Name: noninvasive assessment of cardiac abnormalities in experimental autoimmune myocarditis by magnetic resonance microscopy imaging in the mouse
Uploaded: 2014-06-20T15:00:00.000+00:00
Duration: 12 min 24 s
Description: Watch this Scientific Journal Video about Noninvasive Assessment of Cardiac Abnormalities in Experimental Autoimmune Myocarditis by Magnetic Resonance Microscopy Imaging in the Mouse at JoVE.com

This work was supported by the National Institutes of Health (HL114669). CM is a recipient of a postdoctoral research fellowship grant awarded by the Myocarditis Foundation, NJ.

ETHICS STATEMENT:
All animal procedures were conducted in accordance with the guidelines for the Care and Use of Laboratory Animals and approved by the University of Nebraska-Lincoln, Lincoln, NE.
1. Induction of Experimental Autoimmune Myocarditis
<ol>
	<li>Prepare the peptide solution by dissolving Myhc-&#945; 334-352 in 1x phosphate-buffered saline to a final concentration of 2 mg/1.5 ml.</li>
	<li>Prepare the pertussis toxin (PT) by adding 1 ml of sterile 1x PBS to a vial containing 50 &#956;g of lyophilized PT to obtain a stock concentration of 50 ng/&#956;l. Take 20 &#956;l of the stock into a sterile 1.5 ml tube and add 980 &#956;l of sterile 1x PBS to obtain a working concentration of 1 ng/&#956;l.</li>
	<li>Prepare complete Freund&#8217;s adjuvant (CFA) by adding 40 mg of Mycobacterium&#160;tuberculosis H37Rv (MTB) extract to 10 ml of CFA to obtain a final concentration of 5 mg/ml.</li>
	<li>Prepare the peptide-CFA emulsion. NOTE: For EAM induction, the peptide-CFA emulsion was administered in 150 &#956;l containing 100 &#181;g of Myhc-&#945; 334-352 per animal. For example, to immunize ten mice use 1.5 ml of peptide-CFA emulsion containing 1 mg of Myhc-&#945; 334-352.
	<ol>
		<li>To prepare 1.5 ml of emulsion, aliquot 750 &#956;l of Myhc-&#945; 334-352 peptide solution in one 1.5 ml tube, and CFA supplemented with MTB into another 1.5 ml tube. Using a 3&#160;ml luer-lok syringe, draw the peptide solution first, followed by the CFA/MTB extract.</li>
		<li>Attach the syringe to a 3-way stopcock and connect the other outlet of the stopcock to an empty 3&#160;ml syringe. Adjust the patency of the stopcock so the peptide-CFA mixture flows from one syringe to the other with reasonably good resistance.</li>
		<li>Mix by pushing the contents from one syringe to the other repeatedly for ~1 min and then place the entire assembly on ice for ~3 min. Repeat this procedure a minimum of 10x.</li>
		<li>Determine the consistency of the emulsion by carefully placing a tiny droplet on still water in a 100 ml glass beaker. The droplet is not expected to disperse in water. If it does, continue mixing until the desired consistency is achieved.</li>
		<li>Transfer the contents of emulsion from 3 ml syringes into 1&#160;ml luer-lok syringes by replacing one of the two 3-ml syringes attached to the stopcock with the 1-ml syringe, and attach a 27 &#189; G needle to the 1&#160;ml syringe.</li>
	</ol>
	</li>
	<li>Inject 150 &#181;l of the peptide-CFA emulsion in split doses subcutaneously in the two inguinal regions of six-to-eight-week-old female A/J mice (~75 &#181;l each).</li>
	<li>Administer 100 &#956;l of PT suspension (100 ng) intraperitoneally to each animal on day 0 and day 2 postimmunization.</li>
	<li>Repeat the immunization procedure on day 7 by administering 150 &#181;l of peptide-CFA emulsion in split doses subcutaneously into either side of the sternum (~75 &#181;l each). Prepare this emulsion fresh as above. Then, on day 21, subject the animals to MRM imaging, see step 3.</li>
</ol>
2. Animal Handling
<ol>
	<li>Place each mouse in an anesthetic induction chamber containing 2% isoflurane-air mixture with a heating pad placed underneath to maintain warmth, and transfer the animal to a specially designed animal holder (Figure&#160;1).</li>
	<li>Immobilize the animal in the prone position on the animal holder so its snout fits into the nose cone to maintain anesthesia (Figure&#160;1). Secure the head of the mouse with a bite-bar attached to the front teeth of the mouse.</li>
	<li>Turn on the air blow heater with its outlet hose inserted in the scanner&#8217;s vertical bore to maintain the animal&#8217;s body temperature during the experiment.</li>
	<li>Maintain anesthesia at 0.5 to 2% isoflurane with a flow rate of 2 ml/min during the entire imaging session. Confirm anesthesia using the toe pinch method, no movement expected.</li>
	<li>Set up a pneumatic pillow sensor, mouse tail/ankle fiber optic pulse oximetry sensor, and rectal temperature probe to monitor respiration, heart rate and body temperature, respectively (Figure&#160;1). Note: Cardiac gating is performed through Pulse Oximetry, which allows noninvasive monitoring of arterial blood oxygen saturation. The pulse oximetry sensor should be attached to the left ankle, and the foot should be secured with a thread loop and taped to maintain the ankle in the sensor. MRM imaging is achieved by gating the respiration and cardiac signals without the use of any contrast agents.</li>
</ol>
3. Image Acquisition
<ol>
