cardiac dysfunction.
Coronary embolization models have been used in several LVAD studies and are useful to device research when an ischemic, naturally progressing HF is desired. Goldstein et al.8 and Monreal and Gerhardt51 used a chronic sheep embolization model and demonstrated reductions in myocardial oxygen consumption and alterations in arteriovenous electrolytes during acute partial LVAD support. Bartoli et al.56 implanted the HeartMate II (Thoratec, Pleasanton CA) and HeartWare ventricular assist device (HeartWare, Miami Lakes FL) in a chronic calf embolization model and found that pulsatile LVAD maintained normal hemodynamics better than continuous LVAD. Ghodsizad et al.57 used a chronic sheep embolization model and demonstrated successful off-pump implantation of the HeartWare MVAD. Kishimoto et al.78 and Umeki et al.79 implanted the Sun Medical EVAHEART (Nagano, Japan) in an acute goat embolization model and showed that using a delayed copulse drive mode provided forward flow with animation of the aortic valve.
Coronary ligation is a popular means of inducing MI and can be performed on a selected coronary artery or arteries to infarct targeted regions of myocardium or specifically induce ischemic-based mitral valve dysfunction.80,81 Depending on how proximal the ligation is or the absence of coronary collaterals, the resultant infarct can be very large (as demonstrated by Markovitz et al.)52; this may be useful to investigators studying LVAD support in the acute timeframe post-MI or post-PCI. In pigs and sheep, coronary ligation yields infarcts with sharp, distinct border zones, which are useful for studies examining differences between infarcted myocardium, border zone regions, and remote tissue areas within the same heart.82 Kim et al.38 reported that coronary ligation resulted in less-pronounced hemodynamic and myocardial alterations in dogs when compared with sheep, possibly because of collateral vessels maintaining perfusion to myocardium distal to the ligation.
Although coronary ligation results in an immediate ischemic insult to the heart, placement of an ameroid constrictor around a coronary artery results in gradual occlusion over a period of time owing to hygroscopic compressive expansion of the constrictor.83 The placement of coronary ligation materials and ameroid constrictors requires invasive surgical procedures. The thoracotomy or sternotomy performed results in a significant chest incision, which requires attentive care of the animal to minimize pain, stress, and the potential for infection and disrupts the chest cavity and pericardium for future device implants.
In contrast to external constriction and to avoid major thoracic surgery, coronary flow can be occluded internally via balloon catheters passed minimally invasively into the coronary artery of choice under fluoroscopic guidance. This technique has been used to establish large animal models of ischemic HF for preclinical gene therapy84 and infarct mapping.85
Coronary ligation has been used to establish animal models for device research. Yoshitake et al.86 implanted the Impella LD (Abiomed, Danvers, MA) alone and in combination with beta-blockade in pigs that underwent coronary ligation (and then reperfusion) just before device implants and demonstrated reductions in infarct size and mortality rate. To investigate cardiac unloading and recovery potential from ventricular defibrillation, Kawashima et al.87 used a dog model of acute coronary artery ligation followed by either ECMO or Impella LD (Abiomed) support and reported superior reductions in pressure-volume area with Impella support. Meyns et al.88 investigated the influence of Impella (Abiomed) support on infarct size in healthy sheep that underwent coronary ligation and reperfusion and demonstrated reductions in both myocardial oxygen consumption and infarct size. Clark et al.89 used a sheep model of coronary ligation performed at the time of LVAD implant and demonstrated feasibility and efficacy of AB-180 implantation (CardiacAssist, Pittsburgh, PA).
