IN VIVO MRI OF MOUSE HEART AT 11.7 T: MONITORING OF STEM‐CELL THERAPY FOR MYOCARDIAL INFARCTION AND EVALUATION OF CARDIAC HYPERTROPHY DISSERTATION Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University By Aditi C. Kulkarni, M. Sc. The Ohio State University 2008 Dissertation Committee Approved by Periannan Kuppusamy, PhD, Adviser Petra Schmalbrock, PhD ‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐‐ Hiranmoy Das, PhD Adviser Graduate Program in Biophysics
Copyright by Aditi C. Kulkarni 2008 ABSTRACT Cardiovascular disease (CVD) is one of the major causes of morbidity and mortality in the western world. It accounts for more than a third of the deaths in the United States and is a serious cause of concern. Early detection of the abnormal cardiovascular conditions may help in their diagnosis and treatment, thus, reducing the mortality associated with them. Visualization of the heart and the blood vessels may help in the detection and diagnosis of these diseases in their early stage of development. Many techniques such as X‐ray computed tomography (CT), angiography, magnetic resonance imaging (MRI), echocardiography and nuclear imaging are used for cardiac imaging in the clinic. Among these, MRI stands out due to its advantages such as noninvasiveness, high spatial and temporal resolution, and repeatability and reproducibility of the measurements. It can also be used to ii
obtain functional data from the structural information of the heart, making it a valuable diagnostic tool for various cardiac pathologies. This dissertation reports the development of cardiac MR imaging methods and its application to cardiac disease models at high magnetic field (11.7 T). Mouse was chosen as the animal model for studying cardiac disease due to several reasons including anatomical similarity of mouse and human hearts, similarity of murine cardiovascular disorders to human, and ease of genetic manipulation in mouse. The development of a cardiac MRI (CMRI) method for mouse at high magnetic field (11.7 T) was the main objective of this dissertation. Optimization of hardware and software parameters was performed to obtain images of the mouse heart, which is characterized by its small size and very fast motion. Methods for data analysis were developed to obtain functional data (ejection fraction, stroke volume, cardiac output etc) from the anatomical and cine‐images of the mouse heart. M‐mode echocardiography (ECHO) is one of the most widely used methods for functional analysis of the heart. Therefore, CMRI was compared to ECHO for functional analysis of healthy and infarcted mouse heart. Although the absolute iii
values of functional parameters obtained from the ECHO and CMRI were different, they showed similar trends over time. In general, there was agreement in the ejection fraction measurements made obtained from the two methods. After development and validation of the method, CMRI was employed for structural and functional evaluation of mouse heart after myocardial infarction (MI). MI was induced in the mice by permanent ligation of the left coronary artery. CMRI was able to detect the structural changes in the MI heart. It also showed a significant decrease in the function of mouse heart after MI. Another application of CMRI was to assess the cardiac hypertrophy in a transgenic mouse model of chronic hypertension. The hypothesis was that chronic hypertension, created by the transgenic model, leads to cardiac hypertrophy. Previous studies using ECHO were not able to substantially support this hypothesis. However, CMRI of these transgenic mice detected the decrease in the cardiac function and increase in the wall of the left ventricle, thus validating the model itself. Thus, CMRI proved to be a valuable and unique diagnostic tool for assessing cardiac hypertrophy. iv
Cardiomyoplasty, or implantation of stem cells into the infarcted heart muscle, is emerging as a promising approach for cardiac therapy. Different types of stem cells have been extensively studied for cardiac tissue regeneration, but very few studies have investigated the fate of transplanted stem cells. In this study, CMRI was used to monitor labeled stem cells that were transplanted in the mouse heart after MI. Mouse mesenchymal stem cells were labeled with superparamagnetic iron oxide particles (SPIO), before transplantation in the mouse heart. An optimized CMRI method was developed and tested in vitro, as well as, ex vivo. CMRI enabled the detection of the cells in vivo and their visualization for up to 4 weeks. In conclusion, in this dissertation the development and validation of cardiac MRI methods for imaging mouse heart at high magnetic field (11.7 T) have been performed successfully. These methods were applied to the evaluation of myocardial infarction and cardiac hypertrophy in mouse and the monitoring of labeled stem cells after their transplantation in the infarcted heart. v
To Aai-Baba
For believing in me, always…
vi
ACKNOWLEDGMENTS This dissertation work is the culmination of twenty‐odd years of educational journey. I would like to begin by thanking all who have been directly or indirectly associated it and have influenced me. Although I may not be able to mention all the names, I would like to thank you all for helping me work towards my goal. Firstly, I would like to express my sincere gratitude towards my adviser Dr. P Kuppusamy for giving me the opportunity to work on this project. Despite his myriad responsibilities, he was always available for his students and ensured that we received all the resources and support needed for our projects. I have learnt a lot from him, on professional as well as personal levels. It’s a testimony to his deep sense of responsibility, genuine caring attitude and confidence in his students that I am completing my dissertation work in the field of magnetic resonance imaging. I would like to thank my dissertation committee members Prof. Petra Schmalbrock and Prof. Hiranmoy Das for their help and support. I am thankful to Prof. Jay L Zweier and Prof. Chandan K Sen for being a part of my candidacy examination committee. Special thanks to Prof. Schmalborck for her staunch support during some rough patches which would have been difficult to cross without her on my side. vii
I am thankful to Dr. Mahmood Khan for his help with the animal models. He helped me focus my project work and always had a few words of encouragement. His enthusiasm is infectious. I would also like to thank our collaborators, Dr. Hamdy Hassanain and Mahmood Awad for their help with the transgenic animal model. Dr. Anna Bratasz has the sole credit for making me animal‐ and EPR‐literate. She is the best lab instructor ever! Thanks to Dr. Surya Gnyawali for guiding me when I was lost in the jungle of cardiac MRI optimization. Nancy Trigg and Brian Rivera were my genies who provided me with whatever I needed whenever I needed it. I am especially grateful to M. Lakshmi Kuppusamy for making Columbus home away from home. Her love and care, not to forget the scrumptious idli‐dosas, have saved my mother countless hours of worry. Overall, I was fortunate to be a part of a happy and nourishing workgroup that helped me grow as a researcher and as a person. Thank you all! This journey would have been very lonely without my friends. I am thankful to all my friends for their love and support. Thanks to Deepti, Guru, Jonathan, Shabnam, Simi, Soma and Vinh for making Columbus and grad school an enjoyable experience. I owe a huge debt of gratitude to Deepti, Anamika, Subhangi and Aparajita. Even after being with me physically and (more often than not) telephonically for over a decade, they still consider me a close friend. If that is not the proof of their courage and patience, I don’t know what is! viii
I would also like to thank my mentors back home who helped shape my mind. I am grateful to Prof. K. C. Sharma, Prof. P. N. Sen and Dr. A. P. Kesarkar for taking me under their wing and easing me into research. I am blessed with a wonderful family. Their unconditional love and unwavering support has often humbled me. They were with me every step of the way, especially when the road got a bit too bumpy. I am grateful to be a part of their life. Abhijit has been the lighthouse of this journey. No part of this would ever have happened without him in my life. Thank you, for not letting me give up when I was tempted to take the easy way out, and giving me the strength and courage to continue. And lastly,’Too karta, mee tar karaN; tujhech deNe, tulach arpaN’. ix
VITA December 1979...................................................Born – Pune, India 2000 .....................................................................B.Sc., Physics, University of Pune, Pune, India 2002 .....................................................................M.Sc., Physics, University of Pune, Pune, India 2005 ‐ present .....................................................Graduate Research Associate, The Ohio State University, Columbus, Ohio PUBLICATIONS Kulkarni AC, Bratasz A, Rivera B, Krishna MC, Kuppusamy P. Redox Mapping of Biological Samples Using EPR Imaging. Israel J. Chem. 2008; 48(1): 27‐31 Kulkarni AC, Kuppusamy P, Parinandi N. Oxygen, the lead actor in the pathophysiologic drama: enactment of the trinity of normoxia, hypoxia, and hyperoxia in disease and therapy. Antioxid Redox Signal. 2007 Oct;9(10):1717‐30. Bratasz A, Kulkarni AC, Kuppusamy P.A highly sensitive biocompatible spin probe for imaging of oxygen concentration in tissues. Biophys J. 2007 Apr 15;92(8):2918‐25.. Raut JS, Bhattad P, Kulkarni AC, Naik VM. ʺMicro‐potteryʺ‐‐marangoni effect driven assembly of amphiphilic fibers. Langmuir. 2005 Jan 18;21(2):516‐9. FIELDS OF STUDY Major Field: Biophysics x
TABLE OF CONTENTS Abstract ................................ .............................................................................................ii Dedication ……………………………………………………………………………....vi Acknowledgments...…………………………………………………………………..vii Vita …………………………………………………………………………………….....x List of Tables...………………………………………………………………………...xvi List of Figures.………………………………………………………………………..xvii Chapters: 1.
Introduction ........................................................................................................................... 1
1.1
Heart: Structure and function .................................................................................... 1
1.2
Choosing mouse model .............................................................................................. 3
1.3
Need for cardiac imaging ........................................................................................... 5
1.4
Cardiac imaging techniques....................................................................................... 5
1.5
Literature review for CMRI in mouse ...................................................................... 9
1.6
List of abbreviations .................................................................................................. 13
1.7
Overview of the dissertation.................................................................................... 14
2.
Cardiac magnetic resonance imaging .............................................................................. 15
2.1
Introduction................................................................................................................ 15
2.2
Protocol for cardiac imaging of mouse................................................................... 17
2.2.1
MR scanner used for mouse cardiac imaging ................................................... 17
xi
2.2.2
Animal preparation and set up ........................................................................... 18
2.2.3
Gating strategies for triggered cardiac imaging ............................................... 19
2.2.4
Cardiac MR data acquisition – choice of pulse sequence ................................ 24
2.2.5
Procedure for getting short‐axis images of mouse heart ................................. 29
2.2.6
MR Image processing to get cardiac functional parameters ........................... 31
2.2.7
Statistical methods used for data analysis ......................................................... 34
2.3
Results and discussion .............................................................................................. 35
2.4
Summary..................................................................................................................... 36
3.
Comparison of MRI and echocardiography to assess cardiac function in a mouse model of myocardial infarction................................................................................................. 37
3.1
3.1.1
Motivation for the study ...................................................................................... 37
3.1.2
Cardiac functional analysis by echocardiography ........................................... 38
3.1.3
Study design .......................................................................................................... 39
3.2
Introduction................................................................................................................ 37
Materials and Methods ............................................................................................. 40
3.2.1
Animals................................................................................................................... 40
3.2.2
Induction of myocardial infarction..................................................................... 41
3.2.3
Cardiac MR imaging............................................................................................. 42
3.2.4
Image processing................................................................................................... 42
3.2.5
Echocardiography measurements ...................................................................... 43
3.2.6
Statistical analysis.................................................................................................. 43
3.3
Results ......................................................................................................................... 43
3.4
Discussion................................................................................................................... 49
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3.5
Summary and Conclusion ........................................................................................ 51
4.
Structural and functional evaluation of mouse heart after myocardial infarction .... 52
4.1
Introduction................................................................................................................ 52
4.1.1
Myocardial infarction ........................................................................................... 52
4.1.2
Left coronary artery ligation model for MI ....................................................... 53
4.1.3
Study design .......................................................................................................... 55
4.2
Materials and Methods ............................................................................................. 55
4.2.1
Animals................................................................................................................... 55
4.2.2
Induction of myocardial infarction..................................................................... 55
4.2.3
Oxygen measurement (pO2) using EPR oximetry ............................................ 55
4.2.4
Cardiac MR imaging............................................................................................. 56
4.2.5
Image processing................................................................................................... 56
4.2.6
Visualization of fibrosis........................................................................................ 56
4.2.7
Statistical analysis.................................................................................................. 57
4.3
Results ......................................................................................................................... 57
4.4
Discussion................................................................................................................... 64
4.5
Summary ad Conclusion .......................................................................................... 66
5.
Noninvasive assessment of cardiac function in a transgenic model of cardiac hypertrophy ................................................................................................................................. 67
5.1
Introduction................................................................................................................ 67
5.1.1
Cardiac hypertrophy............................................................................................. 67
5.1.2
CMRI as a tool for the detection and diagnosis of cardiac hypertrophy....... 69
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5.1.3
Transgenic animal model of cardiac hypertrophy ........................................... 70
5.1.4
Study design .......................................................................................................... 72
5.2
Materials and Methods ............................................................................................. 72
5.2.1
Animals................................................................................................................... 72
5.2.2
Cardiac MR imaging............................................................................................. 73
5.2.3
Image processing................................................................................................... 73
5.2.4
Echocardiography studies.................................................................................... 73
5.2.5
Statistical analysis.................................................................................................. 74
5.3
Results ......................................................................................................................... 74
5.4
Discussion................................................................................................................... 81
5.5
Summary and Conclusion ........................................................................................ 83
6.
In vivo monitoring of SPIO‐labeled stem cells transplanted in infarct mouse heart . 84
6.1
6.1.1
Stem‐cell therapy for cardiac repair and regeneration .................................... 84
6.1.2
Stem‐cell tracking using MRI .............................................................................. 86
6.1.3
Study design .......................................................................................................... 88
6.2
Introduction................................................................................................................ 84
Materials and Methods ............................................................................................. 88
6.2.1
Animals................................................................................................................... 88
6.2.2
Culturing of MSCs ................................................................................................ 88
6.2.3
Labeling of MSCs with SPIOs.............................................................................. 89
6.2.4
Myocardial infarction and cell transplantation................................................. 89
6.2.5
In vitro MR imaging .............................................................................................. 89
6.2.6
Ex‐vivo imaging ..................................................................................................... 89
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6.2.7
In‐vivo imaging ...................................................................................................... 90
6.2.8
Prussian blue staining........................................................................................... 90
6.3
Results ......................................................................................................................... 90
6.4
Discussion................................................................................................................... 95
6.5
Summary and Conclusion ........................................................................................ 98
7.
Summary .............................................................................................................................. 99
Bibliography .............................................................................................................................. 102
xv
LIST OF TABLES Table 1.1 Comparison of cardiac imaging modalities.............................................................. 9
Table 1.2 List of frequently used abbreviations ...................................................................... 14
Table 3.1 Cardiac functional parameters obtained from CMRI and ECHO ....................... 46
Table 5.1: Types of cardiac hypertrophy and their effects on the cardiac tissue .............. 69
Table 5.2: Cardiac functional parameters obtained from CMRI and echocardiography.. 77
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LIST OF FIGURES Figure 1.1 Anatomy of human heart .......................................................................................... 2
Figure 1.2 Anatomical similarities of postnatal mouse and human heart ............................ 4
Figure 1.3 Cardiac imaging modalities ...................................................................................... 8
Figure 2.1 Steps in the development and optimization of CMR imaging protocol........... 16
Figure 2.2 MR scanner used for mouse cardiac imaging ...................................................... 17
Figure 2.3 Monitoring of heart rate and respiration of the mouse....................................... 19
Figure 2.4 Small animal monitoring and gating system used for cardiac imaging ........... 21
Figure 2.5 Additional measures to improve signal‐to‐noise ratio of the ECG signal........ 22
Figure 2.6 Quality of ECG signal and optimization of detection parameters .................... 23
Figure 2.7 Timing diagram of FLASH sequence used for cardiac imaging........................ 26
Figure 2.8 Acquisition of cine‐loop images of the heart ........................................................ 27
Figure 2.9 Unsegmented k‐space filling of an ECG‐triggered GRE sequence.................... 28
Figure 2.10 Protocol for obtaining short‐axis cardiac images of mouse.............................. 30
Figure 2.11 Regions of interests (ROIs) with and without papillary muscles .................... 32
Figure 2.12 Manual planimetry of mouse heart for functional analysis ............................. 33
Figure 2.13 Choice of gating for image acquisition................................................................ 35
Figure 2.14 Bright‐blood images of mouse heart at end‐diastolic phase ............................ 36
Figure 3.1 Typical M‐mode echocardiogram of a mouse heart............................................ 39
Figure 3.2 Slice positioning in CMRI and ECHO ................................................................... 40
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Figure 3.3 CMR images of mouse hearts at 4 weeks post‐MI............................................... 44
Figure 3.4 M‐mode echocardiograms of Control and MI mice at 4 weeks post‐MI .......... 45
Figure 3.5 Comparison of EF of control and MI groups........................................................ 45
Figure 3.6 Correlations between EDV and ESV measurements of ECHO and MRI ......... 46
Figure 3.7 Comparison of mid‐papillary MRI with ECHO................................................... 47
Figure 3.8 Comparison of global MRI with ECHO ................................................................ 48
Figure 3.9 Reproducibility of EF measurements by ECHO and MRI.................................. 48
Figure 4.1 Myocardial infarction as a result of occlusion in the left coronary artery........ 53
Figure 4.2 Ligation of the left coronary artery in mice. ......................................................... 54
Figure 4.3 Measurement of myocardial oxygenation using EPR oximetry ........................ 58
Figure 4.4 Bright‐blood images of control and MI heart as a function of distance from the apex......................................................................................................................................... 59
Figure 4.5 Structural changes in mouse heart after myocardial infarction......................... 59
Figure 4.6 LV remodeling in the mouse heart 4 weeks after the surgery ........................... 60
Figure 4.7 Cine images of the cardiac cycle of control and MI heart................................... 61
Figure 4.8 Progressive loss of cardiac function after MI ....................................................... 62
Figure 4.9 Variability in the extent of remodeling in MI....................................................... 63
Figure 4.10 Histological assessment of myocardial infarction in mouse heart .................. 63
Figure 5.1 Transgenic mouse model of cardiac hypertrophy used in this study............... 70
Figure 5.2 MR images of transgenic (mutant RacD overexpressed) and control mouse hearts............................................................................................................................................. 74
xviii
Figure 5.3 Short‐axis images of hearts of control and transgenic mouse hearts at end‐
diastolic and end‐systolic states................................................................................................ 75
Figure 5.4 Mid‐ventricular short‐axis images of control and transgenic mouse hearts... 76
Figure 5.5 Cardiac functional parameters computed from MRI and ECHO of mouse hearts............................................................................................................................................. 78
Figure 5.6 Comparison of LV width and EF obtained from MRI and echocardiography of mouse hearts................................................................................................................................ 79
Figure 5.7 Effect of age on ejection fraction, LV mass and body weight ............................ 80
Figure 6.1 Confirmation of labeling of MSCs with SPIOs.................................................... 91
Figure 6.2 In vitro images of SPIO‐labeled stem cells............................................................. 91
Figure 6.3 Ex vivo images of mouse heart after transplantation of the labeled cells ......... 92
Figure 6.4 Locating the SPIO‐labeled stem cells in mouse heart.......................................... 93
Figure 6.5 Monitoring the SPIO‐labeled stem cells in mouse heart (Short‐axis images) . 94
Figure 6.6 Monitoring the SPIO‐labeled stem cells in mouse heart (Long‐axis images) . 95
Figure 6.7 Prussian blue staining 4 weeks after stem‐cell transplantation ......................... 96
xix
CHAPTER 1 INTRODUCTION 1.
