This work was supported by NIH grants HL111121 (JDM) and TR000954 (JDM).

1. Prepare the Materials and Equipment (Table 1 and Figure 1)
<ol>
	<li>Turn on the ultrasound machine. Enter the animal ID, date, and time (for serial imaging experiments) and other relevant information.</li>
	<li>Use a high-frequency ultrasound transducer, 40 MHz for imaging mice less than ~20 g or 30 MHz for mice greater than ~20 g.</li>
	<li>Connect the platform to the electrocardiogram (ECG) monitor for ECG gating of imaging for certain modalities. 
	NOTE: Critically, this also allows for the instantaneous ca...

<ol>
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M., 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=Recommendations+for+cardiac+chamber+quantification+by+echocardiography+in+adults:+an+update+from+the+American+Society+of+Echocardiography+and+the+European+Association+of+Cardiovascular+Imaging.">Recommendations for cardiac chamber quantification by echocardiography in adults: an update from the American Society of Echocardiography and the European Association of Cardiovascular Imaging.</a> Eur Heart J Cardiovasc Imaging. 16, 233-270 (2015).</li><li>Devereux, R. B., Reichek, N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Echocardiographic+determination+of+left+ventricular+mass+in+man.+Anatomic+validation+of+the+method.">Echocardiographic determination of left ventricular mass in man. Anatomic validation of the method.</a> Circulation. 55, 613-618 (1977).</li><li>Ommen, S. 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G., Lindsey, M. L., Lange, R. A., Aune, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+Assessment+in+Pediatric+Mice:+Strain+Analysis+as+a+Diagnostic+Measurement.">Cardiac Assessment in Pediatric Mice: Strain Analysis as a Diagnostic Measurement.</a> Echocardiography. 31, 375-384 (2014).</li><li>Ferferieva, V., 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=Assessment+of+strain+and+strain+rate+by+two-dimensional+speckle+tracking+in+mice:+comparison+with+tissue+Doppler+echocardiography+and+conductance+catheter+measurements.">Assessment of strain and strain rate by two-dimensional speckle tracking in mice: comparison with tissue Doppler echocardiography and conductance catheter measurements.</a> Eur Heart J Cardiovasc Imaging. 14, 765-773 (2013).</li><li>Fine, N. 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Induction of anesthesia
Proper induction and maintenance of anesthesia is critical for the accurate assessment of changes in heart valve and cardiac function in mice. Given the rapid induction of anesthesia elicited by isoflurane and the relatively long wash-out time of this anesthetic following deep anesthesia, we do not use a stand-alone anesthesia chamber for induction. Instead, as noted in detail above, animals are guided directly to the anesthesia cone, which allows for rapi...

Aging is associated with progressive increases in cardiovascular calcification1. Hemodynamically significant aortic valve stenosis affects 3% of the population over the age of 652, and patients with even moderate aortic valve stenosis (peak velocity of 3-4 m/s) have a 5 year event-free survival of less than 40%3. Presently, there are no effective treatments to slow the progression of aortic valve calcification, and surgical aortic valve replacement is the only available treatment for advanced aortic valve stenosis4.
Studies aimed at gaining a deeper understanding of the mechanisms that contribute to the initiation and progression of aortic valve calcification are a key first step in moving towards pharmacological and non-surgical methods to manage aortic valve stenosis5,6. Genetically-altered mice have played a major role in developing our understanding of the mechanisms that contribute to a variety of diseases and are now coming to the forefront of mechanistic studies aimed at understanding the biology of aortic valve stenosis6,7,8. Unlike other cardiovascular diseases such as atherosclerosis and heart failure-where standard protocols for evaluating vascular and ventricular function are for the most part well-established-there are unique challenges associated with in vivo phenotyping of heart valve function in mice. While recent reviews have provided thorough discussions regarding the advantages and disadvantages to numerous imaging and invasive modalities used to assess valve function in rodents9,10,11, to date, we are not aware of a publication that provides a comprehensive, step-by-step protocol for phenotyping heart valve function in mice.
The purpose of this manuscript is to describe the methods and protocols to phenotype heart valve function in mice. All methods and procedures have been approved by the Mayo Clinic Institutional Animal Care and Use Committee. Key components of this protocol include the depth of anesthesia, the evaluation of cardiac function, and the evaluation of heart valve function. We hope this report will not only serve to guide investigators interested in pursuing research in the field of heart valve disease, but will also start a national and international dialogue related to protocol standardization to ensure data reproducibility and validity in this rapidly-growing field. Importantly, successful imaging using high-resolution ultrasound systems requires a working knowledge of the principles of sonography (and terminology commonly used in sonography), an understanding of the fundamental principles of cardiac physiology, and significant experience with sonography to allow for accurate and time-efficient assessment of cardiac function in rodents.