Pressure overload induces cardiac dysfunction that progresses into compensatory hypertrophy and eventual HF via heightened afterload. The primary technique for creating pressure-overload HF in large animals is aortic constriction/banding, at the supraaortic position (coronary pressure overload),90 the ascending aorta,91,92 or descending aorta.93 An advantage to pressure-overload models is that the constriction can be reversed with either removal of the band or alleviation of the hydraulic constrictor. Although these techniques result in progressive cardiac dysfunction, the initial insult is nonischemic, the effects of pressure overload are being experienced by a healthy heart, and depending on the degree of stenosis created, the physiological impact may be pronounced and immediate. A disadvantage to use this model as a platform for future device research is that it is invasive and requires a thoracotomy to access the aorta; therefore, the animal has already undergone one major survival surgery procedure, and substantial scarring may be present that could limit or alter the surgical approach to LVAD implantation. To our knowledge, no pressure overload animal models have been used for subsequent LVAD implant studies.
Volume overload of the heart is used to simulate the progressive response of the myocardium, vasculature, and neurohormonal system to augmented preload as a result of pump failure or regurgitation and trigger the compensatory progression from acute cardiac dysfunction to hypertrophy to dilated cardiomyopathy. Several experimental techniques have been published with varied degrees of invasiveness, including minimally invasive creation of mitral valve regurgitation by chordae rupture or leaflet damage,94 ligation of the obtuse marginal artery to induce ischemic mitral regurgitation,80 and surgical creation of an arteriovenous fistula (via femoral95 or aortocaval shunt).96 Similar to the disadvantages of pressure-overload models, volume overload is nonischemic, experienced by an otherwise healthy heart and cardiorenal system, and can result in sudden and marked dysfunction. This model has been used as a platform for device investigations. Using a chronic sheep model of volume overload (mitral regurgitation via chordae rupture) as a pilot study, Tuzun et al.58 demonstrated reverse remodeling after 11 weeks of LVAD support (Levitronix CentriMag; Levitronix LLC, Waltham, MA; or HeartWare LVAD; HeartWare).
The need for suitable preclinical HF large animal models is great as the imbalance between available donor hearts coupled with the expanding epidemic of HF requires new paradigms for the clinical care of HF patients. At present, LVAD therapy is generally reserved for New York Heart Association class IV patients; however, as Interagency Registry for Mechanically Assisted Circulatory Support data indicate, an increasing number of patients are undergoing LVAD implantation as DT.97 This trend reflects the advances in medical management and assist device R&D but nevertheless forewarns of a growing future population that heralds the need for continued design optimization and evolving technologies. In vivo testing of an LVAD requires two noteworthy components: 1) an animal comparable in size as to represent an adult human device recipient; 2) a means of recreating the clinical etiology of human HF in that animal. The ideal model should represent a human HF patient as closely as possible, be technically and financially possible to perform and maintain, result in predictable and stable HF that is uniform across animals, and demonstrate the bidirectionality of the remodeling cascade in HF following cardiac unloading. The trend toward smaller and less invasive devices coupled with the potential benefits of partial ventricular unloading in both HF animals8 and human patients98,99 may yield insights into future LVAD designs for the pediatric population or treatment of earlier stage HF. Advances in biomedicine may 1 day lead to the successful creation of transgenic large animal HF models. Thus, refinement of preclinical HF animal models for device research is one step toward the goal of reducing morbidities and improving patient care and quality of life.