Introduction The overall goal of this dissertation was to develop cardiac magnetic resonance methods for imaging mouse heart at 11.7 T. This chapter provides the general introduction to the work done as a part of this dissertation. It begins with an account of heart structure and function. It explains the choice of mouse as an animal model for studying cardiac pathologies. Further, it discusses the need for cardiac imaging and provides a brief review of currently used noninvasive cardiac imaging methods. This is followed by a report on the current status of cardiac MRI in studying mouse heart. An overview of the dissertation is given at the end of this chapter. 1.1
HEART: STRUCTURE AND FUNCTION Human heart is a four‐chambered muscular organ that pumps blood throughout the body. Heart is situated in the thoracic cavity. It is covered by pericardium and surrounded by lungs. Figure 1.1 shows the anatomy of the human heart, illustrating the four chambers of the heart along with the major blood vessels and valves. The right side of the heart has deoxygenated blood while the left side handles oxygenated blood. The atria pump blood into the ventricles while the ventricles pump blood to the lungs (right ventricles) and to the body (left ventricle). The flow of the blood into and out of the ventricles is controlled by various unidirectional valves. 1
In humans, heart beats about 60‐80 times per minute. This motion is involuntary and is controlled by the cardiac electrical conduction system and neural input. The perpetual contraction‐relaxation of the heart is responsible for sustaining a positive blood pressure in the arteries, ensuring adequate supply of blood and nutrients to body organs. The efficiency of cardiac function is measured using many parameters termed as cardiac functional parameters. Figure 1.1 Anatomy of human heart Human heart consists of four chambers (two atria and two ventricles) that are supplied by major blood vessels. The valves (mitral and tricuspid) regulate the blood flow between the chambers and into the blood vessels. Cardiac functional parameters are mainly used for quantifying the pumping capacity of the heart. Since left ventricle (LV) is responsible for blood perfusion to body organs, unless otherwise specified, these parameters are used for describing LV function. The main parameters used for describing function of the heart are obtained from the end‐diastolic volume (EDV) and end‐systolic volume (ESV) of the LV. 2
The functional parameters are: •
End‐diastolic volume (EDV): Volume of the LV chamber when heart is completely relaxed •
End‐systolic volume (ESV): Volume of the LV chamber when the heart is completely contracted •
Stroke volume (SV): Amount of blood pumped out of the LV in one heart cycle •
Cardiac output (CO): Volume of the blood being pumped by the LV in a minute •
Ejection fraction (EF): Fraction of the blood pumped out of the ventricle per heart cycle The normal values of these parameters vary with the measurement techniques and the health of the individual. However, there are some general guidelines to help the physicians determine the efficiency of cardiac function. For instance, the normal value of EF is considered between 60‐80 %. Most of the cardiac pathologies can be diagnosed on the basis of decreased efficiency of the heart that is reflected in the functional parameters. To study cardiac disease and their effect on the function of the heart, many animal models, such as sheep, pigs, dogs, have been used. Recently small animal models such as rats and mice are being used for these studies. For this dissertation work, mouse model was chosen over the others. 1.2
CHOOSING MOUSE MODEL Mouse model has many advantages over other small and large animal models that have been used to study cardiovascular disease [1]. Mice are relatively inexpensive, have small gestation period and a short life‐span. Mice also share many human phenotypes and pathologies. Almost 97% of mouse genes are similar to that of human [2] and it is easy to create transgenic mouse models to study various human diseases. The only disadvantage of using mouse model is the need to miniaturize appropriate physiological techniques due to the small size of mouse organs. 3
Figure 1.2 Anatomical similarities of postnatal mouse and human heart The anatomical similarities of mouse and human hearts can be seen in the scanning electron micrograph of postnatal mouse heart (A) and an image of postnatal human heart (B) Both the hearts are four‐chambered with two atria and two ventricles separated by a septum. Adapted from Wessels et al [3]. Since the advent of transgenic mouse technology, mice have been extensively used to study cardiovascular diseases. Mouse heart is anatomically quite similar to the human heart [3]. Both the species have four‐chambered heart. The two atria are separated by an interarterial septum and the two ventricles are separated by interventricular septum (Figure 1.2). Few of the main differences include the size of the heart and the heart rate. Mouse heart is about 0.2 g whereas human heart weighs about 200‐300 g. The average beating rate of a mouse heart is about 10 ten times that of human. These differences are characterized by allometric equations [4‐8]. Thus, mouse heart can be used as a good model for studying cardiovascular diseases. 4
1.3
NEED FOR CARDIAC IMAGING Cardiovascular disease (CVD) refers to a class of diseases related to the heart and the vascular system. Hypertension, coronary heart disease, heart failure, and stroke, all come under the same umbrella of CVD. One in three American adults (estimated 80,700,000) have one or more types of CVD [9]. It is the number one killer in the western world causing more mortality than cancer, accidents, and diabetes mellitus combined. In 2004, CVD accounted for 1 in every 2.8 deaths in America. This translates to 2400 deaths every day or one death every 37 seconds [9]. Early detection of many of these diseases can help treat the condition and prevent the worsening of the disease. This requires a noninvasive technique that is capable of cardiac functional analysis and is accurate, reliable, and reproducible. Out of the cardiac functional assessment techniques, MRI is most suitable for this purpose. Various cardiac imaging modalities are discussed and compared with cardiac MRI in the following section. 1.4
CARDIAC IMAGING TECHNIQUES Various imaging and non‐imaging techniques are available to study murine cardiology. The non‐imaging methods including electrocardiography, systemic hemodyamics and histology are available for studying murine cardiology. Compared to the imaging modalities, these techniques have many disadvantages. Electrocardiography is a semi‐
quantitative method that assesses the electrical signal conduction in the heart [10, 11]. Many of the CVDs, such as hypertrophy and congestive heart failure, are associated with the changes in the cardiac blood flow or blood pressure. Systemic hemodynamics assesses cardiac disorders that are associated with the changes in the homodynamics. This is done by the measurement of blood pressure and blood flow at the left side of the heart. This is an invasive procedure and does not provide structural information. Histological techniques can be used to assess infarct size, viability of the cardiac tissue, 5
and regenerative cardiac growth [12, 13]. Histological techniques do not provide any functional information and are non‐repeatable. They also require a larger sample size since only one data point per animal can be acquired. Considering the limitations of the aforementioned techniques, noninvasive imaging methods are preferred for cardiac functional measurements. Last century has seen development of many cardiac imaging techniques. Some of them were based on already known techniques (angiography, CT) while some were innovative (MRI, ultrasound). Recent advances have enabled the use of these modalities for small‐animal imaging. To apply these techniques for imaging of small animals, many challenges need to be overcome. For instance, the overall size, physiologically relevant volume of the animal, and the required resolution need to be taken into account while choosing a modality for small‐animal imaging [14]. Echocardiography (ECHO) is ultrasonography of the heart. It is a widely used noninvasive method for cardiac functional analysis. It differentiates various structures of the heart based on their ability to reflect ultrasound waves which depends on different tissue density [15‐18]. Clinical echocardiography can provide information about a number of parameters such as cardiac wall motion, pumping efficiency, LV wall mass, damage to cardiac tissue, etc [17, 19‐21]. Using Doppler mode, it is possible to obtain velocity information from the ECHO data which helps in the diagnosis of blood‐flow abnormalities that are associated with regurgitation [15, 17]. It can also detect changes in the left‐ventricular chamber diameter and wall thickness that are often seen in hypertrophy and dilated cardiomyopathy [15, 20]. Echocardiography is noninvasive, has short examination times (up to 10 minutes), can be performed without anesthesia and has no known side‐effects [17, 19, 22]. The major limitation in two‐dimensional echocardiography of mouse is its relatively suboptimal resolution, requirement of a skilled operator and necessity of geometric assumptions for cardiac functional measurements [17, 20]. 6
Computed tomography (CT) is an X‐ray based noninvasive technique that can provide 3‐D view of the subject’s anatomy [23]. Although initially computed tomography was used for brain imaging, now it is used for various applications including cardiovascular imaging. There are many challenges in CT imaging of small animals such as requirement of high spatial and temporal resolution, longer scan times and consequently, possibility of more radiation exposure. In cardiac imaging of small animals, an additional degree of complexity is added due to cardiac and respiratory motion of the animals. Several strategies have been employed to overcome these challenges and image small animals [24]. To compensate for cardiac and respiratory motion, prospective as well as retrospective gating has been employed for heart imaging [25‐28]. Recently cine‐CT has been used for obtaining cardiac functional parameters and studying the remodeling in myocardial infarction model [26, 27, 29, 30]. The advantages of micro‐CT include fast scan times and 4‐D images of the heart. Major drawback of this technique is the exposure of the animal to ionizing radiation that limits the acquisition time of longitudinal scans. Another limitation of the method is inherent poor soft‐tissue contrast of the images which sometimes necessitates the use of contrast agents compounding the risk factor [16, 23, 24]. Nuclear cardiology is a noninvasive cardiac molecular imaging technique [16, 31‐
35]. It uses special radioactive molecules that can be imaged to assess the perfusion [16]. It is a class of imaging techniques that includes scintigraphy, positron emission tomography (PET) and single photon emission computed tomography (SPECT). Nuclear cardiology is classified in two groups, direct emission of photons (scintigraphy and SPECT) and indirect emission of photons. In direct emission of photons, photons are emitted one at a time during radioactive decay and are similarly detected by the gamma camera [34, 36, 37]. The other group of radionuclides emits positron which interacts with electrons to emit two gamma photons. These photons travel in opposite directions and are detected by two gamma cameras facing each other [35, 38, 39]. Both, SPECT and PET 7
have been used for LV function assessment [37, 40‐44], infarct size measurement [45] and monitoring LV dilation in mice [46]. The advantages of using nuclear imaging techniques are high sensitivity, ability of cell tracking, and monitoring metabolic activities. The major limitation is the possible risk of radioactive tissue damage. The successful application of nuclear imaging depends on the development, choice and availability of appropriate probe molecules. Figure 1.3 Cardiac imaging modalities X‐ray (A) computed tomography (B), nuclear scintigraphy (C) and MRI (D) can be used for cardiac imaging. MRI is the only technique that does not utilize potentially harmful high‐energy part of the electromagnetic spectrum. Courtesy Yale University School of Medicine [47]. Recently MRI has emerged as a promising modality for cardiac imaging of small animals[17, 48, 49]. Unlike CT and nuclear cardiology, MRI uses non‐ionizing part of the electromagnetic spectrum (Figure 1.3), thus eliminating the risk of radiation exposure and possible tissue damage. It is capable of tomographic acquisition high‐resolution images of mouse heart and does not require any geometrical assumption [49, 50]. It has been extensively used for assessment of LV function, estimation of LV mass and tracking cell therapy in small animal models [49‐59]. Cardiac MRI (CMRI) is noninvasive and 8
reproducible. It can acquire images with high spatial and temporal resolution and can detect small structural and functional changes in the heart. Therefore, cardiac MRI was used for studying mouse heart in this study. The comparison of cardiac MRI with other cardiac imaging techniques is provided in Table 1.1. Feature MRI CT ECHO PET/SPECT Radiowaves Protons X‐rays ‐‐ Ultrasonic waves ‐‐ Soft tissue contrast Risks Excellent Claustrophobia Good None Contrast agent Gadolinium chelates, iron oxide particles No Relatively expensive Variable 30‐60 min Not good Ionizing radiation Barium, iodine Microbubbles Gamma rays Radioactive molecules Not good Possible radiation injury ‐‐ No Cheaper Yes Cheapest Wide Few minutes Wide 10 ‐15 minutes Radiation used Probes Portability Cost Availability Acquisition time No Relatively expensive Variable 20‐30 minutes Table 1.1 Comparison of cardiac imaging modalities 1.5
LITERATURE REVIEW FOR CMRI IN MOUSE Some of the initial reports of human CMRI came out in the eighties using very low‐field imaging systems (0.35T) [60]. These studies were done with spin‐echo sequence using 7‐ mm slices and made assumptions regarding the geometry of the left ventricle to calculate functional parameters such as ejection fraction. Since then CMRI has made a lot of progress in terms of hardware, software and image processing to be able to image tiny and fast‐beating mouse hearts. Compared to the human heart, mouse heart is very small (~ 0.2 g) and beats almost 10 times as fast. Therefore, various strategies had to be developed to acquire high spatial and temporal resolution images of murine hearts. 9
CMRI has been extensively used to image mouse hearts, investigate various cardiac pathological disorders and had emerged as one of the most accurate methods to assess the functional parameters of a murine hearts [49, 57, 61‐65]. Various MR methods are employed for functional imaging of the human heart, namely, ECG‐gated cine‐imaging, use of steady‐state free precession and spoiled gradient echo sequences, MR tagging and velocity‐encoded contrast MRI, and delayed hyperenhancement using gadolinium chelates [66, 67]. Many of these methods have been adapted for use in murine models. Some of the reports published in the 90s [64, 68] were the beginning of extensive field of mouse CMRI. This section is aimed at providing the overview of current techniques and applications in cardiac MRI. MRI acquisition parameters Field strength To get adequate signal from small‐sized mouse heart, CMRI in mice has been done using ≥4.7 T non‐clinical scanners [48, 56, 69, 70]. Recently there have been some studies using higher fields such as 11.7 T [62, 71] and 17.6 T [72]. However, the availability of high‐field scanners is limited. Thus, researchers have developed methods for CMRI of mice and rats using 1.5 T and 3 T clinical scanners [73]. Another variable in terms of the scanner is the orientation of the magnet bore. Most of the studies have been done using horizontal bore magnets since there were some concerns regarding the physiological impact of the vertical bore on the animals. Studies by Wiesmaan et al and Schneider et al have established that high‐field vertical scanners have no adverse effect on the physiology and functional parameters of murine heart [74, 75] Gating strategies Due to the perpetual motion of the heart, gating strategies are indispensible to get an image with good signal‐to‐noise ratio. Depending on the scanner and the position of the animal, cardiac or a combination of cardiac and respiration gating is used. Cassidy et al 10
have compared various gating strategies for their performance at 11.7 T [76]. The gating can be prospective or retrospective [77, 78] depending upon the requirement and availability of retro‐gating software. Heijman et al have compared pro‐ and retro‐gating strategies and found that with appropriate settings, the image quality is comparable [79]. To avoid the electrical noise due to the rapid switching of gradients during data acquisition, a fiber‐optic based method was suggested by Brau et al [80]. Pulse sequences Most of the studies used for mouse heart imaging use T1‐weighted gradient echo sequences for acquisition of cardiac images. Two of the most frequently used sequences are steady‐state free precession gradient echo and spoiled gradient echo [16, 81]. There are no systematic studies comparing the various sequences used for CMRI of mouse. MR data acquisition techniques Cine MRI Cine MRI refers to acquisition of cine‐loop images of cardiac cycle and using that data for analysis of structural and functional parameters of mouse heart. These studies generally use bright‐blood multi‐phase spoiled gradient echo sequences [49, 82]. Black‐
blood imaging sequence has also been used for the measurement of LV mass and volume [83]. Many studies have assessed LV mass [57, 61, 64, 84, 85] as well as LV global function in mice using cine imaging [49, 62, 65, 82]. Strain and velocity imaging Strain and velocity imaging are tissue tracking methods that can be used to assess intramyocardial wall motion, velocity, strain, strain rate, and torsion [66, 67]. One of the first reports of tissue‐tracking methods in mice was by Henson et al in 2000 which compared the LV torsion in mouse and human [86]. Another report in 2002 assessed the regional wall motion in mice [87]. The major limitation of covering only 80% of the heart 11
cycle was subsequently overcome which paved way for increase in the use of this technique [50, 83]. These methods include myocardial tagging [88, 89], velocity‐encoded phase‐contrast imaging [90, 91], displacement encoding with stimulated echoes (DENSE) [92, 93], and harmonic phase (HARP) analysis [89]. The limitation of tissue‐
tracking studies is labor intensive and time consuming data analysis that is involved in getting the wall motion and strain data [66]. Myocardial perfusion and blood volume Blood perfusion in myocardium is critical for optimum functioning of the heart. Quantification of myocardial perfusion may also prove important in determination of infarct size and severity after myocardial infarction. Myocardial perfusion methods quantify the perfusion of myocardium by magnetic spin labeling of endogenous water protons [94]. Although most of the work in this area has been done using isolated hearts, few in vivo studies have been reported [67, 94, 95]. Delayed enhancement MRI Delayed enhancement technique is used to visualize infarct in mouse hearts after MI. The principle of this technique is delayed hyperenhancement of the infarcted myocardium compared to the viable tissue. These studies are generally done using intravenous or intraperitoneal injection of gadolinium chelates such as Gd‐DTPA. A double inversion recovery sequence is used to nullify the signal from blood which helps visualize the hyperintese area in the myocardium. This technique has been standardized in mice and has been extensively used for studying infarct size in mice [58, 67, 71, 96] Applications in cardiac pathologies and treatments CMRI is used for studying various cardiac pathologies [67]. Few of the highly studied cardiac pathologies include myocardial infarction [49, 62, 65, 71, 75, 92, 93, 96, 97], cardiomyopathy [58, 70, 98], and cardiac hypertrophy [57, 61, 64]. CMRI has also been 12
used to study mouse hearts in normal and stressed conditions [65, 99, 100]. Another application of CMRI is studying effect of cardiomyoplasty using iron oxide for cell labeling [56, 69, 101‐103]. 1.6
LIST OF ABBREVIATIONS Table 1.2 gives the list of abbreviations that will be frequently used in this dissertation. Abbreviation Full form CMRI Cardiac magnetic resonance imaging CO Cardiac output DICOM Digital Imaging and COmmunications in Medicine ECG Electrocardiogram ECHO Echocardiography EDV Left‐ventricular end‐diastolic volume EF Ejection fraction ESV Left‐ventricular end‐systolic volume FLASH Fast Low Angle Shot LV Left ventricle MI Myocardial infarction MRI Magnetic resonance imaging NEX Number of excitations or number of averages SEM Standard error of the mean SNR Signal‐to‐noise ratio SV Stroke volume TE Echo time TR Repetition time 13
Table 1.2 List of frequently used abbreviations 1.7
OVERVIEW OF THE DISSERTATION Chapter 2 discusses the development of CMRI protocol used in this study. It also provides the procedure of obtaining cardiac functional parameters from anatomical MR images of the heart in detail. Chapter 3 compares M‐mode echocardiography and MRI for cardiac functional assessment in mouse. Chapter 4 and chapter 5 discuss the application of CMRI for two different models of cardiac pathophysiology, myocardial infarction and cardiac hypertrophy, respectively. Both the chapters include a brief review of the pathophysiology, introduction to the animal model that is used for the study and current status of MRI in studying the particular cardiac disorder in mouse model. Chapter 6 presents the application of CMRI in monitoring iron oxide labeled stem cells that have been injected in myocardium. It discusses the approach of using stem cell therapy, brief introduction to small paramagnetic iron oxide (SPIO) contrast agents for MRI and current status of MRI in using SPIO for stem cell applications. Chapter 7 summarizes the results presented in the previous chapters. 14
CHAPTER 2 CARDIAC MAGNETIC RESONANCE IMAGING 2.