	<li>After preparing the animal (Figure&#160;1), place the mouse in the center of the MRM scanner with the heart positioned in the center of the field of view (FOV), where magnetic field homogeneity is maximum. NOTE: A wide-bore (89 mm)&#160;9.4 T vertical-bore magnet equipped with triple axis gradients of 100 G/cm and a 4 cm radio-frequency (RF) imaging coil is to be used to obtain high-resolution three-dimensional (3D) images. Note: Be sure to non-magnetic supplies when using an MRI scanner.&#160;</li>
	<li>Run the imaging interface and select &#8220;New Study&#8221; from the &#8220;Study Options&#8221; menu. Type &#8220;mtune&#8221; at the command bar and run it to pull up the &#8220;Tune GUI&#8221;. Then select &#8220;Start Probe Tune&#8221; and click on &#8220;Autoscale&#8221; button. The Tune GUI will change to show the RFsignal. Use the remote tune/match knobs at the end of the coil to tune the RF coil to the proton resonance frequency (400 MHz). On the &#8220;Start&#8221; tab go to the &#8220;Prescan&#8221; page to run the frequency and power calibration by clicking on the related buttons. Hit the XYZ (quick) button on the &#8216;Shim&#8221; tab of &#8220;Study&#8217; tab to pull up the shimming page. Go to the shim page, select all of the iterations, and hit the shim button to perform auto shimming.</li>
	<li>Select a scout sequence from the &#8220;Study&#8221; tab of the imaging interface to localize the mouse heart. On the &#8220;Acquire&#8221; tab change the FOV to 35 mm2 and keep the machine&#8217;s default settings. Click Start to run the sequence; adjust the position of the animal holder if the heart is not in the center of FOV, retune the RF coil and obtain the scout images again.</li>
	<li>Click the &#8220;GEMS&#8221; sequence button on the &#8220;Study&#8221; tab then enter the acquisition parameters on the corresponding &#8220;Acquire&#8221; tab to obtain two orthogonal planes along the short axis and long axis of the heart20,21. NOTE: Typical acquisition parameters for a gradient-echo scan are: slice thickness, 1 mm; repetition time (TR), 200 msec; echo time (TE), 2.67 msec (minimum); flip angle, 25&#176;; inplane matrix size, 128 x 128; FOV, 22 mm2; number of acquisitions, 4; and an approximate acquisition time of 1 min&#160;and 30 sec.</li>
	<li>On the &#8220;Adv&#8221; tab select the &#8220;CINE&#8221; sequence to collect the pulse oximetry-gated short-axis cine MR images in order to measure the LV anatomic and functional parameters. Adjust the position and angle of the imaging slices based on the long axis view of the heart by mouse hovering and dragging. Enter the following imaging parameters on the &#8220;Acquire&#8221; tab to obtain the gradient echo cine sequence: TR, 500 msec; TE, 5 msec; FOV, 22 mm2; acquisition matrix, 256 x 256; slice thickness, 1 mm; number of acquisitions, 8; number of frames, 6; and an acquisition time of ~17 min. Click start to begin the acquisition.</li>
	<li>Convert the acquired images to DICOM format using the &#8220;I/O&#8221; tab of the imaging software and transfer the corresponding files to the data center for processing.</li>
	<li>At the end of imaging procedure, allow the mice to recover from anesthesia within the filter-top cages. Do not leave the mice unattended until they regain sufficient consciousness to maintain sternal recumbency and keep them for further studies as needed.</li>
</ol>
4. Data Analysis of Cardiac Magnetic Resonance Microscopy Imaging
<ol>
	<li>Use Segment software to analyze the anatomic and functional parameters of the LV19. By loading the DICOM format cine images into the software using the &#8220;Open Image File(s)&#8221; submenu of the &#8220;File&#8221; menu. On the GUI select &#8216;MRGE&#8217; as the imaging technique, &#8216;Cine&#8217; as the image type and &#8216;short-axis mid-ventricular&#8217; as the image view plane.</li>
	<li>Specify the time frames to be used for end-diastole and end-systole through the &#8220;Set Current Timeframe to End Diastole&#8221; and &#8220;Set Current Timeframe to End Systole&#8221; submenus of the &#8220;Edit&#8221; menu respectively.</li>
	<li>First click the &#8220;LV&#8221; command button and then the &#8220;ENDO&#8221; and &#8220;EPI&#8221; command buttons on the bottom-right panel to manually draw the left ventricle endocardium and epicardium respectively. Remove the papillary muscles by hitting the corresponding command button in order to increase the accuracy of the calculations.</li>
	<li>Read the quantified LV parameters such as diastolic volume, systolic volume, stroke volume, and ejection fraction on the top-right panel. Click the &#8220;Misc&#8221; command button and then the &#8220;Measurement Caliper&#8221; command button to measure the LV parameters such as wall thickness, and ventricular diameter22.</li>
	<li>Click the &#8220;Save Both Image Stacks and Segmentation&#8221; submenu of the &#8220;File&#8221; menu to save the images, including the segmentations, in &#8216;.mat&#8217; format to reprocess the images as needed.</li>
</ol>

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No conflicts of interests were declared.