1. Roger VL, Go AS, Lloyd-Jones DM, et al.American Heart Association Statistics Committee and Stroke Statistics Subcommittee. Executive summary: Heart disease and stroke statistics–2012 update: A report from the American Heart Association. Circulation. 2012;125:188–197
2. Wilhelm MJ, Ruschitzka F, Falk V. Destination therapy—Time for a paradigm change in heart failure therapy. Swiss Med Wkly. 2013;143:w13729
3. Yoshioka D, Toda K, Sakaguchi T, et al. Initial report of bridge to recovery in a patient with DuraHeart LVAD. J Artif Organs. 2013;16:386–388
4. Schweiger M, Potapov E, Vierecke J, Stepanenko A, Hetzer R, Krabatsch T. Expeditious and less traumatic explantation of a heartware LVAD after myocardial recovery. ASAIO J. 2012;58:542–544
5. Birks EJ, George RS. Molecular changes occurring during reverse remodelling following left ventricular assist device support. J Cardiovasc Transl Res. 2010;3:635–642
6. Drakos SG, Kfoury AG, Selzman CH, et al. Left ventricular assist device unloading effects on myocardial structure and function: Current status of the field and call for action. Curr Opin Cardiol. 2011;26:245–255
7. Puwanant S, Hamilton KK, Klodell CT, et al. Tricuspid annular motion as a predictor of severe right ventricular failure after left ventricular assist device implantation. J Heart Lung Transplant. 2008;27:1102–1107
8. Goldstein AH, Monreal G, Kambara A, et al. Partial support with a centrifugal left ventricular assist device reduces myocardial oxygen consumption in chronic, ischemic heart failure. J Card Fail. 2005;11:142–151
9. Topilsky Y, Hasin T, Oh JK, et al. Echocardiographic variables after left ventricular assist device implantation associated with adverse outcome. Circ Cardiovasc Imaging. 2011;4:648–661
10. Weaver ME, Pantely GA, Bristow JD, Ladley HD. A quantitative study of the anatomy and distribution of coronary arteries in swine in comparison with other animals and man. Cardiovasc Res. 1986;20:907–917
11. Sahni D, Kaur GD, Jit H, Jit I. Anatomy and distribution of coronary arteries in pig in comparison with man. Indian J Med Res. 2008;127:564–570
12. Trumble DR, Magovern JA. Comparison of dog and pig models for testing substernal cardiac compression devices. ASAIO J. 2004;50:188–192
13. Dondelinger RF, Ghysels MP, Brisbois D, et al. Relevant radiological anatomy of the pig as a training model in interventional radiology. Eur Radiol. 1998;8:1254–1273
14. Besoluk K, Tipirdamaz S. Comparative macroanatomic investigations of the venous drainage of the heart in Akkaraman sheep and Angora goats. Anat Histol Embryol. 2001;30:249–252
15. Hill AJ, Iazzio PAIaizzo PA. Comparative Cardiac Anatomy, Handbook of Cardiac Anatomy, Physiology, and Devices. 2009 New York Springer
16. Michaelsson M, Ho SYMichaelsson M, Ho SY. Introduction: Normal hearts—A comparison, Congenital Heart Malformations in Mammals. 2000 London Imperial College Press
17. Naimark WA, Lee JM, Limeback H, Cheung DT. Correlation of structure and viscoelastic properties in the pericardia of four mammalian species. Am J Physiol. 1992;263(4 pt 2):H1095–H1106
18. Layton KF, Kallmes DF, Kaufmann TJ. Use of the ulnar artery as an alternative access site for cerebral angiography. AJNR Am J Neuroradiol. 2006;27:2073–2074
19. Sands MP, Rittenhouse EA, Mohri H, Merendino KA. An anatomical comparison of human pig, calf, and sheep aortic valves. Ann Thorac Surg. 1969;8:407–414
20. Ozbag D, Gumusalan Y, Demirant A. The comparative investigation of morphology of papillary muscles of left ventricle in different species. Int J Clin Pract. 2005;59:529–536