Cardiac magnetic resonance imaging This chapter focuses on the development and optimization of functional cardiac MR (CMR) imaging methods for mouse heart. Functional cardiac imaging refers to cine‐loop anatomical images of the heart that are used to derive cardiac functional information. It includes a detailed discussion of animal preparation and setup, cardiac gating strategies, choice of MR acquisition sequence, and the procedure to obtain a short‐axis image of mouse heart. Analysis of cine‐loop images of the heart to get functional parameters such as ejection fraction and cardiac output are described in the later sections, followed by a brief description of the statistical methods used for data analysis. 2.1
INTRODUCTION Perpetual motion of the heart poses a great challenge for any cardiac imaging methods. This difficulty is compounded in mouse due to the small size of its heart and the high heart rate. This necessitates adaptation of the existing methods and development of special strategies to obtain a good quality, artifact‐free MR image of a mouse heart. This requires optimization of various parameters such as choice of anesthesia, appropriate ECG‐gating strategies and development & optimization of appropriate data acquisition sequences. Further, obtaining functional parameters form the anatomical data requires the use of manual as well as software‐based image processing methods. Most clinical scanners, operating at lower strength such as 1.5 T, are equipped with pre‐existing 15
acquisition sequences that require very less operator input. Research scanners generally do not have pre‐optimized methods for specialized applications and have the need of optimization for a specific application and animal model. The aim of this study was to obtain cardiac images in mouse model at 11.7 T. This necessitated the adaptation of the pre‐existing imaging sequences for the higher field strength and smaller size of mouse heart. This chapter describes this development as well as optimization in detail. Figure 2.1 Steps in the development and optimization of CMR imaging protocol Various aspects of the MR protocol are seen in the above schematic. Development of a CMR imaging protocol involves optimization of each of these aspects to the needs of the scanner strength, intended application and animal model of the study. Adapted from [104]. 16
2.2
PROTOCOL FOR CARDIAC IMAGING OF MOUSE Cardiac imaging protocol refers to all the aspects involved in imaging a mouse heart using MRI, including animal preparation, gating strategies, parameter optimization, and data handling [104]. A schematic of this is shown in Figure 2.1. The following section discusses each of the above aspects in depth. 2.2.1 MR scanner used for mouse cardiac imaging Figure 2.2 MR scanner used for mouse cardiac imaging 11.7 T (500 MHz) MRI system and its operator console are seen in the figure. This is a vertical bore scanner capable of imaging small animals. In this study, cardiac imaging of mouse was done at 11.7 T (500 MHz) scanner (Bruker Biospin Inc, Billerica, MA). This system consists of a superconducting magnet with a vertical bore. It is equipped with two different gradient sizes and has maximum gradient strength of 1000 mT/m. For mouse cardiac imaging, a birdcage coil with 30 mm diameter was used for both transmitting and receiving the signal. Imaging at higher field strength offers higher sensitivity and spatial resolution but also has the limitations of increased susceptibility effects, sample noise and shorter transverse relaxation times [104]. This requires adaptation of pre‐existing sequences to get the same image quality as lower fields in comparable acquisition time. As 11.7 T is 17
one the highest strength fields available for imaging, extensive optimization of sequence parameters was needed to get good quality images of mouse heart. The details of sequence optimization are discussed in section 2.2.4. 2.2.2 Animal preparation and set up Choice Of anesthesia Animal preparation refers to anesthetizing the animals and preparing them to be placed in the MR scanner. This can greatly influence the quality of the results. The first crucial step is the choice of anesthesia. The anesthesia needs to be easy to administer, reproducible and having quick onset and recovery. It also needs to have low toxicity and small effect on respiratory and cardiac rate [105]. A detailed discussion of effects of injectable and inhaled anesthesia can be found in Roth et al. [105]. Isoflurane was chosen for this study due to the ease of control of the level and anesthesia level and minimal depression in cardiac function [106]. The animals were anaesthetized with 2.5% isoflurane mixed with 1 liter/min carbogen (95% O2 and 5% CO2) and maintained with 1‐
1.5% isoflurane. The quantity of isoflurane in the mixture was often adjusted based on the vital signs of the animal. Physiological monitoring and restraining For monitoring the physiological parameters (heart rate, respiration and body temperature) of the animal a small animal monitoring system (Model 1025, Small Animals Instruments, Stony Brook, NY) was used. ECG leads were secured to the right forepaw and left hindpaw of the animal (Figure 2.3A). A pneumatic pillow was attached to the abdomen of the animal for monitoring respiration (Figure 2.3B). The core body temperature of the animal was monitored with a rectal thermometer and maintained at 37±0.5°C using a warm air blower. Wires of the ECG electrodes were taped to the animal bed to avoid motion due to the air flow. Animal was placed in the animal bed in prone position and restrained using medical tapes. This restraining is especially important 18
since the imaging was done in a vertical bore. Failure to properly secure the animal to its bed led to slight change in animal’s position during imaging. This position few mm variation in the animal placement resulted in a relatively greater change in mouse heart position since the heart is about 1 cm in length which lead to reacquisition of complete dataset. After securing on the bed, the animal was placed in a 30‐mm birdcage coil and the coil was positioned in the magnet such that heart of the animal was at the isocenter of the magnet. Figure 2.3 Monitoring of heart rate and respiration of the mouse A. Figure shows ECG‐leads attached to the front and hind paws of the mouse. The leads are secured using surgical tape. B. Arrow indicates the pneumatic pillow used as respiration sensor for the mouse. 2.2.3 Gating strategies for triggered cardiac imaging Cardiac or ECG‐gating is indispensible for mouse imaging because of the high heart rate of mouse: 500 – 600 beats per minute or 8‐10 beats per second. Gating reduces motion artifacts and enables image acquisition at the same points in the cardiac cycle. This is 19
crucial for any kind of analysis that requires processing of end‐diastolic (ED) or end‐
systolic (ES). Without gating, it will be very difficult, if not impossible, to discern the phased in cardiac cycle. Therefore, to acquire high resolution data capable of rendering cardiac functional parameters, appropriate gating strategy is essential. Retrospective and prospective gating Different options are available for gating. The first choice is between retrospective and prospective gating. Retrospective gating involves collection of image data along with physiological data. This can be thought of as image data having time‐stamp of corresponding phase of the cardiac cycle. Once the dataset it acquired, the projection data are sorted according to their respective ‘time‐stamp’ and image is reconstructed. In prospective gating sequence, image data collection corresponds to the R‐wave of the cardiac cycle. With retrospective gating, it data can be acquired acquisition at all time points in the cardiac cycle and may enable imaging more than one mouse at the same time. The advantage of using prospective gating over retrospective is greater control over the motion of the heart. With the retrospective gating software available with the 11.7 T system, it was observed that although the images had higher SNR, considerably longer scan times were needed to obtain images having resolution similar to the prospective‐gating sequence. This would increase the acquisition time for one animal to almost 90 minutes which was not desirable. Therefore, prospective gating was used in this study. Cardiac gating setup For prospective gating, MR compatible small animal monitoring and gating system (Model 1025, Small Animal Instruments, Stoney Brook, NY) was used (Figure 2.4). It system was also capable of tracking the respiratory rate and temperature of the animal. This gating system consisted of data acquisition modules located near the animal and control/gating module connected to a computer near the operator console. The electrical 20
signal from the ECG‐leads secured to the animal was converted to optical form using a light emitting diode and then transmitted to the control unit on an optic fiber. Optical transmission of the signal from the magnet to the workstation removed the possibility of electrical interference from the surrounding. The optical signal was converted back to electrical form at the workstation. The electrical signal was displayed on the computer where the operator could control the signal by varying the gating parameters. Figure 2.4 Small animal monitoring and gating system used for cardiac imaging All the components of the small animal monitoring and gating system are shown. Respiration module (A), control module (B), acquisition module (C), battery (D), thermometer (E), respiratory sensor (F), ECG‐leads (G) and computer interface (H). Photo adapted from SAI website. Optimization of SNR of the ECG trace Obtaining a good ECG trace was dependent on many variables such as choice of electrodes, quality of the contact of the electrodes with the skin of the animal, electrical interference from the surrounding etc. Two types of the electrodes were evaluated for their performance: subdermal (Figure 2.5 B) and surface electrodes (Figure 2.5 C). 21
Surface electrodes were observed to be more susceptible to electrical noise and physical disturbances like respiratory motion resulting in a noisy trace with low amplitude of the R‐wave. Subdermal electrodes were observed to be more compatible with the current setup providing strong ECG trace with a clean and stable baseline (Figure 2.6C). To facilitate the conduction of electrical signal between the animal skin and the leads, electrolytic gel was applied to the needles. Interference due to imaging gradients was observed as a result of the initial electrical detection of the ECG signal. To reduce this noise the lead wires were coiled together and taped to the animal bed. To filter spurious electrical noise and interference due to gradients, the acquisition module of the gating system was enclosed in the copper case (Figure 2.5 A). Figure 2.5 Additional measures to improve signal‐to‐noise ratio of the ECG signal A. Acquisition module was enclosed in a copper cylinder to filter out spurious electrical noise. B. Subdermal ECG electrodes C. Surface ECG electrodes. Subdermal ECG electrodes were observed be less susceptible to electrical interference compared to the surface electrodes. When used with the electrolytic gel application, subdermal leads produced a stronger and more stable ECG signal. 22
Figure 2.6 Quality of ECG signal and optimization of detection parameters ECG trace with high signal‐to‐noise ratio (SNR) is essential for an artifact‐free and high resolution mouse heart image. The panel to the right is a magnified image of the ECG trace displayed on the computer. As seen in the figure, R‐wave is the wave with highest amplitude. P‐
wave is the line with second highest amplitude. Blocking‐period, shown as the red line, is the interval for which the system does not generate a triggering signal. R‐R interval is termed as period which decided the repetition time of the MR acquisition sequence. A. This panel shows noisy and low‐amplitude ECG trace that was generally observed in the animals with myocardial infarction. This trace gave rise to erroneous triggering due to difficulties in R‐wave detection. B. ECG signal with unstable baseline and strong respiration interference was seen when ECG leads were not secured properly and the lead wires were not taped to the animal bed. This usually lead to irregular R‐wave detection and consequently, to reduction in image quality. C. ECG trace with high amplitude and good SNR. High amplitude of R‐wave enabled regular triggering despite interference due to respiratory motion. Blocking period had to be adjusted close to the period to avoid triggering of the sequence off the P‐wave. 23
The ECG trace of animals with myocardial infarction was often observed to have more noise and weaker R‐wave (Figure 2.6 A). Noisy trace refers to an ECG spectrum where noise levels are of comparable magnitude to the R‐wave hampering the detection of the true R‐wave. This problem was solved to a great extent by application of the electrolytic gel. Despite the gel, ECG quality was observed to poor in some animals (Figure 2.6A, 2.6B). In such cases, R‐wave detection parameters were appropriately adjusted to enable R‐wave detection. In addition to that, the value of blocking interval (Figure 2.6) of the detection software was kept just very close to the R‐R period of the heart cycle. Surprisingly, it was observed that keeping the blocking interval close to R‐R period was also crucial in the case of a strong ECG‐trace (Figure 2.6 C). Strong ECG‐
trace usually had a prominent P‐wave which could be detected as trigger signal if value of the blocking period was not adjusted according to the R‐R period of the cycle. 2.2.4 Cardiac MR data acquisition – choice of pulse sequence A pulse sequence is a sequence of RF pulses applied to the sample during a MRI study. A simple pulse sequence consists of an excitation pulse, three orthogonal gradients and a refocusing pulse or gradient that forms the echo. The time of between excitation pulse and echo is termed as echo time (TE). Time required for one complete cycle of the pulse sequence events is known as repetition time (TR). Degrees of complexities in the form of additional events are added to the pulse sequence based on the application and capability of the gradients used. All these events involved in a pulse sequence are represented on a diagram called pulse timing diagram. Two examples of very common pulse sequences are spin echo and gradient echo. The following section will discuss the choice of pulse sequence for CMR and the advantages of using this sequence. Bright‐blood imaging sequence for cardiac imaging Bright‐blood sequence, as the name suggests, shows blood with brighter intensity compared to the myocardium and vessel wall. When used for cardiac imaging, bright‐
24
blood sequence can provide structural as well as functional information. The high contrast between blood and cardiac tissue of a bright‐blood cardiac image enables accurate and reproducible functional assessment of the heart. It can also be utilized in magnetic resonance angiography for visualization of blood vessels in the body. Bright‐blood sequences are typically variations of gradient‐recalled echo (GRE) pulse sequence. GRE sequence employs smaller, non‐90°, flip angles that enable shorter TR, effectively reducing the scan time. Due to higher component of residual transverse magnetization, GRE uses a refocusing gradient instead of the 180° pulse that is used in spin echo sequence. The advantages of GRE include faster scan times, capability of imaging flowing blood (MR angiography), and possibility of 3D imaging in reasonable time. The major limitations of GRE include magnetic susceptibility artifacts, reduced SNR due to smaller flip angles, and chemical shift effects at the interface of water and fat. With the use of appropriate parameters, GRE sequence can be optimized to acquire heart images with high blood‐tissue contrast with shorter acquisition time, making it suitable for cardiac functional imaging [107, 108]. The example of routinely used GRE sequences are fast low angle shot (FLASH), fast imaging with steady‐state precession (FISP) and steady‐state free precession (SSFP). The major difference in these sequences is the treatment of steady‐state transverse magnetization at the end of each cycle. Spoiling or maintaining the steady‐state determines whether T1, T2 or T2* effects will dominate. The choice of the type of GRE sequence depends on the application. FLASH ‐ sequence of choice In this study, FLASH sequence was chosen for cardiac imaging. The pulse sequence diagram is shown in Figure 2.7. FLASH has a variable spoiler gradient at the end of each echo. The spoiler gradient eliminates the steady‐state transverse magnetization and leaves the longitudinal magnetization undisturbed. Thus, with short flip angles it is possible to obtain T1‐weighted images with this sequence. The spoiler may increase magnetic susceptibility artifacts and dephasing of spins due to B0 inhomogeneities. 25
These disadvantages may be more apparent at high fields like 11.7 T. To suppress the susceptibility artifacts short‐TEs (~1.5 ms) were used during imaging. Also, manual shimming adjustments were often used to fine tune the shimming results obtained from automatic shimming procedure. In this sequence blood shows flow related signal enhancement [107]. Flow‐related signal enhancement refers to unsaturated blood appearing brighter in intensity compared to the surrounding saturated tissue. This is seen in the first (entry) slice that blood flows in. In a multi‐slice or multi‐frame sequence, flow‐related enhancement can be seen in every slice if the echo time is short and slices are acquired in sequential mode. Employing this strategy, we acquire multi‐frame bright‐blood images of heart that can be combined to form a cine loop. Figure 2.7 Timing diagram of FLASH sequence used for cardiac imaging Figure shows the timing diagram of the ECG‐triggered FLASH sequence for cardiac imaging of mouse heart. As seen in the pulse diagram, steady‐state magnetization in the transverse plane is spoiled after each echo is obtained. The spoiling combined with low flip angles enables faster data acquisition. 26
Acquiring cine‐loop images of the heart Cine‐loops of heart are nothing but a sequential series of heart images obtained at various time‐points in the cardiac cycle. This is illustrated in Figure 2.8. The temporal resolution of the functional study of the heart is dependent on the number of frames acquired per cardiac cycle. In the current study, 16 frames were obtained in each cardiac cycle. The heart rate of the mouse was between 400‐500 corresponding to R‐R period of 120‐150 ms. With 16 frames, the temporal resolution of this study was about 8‐10 ms. Cine‐loop images with this temporal resolution were found to be adequate for functional analysis of the heart. Care was taken to adjust the TR of the sequence to cover the complete heart cycle. Figure 2.8 Acquisition of cine‐loop images of the heart Cardiac images were obtained at different stages of the cardiac cycle and then coupled together to form cine‐loops. Data acquisition began 1 ms after the R‐wave detection, thus, making the first frame end‐diastolic frame. Total 16 frames were obtained per cardiac cycle. TR of the FLASH sequence was kept 10‐15 ms less than the cardiac cycle so that it covered the complete cardiac cycle but accounted for the variability in the R‐R period. For the above mentioned method of cine‐loop imaging of heart, two approaches for data acquisition can be used: segmented k‐space and unsegmented k‐space. Segmented k‐
space approach refers to filling more than one k‐space lines per frame in one R‐R 27