This study describes the MRM procedure and its utility as a non-invasive tool to ascertain cardiac abnormalities in mice affected with autoimmune myocarditis. Since the histologic features of EAM resemble postinfectious myocarditis of humans, mouse models are commonly employed to delineate the immune mechanisms of myocardial injuries23-25. However, the animals affected with myocarditis appear clinically normal, and the diagnosis is made based on histology at the termination of experiments7. The animals are usually sacrificed on day 21 postimmunization. Assessing the disease process in this way at single time points limits use of these models, especially in pharmaceutical research, where monitoring of disease progression in response to treatments is a critical requirement.
To ascertain cardiac abnormalities in living animals, use of non-invasive modalities like MRM is helpful. The MRM technique that described here offers an advantage of obtaining the structural and functional characteristics of hearts without the need to use contrast agents. However, this technique requires acquisition of high resolution 3D anatomical images in strong magnetic fields. Nonetheless, once the images are acquired, functional parameters such as LV end-diastolic volumes and ejection fractions can be analyzed later using commercially available software, without the need to further assemble the MRM apparatus. As shown in Figure&#160;2, MRM examination of animals immunized with Myhc-&#945; 334-352 revealed LV wall thickness to be greater than in healthy mice (Figure&#160;2B), with a corresponding decrease in functional outputs (LV end-diastolic volumes and ejection fractions; Figure&#160;2C). As expected, hearts from immunized, but not healthy, animals had inflammatory infiltrates (Figure&#160;2D). Thus, findings from the MRM technique and histology corroborate each other.
Nonetheless, to obtain reproducible results by MRM, the following three factors need to be addressed. (a) Animals should be placed in the MRM scanner so hearts are positioned in the center of the magnet to expose them to the magnetic field with maximum homogeneity. (b) Motion artifact is a concern in live animal experiments. To diminish image blurring due to respiration and cardiac movements, pulse oximetry and respirometry were used to gate MRM acquisition&#8212;that is, to acquire discrete image signals at specific time points within the respiratory and cardiac cycles&#8212;which required the use of an animal monitoring system. (c) Acquisition of high-resolution 3D cardiac images is a critical requirement to enable detailed analysis of cardiac abnormalities. To obtain images in all three dimensions, and to increase signal-to-noise ratio, it is important to design more sensitive imaging coils specific to MRM that permit accurate and complete capturing of images in strong magnetic fields in a short-axis orientation within the animal scanners.
In conclusion, development of an MRM technique to evaluate cardiac abnormalities in live animals is challenging. This is especially true for mice, because of their smaller heart sizes (~1/2,000 the mass of a human heart) and higher heart rates (~600 beats per min) compared to humans26. Nonetheless, once developed and validated, the MRM technique can be used to compare the anatomical and functional changes of hearts between healthy and sick animals. Thus, the MRM technique could serve as a valuable, non-invasive tool to assess the longitudinal progression of inflammatory cardiac pathologies related to both the acute and chronic nature of the disease process, and to monitor responses to therapies in living animals.

Heart failure is the leading cause of deaths, and myocarditis is one predominant cause of heart failure in young adolescents1. Most patients affected with myocarditis remain asymptomatic and the disease is spontaneously resolved2. However, 10-20% of those affected can develop chronic disease, leading to dilated cardiomyopathy (DCM)3. Various animal models have been developed to study the immune pathogenesis of myocarditis. The disease can be induced in myocarditis-susceptible A/J and Balb/c mice by immunizing the animals with cardiac myosin heavy chain (Myhc)-&#945; or its immunodominant peptide fragments or by infecting with pathogens like coxsackievirus B3 4-9. This study involves Myhc-&#945; 334-352-induced myocarditis in A/J mice. Despite showing myocardial infiltrations, the myocarditis-affected animals appear clinically normal; diagnosis is based on histological evaluation of hearts for inflammation7 and echocardiography10.
Magnetic resonance microscopy (MRM) is a commonly used method to obtain cardiovascular imaging with high resolution three-dimensional planes, permitting assessment of functional details to the level of minute blood vessels (up to 10 &#956;m diameter), but this level of resolving power is not achievable with the routine magnetic resonance imaging (MRI) scanning procedure, in which, the resolution is generally obtained up to 1 mm11-14. MRM offers an advantage as it permits acquisition of high-resolution images and also to derive performance parameters in the early time points of disease process14. Clinically, MRM imaging has been extensively applied to study the functional parameters of diseased heart, lung or brain15-17. In this study, the use of an MRM technique as a non-invasive tool to ascertain cardiac abnormalities in A/J mice affected with autoimmune myocarditis is shown. Specifically, the MRM imaging allows quantification of functional parameters such as left ventricular (LV) end-diastolic volume and ejection fraction (EF) with reasonable accuracy18. The definitions of respective parameters are: LV end diastolic volume, volume of blood in the left ventricle at the end of diastolic cycle, and ejection fraction, stroke volume/end-diastolic volume. The data analysis is performed using the freely available Segment software developed for processing DICOM-compliant cardiovascular images acquired by magnetic resonance scanners19. The data revealed an increase in thickness of the LV wall in the myocarditic animals, corresponding to a decrease in LV end-diastolic volume, stroke volume and ejection fraction, as compared with these functional parameters in healthy mice.