<table border="0" cellpadding="0" cellspacing="0" width="1906">
	<tbody>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">High resolution ultrasound machine</td>
			<td style="width:453px;">VisualSonics, Fujifilm</td>
			<td style="width:293px;">Vevo 2100&#160;</td>
			<td style="width:467px;"></td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;width:693px;">Isoflurane diffuser (capable of delivering 1 % to 1.5 % isoflurane mixed with 1 L/min 100% O2</td>
			<td>VisualSonics, Fujifilm</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr height="68">
			<td height="68" style="height:68px;width:693px;">Transducers for small mice (550D) or larger mice (400)</td>
			<td>MicroScan, VisualSonics, Fujifilm</td>
			<td>MS 550D, MS 400</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Animal platform</td>
			<td>VisualSonics, Fujifilm</td>
			<td>11503</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Advanced physiological monitoring unit</td>
			<td>VisualSonics, Fujifilm</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Isoflurane</td>
			<td>Terrell</td>
			<td>NDC 66794-019-10</td>
			<td></td>
		</tr>
		<tr height="74">
			<td height="74" style="height:74px;width:693px;">Nose cone and tubing connected to isoflurane diffuser and 100% O2</td>
			<td>Custom Engineered in-house</td>
			<td>--</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Hair razor</td>
			<td>Andis Super AGR+ vet pack clipper</td>
			<td>AD65340</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Ultrasound gel</td>
			<td>Parker Laboratories</td>
			<td>REF 01-08</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Electrode gel&#160;</td>
			<td>Parker Laboratories</td>
			<td>REF 15-25</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Adhesive tapes</td>
			<td>Fisher Laboratories</td>
			<td>1590120B</td>
			<td></td>
		</tr>
		<tr height="34">
			<td height="34" style="height:34px;width:693px;">Paper towels</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

echocardiographic approaches and protocols for comprehensive phenotypic characterization of valvular heart disease in mice

The aim of this manuscript and accompanying video is to provide an overview of the methods and approaches used for imaging heart valve function in rodents, with detailed descriptions of the appropriate methods for anesthesia, the echocardiographic windows used, the imaging planes and probe orientations for image acquisition, the methods for data analysis, and the limitations of emerging technologies for the evaluation of cardiac and valvular function. Importantly, we also highlight several future areas of research in cardiac and heart valve imaging that may be leveraged to gain insights into the pathogenesis of valve disease in preclinical animal models. We propose that using a systematic approach to evaluating cardiac and heart valve function in mice can result in more robust and reproducible data, as well as facilitate the discovery of previously underappreciated phenotypes in genetically-altered and/or physiologically-stressed mice.

Examples of images that are routinely obtained from animal cardiac ultrasound imaging are included in this manuscript. An illustration of transducer placement on the animal&#39;s chest is provided to give the reader a clear understanding of where the transducer is positioned to obtain the images as described. A photograph of the ultrasound laboratory set-up is also included to emphasize the importance of the proper equipment, particularly the ultrasound transducer to be used and the metho...

Watch this Scientific Journal Video about Echocardiographic Approaches and Protocols for Comprehensive Phenotypic Characterization of Valvular Heart Disease in Mice at JoVE.com

Echocardiographic Approaches and Protocols for Comprehensive Phenotypic Characterization of Valvular Heart Disease in Mice

This protocol provides a detailed description of the echocardiographic approach for comprehensive phenotyping of heart and heart valve function in mice.