interval. Sequences using segmented k‐space approach have short TE (~ 1.5 ms) and short TR (8‐10 ms) to get high‐resolution images in short acquisition time. This approach is possible and practical for human heart imaging since the heart rate is low and R‐R period is relatively longer. It enables single breath‐hold acquisition of cardiac images ensuring good SNR and temporal resolution. However, the heart rate of mouse is almost 10 times that of human heart, yielding an R‐R period of 120‐160 ms while under anesthesia. Thus, the high temporal resolution (8 – 10 ms each for 16 frames) desired for functional analysis severely limits the application of segmented k‐space approach in mice. Figure 2.9 Unsegmented k‐space filling of an ECG‐triggered GRE sequence This example illustrates the filling of k‐space for 8 frames acquired in the sequence. One line each of the all frames is acquired with each heart beat. The total number of heart beats required to complete the image acquisition depends on the number phase‐encoding steps. Since the sequence used in this study has 192 phase‐encoding steps, 192 heart beats are required to completely acquire the required number of frames. (Adapted from [81]) Considering all the above, unsegmented k‐space approach was used in this study (Figure 2.9). Imaging was done with cardiac‐triggered 2‐D FLASH sequence that acquired one k‐space line per frame in one cardiac cycle. The scan time for this approach was longer than the segmented approach but it achieved the desired temporal 28
resolution. It also covered the cardiac cycle completely, ensuring the accuracy of the functional measurements done on the cine‐loop images. 2.2.5 Procedure for getting short‐axis images of mouse heart This study consisted of tracking the changes in heart function of mice up to 4 weeks or more. Functional analysis of the heart is done based on the measurement done on the short‐axis cine‐loop images. For the analysis to be accurate, few things have to be followed while acquiring the data. First, the short‐axis images have to be obtained along the true short‐axis of the heart. A slight variation in the angle of the slice can change the cross‐section of the heart chamber and lead to erroneous area calculations. Secondly, complete cardiac cycle has to be covered during imaging otherwise a missing a part of the cycle will lead to misinterpretation of the systole and diastole of the heart. Thirdly, it is important to ensure that the left ventricle is completely imaged else there will be errors in the calculation of left ventricular volumes. Therefore, a procedure was developed to eliminate variability in the acquired data irrespective of the used animal or the time point in study. First part of the procedure was to make certain that the animals were in relatively same position inside the magnet. This was accomplished by means of markers on the animal bed and the resonator. It was followed by manual tuning and matching of the RF frequency. After tuning, a combination of automatic and manual volume shimming was usually performed to get a proton line with full‐width‐at‐half‐maximum (FWHM) linewidth of less than 600 Hz. This was an important step in the procedure to ensure good SNR of the acquired images. Figure 2.10 shows the flowchart for getting the short‐axis images after the initial adjustments. After a tri‐pilot scan, a low‐resolution localizer scan was performed to locate the heart of the animal. This was followed by two mutually perpendicular scout scans to obtain the long‐axis of the heart. This was a crucial step in the imaging procedure since the geometry of the short‐axis images was adjusted based on the scout 29
scans. After the scouts, double‐oblique axial images, that were perpendicular to the line between the base of the aorta and the apex of the heart, were obtained based on these two scout scans. Figure 2.10 Protocol for obtaining short‐axis cardiac images of mouse Figure shows the flowchart for getting double‐oblique short‐axis images. After tri‐pilot, low‐
resolution localizer images were obtained to locate the heart. These images were used to get four‐
chamber and two‐chamber images of the heart. As discussed in the previous section, 16 frames with short TE (1.6 ms) were acquired in each cardiac cycle. These frames could be looped to generate a cine‐lopp that simulates the cardiac motion during the cycle. For heart rates of 400‐500 beats per 30
minute, the TR value of the imaging sequence was between 130‐160 ms and it was kept 5‐10 ms less than the R‐R interval to make sure that no part of heart cycle is missed during imaging. 6‐8 slices of thickness 1 mm were acquired ensuring that the left‐
ventricle was completely imaged. In‐plane resolution of 117 μm was obtained with a field of view of 3×3 cm and matrix size of 256×192 (zero‐filled to 256×256). With 6 averages, this gave cine‐loop cardiac images of desired spatial and temporal resolution. With the above parameters, for a TR of 150 ms, the acquisition time for one short‐axis cardiac cine‐slice was about 3 minutes (192 × 150 ms × 6). The total acquisition time, including initial adjustments (20 min) and acquisition of cine‐slices (25 min) was up to 45 minutes per animal. With the initial adjustments, the total scan time for one animal was upto 2.2.6 MR Image processing to get cardiac functional parameters To begin with, all the images were converted to DICOM (Digital Imaging and COmmunications in Medicine) format using Bruker ParaVision 4.0 software (Bruker Biospin, Billerica, MA) and later used for analysis of ejection fraction and LV wall analysis. A few commercially available software packages, tested for their applicability for analysis of mouse images, were found to be better suited for high SNR clinical images. For mouse images have low SNR coupled compared to the clinical images, especially with highly developed infarct in later stages (week 4) of the study. It was very difficult to get accurate estimates for chamber volumes using the automated functions available in these software packages. Therefore, all the data were manually analyzed using ImageJ (NIH, Bethesda, MD) software. Selection of the region of interest For any functional analysis, it is required to measure the area of the left ‐ventricular chamber at end‐diastolic (ED) and end‐systolic (ES) phases. There are two ways of 31
outlining the endocardial contours at ED and ES phases. The contours can be drawn with the area of papillary muscles or excluding the papillary muscles and trabeculae. This is especially important for the mid‐ventricular slices (Figure 2.11). Both these methods were compared to assess the accuracy of one approach over the other. No significant difference was found in the measurements done using either approach. Therefore, for the sake of simplicity and expediency of the analysis, all the measurements were performed with the area of papillary muscles included. Figure 2.11 Regions of interests (ROIs) with and without papillary muscles Figure shows endocardial contours drawn on the end‐diastolic phase image of mouse heart. A Area of papillary muscles is included A while B shows the contour without the muscle area. Analysis of stroke volume, ejection fraction and cardiac output For functional analysis using the cine‐loop images, ED and ES frames were determined by visual inspection. Since there was a 1‐ms trigger delay after R‐wave detection, ED frame was the first frame obtained. ES frame was the frame with smallest LV blood‐pool or lumen area. In the cases where it was difficult to judge the difference visually, ES was determined by comparison of area of the respective frames. Epicardial area was measured only in the ED frame. Epicardial area was measured in both ED and ES frames by manual planimetry of the lumen of the LV chamber (Figure 2.12). 32
Figure 2.12 Manual planimetry of mouse heart for functional analysis For obtaining cardiac functional parameters, endocardial and epicardial areas of the heart are obtained by tracing the borders of left ventricle. A. Total area of the left ventricle B. Area of the blood pool in the left ventricle at the end of diastole termed as end‐diastolic area (EDA) C. Area of the blood pool in the left ventricle at the end of systole termed as end‐systolic area (ESA). EDA and ESA both include the area of the papillary muscles. An average of three such measurements was taken for further calculations. Simpson’s rule algorithm for volume determination (summation of areas from all the slices multiplied by slice thickness, Appendix I) was used to get LV end‐diastolic volume (LVEDV or EDV) and LV end‐systolic volume (LVESV or ESV). It is represented as follows [109] LVEDV = ∑ Apex (end − diastolic area )× (Slice thickness ) Base
LVESV = ∑ Apex (end − systolic area)× (Slice thickness) Base
Left ventricular stroke volume (SV), ejection fraction (EF) and cardiac output (CO) are calculated from EDV and ESV as follows SV = LVEDV − LVESV ⎛ LVEDV − LVESV ⎞
EF = ⎜
⎟ × 100 LVEDV
⎠
⎝
33
CO = SV × Heart Rate Heart rate was noted for each slice and the average of heart rate values was used for calculation of cardiac output. Unless otherwise specified, all volumes are expressed in ml, ejection fraction in % and cardiac output is expressed in ml/min. Analysis of LV mass In MRI, LV mass is calculated based on the volume of LV tissue that is obtained from the short‐axis images. LV tissue volume was calculated as follows LV tissue volume = (ED Endocardia l area − ED Epicardial area ) × slice thickness
From LV tissue volume, LV mass was calculated as LV mass = LV tissue volume × specific gravity of myocardium Where specific gravity of the myocardium = 1.055 g/cm3 [49]. Analysis of LV wall thickness LV wall thickness was measured manually using ImageJ software. LV thickness was measured in 8 radial directions in the end‐diastolic frame. These measurements were averaged to give the mass of the left ventricle. 2.2.7 Statistical methods used for data analysis All the data in this study were assumed to have normal distribution and were analysed statistically for each experiment. Student’s t‐test was used for comparison of means of two groups. For comparison of two methods of cardiac functional measurement, Bland‐
Altman analysis was performed. Microsoft excel™ and GrandPad Prism™ 4 were used for statistical analysis. 34
2.3
RESULTS AND DISCUSSION Figure 2.13 Choice of gating for image acquisition For mouse heart imaging, gating is required to get an image with good resolution and SNR (A). Gating can be done using heart motion (ECG‐based) alone (B) or heart motion in combination with respiratory gating (C) to reduce the artifacts due to respiratory motion. Heart images obtained with a combination of cardiac and respiratory did not show any significant improvement in quality compared to the ones obtained with only cardiac gating. Therefore, all the images in this study were acquired with just cardiac gating. Using ECG‐triggered FLASH sequence, cardiac images with good signal‐to‐noise ratio and high resolution were obtained. As it can be seen in Figure 2.13, images were obtained with no gating, only cardiac gating, and a combination of cardiac and respiratory gating. Additional respiratory gating did not improve the quality of the cardiac images. Therefore, in the interest of time, only cardiac imaging was used for data acquisition. It was possible to clearly see the left and right ventricles, interventricular septum and papillary muscles in the images (Figure 2.9). ECG‐gating and manual shimming ensured images with minimal motion and susceptibility artifacts, respectively. 35
Figure 2.14 Bright‐blood images of mouse heart at end‐diastolic phase Good contrast between blood (bright) and tissue (dark) is obtained in the images acquired with ECG‐triggered FLASH sequence. Short‐axis image shows left and right ventricles, septum and papiallry mucles. In the four‐chamber image, right and left atria and aorta is also clearly seen. LA: Left atrium; LV: left ventricle; RA: Right atrium; RV: Right ventricle; septum: interventricular septum. 2.4
SUMMARY This chapter forms the basis of cardiac MR method that is used for imaging of mouse heart in this study. It gave an account of the development and optimization of this functional CMR method. It included a detailed description of the animal preparation, cardiac gating strategies, choice of MR acquisition sequence, and the procedure to obtain a short‐axis image of mouse heart in detail. Also, a section on analysis of short‐axis cine images to get the functional parameters for heart was included. Using this imaging method, it was possible to get mouse heart images with desired spatial and temporal resolution and minimum artifacts. 36
CHAPTER 3 COMPARISON OF MRI AND ECHOCARDIOGRAPHY TO ASSESS CARDIAC FUNCTION IN A MOUSE MODEL OF MYOCARDIAL INFARCTION 3.
Comparison of MRI and echocardiography to assess cardiac function in a mouse model of myocardial infarction This chapter focuses on the comparison of cardiac MRI (CMRI) and M‐mode echocardiography (ECHO) for assessment of cardiac function. Control and infarcted mice were subjected to CMRI and ECHO measurement before and after myocardial infarction surgery. The functional parameters obtained from the two modalities were compared. 3.1
INTRODUCTION 3.1.1 Motivation for the study Echocardiography is the most widely used technique for cardiac functional assessment in the clinic and experimental animal models. CMRI is emerging as an alternative noninvasive technique for structural and functional assessment of cardiac function. Many comparative studies between ECHO and CMRI in human have been reported and have found the two techniques to compare favorably [71, 110, 111]. There have been very few such studies done in rodents, especially in mice [59, 112, 113]. Therefore, this study was undertaken to compare cardiac MRI and M‐mode echocardiography for 37
cardiac functional assessment in mice. 3.1.2 Cardiac functional analysis by echocardiography Echocardiography works on the principle of reflection of ultrasonic waves from different tissue in the body. Based on different properties of the reflected wave, echocardiography is classified into three types [15, 16]. A‐mode echocardiography displays the amplitude of the reflected wave. B‐mode echocardiography uses the brightness of the wave for visualization. The third mode, mono‐axial (1‐D) or M‐mode, displays the changes in the structure of the heart as a function of time. ECHO can also be used for 2‐D or 3‐D imaging of hearts. Since the late 1990s, ECHO has been extensively used for the assessment of cardiac functional parameters such as ejection fraction, stroke volume, fractional shortening and for estimation of the left ventricular mass in mice [12, 20, 22, 113, 114] A typical echocardiograph is shown in Figure 3.1. M‐mode ECHO measures the diameters of the heart chambers at systole and diastole. A geometrical assumption, such as the ellipsoid method or area‐length method, is used to compute the cardiac chamber volumes from the chamber diameters [111, 115]. M‐mode ECHO is capable of fast data acquisition and has good temporal resolution. It can be used for analysis of abnormalities in cardiac structure as well as function. The limitations of M‐mode ECHO include the need for a skilled operator [51, 110] and geometric assumptions about cardiac chambers that may lead to erroneous estimation of functional parameters [111, 115]. This is, particularly, a greater concern in the case of an asymmetrical left ventricle, typical of myocardial infarction [51, 110, 113]. 38
Figure 3.1 Typical M‐mode echocardiogram of a mouse heart M‐mode echocardiogram has time on the horizontal axis while the vertical axis shows the distance from the transducer to the object that reflected the sound wave. A 2‐D ECHO image is used as the guide for positioning of the transducer. The anterior (A) and posterior (P) walls of the LV chamber are visualized and are used to measure the inner diameter of the chamber at diastole (LVIDd) and systole (LVIDs). A geometrical assumption is used to compute the end‐diastolic and end‐systolic volumes from the diameter measurements and the data is displayed in the left corner of the echocardiograph. 3.1.3 Study design This study aimed at comparing M‐mode echocardiography and CMRI for cardiac functional assessment in mice. Control and MI groups (n=4, each) were measured before the myocardial infarction (MI) surgery (week 0) and 4 weeks after the surgery. ECHO and CMRI were performed on the animals within a period of 5 hours in between the measurements. Also, control group was measured at one additional time point and the data were used for reproducibility analysis. From ECHO and CMRI measurements, end‐
diastolic volume (EDV), end‐systolic volume (ESV), stroke volume (SV) and ejection fraction (EF) of the left‐ventricle were obtained. Since ECHO measures a single point of the heart at mid‐papillary level, EF values were obtained from the mid‐papillary MRI slice for comparison with ECHO. This EF was called mid‐papillary EF while the EF 39
obtained from complete LV volume was termed as global EF. Figure 3.2 illustrates the slice orientations for each modality. The measurements were compared using correlation‐regression as well as Bland‐Altman method. Figure 3.2 Slice positioning in CMRI and ECHO CMRI acquires 6‐8 slices along the LV, covering the whole volume of the heart. The summation of the area of all the slices is used to estimate EDV and ESV. ECHO used a mid‐papillary slice measurement for estimation of EDV and ESV. For comparison, EF was also computed from a mid‐papillary MRI slice (arrow) which was estimated to be in the similar position as that of the ECHO slice. 3.2
MATERIALS AND METHODS 3.2.1 Animals MRI was performed on C57BL mice. Four animals, from the control and MI group each, weighing 30‐35 g and ages between 8‐10 weeks, were imaged. All animal protocols used were approved by the Ohio State Institutional Laboratory Animal Care and Use Committee (ILACUC). 40
3.2.2 Induction of myocardial infarction C57BL male mice (weighing 25–30 g) were anesthetized with a mixture of ketamine (55 mg/kg) and xylazine (15 mg/kg) that was injected intraperitoneally. The intubation tube was made of a 20‐gauge intravenous catheter attached to a connector. A modified Y‐
shaped connector was used to attach the mouse to the ventilator. The ventilator was set at a rate of 120 breaths/min with a tidal volume of 250 μl (Harvard Apparatus, Holliston, MA). The body temperature of the mice was maintained at 37 ± 1°C with the help of an isothermal heating pad (Braintree Scientific, Braintree, MA). All the procedures were performed with the approval of the Institutional Animal Care and Use Committee of the Ohio State University and conformed to the Guide for the Care and Use of Laboratory Animals (National Institutes of Health Publication No. 86‐23, Revised 1996). MI in the mice was created by permanently occluding the left coronary artery (LCA) using the following procedure. An oblique 8‐mm incision was made 2 mm away from the left sternal border towards the left armpit. The chest cavity was opened with scissors by a small incision (5 mm in length) at the level of the third or fourth intercostal space 2–3 mm from the left sternal border. The LCA was visualized as a pulsating bright red spike running through the midst of the heart wall from underneath the left atrium toward the apex. The LCA was ligated 1–2 mm below the tip of the left auricle using a tapered needle. An 8‐0 polypropylene ligature was passed underneath the LCA, and a double knot was made to occlude the LCA. Occlusion was confirmed by the sudden change in color (pale) of the anterior wall of the left ventricle. The chest cavity was closed by bringing together the third and fourth ribs with one 6‐0 polypropylene silk suture. The layers of muscle and skin were closed with a 5‐0 polypropylene suture. After LCA ligation, an ST elevation on ECG and a color change in the LV myocardium were observed in all mice. 41