<table><tbody><tr><td>Myhc-a 334-352 (DSAFDVLSFTAEEKAGVYK)</td><td>Neopeptide, Cambridge, MA</td><td></td><td>Store at 4 &amp;#176;C</td></tr><tr><td>CFA</td><td>Sigma Aldrich, St Louis, MO</td><td>5881</td><td>Store at 4 &amp;#176;C</td></tr><tr><td>MTB H37Rv extract</td><td>Difco Laboratories, Detroit, MI</td><td>231141</td><td>Store at 4 &amp;#176;C</td></tr><tr><td>PT</td><td>List Biologicals Laboratories, Campbell, CA</td><td>181</td><td>Store at 4 &amp;#176;C</td></tr><tr><td>1x PBS</td><td>Corning, Manassas, VA</td><td>21-040-CV</td><td>Store at 4 &amp;#176;C</td></tr><tr><td>Isoflurane</td><td>Piramal Healthcare, Mumbai, India</td><td>NDC66794-013-25</td><td></td></tr><tr><td>Female A/J mice</td><td>Jackson Laboratories, Bar Harbor, ME</td><td>646</td><td></td></tr><tr><td>Luer-lok sterile 1 ml syrringe</td><td>BD, Franklin Lakes, NJ</td><td>309628</td><td></td></tr><tr><td>Luer-lok sterile 3 ml syrringe</td><td>BD, Franklin Lakes, NJ</td><td>309657</td><td></td></tr><tr><td>Sterile needle, 18 G</td><td>BD, Franklin Lakes, NJ</td><td>305195</td><td></td></tr><tr><td>Sterile needle, 27 1/2 G</td><td>BD, Franklin Lakes, NJ</td><td>305109</td><td></td></tr><tr><td>3-way stopcock</td><td>Smiths Medical ASD, Inc. Dublin, OH</td><td>MX5311L</td><td></td></tr><tr><td>Kerlix gauze bandage rolls</td><td>Covidien, Mansfield, MA</td><td>6720</td><td></td></tr><tr><td>Kimwipes</td><td>Kimberly-Clark Professional, Roswell, GA</td><td>34155</td><td></td></tr><tr><td>Protouch Stockinette</td><td>Medline Industries, Mundelein, IL</td><td>30-1001</td><td></td></tr><tr><td>Sterile surgical scissors and forceps</td><td>INOX tool Corporation</td><td></td><td></td></tr><tr><td>Micro oven</td><td>GE Healthcare,&amp;nbsp;</td><td></td><td></td></tr><tr><td>ThermiPAQ hot and cold therapy system&amp;nbsp;</td><td>Theramics Corporation, Springfield, IL</td><td></td><td></td></tr><tr><td>Reptile heating lamp</td><td>Energy Savers Unlimited, Inc. Carson, CA</td><td></td><td></td></tr><tr><td>3M Transpore tapes</td><td>Target Corporation, MN</td><td></td><td></td></tr><tr><td>Up and Up Polymyxin B sulfate/Bacitracin/Neomycin sulfate antibiotic ointment</td><td>Target Corporation, MN</td><td></td><td></td></tr><tr><td>North Safety DeciDamp-2PVC foam ear plugs</td><td>North Safety Products, Smithfield, RI</td><td></td><td></td></tr><tr><td>Cotton tipped applicator, 6&amp;rsquo;&amp;rsquo; wooden stem&amp;nbsp;</td><td>Jorgensen Laboratories, Inc. Loveland, CO</td><td></td><td></td></tr><tr><td>Anesthesia induction chamber&amp;nbsp;</td><td>Summit Anesthesia Solutions, Ann Harbor, MI</td><td></td><td></td></tr><tr><td>Summit Anesthesia Support system for regulating flow of anesthesia&amp;nbsp;</td><td>Summit Anesthesia Solutions, Ann Harbor, MI</td><td></td><td></td></tr><tr><td>Specially designed animal holder</td><td>Agilent Technologies, Santa Clara, CA</td><td></td><td></td></tr><tr><td>Bickford&amp;nbsp;Omnicon F/Air&amp;nbsp;anesthesia gas filter unit&amp;nbsp;</td><td>A.M. Bickford, Inc. Wales Center, NY</td><td></td><td></td></tr><tr><td>Pulse-oximeter module, MR compatible small animal monitoring and gating system&amp;nbsp;</td><td>Small Animal Instruments, Inc. Stony Brook, NY</td><td></td><td></td></tr><tr><td>Oxygen cylinder&amp;nbsp;</td><td>Matheson-Tri Gas, North-Central Zone, Lincoln, NE</td><td></td><td></td></tr><tr><td>Gas regulator&amp;nbsp;</td><td>Western Medica, West Lake, OH</td><td></td><td></td></tr><tr><td>Signal breaking module, MR compatible small animal monitoring and gating system</td><td>Small Animal Instruments, Inc. Stony Brook, NY</td><td></td><td></td></tr><tr><td>9.4 T (400 MHz) 89 mm vertical core bore MR scanner</td><td>Agilent Technologies, Santa Clara, CA</td><td></td><td></td></tr><tr><td>4 cm millipede micro-imaging RF coil</td><td>Agilent Technologies, Santa Clara, CA</td><td></td><td></td></tr><tr><td>SAM PC monitor</td><td>Small Animal Instruments, Inc. Stony Brook, NY</td><td></td><td></td></tr><tr><td>Quantitative Medical Image analysis software</td><td>http://segment.heiberg.se;&amp;nbsp; Segment v1.8 R1430,&amp;nbsp; Medviso, Oresunds region, Sweden</td><td></td><td></td></tr><tr><td>Matlab software</td><td>The Mathworks, Inc.&amp;nbsp; Natick, MA</td><td></td><td></td></tr><tr><td>Computer-Unix operating system</td><td></td><td></td><td></td></tr></tbody></table>

noninvasive assessment of cardiac abnormalities in experimental autoimmune myocarditis by magnetic resonance microscopy imaging in the mouse