21. Mueller XM, Tevaearai HT, Jegger D, Tucker O, von Segesser LK. Hemolysis and hematology profile during perfusion: Inter-species comparison. Int J Artif Organs. 2001;24:89–94
22. Plasenzotti R, Stoiber B, Posch M, Windberger U. Red blood cell deformability and aggregation behaviour in different animal species. Clin Hemorheol Microcirc. 2004;31:105–111
23. Goodman SL. Sheep, pig, and human platelet-material interactions with model cardiovascular biomaterials. J Biomed Mater Res. 1999;45:240–250
24. Morris CL, Grandin T, Irlbeck NA. Companion Animals Symposium: Environmental enrichment for companion, exotic, and laboratory animals. J Anim Sci. 2011;89:4227–4238
25. Takaseya T, Fujiki M, Shiose A, et al. Hemodynamic differences between the awake and anesthetized conditions in normal calves. J Artif Organs. 2012;15:225–230
26. Muggenburg BA, Mauderly JL. Cardiopulmonary function of awake, sedated, and anesthetized beagle dogs. J Appl Physiol. 1974;37:152–157
27. Rankin JS, Conger JL, Tuzun E, et al. In vivo
testing of an intra-annular aortic valve annuloplasty ring in a chronic calf model. Eur J Cardiothorac Surg. 2012;42:149–154
28. Bartoli CR, Sherwood LC, Giridharan GA, et al. Bovine model of chronic ischemic cardiomyopathy: implications for ventricular assist device research. ASAIO J. 2013;59:221–229
29. Koenig SC, Litwak KN, Giridharan GA, et al. Acute hemodynamic efficacy of a 32-ml subcutaneous counterpulsation device in a calf model of diminished cardiac function. ASAIO J. 2008;54:578–584
30. Tuzun E, Narin C, Gregoric ID, Cohn WE, Frazier OH. Ventricular assist device outflow-graft site: effect on myocardial blood flow. J Surg Res. 2011;171:71–75
31. Butler KC, Maher TR, Borovetz HS, et al. Development of an axial flow blood pump LVAS. ASAIO J. 1992;38:M296–M300
32. Kameneva MV, Watach MJ, Litwak P, et al. Chronic animal health assessment during axial ventricular assistance: Importance of hemorheologic parameters. ASAIO J. 1999;45:183–188
33. Farrar DJ, Bourque K, Dague CP, Cotter CJ, Poirier VL. Design features, developmental status, and experimental results with the HeartMate III centrifugal left ventricular assist system with a magnetically levitated rotor. ASAIO J. 2007;53:310–315
34. Slaughter MS, Giridharan GA, Tamez D, et al. Transapical miniaturized ventricular assist device: Design and initial testing. J Thorac Cardiovasc Surg. 2011;142:668–674
35. Kihara S, Yamazaki K, Litwak KN, et al. In vivo
evaluation of a MPC polymer coated continuous flow left ventricular assist system. Artif Organs. 2003;27:188–192
36. Chee HK, Tuzun E, Ferrari M, et al. Baseline hemodynamic and echocardiographic indices in anesthetized calves. ASAIO J. 2004;50:267–271
37. Scheuer J. Effects of physical training on myocardial vascularity and perfusion. Circulation. 1982;66:491–495
38. Kim WG, Shin YC, Hwang SW, Lee C, Na CY. Comparison of myocardial infarction with sequential ligation of the left anterior descending artery and its diagonal branch in dogs and sheep. Int J Artif Organs. 2003;26:351–357
39. Abuchaim DC, Spera CA, Faraco DL, Ribas Filho JM, Malafaia O. Coronary dominance patterns in the human heart investigated by corrosion casting. Rev Bras Cir Cardiovasc. 2009;24:514–518
40. Andreini D, Mushtaq S, Pontone G, et al. Additional clinical role of 64-slice multidetector computed tomography in the evaluation of coronary artery variants and anomalies. Int J Cardiol. 2010;145:388–390
41. Oliviera CLS, David GS, Carvalho MO, et al. Anatomical indicators of dominance between the coronary arteries of dogs. Int J Morphol. 2011;29:845–849
42. Li W, Xu K, Zhong H, Ni Y, Bi Y. A new unibody branched stent-graft for reconstruction of the canine aortic arch. Eur J Vasc Endovasc Surg. 2012;44:139–144