3.2.3 Cardiac MR imaging CMRI was performed using 11.7 T MRI scanner (Bruker Biospin, Billerica, MA). The animals were anaesthetized with 2.5% isoflurane, mixed with 1 liter/min carbogen (95% O2 and 5% CO2) and maintained with 1‐1.5% isoflurane. For monitoring the physiological parameters (heart rate, respiration and body temperature) of the animal, a small animal imaging system (Model 1025, Small Animals Instruments, Stony Brook, NY) was used. ECG leads were secured to the right forepaw and left hindpaw of the animal. A pneumatic pillow was attached to the abdomen of the animal for monitoring respiration. The core body temperature of the animal was monitored with a rectal thermometer and maintained at 37±0.5°C using a warm air blower. The animal was placed in a 30‐mm birdcage coil and the coil was positioned in the magnet such that the heart of the animal was at the isocenter of the magnet. Long‐axis scout images were used as reference to get double‐oblique short‐axis slices (parameters: TR/TE: 4.6/1.4 ms; matrix: 256×192 zero‐filled to 256×256; FOV: 3×3 cm; slice thickness: 1 mm; Number of frames: 12; NEX: 4). 6‐8 axial slices were acquired to completely cover the left ventricle of the animal being studied. ECG‐gated cine images of the heart were acquired using a white‐blood FLASH sequence (parameters: TR/TE: 130/1.4 ms; matrix: 256×192, zero‐
filled to 256×256; FOV: 3×3 cm; slice thickness: 1 mm; Number of frames: 16; NEX: 6). Cardiac MR imaging (CMRI) was performed on control and MI mice for up to 4 weeks (week 0, 2, and 4). 3.2.4 Image processing MRI images were converted to DICOM format using Bruker ParaVision 4.0 software and their quantitative analysis was done using ImageJ software (NIH, Bethesda, MD). Endocardial and epicardial contours were manually traced at end‐systole and end‐
diastole for each slice to get left ventricular (LV) end systolic (ESA) and LV end diastolic area (EDA). The respective areas were multiplied was slice thickness to get end‐systolic volume (ESV) end‐diastolic volume (EDV). ESV and EDV were used to get stroke 42
volume (SV) and ejection fraction (EF) [49, 71]. Cardiac output was computed as the product of EF and heart rate measured at the time of image acquisition. Refer to section 2.2.6, for a detailed description of cardiac functional parameter calculations. 3.2.5 Echocardiography measurements M‐mode echocardiography (ECHO) was performed with a GE Vivid7 echocardiography system and intraoperative epicardial probe (Model i13L; frequency 14 MHz). Animals were anaesthetized using 1.5% isoflurane mixed with 1 liter/min carbogen (95% O2 and 5% CO2). LV tracings were obtained with a 2‐D short‐axis view as guide. The measurements were performed on both the groups for week 0 (pre‐surgery), 2 and 4. End‐diastolic and end‐systolic diameters of the left ventricle were measured using American Society Echocardiography leading‐edge method [116]. LV width and EF were calculated from the measured parameters. 3.2.6 Statistical analysis Data were presented as mean ± standard error of mean (SEM). A two‐tailed Student’s t‐
test was used to determine the significance. A p‐value of less than 0.05 was termed to be significant. Bland‐Altman analysis was used for comparing CMRI and ECHO data. The difference in the methods was represented as bias ± SD. 3.3
RESULTS The goal of this study was to compare the cardiac functional measurements made using cardiac MRI and M‐mode echocardiography. Cardiac measurements were performed on control and MI groups, before the surgery and in week 2 and 4, after the surgery. MR images obtained at week 4 post‐MI demonstrated considerable remodeling in the MI heart. Increased chamber diameters, leading to increased EDV and ESV, were observed in the MR images of the MI mice (Figure 3.3). The difference between EDV and ESV of the MI heart was observed to be small, indicating the decrease in contractility compared 43
to controls. Similar observations could be made from the echocardiograms obtained at the end of week 4 (Figure 3.4). ECHO clearly showed increase in the chamber diameters and hypokinesis of the anterior wall of the left‐ventricle of the MI heart. Control hearts, on the other hand, demonstrated stronger wall motion. Figure 3.3 CMR images of mouse hearts at 4 weeks post‐MI The MRI images of MI mice show severe ventricular remodeling in the MI heart 4‐weeks post‐MI. Compared to control heart, MI heart was larger in size. Also, the end‐diastolic and end‐systolic volumes were greater considerably increased in the MI heart. The values of EF obtained from the MRI and ECHO data supported the above observations (Figure 3.5). MRI EF values were obtained from total LV (global EF) volume as well as the mid‐papillary slice of LV (mid‐papillary EF). All 3 approaches showed decline in the EF of MI groups 4 weeks after the surgery whereas the EF of control group remained stable over the study period. Mid‐papillary MRI showed significant differences when compared with ECHO for both the time points (week 0 and 4) of the control group measurement. Global EF was observed to be significantly different from ECHO for week 0 measurement of control. No significant differences were observed in EF values for the MI group. 44
Figure 3.4 M‐mode echocardiograms of Control and MI mice at 4 weeks post‐MI Increased EDV and ESV of the MI heart could be observed in the ECHO. Also, ECHO showed almost no movement of the anterior LV wall (top end of the arrows), compared to the control heart. The chamber diameter measurements were used to obtain SV and EF. Figure 3.5 Comparison of EF of control and MI groups EF obtained from global MRI, mid‐papillary slice MRI and ECHO was compared from control and MI group. EF obtained from all 3 methods for MI group showed a significant decline at week 4, whereas control EF remained relatively stable. The values are expressed at mean ± SEM. For control group, mid‐papillary MRI and ECHO showed significant difference (∗ p < 0.05, n=4) for both, week 0 and week 4. Global MRI and ECHO were observed to significantly different (# p < 0.05, n=4) for week 0 measurement of control MI. No significant differences were observed in the EF measurements of MI group. 45
MI Control CMRI ECHO CMRI ECHO EDV (mL) 0.067 ± 0.003 0.153 ± 0.027 0.098 ± 0.016 0.328 ± 0.083 ESV (mL) 0.021 ± 0.002 0.040 ± 0.007 0.058 ± 0.013 0.220 ± 0.051 SV (mL) 0.046 ± 0.002 0.113 ± 0.020 0.040 ± 0.003 0.108 ± 0.031 EF (%) 68.47 ± 2.52 74.16 ± 0.72 39.89 ± 3.05 31.97 ± 1.15 Table 3.1 Cardiac functional parameters obtained from CMRI and ECHO The cardiac functional parameters were obtained in week 4 after MI. The values are represented as mean ± SEM (n=4). EDV: Left ventricular end‐diastolic volume; ESV: Left ventricular end‐
systolic volume; SV: stroke volume; EF: Ejection fraction. Figure 3.6 Correlations between EDV and ESV measurements of ECHO and MRI EDV and ESV measurements, obtained 4 weeks post‐MI, are shown. For both the parameters, ECHO is observed to overestimate the volumes compared to MRI. More variation is observed in MI group compared to the control group for both EDV and ESV. Table 3.1 summarizes the cardiac functional parameters obtained from CMRI (global) and ECHO in the 4th week after MI. As it can be observed from the table, ECHO overestimated the cardiac functional parameters in both the groups. The difference was 46
more noticeable in the estimates of EDV, ESV and SV. Figure 3.6 shows the correlation plots for EDV and ESV measurements by ECHO and MRI for control and MI groups, in the 4th week after MI. The line of best fit did not pass through origin for EDV or ESV. Average ECHO volumes were observed to be approximately 3‐fold of MRI measurements. Volume measurements were not done for the mid‐papillary MRI slice. EF values obtained from MRI (global and mid‐papillary) were compared with the ones from ECHO using Bland‐Altman analysis. This analysis generated a plot of average EF (on the x‐axis) and the difference between the two measurements (on the y‐
axis). Figure 3.7 shows the comparison between mid‐papillary EF and ECHO EF. The difference, in the methods, obtained from Bland‐Altman analysis was ‐4.11±10.66% (Figure 3.7A). From the correlation plot, it can be seen that most of the data points do not lie on the line with slope 1(r2 = 0.77) (Figure 3.7B). Similarly, Figure 3.8 shows the comparison between global MRI and ECHO for EF measurement. Bland‐Altman analysis showed a difference of 1.08±9.89% (Figure 3.7A). Very few data points were on the line of agreement (r2 = 0.85). Figure 3.7 Comparison of mid‐papillary MRI with ECHO A. Bland‐Altman plot for the comparison of EF. The difference between EF measurements using mid‐papillary slice and ECHO was found to be ‐4.11 ± 10.66 %. The degree of error was not found to be proportional to EF. B. Correlation plot for comparison of EF. 47
Figure 3.8 Comparison of global MRI with ECHO A. Bland‐Altman plot for the comparison of EF. The difference between EF measurements using mid‐papillary slice and ECHO was found to be 1.08 ±9.89%. The degree of error was not found to be proportional to EF. B. Correlation plot for comparison of EF. Figure 3.9 Reproducibility of EF measurements by ECHO and MRI Three measurements were performed on control mice with ECHO and MRI at three different time points. Two of the three measurements (each) were analyzed for reproducibility using Bland‐Altman analysis. ECHO (difference 1.29 ± 3.36) had less variation compared to MRI (‐3.84 ± 4.08). Different EF measurements performed on control mice are shown in the correlation plot. All the measurement were observed to lie in the upper half of the plot, indicating overestimation of EF by ECHO. 48
ECHO and global MRI were also compared for their reproducibility in EF measurements (Figure 3.7). Three measurements were done on the mouse in the control group at different time points and the EF values were obtained using MRI and ECHO. Bland‐Altman analysis showed less variation in ECHO (difference 1.29± 3.36) compared to MRI (‐3.84±4.08). Correlation plot showed that the EF values were overestimated by ECHO and that there was no significant variability between the two methods. 3.4
DISCUSSION The primary focus of this study was to compare MRI and M‐mode echocardiography for their measurement of cardiac functional parameters. M‐mode echocardiography and CMRI are both widely used for cardiac functional measurements. Comparative studies between MRI and ECHO have been done in humans [117‐119] and in rats [59, 112]. This is the first comparative study that compares the two methods in mice. Cardiac functional measurements performed on control and MI mice showed that there was general agreement in the measurement of EF by both the modalities. Although, the cardiac functional parameters from ECHO and CMRI showed similar trends, the absolute values were different. ECHO values were overestimated compared to CMRI. Also, ECHO showed less variability in EF measurement than MRI. Measurements were made for control and MI group mice before the surgery and 4 weeks after the surgery. ECHO and MRI measurements were performed on the mice on the same day. In the case of some animals, the interval between ECHO and CMRI was up to 5 hours. This time difference might be one of the reasons of the discrepancy in the absolute values of the cardiac functional parameters obtained from ECHO and CMRI. To get accurate measurements of the chamber volumes, it is required that the measurement of chamber diameter be along the short‐axis of the heart. In CMRI, several short‐axis slices are acquired to get LV volume (Figure 3.2). In the case of M‐mode 49
ECHO, a single point, generally at the level of the papillary muscle, was measured to get end‐systolic and end‐diastolic diameters. A geometric assumption was required to get EDV and ESV from these values [111, 115]. Also, the positioning of the ECHO transducer was critical to get proper short‐axis measurement. It was observed by Stuckey et al. that a variation of 1.5 mm in the transducer position lead to almost 20% variation in the EF of MI rat heart [112]. Due to smaller heart size and greater heart rate, the positioning of the transducer was even more critical in mice. Although a 2‐D image was used to orient the transducer along the short‐axis of heart, perfect short‐axis acquisition was not always possible. The geometric assumptions for volume measurement, coupled with the uncertainty in the positioning of the transducer could explain the unusually high EDV and ESV values obtained from ECHO measurements. EF, being a normalized parameter, did not reflect the error in the volume measurements. Therefore, EF was used to compare the methods and to analyse the reproducibility of measurement. For a relevant comparison with ECHO, EF was calculated using only the mid‐
papillary slice of the left ventricle (termed as mid‐papillary EF) in MRI (Arrow in Figure 3.2). Since the data was collected from a position similar to that of ECHO, the EF measurements from mid‐papillary slice were expected to be similar to that of ECHO. Surprisingly, significant differences were observed in the EF measurements using these two methods. This could be explained by the difference in the positions and angular orientations of the mid‐papillary MRI slice compared to ECHO measurement slice. Bland‐Altman analysis of both the methods for EF measurements showed that there was general agreement between the EF values obtained from CMRI and ECHO. Since the EF values were observed to lie in the 95% agreement interval, the values obtained from ECHO and MRI could be used interchangeably. However, higher difference was observed in the comparison of mid‐papillary EF with ECHO (‐4.11) than that of global EF (1.08). This indicated that global MRI was a better method of EF estimation than mid‐papillary MRI. The reproducibility analysis showed less variation 50
in ECHO measurements compared to CMRI. Overall, both the modalities were observed to be equally reproducible. This study is the first step in a rigorous comparative analysis of CMRI and ECHO that can be performed in mice with a few more changes to the study design. One limitation of this study was the small number of animals in each group (n=4). For more accurate statistical analysis, a study with more number of animals that will reduce intra‐
method variability will be needed. Also, after 4 weeks of MI, a bipolar distribution was observed in the EF values of the mice. A broader distribution will help achieve more accurate correlation analysis. Therefore, it might be beneficial to measure the cardiac parameters at an earlier stage of remodeling, where different EF values are more probable. Due to logistical concerns, the ECHO and CMRI measurements were performed with an inertval of up to 5 hours between them. Any inter‐method variability will be reduced if the measurements are performed in succession. 3.5
SUMMARY AND CONCLUSION The goal of this study was to compare CMRI and M‐mode echocardiography for cardiac functional assessment. The study was performed on control and MI mice. Cardiac functional parameters such as end‐diastolic, end‐systolic, stroke volume, and ejection fraction of the left ventricle were computed before the surgery and 4 weeks after the surgery. Echocardiography was observed to overestimate volume measurements compared to CMRI. Overall, there was agreement in the measurements of ejection fraction using CMRI and echocardiography. 51
CHAPTER 4 STRUCTURAL AND FUNCTIONAL EVALUATION OF MOUSE HEART AFTER MYOCARDIAL INFARCTION 4.
Structural and functional evaluation of mouse heart after myocardial infarction The primary focus of this chapter is to study the structural and functional changes in a mouse heart after myocardial infarction (MI). The mice were imaged for up to 4 weeks after myocardial infarction to monitor the structural changes in the MI heart. Cardiac functional parameters were obtained from the acquired data and compared with controls to evaluate the changes in cardiac function after MI. 4.1
INTRODUCTION 4.1.1 Myocardial infarction Heart attack or myocardial infarction (MI) leads to rapid development of myocardial necrosis as a result of disruption of blood flow to a part of the heart muscle. The loss of blood flow leads to an imbalance in the supply and demand of oxygen and other nutrients, causing injury or death of the myocardial tissue. The most frequent causes for the disruption of the blood flow are the rupture of the coronary plaque, arterial spasms and thrombus formation. Myocardial infarction initiates ventricular remodeling that includes changes in heart structure, geometry and function. Though initially an adaptive response, chronic remodeling ultimately leads to heart failure. 52
Anterior descending branch of the left coronary artery (LCA) is prone to atherosclerotic plaque formation. LCA is the primary artery supplying blood to the anterior part of the left ventricle (LV) in mice, Thus, occlusion of LCA causes an infarct in the LV muscle, eventually leading to impaired function of the heart (Figure 4.1). Thus, LCA ligation of mouse heart provides an appropriate model for understanding various aspects of myocardial infarction in human heart, and for studying possible surgical and drug interventions. Figure 4.1 Myocardial infarction as a result of occlusion in the left coronary artery Left coronary artery (LCA) and its branches supply blood to the anterior part of the left ventricle of the heart. Occlusion of LCA, due to the deposition of atherosclerotic plaque, thrombosis, or vascular spasm, causes disruption of the blood supply to the anterior part of the heart. This leads to injury and death of the cardiac tissue (myocardial infarction). The damaged tissue is shown in purple. 4.1.2 Left coronary artery ligation model for MI Surgical models for myocardial infarction were developed in an effort to closely replicate the progression of the condition in humans [120]. These models include occlusion of the coronary artery using ring or U‐shaped occluders, use of ameloid 53
constrictors, balloon inflation, and coronary artery ligation [121]. Out of these, due to the size of the heart and the coronary arteries, artery ligation is preferred in laboratory rodents. Coronary artery ligation procedure in mice was first mentioned in 1978 but was seldom used until the advent of transgenic mouse technologies [122]. Since then, coronary artery ligation is one of most popular methods for myocardial infarction. The coronary artery ligation procedure in mice similar to the ones used in rat but involves the use of a microscope due to the relatively small size of mouse LCA. The surgical procedure involves left throroctomy of the anesthetized animal to get to the heart, temporary deflation of the lung, identification and ligation of the LCA using a fine suture (Figure 4.2). After ligation, the lung is reinflated and the chest cavity is closed [122]. LCA ligation in mice being a complicated procedure, the mortality associated with it is 37‐50% [121]. Figure 4.2 Ligation of the left coronary artery in mice. 1. The ventilated mouse is kept on a heated pad. 2. Left side of the chest is opened to have access to the left ventricle. 3 & 4. One main branch of the left coronary artery is ligated with a suture. Arrow points to the pale ischemic area. Adapted from Klocke et al. [121] 54
4.1.3 Study design The study consisted of two groups, control (n=6) and MI (n=5). Both the groups were imaged before surgery (week 0) and up to 4 weeks after the surgery, to monitor the ventricular remodeling after MI. The short‐axis cardiac images were analyzed to obtain cardiac functional parameters such as end‐diastolic volume (EDV), end‐systolic volume (ESV), stroke volume (SV), ejection fraction (EF) and cardiac output (CO). EPR oximetry was used for measurement of myocardial oxygenation. Histological analysis was performed to confirm the fibrosis in the left‐ventricular wall due to MI. Statistical analysis was performed to evaluate the changes in MI heart as a function of time. 4.2