The aim of this manuscript and accompanying video is to provide an overview of the methods and approaches used for imaging heart valve function in ...

The overall goal of this echo protocol is to show methods and approaches used to image heart valve function in rodents. The methods we'll be showing you today can answer key questions in the field of heart disease, with a specific emphasis on evaluating the heart as integrated unit under conditions of heart valve disease. The implications of this approach are quiet profound for the field of heart valve diseases because the comprehensive echocardiographic evaluation of the heart will not only allow for detection of valvular dysfunction in multiple heart valves, but will also allow investigators to understand the complex interplay between cardiac function and various measures of heart valve function.Though this method can be leveraged to characterize animal models in which heart valve disease is our primary defect, it can also be applied to other systems where heart valve dysfunction may appear secondary to other insults such as heart failure, or a thoracic aortic aneurysm. Thus, it is our recommendation that comprehensive cardiac and heart valve phenotyping should be routinely performed in most research studies. We first had the idea to film our approach to comprehensive cardiac and heart valve phenotyping after attending a number of meetings where there was a profound variability in the methods and protocols used to assess heart valve function.It is our hope that this manuscript and the accompanying video spurs discussion in the field regarding the best methodologies and standards to follow, which ultimately aligns with recent initiatives from the National Institutes of Health to improve rigor and reproducibility of research studies. The main advantage of echocardiography is that it is able to noninvasively assess morphology and function of the heart in real time. It is easy to perform, and can be performed serially.Thus, it is most ideal in serial follow up studies. Generally, individuals new to cardiac and heart valve imaging will struggle due to logistical challenges associated with administration and titration of anesthesia, as well as the technical challenges associated with the very small anatomic windows that can be used for echocardiography and the need to precisely identify cardiac structures and probe orientations while imaging. We hope that visual demonstration of our approach accelerates the rate at which investigators can learn to conduct echocardiography in mice, and will ultimately serve as a platform to discuss best practices for small animal imaging.To begin, turn on the ultrasound machine equipped with a high frequency ultrasound transducer. Enter the ID of the animal to be measured, along with date, time, and any other relevant information. Preheat the platform to 37 degrees celsius, and then gently pick up the mouse by it's tail, and firmly hold the animal at the nape of it's neck.Guide the nose of the animal into the nose cone and initiate anesthesia as described in the accompanying text protocol. Once sedated, quickly and accurately lay the animal on the platform in a supine position, making sure that the forefeet and hindfeet lie on the ECG sensors of the platform. Use adhesive tape to secure the animal on all four limbs, to stabilize the head in the nose cone apparatus, and to stabilize the tail.Next, check the heart rate using the ECG sensors in the platform, or using an external device. Ensure that the baseline heart rate is between 600 to 700 beats per minute. Monitor the heart rate and ensure that it does not fall below 450 beats per minute under any circumstances.In addition, monitor the body temperature using a rectal thermometer and keep the temperature between 36.5 degrees celsius and 38 degrees celsius. As long as the vitals are stable, use an electric clipper designed for use with fine hair to shave off the hair from the mouse's chest. Then, take a damp paper towel and wipe the chest to remove any remaining hairs.With the animal securely fastened on the platform, and the head facing away, tilt the table 15 to 20 degrees to the left to bring the heart closer to the chest wall. Then, apply a generous amount of ultrasound gel on the transducer or directly on the animal's chest. Position the transducer parasternally about 90 degrees perpendicular with the long axis of the heart so that the image index marker of the transducer is pointing posteriorly.While in two DB mode slide the transducer's defilade until the aortic valve comes into view along the short axis. Next, rotate the transducer clockwise until the image index marker points cadad. This is the parasternal long axis view.Observe the aortic route, aortic valve, left ventricular outflow tract, mitral valve, left atrium, and part of the right ventricular outflow tract on the image display. Measure the largest atrial posterior dimension of the aorta using the electronic caliper associated with the measurement tool that is embedded in the machine. Next, reduce the image width so that only the aortic valve is on the image display by adjusting the image width button in the control panel.Position the M mode line of interrogation where it intersects the tips of the aortic valve to accurately assess aortic valve cusp