43. Lipovetsky G, Fenoglio JJ, Gieger M, Srinivasan MR, Dobelle WH. Coronary artery anatomy of the goat. Artif Organs. 1983;7:238–245
44. Ohlerth S, Becker-Birck M, Augsburger H, Jud R, Makara M, Braun U. Computed tomography measurements of thoracic structures in 26 clinically normal goats. Res Vet Sci. 2012;92:7–12
45. Ogeng’o JA, Malek AA, Kiama SG. Structural organisation of tunica intima in the aorta of the goat. Folia Morphol (Warsz). 2010;69:164–169
46. Leroux AA, Farnir F, Moonen ML, Sandersen CF, Deleuze S, Amory H. Repeatability, variability and reference values of pulsed wave Doppler echocardiographic measurements in healthy Saanen goats. BMC Vet Res. 2012;8:190
47. Crick SJ, Sheppard MN, Ho SY, Gebstein L, Anderson RH. Anatomy of the pig heart: Comparisons with normal human cardiac structure. J Anat. 1998;193(pt 1):105–119
48. van den Wijngaard JP, Schulten H, van Horssen P, et al. Porcine coronary collateral formation in the absence of a pressure gradient remote of the ischemic border zone. Am J Physiol Heart Circ Physiol. 2011;300:H1930–H1937
49. Patterson RE, Kirk ES. Analysis of coronary collateral structure, function, and ischemic border zones in pigs. Am J Physiol. 1983;244:H23–H31
50. Goldstein AH, Pacella JJ, Clark RE. Predictable reduction in left ventricular stroke work and oxygen utilization with an implantable centrifugal pump. Ann Thorac Surg. 1994;58:1018–1024
51. Monreal G, Gerhardt MA. Left ventricular assist device support induces acute changes in myocardial electrolytes in heart failure. ASAIO J. 2007;53:152–158
52. Markovitz LJ, Savage EB, Ratcliffe MB, et al. Large animal model of left ventricular aneurysm. Ann Thorac Surg. 1989;48:838–845
53. Schmitto JD, Burkhoff D, Avsar M, et al. Two axial-flow Synergy Micro-Pumps as a biventricular assist device in an ovine animal model. J Heart Lung Transplant. 2012;31:1223–1229
54. Ostadal P, Mlcek M, Holy F, et al. Direct comparison of percutaneous circulatory support systems in specific hemodynamic conditions in a porcine model. Circ Arrhythm Electrophysiol. 2012;5:1202–1206
55. Jassawalla JS, Daniel MA, Chen H, et al. In vitro
and in vivo
testing of a totally implantable left ventricular assist system. ASAIO Trans. 1988;34:470–475
56. Bartoli CR, Giridharan GA, Litwak KN, et al. Hemodynamic responses to continuous versus pulsatile mechanical unloading of the failing left ventricle. ASAIO J. 2010;56:410–416
57. Ghodsizad A, Kar BJ, Layolka P, et al. Less invasive off-pump implantation of axial flow pumps in chronic ischemic heart failure: Survival effects. J Heart Lung Transplant. 2011;30:834–837
58. Tuzun E, Bick R, Kadipasaoglu C, et al. Modification of a volume-overload heart failure model to track myocardial remodeling and device-related reverse remodeling. ISRN Cardiol. 2011;2011:831062
59. Meyns B, Siess T, Nishimura Y, et al. Miniaturized implantable rotary blood pump in atrial-aortic position supports and unloads the failing heart. Cardiovasc Surg. 1998;6:288–295
60. Shinbane JS, Wood MA, Jensen DN, Ellenbogen KA, Fitzpatrick AP, Scheinman MM. Tachycardia-induced cardiomyopathy: A review of animal models and clinical studies. J Am Coll Cardiol. 1997;29:709–715
61. Carlyle PF, Cohn JN. A nonsurgical canine model of chronic left ventricular myocardial dysfunction. Am J Physiol. 1983;244:H769–H774
62. Bartoli CR, Brittian KR, Giridharan GA, Koenig SC, Hamid T, Prabhu SD. Bovine model of doxorubicin-induced cardiomyopathy. J Biomed Biotechnol. 2011;2011:758736
63. Litwak KN, McMahan A, Lott KA, Lott LE, Koenig SC. Monensin toxicosis in the domestic bovine calf: A large animal model of cardiac dysfunction. Contemp Top Lab Anim Sci. 2005;44:45–49