MATERIALS AND METHODS 4.2.1 Animals MRI was performed on C57BL mice. Two groups, control (n=6) and MI (n=5), weighing 25‐30 g and ages between 8‐10 weeks, were imaged. All animal protocols used were approved by the Ohio State Institutional Laboratory Animal Care and Use Committee (ILACUC). 4.2.2 Induction of myocardial infarction Myocardial infarction was induced by the ligation of left coronary artery in mice. For detailed procedure, refer to section 3.2.2. 4.2.3 Oxygen measurement (pO2) using EPR oximetry A single injection of OxySpin, an oxygen‐sensitive EPR probe, was performed in the mid‐ventricular region of the mice (n=5) immediately after LCA ligation. Measurements of myocardial oxygenation were performed noninvasively using an L‐band in vivo EPR spectrometer (L‐band, Magnettech) equipped with automatic coupling and tuning controls for measurements in beating hearts. Mice, under anesthesia (2% isoflurane), 55
were placed in a right‐lateral position with their chest close to the loop of the surface coil resonator. EPR spectra were acquired as single 30‐s scans. The instrument settings were as follows: incident microwave power, 4 mW; modulation amplitude, 180 mG, modulation frequency, 100 kHz; and receiver time constant, 0.2 s. The peak‐to‐peak width of the EPR spectrum was used to calculate PO2 using a standard calibration curve [123]. The measurements of myocardial infarction were performed before LCA ligation, on the day of ligation (Day 1) and for weeks after the ligation (Day 28). 4.2.4 Cardiac MR imaging Cardiac‐gated cine images of the heart were acquired using a white‐blood FLASH sequence (parameters: TR/TE: 130/1.4 ms; matrix: 256×192, zero‐filled to 256×256; FOV: 3×3 cm; slice thickness: 1 mm; Number of frames: 16; NEX: 6). Cardiac MR imaging (CMRI) was performed on control and MI mice for up to 4 weeks (week 0, 2, and 4). For detailed MR imaging procedure, refer to section 3.2.3. 4.2.5 Image processing For detailed description of cardiac functional parameter calculations, please refer to section 2.2.6 and section 3.2.4. 4.2.6 Visualization of fibrosis At the end of experimental period, animals were euthanized with an overdose of pentobarbital sodium. The hearts were removed immediately and attached to a Langendorff apparatus. Triphenyltetrazolium chloride (1.5%) solution was injected down the side arm of the aortic cannula and infused into the coronary circulation. Once the hearts were stained dark red, they were removed, weighed, and frozen. The following day, the hearts were defrosted and sliced into 1‐mm sections parallel to the atrioventricular groove and then fixed in 10% buffered formalin for overnight. For measurement of fibrosis area, after measurement of the infarct size, heart tissues were fixed with 10% buffered formalin and embedded in paraffin. Paraffin sections were 56
stained with Massonʹs trichrome stain. Images of the LV area of each slide were prepared by Nikon Model C‐PS (objective X20) with Spot Insight camera (Diagnostic). 4.2.7 Statistical analysis The data were represented as mean ± standard error of mean (SEM). A two‐tailed Student’s t‐test was performed to determine the significance. A p‐value of less than 0.01 was termed to be significant. 4.3
RESULTS This study focused on monitoring and evaluating the changes in mouse heart after permanent occlusion of left coronary artery. Cardiac‐gated images were acquired for control and MI groups at different time points after MI to observe the structural and functional changes in MI heart. EPR oximetry and histology were used for the confirmation of myocardial infarction in the mouse hearts. Shortage of oxygen is one on the main causes of myocardial infarction. To confirm the success of LAD ligation and progress of infarction, myocardial oxygenation was determined using EPR spectroscopy. Oxygen‐sensitive probe LiNc‐BuO was injected in the mid‐myocardium and the oxygen partial pressure (pO2) was measured using a surface resonator (Figure 4.3A). There was significant drop in the pO2 of the myocardium after LCA ligation and pO2 remained in the ischemic range even after 4 weeks after the surgery (Figure 4.3B), indicating myocardial ischemia. Cardiac MR images of the control and MI hearts were acquired before MI (week 0) and for up to 4 weeks post‐MI. The images were acquired such that LV area was completely covered. Figure 4.4 shows the representative images of the heart from base (slice # 1) to the apex (slice # 6). The structural changes in the MI heart were more apparent in the slices acquired towards the apex of the heart indicating the location of the infarct. 57
Figure 4.3 Measurement of myocardial oxygenation using EPR oximetry A. LiNc‐BuO, oxygen sensitive probe, was implanted in the mouse heart. Mice were anesthetized using 2% isoflurane and the surface coil resonator was placed on the chest of the mice. Myocardial oxygenation was measured before LCA ligation, on the day of the surgery and 4 weeks after the surgery. B. LCA ligation decreased myocardial infarction to about 15% of the pre‐
ligation value (p < 0.01, compared to the pre‐ligation value). The time course of structural changes in the MI heart is shown in Figure 4.5. Within a week of MI, the mouse heart underwent prominent structural changes including increase in the LV chamber size and change in the shape of the heart. Between week 1 and week 3, no further changes were noticed. Week 4 showed further increase in the size of the heart. Therefore, datasets from week 2 and week 4 were chosen for functional analysis. The details of the ventricular remodeling were clearly seen in the images that were acquired 4 weeks after MI (Figure 4.6). Compared to control, the short‐axis images of MI heart showed the increase in the end‐diastolic (red arrow) and end‐systolic (blue arrow) volume. Changes in the heart shape and size were evident in the long‐axis images of MI heart. The thickness of the LV wall of MI hearts was observed to have considerably decreased compared to that of controls (green arrow). 58
Figure 4.4 Bright‐blood images of control and MI heart as a function of distance from the apex Mid‐papillary short‐axis images of a mouse heart acquired before and after MI. Slices from base (slice # 1) to the apex (slice # 6) were examined for structural changes in the MI heart. The structural and functional changes in the MI heart were especially visible in the slices towards the apex (slice # 4, 5, 6) indicating the possible infarct region. Figure 4.5 Structural changes in mouse heart after myocardial infarction Short‐axis and long‐axis images of mouse heart, before (week 0) and after (weeks 1‐4) MI, are shown. There were noticeable changes in the chamber size and heart shape of the mouse after MI. The majority of the changes happened in the first week after the surgery. From week 1 to week 3, the changes in the volume were not significant. Week 4 images showed further perceptible changes in the shape of the heart. 59
Figure 4.6 LV remodeling in the mouse heart 4 weeks after the surgery Images of control and MI heart at end‐diastolic and end‐systolic stages of the cardiac cycle clearly indicate the structural and functional remodeling in MI heart. Compared to controls, MI hearts showed increase in the end‐diastolic diameter (red arrow) indicating dilation of the heart. The increase in the end‐systolic diameter (blue arrow) was indicative of loss in contractility of MI heart. The change in the overall shape of MI heart was visible in the long‐axis images at 4‐week post‐MI. These images also showed the thinning of left‐ventricular wall (green arrow) of the MI heart compared to control. In addition to this, long‐axis cine‐loop images (Figure 4.7) showed the decrease in the contraction of the lower part of the MI heart due to the presence of scar tissue. Short‐axis cine‐images were acquired for the functional analysis of control and MI mice. Each cine image consisted of 16 frames that were obtained 8‐10 ms apart. End‐diastolic (frame # 1) and end‐systolic (frame # 11) frames were identified and used for functional measurements. The decrease in the contractility of the heart was evident in these short‐
axis cine images. 60
Figure 4.7 Cine images of the cardiac cycle of control and MI heart Mid‐papillary short‐axis cine‐loop images of control and mouse hearts are seen. Each cine images consisted of 16 images acquired with a temporal resolution of 8‐10 ms. End‐diastolic (#1) and end‐systolic (#11) frames were identified for further functional measurements. Global parameters of cardiac volume (EDV, ESV) and function (SV, CO, EF) were computed from the cine‐loop images. These factors signaled to the worsening of cardiac function after MI. Significant increase was observed in EDV and ESV after MI (Figure 4.8A, 4.8B). EDV showed an increase of 87% in week 2 (p value: 0.003) and 93% in week 4 (p value: 0.004) over the respective controls. The change in ESV was more striking with 240% increase in week 2 (p value: 0.001) and 264% increase in week 4 (p value: 0.001). SV did not show any significant difference compared to the control (Figure 4.5C). CO showed a decrease of ~ 35% post‐MI but it was not significant compared to the respective control (Figure 4.8D). Ejection fraction is an important indicator of cardiac function (Figure 4.6E). Compared to the respective control as well as to MI baseline value (MI week 0), EF showed significant decrease in week 2 (p‐value < 0.0001) and week 4 (p value < 0.0001). 61
Figure 4.8 Progressive loss of cardiac function after MI Global parameters of LV volume and function were measure in controls (n=6) and MI (n=4), before (week 0) and after (week 2, 4) MI. MI hearts showed a significant increase in EDV and ESV (p < 0.01) and a significant decrease in EF (p < 0.001). SV and CO did not show any significant change. 62
Figure 4.9 Variability in the extent of remodeling in MI Due to the inherent difficulty and complications of the LAD ligation microsurgery, variability was observed in the extent of remodeling in MI hearts. Some MI hearts were observed to have twice as much chamber volumes as the others. This led to high variability in the estimation of cardiac parameters. Figure 4.10 Histological assessment of myocardial infarction in mouse heart Mason trichrome staining of the MI hearts showed the thinning of the LV wall and the presence of fibrosis (arrows) indicating the presence of scar tissue at the end of week 2 and week 4. Control heart showed a uniform LV wall with no fibrosis. 63
Compared to control, more variability was seen in the MI group (Figure 4.9). One subgroup had EF in the range of 8‐14 % while the other had 25‐40% EF. Echocardiography measurements (data not shown) showed the ejection fraction to be ~ 30% for the both subgroups. At end of 4 weeks, mouse hearts were isolated and stained using Masson trichrome stain. The histological analysis showed fibrosis at the location of scar tissue which is an indication of necrosis as a result of myocardial infarction (Figure 4.10). The thinning of the LV walls was also clearly seen, supporting the data from CMR images. 4.4
DISCUSSION Most of the mouse CMRI studies have been done at ≤ 7T. This is one of the very few CMRI studies of LCA ligation mouse model of MI at 11.7T scanner [62, 75]. The high spatial and temporal resolution of obtained using the 11.7 T scanner facilitates the structural and functional analysis of MI in mouse heart. For this study, the imaging protocols were optimized for optimal image quality. The spatial resolution was 117 µm × 156 µm was adequate for detecting the structural changes in the mouse heart. Consistent with earlier reports, LV chamber dilation and LV wall thinning were observed in the anterior apical region of the heart [49]. Shortage of oxygen in the infarct and peri‐infarct region is one of the characteristics of myocardial infarction. To measure the oxygen levels and confirm the occurrence of myocardial infarction, noninvasive measurement of myocardial infarction was performed using EPR oximetry. The pO2 values obtained in this study were in agreement with the previous reports [12, 13, 124] and confirmed the low oxygen levels that are associated with myocardial infarction. Myocardial infarction triggers remodeling of the ventricles that includes thinning of LV wall, cardiac hypertrophy, and impaired cardiac function. In this study, CMRI was able to detect the changes in the structure as well as the function of the heart. The anatomical changes associated with ventricular remodeling were easily detected with 64
the bright‐blood images of the heart. These changes were confirmed with the histological images showing the LV wall thinning and fibrosis. Cine images provided data necessary for the quantification of the structural changes such as chamber dilation. As seen in the results, most of the structural changes took place in the initial 7 day period after the LAD ligation surgery. This was also reflected in the functional parameters. MI hearts showed up to 93% increase in EDV and up to 240% increase in ESV over controls. Consequently, there was more than 50% decrease in the EF of MI hearts compared to the controls. This is consistent with the previous reports of functional parameters in mouse heart [62, 71, 75, 125]. A few observed differences may be due to the difference in the MI model since most of the reported mouse MI studies have used ischemia/reperfusion model, not permanent coronary ligation. The functional parameters of MI hearts showed a higher variability compared to the controls. This was due to the variability in the remodeling of MI heart (Figure 4.9). Two different ‘extents’ of remodeling were seen in the mouse model revealed by the different sizes of the MI heart, creating two subgroups within the MI group. One subgroup had EF in the range of 8‐14 % while the other had 25‐40% EF. This led to a wide range of values for cardiac functional parameters. Thus, the variability in functional parameters of the MI heart stems from the inherent differences in the cardiac function of the MI heart, not from errors in the data acquisition or analysis. It should also be noted that echocardiography measurements were unable to discern the difference between the ejection fractions of the two subgroups within MI animal group. This indicates that MRI is more suitable to assess the severity of the infarction which is indicated by the extent of remodeling. This study could have been benefitted with infarct‐size measurements delayed hyperenhancement studies using gadolinium‐based contrast agents. Unfortunately, thinning of the scar tissue and permanent occlusion of the artery led to complications in 65
terms of time required for delayed hyperenhancement. This prevented acquisition of good quality images and measurement of infarct size. 4.5
SUMMARY AD CONCLUSION This study applied cardiac MRI for structural and functional evaluation of mouse hearts after myocardial infarction. The animals were imaged for up to 4 weeks after MI. The compensatory changes in the structure of the heart (remodeling) were clearly observed in the MR images. CMRI was also able to analyze the functional parameters to evaluate the changes in the heart function post‐MI. The MI hearts showed increase in their shape and size, especially the size of the LV chamber. The cardiac function showed significant decrease after MI. Thus, CMRI was able to monitor the structural and functional changes in mouse heart post‐MI. 66
CHAPTER 5 NONINVASIVE ASSESSMENT OF CARDIAC FUNCTION IN A TRANSGENIC MODEL OF CARDIAC HYPERTROPHY 5.
Noninvasive assessment of cardiac function in a transgenic model of cardiac hypertrophy This chapter focuses on the study of the structural and functional changes in a transgenic mouse model of cardiac hypertrophy. The control and transgenic mice were imaged using CMRI to obtain structural and functional information. The cardiac functional parameters were also measured using M‐mode echocardiography (ECHO). The data were used to assess the transgenic model of cardiac hypertrophy. 5.1
INTRODUCTION 5.1.1 Cardiac hypertrophy Cardiac hypertrophy refers to the enlargement of the heart and thickening of the heart muscle, more commonly that of the left ventricle. It is an adaptive response to the increased stress on cardiac walls due to volume or pressure overload. Heart, a terminally differentiated organ, adapts to stress by means of increasing its muscle mass using hypertrophic remodeling. There are three major types of cardiac hypertrophy: developmental, physiological, and pathological [126]. Developmental hypertrophy is normal growth of heart since birth till adulthood [127]. Healthy or physiological cardiac hypertrophy occurs as a response to chronic exercise or pregnancy. This is believed to be 67
an adaptive response that leads to better functioning of heart. Chronic volume overload (aortic valve regurgitation, dilated cardiomyopathy) or chronic pressure overload (hypertension, aortic valve stenosis) leads to pathological cardiac hypertrophy. Unlike physiological/healthy hypertrophy, this condition is associated with thickening of the heart muscle leading to increase in heart mass, accumulation of scar tissue and decrease in heart efficiency. Although cardiac hypertrophy is an adaptive response in its initial stages, chronic condition is an independent risk factor for heart disease. It can lead to diastolic dysfunction and further, to congestive heart failure. There are two types of hypertrophies as far as the growth of cardiac muscle is concerned. The first type, called concentric hypertrophy, occurs when there is pressure overload. In this type, new sarcomeres are added in‐parallel with the existing ones, Thus, there is no great increase in the radius of the ventricle but the thickness of the ventricular wall increases. This enables the ventricle to generate higher forces while maintaining normal wall stress. The ventricle becomes stiff leading to diastolic dysfunction. In the second type, referred to as eccentric hypertrophy, new sarcomeres are added in‐series with the existing ones leading to increase in diameter as well as wall thickness of the ventricle. This occurs when there is both pressure and volume overload leading to chamber dilation and further, to systolic dysfunction. Salient features of the cardiac hypertrophies are summarized in Table 5.1 Traditionally, echocardiography has been the method of choice for the diagnosis of cardiac hypertrophy, specifically left‐ventricular hypertrophy, as it can measure the thickness of the heart wall. In many cases, abnormalities in the electrical signal of hypertrophied heart can be observed using electrocardiogram (ECG), enabling the use of ECG as a screening tool. Chest X‐ray/ CT can also be used to detect cardiac enlargement. Cardiac MRI (CMRI) is emerging as a promising tool for diagnosing cardiac hypertrophy. 68
Physiological hypertrophy Pathological hypertrophy Causes Chronic exercise, pregnancy Chronic hypertension, aortic valve stenosis Muscle thickening Fibrosis Cardiac dysfunction Yes No No Increase in length greater than increases in width Yes Yes Yes Increase in width greater than increase in length Change in myocytes Table 5.1: Types of cardiac hypertrophy and their effects on the cardiac tissue 5.1.2 CMRI as a tool for the detection and diagnosis of cardiac hypertrophy CMRI, due to its high temporal and spatial resolution, is one of the most suitable methods for imaging a hypertrophied heart. It is capable of distinguishing between physiological and pathological hypertrophies [128]. It can also diagnose and differentiate cardiomyopathies in a single study [129]. MRI is advantageous for studying cardiac hypertrophy, because, in addition to precise measurements of wall thickness and mass of the heart, it is also capable of early detection of changes in the myocardial tissue. With the use of delayed enhancement contrast, MRI can be used to differentiate between ischemic and non‐ischemic cardiomyopathy [58, 130]. MRI is also useful in the determination of cardiac functional parameters that may be associated with pathological hypertrophy and can be used to characterize the changes in the structure and function of the heart due to hypertension [131]. Previously many studies have used MRI to image hypertrophied hearts in humans, large animal models like dogs, and murine models [57, 61, 64]. In summary, CMRI can be effectively used for detection and diagnosis of cardiac hypertrophy. 69