separation. In the M mode display of the aortic valve, measure the cusp separation distance using the electronic caliper. While still in the parasternal long axis view of the aortic valve, press the color doppler control key in the control panel to apply color doppler to the region of the aortic valve.Next, press the post wave doppler control key. Using the track ball, place the post wave sample volume in the proximal ascending aorta just above the aortic valve, making sure that the angle between the ultrasound beam and the blood flow is less than 60 degrees by tilting the platform, and or the transducer. Then, measure the peak velocity from the spectral display using the electronic calipers.Start by placing the transducer in the apical position in B mode and positioning the transducer so that it is angled towards the head of the mouse. Observe the right ventricle, left ventricle, right atrium and left atrium, on the image display. From the apical four chamber view bring the mitral valve in focus by reducing the image width.Then, place the M mode cursor across the mitral valve to assess the thickness of the leaflets. Using the apical four chamber view, apply color doppler to image the flow from the left atrium through the mitral valve during diastole. Observe the flow across the mitral valve which is encoded in red.From the apical lung axis view, tilt or point the transducer tip using a rocking motion so that the right ventricle is at the center of the image display. Reduce the image width so that only the right ventricle is visible in the image display. Then, apply color doppler in the region of the tricuspid valve.Next, move the transducer to a modified parasternal long axis position at the level of the aortic valve. Then, tilt the transducer slightly upward to obtain a short axis view of the pulmonic valve. In this view, apply M mode imaging to evaluate the separation distance of the pulmonic valve.Then, apply color doppler in the region of the pulmonic valve to assess for valvular regurgitation and stenosis. In two DB mode, place the transducer in the parasternal short axis position at the level of the papillary muscles to obtain a short axis view of the left ventricle. From this view, press the M mode button located in the control panel.Using the track ball, position the M mode cursor at the center of the left ventricular cavity at the level of the papillary muscles, and obtain M mode images. Then, measure the left ventricular cavity dimension at end diastole where the distance between the interior wall and posterior wall is the largest, and an end systole where the inward motion of both interior and posterior walls is maximal. Next, move the transducer to the apical window, apply color on the mitral valve, and place the sample volume at the tips of the mitral valve leaflets.Measure the peak mitral inflow velocity from the spectral display of pulse waved doppler velocities across the mitral valve. With the sample volume positioned between left ventricle inflow and outflow not the mitral and aortic valve closing and opening signals. Measure the isovolumic relaxation time, isovolumic contraction time, and left ventricular ejection time.Then, perform tissue doppler imaging of the mitral annulus in the apical long axis view. Press the TGI control key and place the sample volume at the medial aspect of the mitral annulus. When finished, remove any excess ultrasound gel from the mouse.Gently remove the tape securing the animal, and turn off the anesthesia. Place the animal on an absorbent paper towel and observe the animal until sternal recumbency is attained. Shown here is an assessment of aortic valve function in a normal mouse versus the aortic valve function in a mouse with calcific aortic valve disease.In the mouse with calcified valves, the cusps are thickened and have increased echogenicity which results in restricted opening during systole. From the apical window a long axis view of the mitral valve is presented. The mitral leaflet thickness can be measured using the M mode line of interrogation.Measurement of the thickness of the mitral valve can be extremely challenging given the thin, poorly echogenic, and rapidly moving leaflets of the normal mitral valve. The color doppler shown in this modified long axis view of the mitral valve shows a mosaic color jet at the mitral valve during systole. The blue color is evidence of mitral valve regurgitation.From the parasternal window, both short and long axis views of the pulmonic valve are obtained. In this image, the M mode line of interrogation is applied across the pulmonic valve, and the valve cusp separation distance can be measured from this view. Once mastered, this technique can be done in 15 to 20 minutes if it is performed properly.While attempting this procedure, it's important to have a basic knowledge of cardiac anatomy and physiology, a working knowledge of the principles and terminology of sonography, and experience in cardiac ultrasound to allow for accurate and time efficient assessment of cardiac function in rodents. After watching this video you should have a good understanding of how to perform cardiac ultrasound in rodents by obtaining images from various imaging planes, and measuring echo doppler parameters that adequately describe cardiac and heart valve function.