64. Devlin G, Matthews K, McCracken G, et al. An ovine model of chronic stable heart failure. J Card Fail. 2000;6:140–143
65. Chatterjee K, Zhang J, Honbo N, Karliner JS. Doxorubicin cardiomyopathy. Cardiology. 2010;115:155–162
66. Kawada T, Yoshida Y, Sakurai H, Imai S. Myocardial Na+ during ischemia and accumulation of Ca2+ after reperfusion: A study with monensin and dichlorobenzamil. Jpn J Pharmacol. 1992;59:191–200
67. Singh IM, Holmes DR Jr. Myocardial revascularization by percutaneous coronary intervention: Past, present, and the future. Curr Probl Cardiol. 2011;36:375–401
68. Hamm CW, Bassand JP, Agewall S, et al.ESC Committee for Practice Guidelines. ESC Guidelines for the management of acute coronary syndromes in patients presenting without persistent ST-segment elevation: The Task Force for the management of acute coronary syndromes (ACS) in patients presenting without persistent ST-segment elevation of the European Society of Cardiology (ESC). Eur Heart J. 2011;32:2999–3054
69. Heusch G, Kleinbongard P, Böse D, et al. Coronary microembolization: From bedside to bench and back to bedside. Circulation. 2009;120:1822–1836
70. Matangi MF, Cohn JN. Beneficial effects of atrioventricular synchrony in dogs with right coronary artery embolization, and complete heart block. Can J Cardiol. 1987;3:144–147
71. Monreal G, Gerhardt MA, Kambara A, Abrishamchian AR, Bauer JA, Goldstein AH. Selective microembolization of the circumflex coronary artery in an ovine model: Dilated, ischemic cardiomyopathy and left ventricular dysfunction. J Card Fail. 2004;10:174–183
72. Schmitto JD, Ortmann P, Ortomann P, et al. Histological changes in a model of chronic heart failure induced by multiple sequential coronary microembolization in sheep. J Cardiovasc Surg (Torino). 2008;49:533–537
73. Sabbah HN, Stein PD, Kono T, et al. A canine model of chronic heart failure produced by multiple sequential coronary microembolizations. Am J Physiol. 1991;260(4 pt 2):H1379–H1384
74. Sakaguchi G, Sakakibara Y, Tambara K, et al. A pig model of chronic heart failure by intracoronary embolization with gelatin sponge. Ann Thorac Surg. 2003;75:1942–1947
75. Herr MD, McInerney JJ, Copenhaver GL, Morris DL. Coronary artery embolization in closed-chest canines using flexible radiopaque plugs. J Appl Physiol (1985). 1988;64:2236–2239
76. Kwiatkowski P, Sai-Sudhakar C, Philips A, Parthasarathy S, Sun B. Development of a novel large animal model of ischemic heart failure using autologous platelet aggregates. Int J Artif Organs. 2010;33:63–71
77. Peukert D, Laule M, Kaufels N, et al. A minimally invasive method for induction of myocardial infarction in an animal model using tungsten spirals. Int J Cardiovasc Imaging. 2009;25:529–535
78. Kishimoto Y, Takewa Y, Arakawa M, et al. Development of a novel drive mode to prevent aortic insufficiency during continuous-flow LVAD support by synchronizing rotational speed with heartbeat. J Artif Organs. 2013;16:129–137
79. Umeki A, Nishimura T, Takewa Y, et al. Change in myocardial oxygen consumption employing continuous-flow LVAD with cardiac beat synchronizing system, in acute ischemic heart failure models. J Artif Organs. 2013;16:119–128
80. Siefert AW, Rabbah JP, Koomalsingh KJ, et al. In vitro
mitral valve simulator mimics systolic valvular function of chronic ischemic mitral regurgitation ovine model. Ann Thorac Surg. 2013;95:825–830
81. Schmitto JD, Mokashi SA, Lee LS, et al. A novel, innovative ovine model of chronic ischemic cardiomyopathy induced by multiple coronary ligations. Artif Organs. 2010;34:918–922