Myocarditis is an inflammation of the myocardium, but only ~10% of those affected show clinical manifestations of the disease. To study the immune events of myocardial injuries, various mouse models of myocarditis have been widely used. This study involved experimental autoimmune myocarditis (EAM) induced with cardiac myosin heavy chain (Myhc)-&#945; 334-352 in A/J mice; the affected animals develop lymphocytic myocarditis but with no apparent clinical signs. In this model, the utility of magnetic resonance microscopy (MRM) as a non-invasive modality to determine the cardiac structural and functional changes in animals immunized with Myhc-&#945; 334-352 is shown. EAM and healthy mice were imaged using a 9.4 T (400 MHz) 89 mm vertical core bore scanner equipped with a 4 cm millipede radio-frequency imaging probe and 100 G/cm triple axis gradients. Cardiac images were acquired from anesthetized animals using a gradient-echo-based cine pulse sequence, and the animals were monitored by respiration and pulse oximetry. The analysis revealed an increase in the thickness of the ventricular wall in EAM mice, with a corresponding decrease in the interior diameter of ventricles, when compared with healthy mice. The data suggest that morphological and functional changes in the inflamed hearts can be non-invasively monitored by MRM in live animals. In conclusion, MRM offers an advantage of assessing the progression and regression of myocardial injuries in diseases caused by infectious agents, as well as response to therapies.

In this report, the utility of MRM technique as a non-invasive modality to determine the structural and functional changes in the hearts of animals affected with EAM is shown. Myocarditis was induced in A/J mice by immunizing the animals with Myhc-&#945; 334-352 in CFA7, and the animals were subjected to MRM experiments on day 21 post-immunization. The MRM imaging was performed on live animals under isoflurane anesthesia at 9.4 T (400 MHz for protons) using an 89&#160;mm vertical-bore magnet equipped with triple axis gradients (maximum strength 100 G/cm). A scout image was acquired to locate and position the mouse heart in the center of the FOV, followed by axial images to obtain the long-axis 4-chamber view. The angle at which the heart was imaged for the 2-chamber view is shown in Figure 2A. Cardiac images were acquired using a 4&#160;cm millipede RF imaging probe with a gradient-echo-based cine pulse sequence. Cardiac function measurements (imaging: LV wall thickness; output: LV end-diastolic volume and ejection fraction) were then analyzed using Segment software. Structural defects in the hearts of EAM-affected mice were evidenced by increase in LV thickness by approximately 1.5-fold (p = 0.018) (Figure&#160;2B&#160;and Table 1), with corresponding decrease in LV end-diastolic volume [18.0 &#177; 4.2 &#181;l vs. 37.5 &#177; 3.5 &#181;l, Figure&#160;2C(i); p = 0.002] and ejection fraction [49.4 &#177; 2.3% vs. 71.5 &#177; 6.0%, p = 0.00066; Figure&#160;2C(ii)] as compared to healthy mice. As expected, histological evaluation of hearts from myocarditic, but not healthy, mice showed multifocal lymphocyte infiltrates, as we have demonstrated previously7; Figure&#160;2D). The data suggest that morphological and functional changes in inflamed hearts can be non-invasively monitored by MRM in live animals.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51654/51654fig1highres.jpg" width="500" /> 
Figure 1. Animal preparation and positioning of probes for acquisition of MRM images from the mouse heart. To acquire images from the heart, the anesthetized mouse is placed in the animal holder specially designed for MRM imaging and connected to the air blow heater to maintain the body temperature. Under continuous anesthesia, the animal is immobilized in the prone position. A pneumatic pillow, fiber optic oximetry sensor and temperature sensor are set up to monitor respiration, pulse and body temperature, respectively, until MRM acquisition of cardiac images is complete. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51654/51654fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51654/51654fig2highres.jpg" width="500" /> 
Figure 2. MRM imaging of mice affected with autoimmune myocarditis reveals cardiac abnormalities. Myocarditis was induced in A/J mice by immunizing the animals with Myhc-&#945; 334-352 in CFA. The animals were subjected to MRM imaging on day 21 postimmunization to evaluate cardiac abnormalities. (A) Position of MRM slicing. The angle at which the heart was sliced for image acquisition is shown. (B) MRM imaging. Short-axis slices of heart were captured using echo-based cine pulse sequence in eight time frames with a TR of 500 msec (TE, 5 msec; flip angle, 20&#176;; number of acquisitions, 4; acquisition matrix, 256 x 256) [arrows: LV wall thickness]. (C) Cardiac output. Cardiac output was measured based on (i) LV end-diastolic volume and (ii) ejection fraction in healthy and myocarditic mice using quantitative medical image analysis with Segment software. Mean SEM values for a group of mice are shown (n = 2 to 5 per group). (D) Histology. Hearts from the above treatment groups were evaluated for inflammation by hematoxylin and eosin staining. Circles: multifocal lymphocytic infiltrations. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/51654/51654fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Animals</td>
			<td>Healthy (n = 3)</td>
			<td>EAM (n = 5)</td>
		</tr>
		<tr>
			<td>Mouse 1</td>
			<td>1.03</td>
			<td>1.48</td>
		</tr>
		<tr>
			<td>Mouse 2</td>
			<td>1.3</td>
			<td>1.59</td>
		</tr>
		<tr>
			<td>Mouse 3</td>
			<td>0.94</td>
			<td>1.44</td>
		</tr>
		<tr>
			<td>Mouse 4</td>
			<td></td>
			<td>2.11</td>
		</tr>
		<tr>
			<td>Mouse 5</td>
			<td></td>
			<td>1.92</td>
		</tr>
		<tr>
			<td>Mean &#177; SEM</td>
			<td>1.09 &#177; 0.1</td>
			<td>1.71 &#177; 0.1</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;Comparison of left ventricular (LV) wall thickness between healthy and experimental autoimmune myocarditis (EAM) mice. Three healthy and five EAM-induced mice were subjected to magnetic resonance microscopy (MRM) imaging on day 21 postimmunization. After acquiring the cardiac images by MRM, the thickness of LV wall was measured using Segment software as described in the protocol. The values displayed in the table represent LV wall thickness in mm.