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15.7K ビュー.  Mayo Clinic. The overall goal of this echo protocol is to show methods and approaches used to image heart valve function in rodents. The methods we'll be showing you today can answer key questions in the field of heart disease, with a specific emphasis on evaluating the heart as integrated unit under conditions of heart valve disease. The implications of this approach are quiet profound for the field of heart valve diseases because the comprehensive echocardiographic evaluation of the heart will not only ...

ビデオ: Echocardiographic Approaches and Protocols for Comprehensive Phenotypic Characterization of Valvular Heart Disease in Mice

Assessment of Cardiac Function and Myocardial Morphology Using Small Animal Look-locker Inversion Recovery (SALLI) MRI in Rats

Small animal magnetic resonance imaging is an important tool to study cardiac function and changes in myocardial tissue. The high heart rates of small animals (200 to 600 beats/min) have previously limited the role of CMR imaging. Small animal Look-Locker inversion recovery (SALLI) is a T1 mapping sequence for small animals to overcome this problem 1. T1 maps provide quantitative information about tissue alterations and contrast agent kinetics. It is also possible to detect diffuse myocardial processes such as interstitial fibrosis or edema 1-6. Furthermore, from a single set of image data, it is possible to examine heart function and myocardial scarring by generating cine and inversion recovery-prepared late gadolinium enhancement-type MR images 1.
	The presented video shows step-by-step the procedures to perform small animal CMR imaging. Here it is presented with a healthy Sprague-Dawley rat, however naturally it can be extended to different cardiac small animal models.

Assessment of Cardiac Function and Myocardial Morphology Using Small Animal Look-locker Inversion Recovery SALLI MRI in Rats

Contrast enhanced cardiac magnetic resonance (CMR) imaging allows comprehensive in vivo assessment of the heart in small animal models of cardiovascular disease. Here we detail the procedures to perform CMR imaging, reconstruction and analysis using Small Animal Lock-Locker Inversion Recovery (SALLI) in rats.

Small animal magnetic resonance  imaging is an important tool to study cardiac function and changes in myocardial  tissue. The high heart rates of small ...

Echocardiographic Assessment of the Right Heart in Mice

Transgenic and toxic models of pulmonary arterial hypertension (PAH) are widely used to study the pathophysiology of PAH and to investigate potential therapies. Given the expense and time involved in creating animal models of disease, it is critical that researchers have tools to accurately assess phenotypic expression of disease. Right ventricular dysfunction is the major manifestation of pulmonary hypertension. Echocardiography is the mainstay of the noninvasive assessment of right ventricular function in rodent models and has the advantage of clear translation to humans in whom the same tool is used. Published echocardiography protocols in murine models of PAH are lacking.
In this article, we describe a protocol for assessing RV and pulmonary vascular function in a mouse model of PAH with a dominant negative BMPRII mutation; however, this protocol is applicable to any diseases affecting the pulmonary vasculature or right heart. We provide a detailed description of animal preparation, image acquisition and hemodynamic calculation of stroke volume, cardiac output and an estimate of pulmonary artery pressure.

This article provides a protocol for the echocardiographic assessment of right ventricular size and pulmonary hypertension in mice. Applications include phenotype determination and serial assessment in transgenic and toxin-induced mouse models of cardiomyopathy and pulmonary vascular disease.

Transgenic and toxic models of pulmonary arterial hypertension (PAH) are widely used to study the pathophysiology of PAH and to investigate potential ...

Implantation of Total Artificial Heart in Congenital Heart Disease

In patients with end-stage heart failure (HF), a total artificial heart (TAH) may be implanted as a bridge to cardiac transplant. However, in congenital heart disease (CHD), the malformed heart presents a challenge to TAH implantation. 
In the case presented here, a 17 year-old patient with congenital transposition of the great arteries (CCTGA) experienced progressively worsening HF due to his congenital condition. He was hospitalized multiple times and received an implantable cardioverter defibrillator (ICD). However, his condition soon deteriorated to end-stage HF with multisystem organ failure. 
Due to the patient's grave clinical condition and the presence of complex cardiac lesions, the decision was made to proceed with a TAH. The abnormal arrangement of the patient's ventricles and great arteries required modifications to the TAH during implantation.
With the TAH in place, the patient was able to return home and regain strength and physical well-being while awaiting a donor heart. He was successfully bridged to heart transplantation 5 months after receiving the device. This report highlights the TAH is feasible even in patients with structurally abnormal hearts, with technical modification.