5.1.3 Transgenic animal model of cardiac hypertrophy Figure 5.1 Transgenic mouse model of cardiac hypertrophy used in this study Rac D gene from corn (Zea Mays) was overexpressed in mouse heart. This gene increases superoxide production that leads to chronic hypertension and further, to pathological cardiac hypertrophy due to pressure overload. Heart images adapted from [127]. Previously, animal models that were employed to study left‐ventricular hypertrophy due to pressure‐overload have used surgical manipulations to constrict the aorta, ablation of the kidney by incision or drug administration, or intravascular volume expansion with high‐salt diet [57]. These models are not good representatives of human hypertensive disorder as hypertension in human is a chronic disorder and usually has some genetic component associated with it [57]. Therefore, transgenic models are better 70
suited for this purpose since they mimic the conditions in human body closely. Further, it is possible to study the effect of a specific gene of interest in the development of hypertension. This study used a transgenic model of hypertension‐induced cardiac hypertrophy developed by Hassanain and co‐workers[132, 133]. Previously many studies have correlated hypertension and cardiac disorders with oxidative stress [134‐
136]. Redox‐sensitive signaling pathways are implicated in cardiac hypertrophy and remodeling [136, 137]. One of the sources of increased production of reactive oxygen species (ROS) in tissue is NADPH oxidase. NADPH oxidase is a multicomponent enzyme complex that consists of the membrane‐bound cytochrome b558, the cytosolic regulatory subunits p47phox and p67phox, and small GTP‐binding protein Rac1, a member of the Rho family of GTPases. Extensive in vitro and in vivo research has indicated the involvement of a protein network that has Rho and Ras family members as key components in cardiac hypertrophy[137‐139]. Rac1 is a part of the signaling pathways that are involved in cardiomyocyte hypertrophy [137, 138]. Rac1 maintains the homeostasis of blood pressure by regulating the production of ROS in phagocytic and non‐phagocytic cells. Thus, mutation/overexpression of Rac1 can lead to hypertension and eventually to cardiac disorders. We have used a Rac1‐overexpressed transgenic mouse model to induce cardiac hypertrophy. This is a unique transgenic model that has overexpression of Rac1 gene from corn in mouse heart (Figure 5.1). Plant Rho/Rac sequences are very similar to the mammalian Rac genes. The ability of the Rac gene to induce superoxude production has been conserved during evolution [132]. This gene was shown to induce maximum superoxide production when expressed in mammalian cells [132]. This transgenic model was accomplished by over‐expresseion of cDNA of a constitutively active mutant of Zea mays (ZmRac D) gene in the heart of transgenic FVB/N mice using a mouse alpha‐
myosin heavy‐chain promoter [132]. Gene expression was confirmed by RT‐PCR and immunohistochemistry of heart tissue of transgenic mice and control littermates. 71
Recently, it has been shown that the transgenic expression of Rac gene causes hypertension in murine models [133]. As chronic hypertension is one of the major causes of cardiac hypertrophy, these mice are eventually expected to develop cardiac hypertrophy. The aim of this study was to establish the capability of MRI for the assessment of cardiac functional parameters in this transgenic cardiac hypertrophy model. 5.1.4 Study design The study consisted of two groups, control (n=5) and transgenic (n=5). Both the groups were imaged using CMRI. The short‐axis cardiac images were analyzed to obtain cardiac functional parameters such as end‐diastolic volume (EDV), end‐systolic volume (ESV), stroke volume (SV), ejection fraction (EF), cardiac output (CO) and left‐ventricular width (LV width). Cardiac functional parameters were also measured using M‐mode echocardiography. Statistical analysis was performed to compare data from the control and transgenic groups. 5.2
MATERIALS AND METHODS 5.2.1 Animals CMRI was performed on a transgenic mouse model developed in Dr. Hassanain’s laboratory that over‐expressed the cDNA of a constitutively active mutant of Zea maize Rac D (Rac D) gene in the heart of transgenic FVB/N mice [140] using a mouse alpha‐
myosin heavy chain promoter [132]. Five animals each of control group and transgenic group (TG), weighing 30‐35 g and ages between 190 – 475 days, were imaged. All animal protocols used were approved by the Ohio State Institutional Laboratory Animal Care and Use Committee (ILACUC). 72
5.2.2 Cardiac MR imaging Cardiac‐gated cine images of the heart were acquired using a white‐blood FLASH sequence (parameters: TR/TE: 130/1.4 ms; matrix: 256×192, zero‐filled to 256×256; FOV: 3×3 cm; slice thickness: 1 mm; Number of frames: 16; NEX: 6). Cardiac MR imaging (CMRI) was performed on control and MI mice for up to 4 weeks (week 0, 2, and 4). For detailed MR imaging procedure, refer to section 3.2.3. 5.2.3 Image processing MRI images were converted to DICOM format using Bruker ParaVision 4.0 software (Billerica, MA) and their quantitative analysis was done using ImageJ software (NIH, Bethesda, MD). To get LV width, LV wall thickness was obtained by taking an average eight radial measurements performed in the end‐diastolic slice. LV mass was calculated by product of the specific gravity of cardiac muscle (1.055 g/cm3) and total volume of the LV tissue. For detailed description of For detailed description of cardiac functional parameter calculations, please refer to section 2.2.6 and section 3.2.4. 5.2.4 Echocardiography studies 2‐Dimensional echocardiography (ECHO) was performed with a GE Vivid7 echocardiography system and intraoperative epicardial probe (Model i13L; frequency 14 MHz). Animal was anaesthetized using 1.5% isoflurane mixed with 1 liter/min carbogen (95% O2 and 5% CO2). LV tracings were obtained with 2‐D short‐axis view as guide. End‐diastolic and end‐systolic diameters of the left ventricle were measured using American Society Echocardiography leading‐edge method [116]. LV width and EF were calculated from the parameters measured. 73
5.2.5 Statistical analysis The data were represented as mean ± standard error of mean (SEM). Two‐tailed Student’s t‐test was performed to determine the significance. A p value of less than 0.05 was termed to be significant. 5.3
RESULTS Figure 5.2 MR images of transgenic (mutant RacD overexpressed) and control mouse hearts. Representative long‐axis images of mouse hearts (two from each group) obtained using ECG‐
triggered FLASH sequence are shown. (parameters: TR/TE: 4.6/1.4 ms; matrix: 256×192, zero‐
filled to 256×256; FOV: 3×3 cm; slice thickness: 1 mm; number of frames: 12; NEX: 4). All four chambers of the heart and aorta can be seen in the images. The transgenic hearts are larger and more round‐shaped when compared to control hearts. 74
A series of 2‐D MR images was acquired along both long and short axes as a function of the cardiac cycle. Long‐axis MR images of the control and transgenic mouse hearts (Figure 5.2) demonstrated the difference in the shape and the size of the heart in both groups. In general, TG hearts were larger in size when compared to the control hearts. Also, the control hearts were more oblong with an elliptical left ventricle whereas TG hearts were observed to be rounder with a blunt apical section. Figure 5.3 Short‐axis images of hearts of control and transgenic mouse hearts at end‐diastolic and end‐systolic states Representative images of mouse heart, from the base of the aorta (slice # 1) to the apex of the heart (slice # 6) are shown. Left ventricle, right ventricle, and interventricular septum are clearly seen in the images. The images were acquired using a FLASH‐cine sequence (parameters: TR/TE: 130/1.4 ms; matrix: 256×192; FOV: 3×3 cm; slice thickness: 1 mm; NEX: 6). The images were cropped for representation purposes. The transgenic hearts are bigger than control hearts. 75
The short‐axis images of the heart (Figure 5.3) were acquired with the long‐axis images as reference and showed the left ventricle, right ventricle, and the interventricular septum. The 1‐mm thick slices were obtained from the base of the aorta (slice # 1) to the apex of the heart (slice # 6) to ensure that the whole left ventricle was adequately covered. These images, coupled with the mid‐ventricular slices shown in Figure 5.4, further illustrated the differences in the size of control and TG hearts. Figure 5.4 Mid‐ventricular short‐axis images of control and transgenic mouse hearts. Zoomed‐in view of representative mid‐ventricular short‐axis images of mouse hearts (two each from control and transgenic group) obtained using ECG‐triggered FLASH sequence (parameters: TR/TE: 130/1.4 ms; matrix: 256×192; FOV: 3×3 cm; slice thickness: 1 mm, NEX: 6). Transgenic hearts show a significantly larger end‐systolic volume compared to the control hearts. No significant difference in the end‐diastolic volume was observed. 76
The MR images were processed to quantify the cardiac functional parameters using ImageJ software. The results showed that the TG and control hearts had similar end‐diastolic volumes (0.050 ± 0.01 ml for control vs 0.060 ± 0.01 mL for TG), which was clearly seen in Table 5.2 and Figure 5.5A. Significant difference was observed in end‐
systolic volume with TG hearts (0.013 ± 0.002 mL) having larger end‐systolic volume compared with control hearts (0.024 ± 0.007 mL). The CMRI results (Table 5.2 and Figure 5.5A) indicated that compared to the control hearts, TG hearts had a significantly lower EF (63.256 ± 3.26 % vs 77.86 ± 2.86 %; p < 0.05) and greater LV wall thickness (1.043 ± 0.026 vs 0.945 ± 0.030 mm; p< 0.05). No significant difference was observed in stroke volume, cardiac output, and LV mass of TG mice compared to that of the control (Figure 5.5A). ECHO CMRI Control TG Control TG EDV (mL) 0.050 ± 0.004 0.060 ± 0.003 0.102 ± 0.013 0.079 ± 0.008 ESV (mL) 0.013 ± 0.001 0.024 ± 0.003 0.020 ± 0.003 0.021 ± 0.002 SV (mL) 0.037 ± 0.003 0.036 ± 0.003 0.086 ± 0.012 0.054 ± 0.008 EF (%) 77.86 ± 2.86 63.25 ± 3.26 83.14 ± 2.86 70.57 ± 2.05 CO(mL/min) 0.013 ± 0.001 0.012 ± 0.001 0.043 ± 0.006 0.027 ± 0.004 LV width (mm) 0.945 ± 0.030 1.043 ± 0.026 0.091 ± .007 0.081 ± 0.01 LV mass (mg) 80.59 ± 1.828 93.95 ± 7.077 ‐ ‐ ‐ ‐ Table 5.2: Cardiac functional parameters obtained from CMRI and echocardiography The values are represented as mean ± SEM (n=5). EDV: Left ventricular end‐diastolic volume; ESV: Left ventricular end‐systolic volume; SV: stroke volume; EF: Ejection fraction; CO: cardiac output; LV width: left ventricular width; LV mass: left ventricular mass. ECHO measurements showed no significant difference in EDV, ESV, cardiac output and LV wall thickness (Table 5.2 and Figure 5.5 B). Stroke volume in TG hearts 77
was observed to be significantly lower compared to control (0.086 ± 0.012 ml vs 0.054 ± 0.008 mL; p < 0.05). Significant difference was also observed in the EF values of the TG and control hearts (70.570 ± 2.05 % vs 83.14 ± 2.86 %; p < 0.05). The EF and EDV values obtained from ECHO were on the higher side compared to the ones from MRI images. Figure 5.5 Cardiac functional parameters computed from MRI and ECHO of mouse hearts. Functional parameters of mouse hearts assessed from MR images and ECHO data. Values are represented as mean ± SEM (n = 5 hearts/group). A. Cardiac parameters computed from MRI data. Compared to control hearts, in transgenic hearts left ventricular end‐systolic volume (ESV), 78
and left ventricular width (LV Width) were significantly greater whereas ejection fraction was significantly lower, indicating cardiac hypertrophy in the transgenic animals. B. Cardiac functional parameters computed from ECHO data. Compared to the control hearts, significant decrease in EF and stroke volume of the transgenic hearts was observed. (∗ p < 0.05, compared with the respective control) Figure 5.6 Comparison of LV width and EF obtained from MRI and echocardiography of mouse hearts. No significant difference in LV width was observed in the MRI and ECHO values for either of the groups. The EF values obtained from MRI are higher than those from ECHO. No significant difference was observed in MRI and ECHO values for either group (∗ p < 0.05, compared to the respective MRI values) Although the trends in the values of cardiac functional parameters obtained from ECHO and MRI images were similar, differences were observed in the actual values. Therefore, the results for EF and LV width obtained from the two techniques were compared using t‐test (Figure 5.6). No significant differences were observed in the EF values obtained using these two methods for both the groups, further supporting the results. In this study, control and TG groups were not age‐matched. Therefore, data analysis was conducted to investigate the effect of age on the cardiac parameters, 79
especially EF and LV mass. The analysis showed no clear correlation between the age of the animal and its EF (Figure 5.7A). The TG mice seemed to have an increase in LV mass with age while control mice did not exhibit that trend (Figure 5.7B). In general, for similar age, control mice were observed to have more body weight compared to the TG ones and TG mice showed an increase in weight with age (Figure 5.7C). No clear correlation between LV mass index (LV mass / body weight) of the animal and age was seen, although there was more variation in the mass index in TG mice compared to controls (Figure 5.7D) Figure 5.7 Effect of age on ejection fraction, LV mass and body weight The plots demonstrate the effect of age on the ejection fraction and LV mass computed from the MRI data of the control and transgenic mice. A. No clear correlation between the age of the animal and its ejection fraction was observed in control as well as transgenic mice. B. In transgenic mice, LV mass seems to increase with age whereas in controls there is no direct correlation. C. Control mice had more body weight than the transgenic ones. In transgenic mice, body weight was observed to increase with age. D. No clear correlation between body LV mass index and age of the mice was seen. Transgenic mice had more variation in body weight and LV mass compared with the control. 80
5.4
DISCUSSION In the present study, high quality, relatively artifact‐free, ECG‐gated cine‐MRI images of the heart were obtained. Good contrast between blood pool and non‐moving tissue was obtained using bright‐blood imaging sequence. This enabled precise measurements of inner and outer diameter of LV, leading to accurate cardiac functional parameter measurements. The long‐axis and short‐axis MR images clearly showed the enlargement of heart as well as change in the heart shape of TG mice indicating the development of hypertrophy. The change in the shape of the hearts, from oblong to rounder, as observed in TG hearts, is more associated with dilated cardiomyopathy. Therefore, possible transition of the TG heart from hypertrophy to dilated cardiomyopathy needs to be further investigated. Hypertrophied hearts, especially the ones with maladaptive or pathological hypertrophy, are prone to lower EF and higher LV wall thickness. In the present study, a significant decrease in the EF of TG hearts was observed. The EF observed for control mice is in agreement with what has been previously observed Schneider et al in am ouse model using a 500 MHz vertical bore system [75]. This, coupled with ECHO measurements obtained in the present study, is in agreement with the decrease in the heart function for TG mice. Another major observation from MRI was increase in the ESV of TG mice (up to 80%) compared to control which was not reflected in the ECHO measurements. Despite similar EDV values for both animal groups (about 20% increase in EDV for TG hearts), no significant difference was observed in the SV values obtained from either of the techniques. There was a significant increase in the ESV values of TG hearts obtained from MRI but no significant difference in the SV values was seen. In contrast, ECHO data indicated significant difference between SV but did not show any increase in ESV of TG mice compared to control. SV is simple subtraction of EDV and ESV. Since 81
both the techniques did not find significant difference in the EDV values for TG and control hearts, it was surprising to find that ESV and SV follow different trends. Increasing the sample size may be required for better understanding of the actual trend. As expected in the hypertrophy model, LV width of TG hearts measured from MR images was found to be larger than that of TG hearts. No significant difference in LV mass obtained from MRI was seen. Cardiac functional parameters obtained from MRI as well as ECHO measurements indicate that the TG mouse hearts were working with less efficiency compared to the control ones. Since the actual values obtained from MRI and ECHO for EF and LV width were different, a comparison of measurements from both the techniques was performed. No significant difference was observed in cardiac functional parameters derived from the two methods for either of the groups. MRI underestimated EDV values compared to ECHO data which led to lower EF values. This does not concur with a comparative study done on rats by Stuckey et al. This study was done in a rat model using a 500 MHz vertical bore system where MRI was found to overestimate EF values compared to ECHO [112]. The difference in this particular case could be because of difference in the species being studied. [A more detailed account of the comparison of these two methods, can be found to section 3.4. Different age group for control and TG mice is a major limitation of this study. Control group was younger (232 ± 60 days) than the TG group (417 ± 72 days). Therefore, it is important to know whether the decrease in EF and thickening of LV wall is an effect of maladaptive hypertrophy or aging. LV mass showed an increase with age in TG animals and four of the five TG animals were significantly older than the control ones. Therefore, it is important to distinguish the effect of age from the effect of hypertension‐induced cardiac hypertrophy. Although in the current study no clear correlation of age or body weight with cardiac functional parameters is observed, it will be useful to compare the results with age‐matched groups. 82
5.5
SUMMARY AND CONCLUSION Cardiac imaging and functional assessment of cardiac hypertrophy in a transgenic mouse model was performed. ECG‐gated cine images of the transgenic and control mouse hearts were used to compute cardiac functional parameters. These results were compared with those obtained from 2‐dimensional ECHO. Transgenic mouse hearts were found to have larger size and different shape compared to the control. Also, significant changes in the left ventricular ejection fraction and left ventricular wall thickness of transgenic mice, indicating hypertension‐induced cardiac hypertrophy, were observed. Echocardiography was unable to measure the increase in the LV width of the transgenic animals. Thus, this study demonstrates the application of high‐
resolution MRI for noninvasive and accurate determination of cardiac parameters in a transgenic mouse model of cardiac hypertrophy. 83
CHAPTER 6 IN VIVO MONITORING OF SPIO‐LABELED STEM CELLS TRANSPLANTED IN INFARCT MOUSE HEART 6.