Watch this Scientific Journal Video about Noninvasive Assessment of Cardiac Abnormalities in Experimental Autoimmune Myocarditis by Magnetic Resonance Microscopy Imaging in the Mouse at JoVE.com

Noninvasive Assessment of Cardiac Abnormalities in Experimental Autoimmune Myocarditis by Magnetic Resonance Microscopy Imaging in the Mouse

This study demonstrates the successful establishment of magnetic resonance microscopy imaging as a non-invasive tool to assess the cardiac abnormalities in mice affected with autoimmune myocarditis. The data indicate that the technique can be used to monitor the disease-progression in live animals.

Myocarditis is an inflammation of the myocardium, but only ~10% of those affected show clinical manifestations of the disease. To study the immune events ...

noninvasive-assessment-cardiac-abnormalities-experimental-autoimmune

Nanyang Technological University

Research

JoVE Journal

Medicine

9.9K Views.  University of Nebraska-Lincoln. This study demonstrates the successful establishment of magnetic resonance microscopy imaging as a non-invasive tool to assess the cardiac abnormalities in mice affected with autoimmune myocarditis. The data indicate that the technique can be used to monitor the disease-progression in live animals.

Video: Noninvasive Assessment of Cardiac Abnormalities in Experimental Autoimmune Myocarditis by Magnetic Resonance Microscopy Imaging in the Mouse

High-frequency High-resolution Echocardiography: First Evidence on Non-invasive Repeated Measure of Myocardial Strain, Contractility, and Mitral Regurgitation in the Ischemia-reperfused Murine Heart

Ischemia-reperfusion (IR) was surgically performed in murine hearts which were then subjected to repeated imaging to monitor temporal changes in functional parameters of key clinical significance. Two-dimensional movies were acquired at high frame rate (8 kHz) and were utilized to estimate high-quality myocardial strain. Two-dimensional elastograms (strain images), as well as strain profiles, were visualized. Results were powerful in quantitatively assessing IR-induced changes in cardiac events including left-ventricular (LV) contraction, LV relaxation and isovolumetric phases of both pre-IR and post-IR beating hearts in intact mice. In addition, compromised sector-wise wall motion and anatomical deformation in the infarcted myocardium were visualized.  The elastograms were uniquely able to provide information on the following parameters in addition to standard physiological indices that are known to be affected by myocardial infarction in the mouse: internal diameters of mitral valve orifice and aorta, effective regurgitant orifice, myocardial strain (circumferential as well as radial), turbulence in blood flow pattern as revealed by the color Doppler movies and velocity profiles, asynchrony in LV sector, and changes in the length and direction of vectors demonstrating slower and asymmetrical wall movement. This work emphasizes on the visual demonstration of how such analyses are performed. 

High-frequency High-resolution Echocardiography: First Evidence on Non-invasive Repeated Measure of Myocardial Strain, Contractility, and Mitral Regurgitation in the Ischemia-reperfused Murine Heart 

High frequency Doppler ultrasound is a novel technology for assessing regional myocardial function. This work presents first evidence demonstrating applicability of this versatile imaging platform for the repeated measure of myocardial strain, dp/dt, and mitral regurgitation in the ischemia-reperfused (IR) murine heart.

Ischemia-reperfusion (IR) was surgically performed in murine hearts which were then subjected to repeated imaging to monitor temporal changes in ...

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 ...

MRI and PET in Mouse Models of Myocardial Infarction

Myocardial infarction is one of the leading causes of death in the Western world. The similarity of the mouse heart to the human heart has made it an ideal model for testing novel therapeutic strategies.
In vivo magnetic resonance imaging (MRI) gives excellent views of the heart noninvasively with clear anatomical detail, which can be used for accurate functional assessment. Contrast agents can provide basic measures of tissue viability but these are nonspecific. Positron emission tomography (PET) is a complementary technique that is highly specific for molecular imaging, but lacks the anatomical detail of MRI. Used together, these techniques offer a sensitive, specific and quantitative tool for the assessment of the heart in disease and recovery following treatment.
In this paper we explain how these methods are carried out in mouse models of acute myocardial infarction. The procedures described here were designed for the assessment of putative protective drug treatments. We used MRI to measure systolic function and infarct size with late gadolinium enhancement, and PET with fluorodeoxyglucose (FDG) to assess metabolic function in the infarcted region. The paper focuses on practical aspects such as slice planning, accurate gating, drug delivery, segmentation of images, and multimodal coregistration. The methods presented here achieve good repeatability and accuracy maintaining a high throughput.

We describe how to perform MRI and PET imaging of the mouse heart. The protocol is tailored to assess treatment efficacy in models of myocardial infarction and heart failure.