This is a case report of a patient with congenitally corrected transposition of the great arteries (CCTGA) who received a total artificial heart (TAH) as a bridge to heart transplant. The TAH was successfully implanted with modifications to accommodate the patient's congenitally malformed heart.

In patients with end-stage heart failure (HF), a total artificial heart (TAH) may be implanted as a bridge to cardiac transplant. However, in congenital ...

Establishment and Characterization of UTI and CAUTI in a Mouse Model

Urinary tract infections (UTI) are highly prevalent, a significant cause of morbidity and are increasingly resistant to treatment with antibiotics. Females are disproportionately afflicted by UTI: 50% of all women will have a UTI in their lifetime. Additionally, 20-40% of these women who have an initial UTI will suffer a recurrence with some suffering frequent recurrences with serious deterioration in the quality of life, pain and discomfort, disruption of daily activities, increased healthcare costs, and few treatment options other than long-term antibiotic prophylaxis. Uropathogenic Escherichia coli (UPEC) is the primary causative agent of community acquired UTI. Catheter-associated UTI (CAUTI) is the most common hospital acquired infection accounting for a million occurrences in the US annually and dramatic healthcare costs. While UPEC is also the primary cause of CAUTI, other causative agents are of increased significance including Enterococcus faecalis. Here we utilize two well-established mouse models that recapitulate many of the clinical characteristics of these human diseases. For UTI, a C3H/HeN model recapitulates many of the features of UPEC virulence observed in humans including host responses, IBC formation and filamentation. For CAUTI, a model using C57BL/6 mice, which retain catheter bladder implants, has been shown to be susceptible to E. faecalis bladder infection. These representative models are being used to gain striking new insights into the pathogenesis of UTI disease, which is leading to the development of novel therapeutics and management or prevention strategies.

The ability to model urinary tract infections (UTI) is crucial in order to be able to understand bacterial pathogenesis and spawn the development of novel therapeutics. This work&#8217;s goal is to demonstrate mouse models of experimental UTI and catheter associated UTI that recapitulate and predict findings seen in humans.

Urinary tract infections (UTI) are highly prevalent, a significant cause of morbidity and are increasingly resistant to treatment with antibiotics. ...

Echocardiographic and Histological Examination of Cardiac Morphology in the Mouse

An increasing number of genetically modified mouse models has become available in recent years. Moreover, the number of pharmacological studies performed in mice is high. Phenotypic characterization of these mouse models also requires the examination of cardiac function and morphology. Echocardiography and magnetic resonance imaging (MRI) are commonly used approaches to characterize cardiac function and morphology in mice. Echocardiographic and MRI equipment specialized for use in small rodents is&#160;expensive and requires a dedicated space. This protocol describes cardiac measurements in mice using a clinical echocardiographic system with a 15 MHz human vascular probe. Measurements are performed on anesthetized adult mice. At least three image sequences are recorded and analyzed for each animal in M-mode in the parasternal short-axis view. Afterwards, cardiac histological examination is performed, and cardiomyocyte diameters are determined on hematoxylin-eosin- or wheat germ agglutinin (WGA)-stained paraffin sections. Vessel density is determined morphometrically after Pecam-1 immunostaining. The protocol has been applied successfully to pharmacological studies and different genetic animal models under baseline conditions, as well as after experimental myocardial infarction by the permanent ligation of the left anterior descending coronary artery (LAD). In our experience, echocardiographic investigation is limited to anesthetized animals and is feasible in adult mice weighing at least 25 g.

Echocardiographic examination is frequently used in mice. Expensive high-resolution ultrasound devices have been developed for this purpose. This protocol describes an affordable echocardiographic procedure combined with histological morphometric analyses to determine cardiac morphology.