82. Wu EX, Wu Y, Nicholls JM, et al. MR diffusion tensor imaging study of postinfarct myocardium structural remodeling in a porcine model. Magn Reson Med. 2007;58:687–695
83. Caillaud D, Calderon J, Réant P, et al. Echocardiographic analysis with a two-dimensional strain of chronic myocardial ischemia induced with ameroid constrictor in the pig. Interact Cardiovasc Thorac Surg. 2010;10:689–693
84. Pleger ST, Shan C, Ksienzyk J, et al. Cardiac AAV9-S100A1 gene therapy rescues post-ischemic heart failure in a preclinical large animal model. Sci Transl Med. 2011;3:92ra64
85. Betensky BP, Jauregui M, Campos B, et al. Use of a novel endoscopic catheter for direct visualization and ablation in an ovine model of chronic myocardial infarction. Circulation. 2012;126:2065–2072
86. Yoshitake I, Hata M, Sezai A, et al. The effect of combined treatment with Impella(®) and landiolol in a swine model of acute myocardial infarction. J Artif Organs. 2012;15:231–239
87. Kawashima D, Gojo S, Nishimura T, et al. Left ventricular mechanical support with Impella provides more ventricular unloading in heart failure than extracorporeal membrane oxygenation. ASAIO J. 2011;57:169–176
88. Meyns B, Stolinski J, Leunens V, Verbeken E, Flameng W. Left ventricular support by catheter-mounted axial flow pump reduces infarct size. J Am Coll Cardiol. 2003;41:1087–1095
89. Clark RE, Walters RA, Hughson S, Davis SA Sr, Magovern GJ. Left ventricular support with the implantable AB-180 centrifugal pump in sheep with acute myocardial infarction. ASAIO J. 1998;44:804–811
90. Ghaleh B, Hittinger L, Kim SJ, et al. Selective large coronary endothelial dysfunction in conscious dogs with chronic coronary pressure overload. Am J Physiol. 1998;274(2 pt 2):H539–H551
91. Yarbrough WM, Mukherjee R, Stroud RE, et al. Progressive induction of left ventricular pressure overload in a large animal model elicits myocardial remodeling and a unique matrix signature. J Thorac Cardiovasc Surg. 2012;143:215–223
92. Moorjani N, Catarino P, Trabzuni D, et al. Upregulation of Bcl-2 proteins during the transition to pressure overload-induced heart failure. Int J Cardiol. 2007;116:27–33
93. Hirsch JC, Borton AR, Albayya FP, Russell MW, Ohye RG, Metzger JM. Comparative analysis of parvalbumin and SERCA2a cardiac myocyte gene transfer in a large animal model of diastolic dysfunction. Am J Physiol Heart Circ Physiol. 2004;286:H2314–H2321
94. Kleaveland JP, Kussmaul WG, Vinciguerra T, Diters R, Carabello BA. Volume overload hypertrophy in a closed-chest model of mitral regurgitation. Am J Physiol. 1988;254(6 pt 2):H1034–H1041
95. Omoto T, Aeba R, Katogi T, Ito T, Kawada S. Effects of arteriovenous shunt on ventricular function in dog. Jpn J Thorac Cardiovasc Surg. 1999;47:116–120
96. Fiorillo C, Nediani C, Ponziani V, et al. Cardiac volume overload rapidly induces oxidative stress-mediated myocyte apoptosis and hypertrophy. Biochim Biophys Acta. 2005;1741:173–182
97. Kirklin JK, Naftel DC, Kormos RL, et al. Fifth INTERMACS annual report: Risk factor analysis from more than 6,000 mechanical circulatory support patients. J Heart Lung Transplant. 2013;32:141–156
98. Klotz S, Meyns B, Simon A, et al. Partial mechanical long-term support with the CircuLite Synergy pump as bridge-to-transplant in congestive heart failure. Thorac Cardiovasc Surg. 2010;58(suppl 2):S173–S178
99. Meyns BP, Simon A, Klotz S, et al. Clinical benefits of partial circulatory support in New York Heart Association Class IIIB and Early Class IV patients. Eur J Cardiothorac Surg. 2011;39:693–698
left ventricular assist device; heart failure; animal model; preclinical
Copyright © 2014 by the American Society for Artificial Internal Organs
Highlight selected keywords in the article text.