Myocardial infarction is one of the leading causes of death in the Western world. The similarity of the mouse heart to the human heart has made it an ...

In Vivo Quantitative Assessment of Myocardial Structure, Function, Perfusion and Viability Using Cardiac Micro-computed Tomography

The use of Micro-Computed Tomography (MicroCT) for in vivo studies of small animals as models of human disease has risen tremendously due to the fact that MicroCT provides quantitative high-resolution three-dimensional (3D) anatomical data non-destructively and longitudinally. Most importantly, with the development of a novel preclinical iodinated contrast agent called eXIA160, functional and metabolic assessment of the heart became possible. However, prior to the advent of commercial MicroCT scanners equipped with X-ray flat-panel detector technology and easy-to-use cardio-respiratory gating, preclinical studies of cardiovascular disease (CVD) in small animals required a MicroCT technologist with advanced skills, and thus were impractical for widespread implementation. The goal of this work is to provide a practical guide to the use of the high-speed Quantum FX MicroCT system for comprehensive determination of myocardial global and regional function along with assessment of myocardial perfusion, metabolism and viability in healthy mice and in a cardiac ischemia mouse model induced by permanent occlusion of the left anterior descending coronary artery (LAD).

In Vivo Quantitative Assessment of Myocardial Structure, Function, Perfusion and Viability Using Cardiac Micro-computed Tomography

This method outlines the use of Quantum Micro-Computed Tomography (MicroCT) to assess cardiac morphology, function, perfusion, metabolism and viability with iodinated contrast agent in mice with experimentally-induced myocardial ischemia. The technique can be applied for non-destructive high-throughput longitudinal in vivo imaging of various animal models of human heart disease.

The use of Micro-Computed Tomography (MicroCT) for in vivo studies of small animals as models of human disease has risen tremendously due to the fact that ...

Bioengineering

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 ...

Quantification of Mouse Heart Left Ventricular Function, Myocardial Strain, and Hemodynamic Forces by Cardiovascular Magnetic Resonance Imaging

Mouse models have contributed significantly to understanding genetic and physiological factors involved in healthy cardiac function, how perturbations result in pathology, and how myocardial diseases may be treated. Cardiovascular magnetic resonance imaging (CMR) has become an indispensable tool for a comprehensive in vivo assessment of cardiac anatomy and function. This protocol shows detailed measurements of mouse heart left ventricular function, myocardial strain, and hemodynamic forces using 7-Tesla CMR. First, animal preparation and positioning in the scanner are demonstrated. Survey scans are performed for planning imaging slices in various short- and long-axis views. A series of prospective ECG-triggered short-axis (SA) movies (or CINE images) are acquired covering the heart from apex to base, capturing end-systolic and end-diastolic phases. Subsequently, single-slice, retrospectively gated CINE images are acquired in a midventricular SA view, and in 2-, 3-, and 4-chamber views, to be reconstructed into high-temporal resolution CINE images using custom-built and open-source software. CINE images are subsequently analyzed using dedicated CMR image analysis software.
Delineating endomyocardial and epicardial borders in SA end-systolic and end-diastolic CINE images allows for the calculation of end-systolic and end-diastolic volumes, ejection fraction, and cardiac output. The midventricular SA CINE images are delineated for all cardiac time frames to extract a detailed volume-time curve. Its time derivative allows for the calculation of the diastolic function as the ratio of the early filling and atrial contraction waves. Finally, left ventricular endocardial walls in the 2-, 3-, and 4-chamber views are delineated using feature-tracking, from which longitudinal myocardial strain parameters and left ventricular hemodynamic forces are calculated. In conclusion, this protocol provides detailed in vivo quantification of the mouse cardiac parameters, which can be used to study temporal alterations in cardiac function in various mouse models of heart disease.

This study describes a comprehensive cardiovascular magnetic resonance imaging (CMR) protocol to quantify the left ventricular functional parameters of the mouse heart. The protocol describes the acquisition, post-processing, and analysis of the CMR images as well as assessment of different cardiac functional parameters.

Mouse models have contributed significantly to understanding genetic and physiological factors involved in healthy cardiac function, how perturbations ...

Quantification of Biventricular Function and Morphology by Cardiac Magnetic Resonance Imaging in Mice with Pulmonary Artery Banding

Right ventricular (RV) function and failure are major determinants of outcome in acquired and congenital heart diseases, including pulmonary hypertension. Assessment of RV function and morphology is complex, partly due to the complex shape of the RV. Currently, cardiac magnetic resonance (CMR) imaging is the golden standard for noninvasive assessment of RV function and morphology. The current protocol describes CMR imaging in a mouse model of RV pressure load induced by pulmonary artery banding (PAB). PAB is performed by placing a 6-0 suture around the pulmonary artery over a 23 G needle. The PAB gradient is determined using echocardiography at 2 and 6 weeks. At 6 weeks, the right and left ventricular morphology and function is assessed by measuring both end-systolic and end-diastolic volumes and mass by ten to eleven cine slices 1 mm thick using a 9.4 T magnetic resonance imaging scanner equipped with a 1,500 mT/m gradient. Representative results show that PAB induces a significant increase in RV pressure load, with significant effects on biventricular morphology and RV function. It is also shown that at 6 weeks of RV pressure load, cardiac output is maintained. Presented here is a reproducible protocol for the quantification of biventricular morphology and function in a mouse model of RV pressure load and may serve as a method for experiments exploring determinants of RV remodeling and dysfunction.