An increasing number of genetically modified mouse models has become available in recent years. Moreover, the number of pharmacological studies performed ...

Echocardiographic Measurement of Right Ventricular Diastolic Parameters in Mouse

Diastolic dysfunction is a prominent feature of right ventricular (RV) remodeling associated with conditions of pressure overload. However, the RV diastolic function is rarely quantified in experimental studies. This might be due to technical difficulties in the visualization of the RV in the apical four-chamber view in rodents. Here we describe two positions facilitating the visualization of the apical four-chamber view in mice to assess the RV diastolic function.
The apical four-chamber view is enabled by tilting the mouse fixation platform to the left and caudally (LeCa) or to the right and cranially (RiCr). Both positions provide images of comparable quality. The results of the RV diastolic function obtained from two positions are not significantly different. Both positions are comparably easy to perform. This protocol can be incorporated into published protocols and enables detailed investigations of the RV function.

Here we describe and compare two positions for obtaining the apical four-chamber view in mice. These positions enable the quantification of the right ventricular function, provide comparable results, and can be used interchangeably.

Diastolic dysfunction is a prominent feature of right ventricular (RV) remodeling associated with conditions of pressure overload. However, the RV ...

Induction and Phenotyping of Acute Right Heart Failure in a Large Animal Model of Chronic Thromboembolic Pulmonary Hypertension

The development of acute right heart failure (ARHF) in the context of chronic pulmonary hypertension (PH) is associated with poor short-term outcomes. The morphological and functional phenotyping of the right ventricle is of particular importance in the context of hemodynamic compromise in patients with ARHF. Here, we describe a method to induce ARHF in a previously described large animal model of chronic PH, and to phenotype, dynamically, right ventricular function using the gold standard method (i.e., pressure-volume PV loops) and with a non-invasive clinically available method (i.e., echocardiography). Chronic PH is first induced in pigs by left pulmonary artery ligation and right lower lobe embolism with biological glue once a week for 5 weeks. After 16 weeks, ARHF is induced by successive volume loading using saline followed by iterative pulmonary embolism until the ratio of the systolic pulmonary pressure over systemic pressure reaches 0.9 or until the systolic systemic pressure decreases below 90 mmHg. Hemodynamics are restored with dobutamine infusion (from 2.5 &#181;g/kg/min to 7.5 &#181;g/kg/min). PV-loops and echocardiography are performed during each condition. Each condition requires around 40 minutes for induction, hemodynamic stabilization and data acquisition. Out of 9 animals, 2 died immediately after pulmonary embolism and 7 completed the protocol, which illustrates the learning curve of the model. The model induced a 3-fold increase in mean pulmonary artery pressure. The PV-loop analysis showed that ventriculo-arterial coupling was preserved after volume loading, decreased after acute pulmonary embolism and was restored with dobutamine. Echocardiographic acquisitions allowed to quantify right ventricular parameters of morphology and function with good quality. We identified right ventricular ischemic lesions in the model. The model can be used to compare different treatments or to validate non-invasive parameters of right ventricular morphology and function in the context of ARHF.

We present a protocol to induce and phenotype an acute right heart failure in a large animal model with chronic pulmonary hypertension. This model can be used to test therapeutic interventions, to develop right heart metrics&#160;or to improve the understanding of acute right heart failure pathophysiology.

The development of acute right heart failure (ARHF) in the context of chronic pulmonary hypertension (PH) is associated with poor short-term outcomes. The ...