To understand the pathophysiology of right ventricular (RV) adaptation to abnormal loading, experimental models are crucial. However, assessment of RV dimensions and function is complex and challenging. This protocol provides a method to perform cardiac magnetic resonance imaging (CMR) as a noninvasive benchmark procedure in mice subjected to RV pressure load.

Right ventricular (RV) function and failure are major determinants of outcome in acquired and congenital heart diseases, including pulmonary hypertension. ...

Induction of Myocardial Infarction and Myocardial Ischemia-Reperfusion Injury in Mice

Acute myocardial infarction is a common cardiovascular disease with high mortality. Myocardial reperfusion injury can counteract the beneficial effects of heart reflow and induce secondary myocardial injury. A simple and reproducible model of myocardial infarction and myocardial ischemia-reperfusion injury is a good tool for researchers. Here, a customizable method to create a myocardial infarction (MI) model and MIRI by precision ligation of the left anterior descending coronary artery (LAD) through micromanipulation is described. Accurate and reproducible ligature positioning of the LAD helps obtain consistent results for heart injury. ST-segment changes can help to identify model accuracy. The serum level of cardiac troponin T (cTnT) is used to assess the myocardial injury, cardiac ultrasound is employed to evaluate the myocardial systolic function, and Evans-Blue/triphenyl tetrazolium chloride staining is used to measure infarct size. In general, this protocol reduces procedure duration, ensures controllable infarct size, and improves mouse survival.

Here we describe a simple and reproducible method that can induce myocardial infarction or myocardial ischemia-reperfusion injury in mice by precision ligation of the left anterior descending coronary artery through micromanipulation.

Acute myocardial infarction is a common cardiovascular disease with high mortality. Myocardial reperfusion injury can counteract the beneficial effects of ...

An Experimental Model of Myocardial Infarction for Studying Cardiac Repair and Remodeling in Knockout Mice

Cardiovascular disease is the most prevalent cause of death in Western countries, with acute myocardial infarction (MI) being the most prevalent form. This paper describes a protocol for studying the role of galectin 3 (Gal-3) in the temporal evolution of cardiac healing and remodeling in an experimental animal model of MI. 
The procedures described include an experimental model of MI with a permanent coronary ligature in male C57BL/6J (control) and Gal-3 knockout (KO) mice, an echocardiography procedure to study cardiac remodeling and systolic function in vivo, a histological evaluation of interstitial myocardial fibrosis with picrosirius red-stained and rhodamine-conjugated lectin-stained sections for studying myocyte hypertrophy by the cross-sectional area (MCSA), and the quantification of infarct size and cardiac remodeling (scar thinning, septum thickness, and expansion index) by planimetry in slices stained with Masson's trichrome and triphenyl tetrazolium chloride. Gal-3 KO mice with MI showed disrupted cardiac remodeling and an increase in the scar thinning ratio and the expansion index. At the onset of MI, myocardial function and cardiac remodeling were also severely affected. At 4 weeks post MI, the natural evolution of fibrosis in infarcted Gal-3 KO mice was also affected. 
In summary, the experimental model of MI is a suitable model for studying the temporal evolution of cardiac repair and remodeling in mice with the genetic deletion of Gal-3 and other animal models. The lack of Gal-3 affects the dynamics of cardiac repair and disrupts the evolution of cardiac remodeling and function after MI.

Here, we describe an experimental model of myocardial infarction, an echocardiography procedure to study cardiac remodeling and function, and procedures for quantifying fibrosis and hypertrophy in picrosirius red-stained and rhodamine-stained sections, as well as the infarct size and expansion index in slices stained with Masson's trichrome.

Cardiovascular disease is the most prevalent cause of death in Western countries, with acute myocardial infarction (MI) being the most prevalent form. ...

Mesoscopic Optical Imaging of Whole Mouse Heart

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition (fibrosis) and cellular misalignment, can affect electrical conduction, introduce electromechanical dysfunctions and, eventually, lead to arrhythmia. Current predictive models of these functional alterations are based on non-integrated and low-resolution structural information. Placing this framework on a different order of magnitude is challenging due to the inefficacy of standard imaging methods in performing high-resolution imaging in massive tissue. In this work, we describe a methodological framework that allows imaging of whole mouse hearts with micrometric resolution. The achievement of this goal has required a technological effort where advances in tissue transformation and imaging methods have been combined. First, we describe an optimized CLARITY protocol capable of transforming an intact heart into a nanoporous, hydrogel-hybridized, lipid-free form that allows high transparency and deep staining. Then, a fluorescence light-sheet microscope able to rapidly acquire images of a mesoscopic field of view (mm-scale) with the micron-scale resolution is described. Following the mesoSPIM project, the conceived microscope allows the reconstruction of the whole mouse heart with micrometric resolution in a single tomographic scan. We believe that this methodological framework will allow clarifying the involvement of the cytoarchitecture disarray in the electrical dysfunctions and pave the way for a comprehensive model that considers both the functional and structural data, thus enabling a unified investigation of the structural causes that lead to electrical and mechanical alterations after the tissue remodeling.

We report a method for mesoscopic reconstruction of the whole mouse heart by combining new advancements in tissue transformation and staining with the development of an axially scanned light-sheet microscope.

Both genetic and non-genetic cardiac diseases can cause severe remodeling processes in the heart. Structural remodeling, such as collagen deposition ...

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