Generation of Human 3D Lung Tissue Cultures (3D-LTCs) for Disease Modeling

Translation of novel discoveries to human disease is limited by the availability of human tissue-based models of disease. Precision-cut lung slices (PCLS) used as 3D lung tissue cultures (3D-LTCs) represent an elegant and biologically highly relevant 3D cell culture model, which highly resemble in situ tissue due to their complexity, biomechanics and molecular composition. Tissue slicing is widely applied in various animal models. 3D-LTCs derived from human PCLS can be used to analyze responses to novel drugs, which might further help to better understand the mechanisms and functional effects of drugs in human tissue. The preparation of PCLS from surgically resected lung tissue samples of patients, who experienced lung lobectomy, increases the accessibility of diseased and peritumoral tissue. Here, we describe a detailed protocol for the generation of human PCLS from surgically resected soft-elastic patient lung tissue. Agarose was introduced into the bronchoalveolar space of the resectates, thus preserving lung structure and increasing the tissue&#39;s stiffness, which is crucial for subsequent slicing. 500 &#181;m thick slices were prepared from the tissue block with a vibratome. Biopsy punches taken from PCLS ensure comparable tissue sample sizes and further increase the amount of tissue samples. The generated lung tissue cultures can be applied in a variety of studies in human lung biology, including the pathophysiology and mechanisms of different diseases, such as fibrotic processes at its best at (sub-)cellular levels. The highest benefit of the 3D-LTC ex vivo model is its close representation of the in situ human lung in respect of 3D tissue architecture, cell type diversity and lung anatomy as well as the potential for assessment of tissue from individual patients, which is relevant to further develop novel strategies for precision medicine.

Generation of Human 3D Lung Tissue Cultures 3D-LTCs for Disease Modeling

Here, we present a protocol for the preparation of agarose-filled human precision-cut lung slices from resected patient tissue that are suitable for generating 3D lung tissue cultures to model human lung diseases in biological and biomedical studies.

Translation of novel discoveries to human disease is limited by the availability of human tissue-based models of disease. Precision-cut lung slices (PCLS) ...

Assessment and Characterization of Hyaloid Vessels in Mice

In the eye, the embryonic hyaloid vessels nourish the developing lens and retina and regress when the retinal vessels develop. Persistent or failed regression of hyaloid vessels can be seen in diseases such as persistent hyperplastic primary vitreous (PHPV), leading to an obstructed light path and impaired visual function. Understanding the mechanisms underlying the hyaloid vessel regression may lead to new molecular insights into the vascular regression process and potential new ways to manage diseases with persistent hyaloid vessels. Here we describe the procedures for imaging hyaloid in live mice with optical coherence tomography (OCT) and fundus fluorescein angiography (FFA) and a detailed technical protocol of isolating and flat-mounting hyaloid ex vivo for quantitative analysis. Low-density lipoprotein receptor-related protein 5 (LRP5) knockout mice were used as an experimental model of persistent hyaloid vessels, to illustrate the techniques. Together, these techniques may facilitate a thorough assessment of hyaloid vessels as an experimental model of vascular regression and studies on the mechanism of persistent hyaloid vessels.

This protocol describes both in vivo and ex vivo methods to fully visualize and characterize hyaloid vessels, a model of vascular regression in mouse eyes, using optical coherence tomography and fundus fluorescein angiography for the live imaging and ex vivo isolation and subsequent flat mount of hyaloid for quantitative analysis.

In the eye, the embryonic hyaloid vessels nourish the developing lens and retina and regress when the retinal vessels develop. Persistent or failed ...

Transthoracic Echocardiographic Examination in the Rabbit Model

Large animal models such as the rabbit are valuable for translational preclinical research. Rabbits have a similar cardiac electrophysiology compared to that of humans and that of other large animal models such as dogs and pigs. However, the rabbit model has the additional advantage of lower maintenance costs compared to other large animal models. The longitudinal evaluation of cardiac function using echocardiography, when appropriately implemented, is a useful methodology for preclinical assessment of novel therapies for heart failure with reduced ejection fraction (e.g. cardiac regeneration). The correct use of this non-invasive tool requires the implementation of a standardized examination protocol following international guidelines. Here we describe, step by step, a detailed protocol supervised by veterinary cardiologists for performing echocardiography in the rabbit model, and demonstrate how to correctly obtain the different echocardiographic views and imaging planes, as well as the different imaging modes available in a clinical echocardiography system routinely used in human and veterinary patients.

Here we describe, step by step, a detailed protocol for performing echocardiography in the rabbit model. We show how to correctly obtain the different echocardiographic views and imaging planes, as well as the different imaging modes available in a clinical echocardiography system routinely used in human and veterinary patients.

Large animal models such as the rabbit are valuable for translational preclinical research. Rabbits have a similar cardiac electrophysiology compared to ...