In vivo monitoring of SPIO‐labeled stem cells transplanted in infarct mouse heart The primary focus of this chapter is the development of a MRI method to monitor the superparamagnetic iron oxide (SPIO)‐labeled mouse mesenchymal stem cells (MSCs), after transplantation in the infarct heart. Mouse MSCs were labeled using SPIOs. MRI methods were developed for in vitro, ex vivo and in vivo imaging of the labeled cells. Labeled were monitored up to 4 weeks after transplantation in infarcted mouse heart. 6.1
INTRODUCTION 6.1.1 Stem‐cell therapy for cardiac repair and regeneration Cardiovascular disease is one of the major causes of morbidity and mortality, accounting for more than half the deaths worldwide. Acute myocardial infarction (MI), leading to heart failure, is one of the major contributors to this statistic. Although many surgical interventions are available to rescue the failing heart, there is no definite cure for this condition. Therefore, any therapeutic approach that will lead to improved cardiac function following MI would be beneficial. Stem‐cell therapy is being pursued as a potential therapeutic strategy for reviving the failing heart [141, 142]. 84
Stem cells are unspecialized cells that have self‐renewal capability and can differentiate into cells of various lineages that have specific functions in the body [141]. They can be classified based on their differentiation capability (totipotent, pluripotent, multipotent) or based on the source (embryonic, adult). Stem cells are unspecialized cells that have self‐renewal capability and can differentiate into cells of various lineages that have specific functions in the body [141]. They can be classified based on their differentiation capability (totipotent, pluripotent, multipotent) or based on the source (embryonic, adult). They are further classified by the source organ or lineage such as mesenchymal stem cells, endothelial progenitor stem cells, skeletal myoblasts etc. Many of these stem cell types are being investigated for their possible therapeutic benefits. Due to the limited regenerative capability of the adult myocardium, the loss of cardiomyocytes owing to any cause such as ischemic heart disease, cardiomyopathies, hypertensive disorder etc, begins an irreversible cascade that results into heart failure [143]. This has sparked an interest in the possibility of cell transplantation therapies that can promote regeneration of cardiac tissue [144]. Stem‐cell therapy for the heart involves transplantation of immature cells into the infarct area for regeneration and revival of the cardiac tissue [145‐149]. The goal of stem‐cell transplantation is to improve cardiac function, induce angiogenesis/myogenesis and to reverse ventricular remodeling. A comparative study of various cell lines used for cell therapy in the heart was reported by van der Bogt et al. [150]. Of the many stem cell types studied for cardiac therapy, skeletal myoblasts are the most widely used in clinical trails in North America [151]. However, they have a few limitations. They can not differentiate into completely functional cardiomyocytes and have limited capability of forming electromechanical junctions with the native cardiomyocytes. These limitations drive the need for a more suitable cell line for use in cardiac therapy. Mesenchymal stem cells (MSCs) are pluripotent adult stem cells that primarily reside in the bone marrow [152]. MSCs are easily obtainable without ethical concerns, are adherent and can be easily expanded in culture. In addition to differentiating into 85
osteoblasts, chondrocytes, adipocytes, neurons, and skeletal muscle cells they also differentiate into vascular endothelial cells and cardiomyocytes [143, 152, 153]. They have been shown to improve cardiac function after transplantation [52, 69, 154, 155]. This indicates a possibility of therapeutic benefits after transplantation of MSCs into the injured myocardium. In this study, mesenchymal stem cells, isolated from the bone marrow of mouse, have been used for transplantation into the infarct heart. 6.1.2 Stem‐cell tracking using MRI Noninvasive methods for cell‐tracking Despite a plethora of studies related to the applications of stem‐cell therapy, not much is known of the fate of the stem cells after they are transplanted in vivo. Therefore, it is very important to develop methods to track the cell in vivo to locate the cells and monitor cell survival and integration. Currently histological analysis is used for cell tracking which is an invasive technique [143, 156]. The noninvasive methods used for cell tracking are positron emission tomography (PET), single photon emission computed tomography (SPECT) optical imaging methods, and MRI [33, 54, 157, 158]. Radionuclide imaging methods (PET and SPECT), although more sensitive than MRI, carry the risk of cell damage due to radiation [101]. Also, the radioactive probes used have short half‐life, thereby limiting long‐term tracking of the cells [101]. Optical imaging methods such as fluorescence have been used for stem‐cell tracking [159]. This technique depends on the transmitted light for detection of the cell which has high susceptibility to attenuation in the tissue and has low penetration depth, limiting its use near the surface. Advantages of MRI for cell‐tracking MRI presents several advantages over other noninvasive cell‐tracking techniques mentioned above. It uses radio waves that are not harmful to the cells and can easily pass through tissue without significant attenuation. MRI can also provide high‐
86
resolution information about the anatomy of the tissue and allows repetitive imaging, which facilitates long‐term tracking. However, for tracking cells using MRI, they have to be labeled using an MRI contrast agent. Two types of MRI contrast agents are generally used for cell labeling, gadolinium chelates and iron oxide particles [53, 101, 160]. Gadolinium‐based agents impart positive contrast (labeled tissue brighter than surrounding) based on T1‐shortening effects. A high concentration of labeled cells is required for detection using MRI. Iron oxide particles produce negative contrast (labeled tissue darker than the surrounding) effect due to the induction of magnetic field inhomogeneity in the surrounding water molecules and shortening T2 and T2* [55]. Negative contrast agents have a few limitations such as similarity of the hypointense region to a signal void due to anatomy and partial volume effects. Therefore, for cardiac applications, this necessitates optimized cardiac gating to obtain high resolution images. SPIO particles for cell tracking The commercially available dextran‐coated iron oxide particles are classified on the basis of their size as superparamagnetic iron oxide particles (SPIO) and ultrasmall SPIO (USPIO). SpiosBoth SPIOs and USPIOs have been used for cell tracking [52, 160]. One disadvantage of the USPIO particles is the requirement of high concentration (per cell) to produce detectable contrast. With each cell division, the concentration of the iron in each cell steadily decreases and finally the daughter cells do not retain enough iron to be detected by MRI. Therefore, for long‐term tracking, larger‐sized SPIOs are more appropriate. Micron size particles have been known to extend the magnetic field distortion up to 50 times their size [161, 162]. This decreases the particle‐concentration required for detection of the cells. One major concern with using SPIOs for cell tracking is the effect of the iron oxide particles on the cells. This is especially important in the case of stem cells since it can affect the differentiation ability of the cell, potentially negating the therapeutic benefit. Previous studies performed on human MSCs [52, 102, 163] have 87
shown that the internalization of these particles does not have a significant effect on the viability, proliferation or differentiation ability of the cells [53, 101, 158]. SPIO particles have been extensively used to monitor stem‐cell therapy in the heart [52, 156, 157, 164]. In this study, SPIO particles (average size – 0.9 µm) were used for labeling mouse MSCs. 6.1.3 Study design The aim of this study was to develop a method for MR imaging iron oxide labeled stem cells. MSCs were isolated from the bone marrow of the mouse and cultured. These MSCs were labeled using SPIO particles. For in vitro studies, agarose based phantoms were designed. Ex vivo studies were performed using hearts that were isolated one hour after the transplantation of the labeled cells. For in vivo studies, labeled‐cells were transplanted in mouse hearts after MI and then the animals were monitored for 4 weeks. 6.2
MATERIALS AND METHODS 6.2.1 Animals Male C57BL mice (n=2) were used in this study. All animal protocols used were approved by the Ohio State Institutional Laboratory Animal Care and Use Committee (ILACUC). 6.2.2 Culturing of MSCs MSCs were cultured from cryopreserved primary mouse mesenchymal stem cells isolated from the bone marrow of CBL57 mice. The cells were characterized to be positive for integrin β1 and CD54. The primary cells were thawed and passaged using Dulbecco’s modified Eagle medium (DMEM) (1X) (low glucose). Accutase, a cell‐
detachment solution containing proteolytic and collagenolytic enzymes, was used for separation of adherent cells. MSCs of passage 2 were used for experiments. The cells were grown at 37°C in 5% CO2 in an air‐humidified environment. 88
6.2.3 Labeling of MSCs with SPIOs SPIOs (Bangs Laboratories, Fishers, IN) (average size: 0.9 μm) containing magnetite cores encapsulated with styrene/divinyl benzene and coated with dragon green fluorescence dye (λEx =480 nm, λEm =520 nm) were used for cell labeling. Mouse MSCs were incubated for 16 h with 15 μl of SPIO/15 ml of medium in a T‐75 flask plated with 1 million cells. After the 16 hour period, the media were changed and a Nikon TE inverted fluorescence microscope equipped with a FITC filter set was used to confirm the presence of SPIOs in the MSCs. 6.2.4 Myocardial infarction and cell transplantation Refer to section 3.2.2 for the description of the surgical procedure. After the surgery, SPIO‐labeled cells were directly injected in the mouse myocardium at multiple sites. Totally, 15 µL of 0.5 million cells were injected in the left ventricular wall of the heart. 6.2.5 In vitro MR imaging Agarose phantoms were made by pouring 2% agarose solution in 3 ml eppendorf tubes. SPIO particle phantom was made by injecting 5 µL of SPIOs having 1µL/ml concentration. Similarly, 5 µL of 0.5 million/ml SPIO‐labeled MSCs were injected into the gel to make the labeled‐cell phantom. Control phantom was made by injecting 5 µL of 1 million/ml unlabeled cells into the agarose gel‐filled tube. The gel was allowed to cool at room temperature.For imaging the phantoms, a FLASH sequence with the following parameters was used: TR /TE: 75/1.4 ms; matrix: 256 × 192 zero filled to 256× 256; FOV: 3 × 3 cm; slice thickness: 1 mm; Number of slices: 5; NEX: 8. 6.2.6 Ex‐vivo imaging Mouse hearts were isolated one hour after surgery. The hearts were fixed with 10% formalin in ethanol solution. 6‐8 h after the treatment with formalin, the hearts were 89
suspended in 2% agarose gel. The same protocol as described above was used for the imaging of the isolated hearts. 6.2.7 In‐vivo imaging For the general procedure of cardiac imaging of mouse, refer to section 3.2.3 For imaging the phantoms, a FLASH sequence with the following parameters was used: TR /TE: 75/1.4 ms; matrix: 256 × 192 zero filled to 256× 256; FOV: 3 × 3 cm; slice thickness: 1 mm; Number of frames: 16; NEX: 8. 6.2.8 Prussian blue staining Efficacy of magnetic labeling was assessed using Prussian blue staining. For Prussian blue staining, the cells were incubated for 30 minutes with 2% potassium ferrocyanide in 6% hydrochloric acid and then counterstained with nuclear fast red. This indicates the presence of intracellular iron and the cellsʹ profile. Iron oxide particles appeared as blue precipitate in the cytoplasm in a cell positive for Prussian blue staining. 6.3
RESULTS SPIO particles were successfully incorporated into MSCs. The internalization was confirmed by fluorescence microscopy (Figure 6.1). After 24‐h incubation with the SPIO particles, cells ere visualized using a fluorescence microscope. Presence of black iron oxide particles (indicated by red arrows) was observed in SPIO‐labeled cells which were not seen in unlabeled particles. In vitro imaging was performed using 2% agarose gel. Control phantoms, with only agarose and unlabled cells, showed uniform signal intensity. Hypointensity was observed in the MR images of the phantoms with SPIO particles and SPIO‐labeled cells, confirming the presence of particles in the cells. Representative axial images are shown in Figure 6.2. 90
Figure 6.1 Confirmation of labeling of MSCs with SPIOs MScs were incubated with SPIO particles for 16 hours for labeling the cells. After 16‐h incubation the cells were washed and imaged using a fluorescence microscope equipped with FTIC filter. The fluorescence microscopy images (100X) confirmed the labeling of MSCs with SPIO particles. Arrows indicate the iron oxide particles that have adhered to the MSCs. Figure 6.2 In vitro images of SPIO‐labeled stem cells In vitro imaging of the SPIO‐labeled MSCs was done by using agarose based phantoms. A. Phantom with only 2% agarose gel B. 10 µL of SPIO particles was injected into semi‐cooled agarose gel and then the gel was allowed to cool at room temperature. C. 2% agarose gel injected with 5 µL of 1 mil unlabeled stem cells. D. 2% agarose gel injected with 5 µL of SPIO‐labeled stem cells (0.5 mil/ml). All the phantoms were imaged using a FLASH sequence (parameters: TR /TE: 75/1.4 ms; spatial resolution: 0.117 × 0.156 × 1 mm; Number of slices: 5; NEX: 8). Control phantoms (A, C) showed uniform signal intensity. Loss in the intensity of the MR signal was observed in the SPIO‐particle phantom (B) as well as the phantom with SPIO‐labeled MSCs (D), confirming the internalization of the probe in MSCs. 91
To establish the feasibility of imaging the transplanted labeled cells in cardiac tissue, ex vivo imaging of isolated heart after the transplantation of labeled MSCs was performed. The hearts were isolated from mice 1 hour after the surgery and then MR imaging was performed. The images show brighter outline of the isolated heart surrounded by agar matrix. A decrease in the signal intensity was observed in the areas corresponding to the injection sites of the labeled cells (Figure 6.3). Dry blood, which is seen along the base of the heart, also appears dark on these images. MRI imaging protocol was optimized to locate the SPIO‐labeled stem cells in the left‐ventricular wall (Figure 6.4). The hypointense area was visible all through the cardiac cycle and was observed to move in‐sync with the heart. This indicated that the source of the negative contrast was embedded in the heart muscle, further confirming that the hypointese area corresponded to the SPIO‐labeled stem cells injected in the left‐
ventricular wall of the mouse heart. Figure 6.3 Ex vivo images of mouse heart after transplantation of the labeled cells Mouse hearts were isolated an hour after the injection of SPIO‐labeled MSCs into the myocardium and treated with formalin (10% solution in ethanol) for 6‐8 hours. After fixation, the hearts were suspended in 2% agarose gel. Ex vivo MR imaging was performed using FLASH sequence (parameters: TR /TE: 75/1.4 ms; spatial resolution: 0.117 × 0.156 × 1 mm; Number of slices: 5; NEX: 8). Low signal intensity regions were observed where the cells were injected in the left‐ventricular wall. 92
To monitor the in vivo behavior of the labeled stem cells transplanted in cardiac tissue, mouse hearts were imaged up to 4 weeks post‐injection. The short‐axis images show right and left ventricle and the interventricular septum (Figure 6.5). The hypointense area corresponding to the labeled stem cells can be seen along the posterior wall of the left ventricle (indicated by the circles). This observation was supported by the long‐axis images (Figure 6.6) of the heart. These images show all four chambers of the heart along with the aorta. A region of hypointensity, corresponding to the site of injection of the labeled cells, can be observed along the LV wall (indicated by the circles). Images taken along both the axes show the presence of the labeled stem cells, for up to 4 weeks. The left‐ventricular wall was observed to have uniform signal intensity in the control images (week 0) that were acquired before the transplantation of the labeled cells. Figure 6.4 Locating the SPIO‐labeled stem cells in mouse heart Mouse heart images taken along short‐axis and long‐axis show the presence of a hypointense region that corresponds to the presence of SPIO‐labeled stem cells injected in the myocardium. The hypointense area was observed in the end‐diastolic as well as end‐systolic frames, confirming that the source of the contrast is embedded in the left‐ventricular wall. 93
Histological studies were carried out at the end of the study period to confirm the presence of stem cells labeled with SPIO particles. Prussian blue staining performed on the heart sections showed the presence of iron oxide particles. Higher magnification images showed that SPIO particles were also seen in the area surrounding the cell nuclei, confirming the presence of labeled stem cells 4 weeks after the transplantation. Figure 6.5 Monitoring the SPIO‐labeled stem cells in mouse heart (Short‐axis images) Short‐axis images of the heart show the right ventricle, left‐ventricle and the interventricular septum. A region of low signal intensity was observed in the left‐ventricular wall of the heart (encircled). The hypointense area corresponded to the injection site of the SPIO‐labeled MSCs and was observed clearly up to 4 weeks. In the pre‐injection (week 0) image, left‐ventricular wall was seen to have uniform signal intensity. 94
Figure 6.6 Monitoring the SPIO‐labeled stem cells in mouse heart (Long‐axis images) Both the atria and the ventricles can be clearly seen in the long‐axis images of the mouse heart. In the pre‐injection (week 0) image, left‐ventricular wall was seen to have uniform signal intensity. A region of low signal intensity was observed in the left‐ventricular wall of the heart (encircled). The hypointense area corresponded to the injection site of the SPIO‐labeled MSCs and was observed clearly up to 4 weeks. 6.4
DISCUSSION This chapter describes the in vivo identification and monitoring of SPIO‐labeled MSCs that were transplanted in mouse heart. Previous studies with mesenchymal stem cells have used human [14, 33] or rat [16, 32] MSCs for therapy. To the best of our knowledge, this is the first study to report the use of MRI to monitor SPIO‐labeled mouse MSCs transplanted in the infarcted mouse heart. The present study established the feasibility of labeling mouse MSCs and then monitoring them in vivo using MRI, for up to 4 weeks. 95
Previously, different sizes and types of iron oxide particles have been used to label stem cells [10, 16, 22, 32, 34]. The choice of the particles usually depends on the particle size, effect of particle internalization on the cells and the efficacy of the labeling. In this study, SPIO particles were chosen to label mouse MSCs. In vitro studies have shown that SPIO particles have little effect on the viability, proliferation, and differentiation potential of MSCs [16, 30, 31]. These studies have primarily been done in human MSCs, not in mouse MSCs. Although the fluorescence microscopy images obtained from labeled mouse MSCs did not show any significant changes in the cellular morphology, a thorough study of the effect of SPIOs on the viability and differentiation of mouse MSCs is warranted to establish the safety of labeling. Figure 6.7 Prussian blue staining 4 weeks after stem‐cell transplantation Prussian blue staining was performed 4 weeks post‐transplantation to confirm the presence of labeled cell in the left‐ventricular wall. A. Arrows show the presence of SPIOs (blue color) in the thinned LV wall. B and C show the magnified images of the LV wall showing the labeled cells. C. SPIOs (blue color) surrounding the cell nuclei (encircled) can be clearly seen confirming the presence of the labeled‐cells 4 weeks after transplantation. The small size and negative contrast effect of the iron oxide particles necessitate the optimization of MR imaging protocols for detection and monitoring of labeled stem cells. The proof‐of‐the‐concept was demonstrated using phantoms that contained labeled and unlabeled cells in agar matrix. In vitro imaging also confirmed the labeling of the cells and ruled out the possibility of hypointensity from unlabeled cells. This 96
established that the labeling concentration was optimal for in vitro detection of the labeled cells. The MRI protocol was further tested for sensitivity by imaging hearts that were isolated after the transplantation of the labeled cells. In the isolated hearts, a hypointense area was clearly observed at the site of cell transplantation. This indicated the presence of labeled cells in cardiac tissue and also established the capability of MRI to detect the cells in a biological matrix. The major challenge in in vivo imaging of labeled stem cells is the need for developing an optimized MR protocol with an appropriate gating setup. Also, it is difficult to distinguish the hypointense areas of labeled stem cells from the surrounding tissue. Appropriate changes were made to the pulse sequence parameters so that the contrast between the labeled cells and the surrounding tissue would be maximized. An optimized imaging protocol with minimization of motion artifacts resulted in images of the heart that showed a clear hypointense area corresponding to the injection sites of the labeled cells. This darker region was easy to distinguish from the otherwise homogenous cardiac tissue. It clearly demonstrated the capability of MRI to image and visualize the SPIO‐labeled mouse MSCs in vivo. Images were acquired to capture all the phases of the cardiac cycle. The labeled cells were observed in all the frames of the cardiac cycle, eliminating the possibility of an artifact generating the hypointensity in the images. Also, the labeled cells were clearly visible up to 4 weeks, indicating the possibility of long‐term monitoring without significant loss in the contrast. Also, good SNR of the MRI images obtained in this study may help concentration of SPIO particles that has been used for cell labeling . Reducing the SPIO concentration may also reduce the adverse effects that the particles may have on the differentiation of mouse MSCs [30]. The optimization of concentration will have to be performed taking into consideration the duration of the intended study and cell division, which can potentially dilute the contrast. One of the main concerns of cell labeling and transplantation studies is the fate of the labeled cells and the labeling agents after their transplantation in the tissue. To 97
confirm the presence of the SPIO particles at the end of the study period, Prussian blue staining was performed. The results indicated that the SPIO particles were present in the LV wall 4‐weeks post‐transplantation. Higher magnification images revealed the presence of cell nuclei confirming that the SPIO particles were associated with the cells. Although this gives a confirmation that cells and particles were both present at the end of the study period, this does not answer the questions regarding the viability or the differentiation of the cells at that time. Staining for SPIOs as well as MSC‐specific cell‐
surface marker may provide some of the answers. 6.5
SUMMARY AND CONCLUSION This study established the feasibility of labeling mouse MSCs with SPIO particles. It reported the development of method for in vitro, ex vivo and in vivo MR imaging of SPIO‐
labeled stem cells. It also demonstrated the capability of cardiac‐gated MRI to locate and monitor SPIO‐labeled MSCs in mouse heart over an extended period. 98
CHAPTER 7 SUMMARY 7.
Summary The development of cardiac MR imaging method for mouse and its application to different models of cardiac pathophysiology is reported. Chapter 1 introduced the need for the development of noninvasive cardiac imaging techniques. After brief overview of anatomy and functional parameters of the heart, the choice of mouse as an animal model for cardiovascular diseases was discussed. Cardiac MRI (CMRI) was compared with other noninvasive cardiac imaging modalities and the advantages of CMRI (noninvasive, capable of high resolution, clinically relevant) were discussed. The chapter ended with the review of current status of CMRI in mouse research. CMRI method development for mouse imaging at 11.7 T was discussed in detail in Chapter 2. It illustrated the optimizations that were performed to overcome the challenges of imaging fast‐beating mouse heart. Detailed discussion of the measures taken to improve cardiac gating was given. Last section was devoted to the functional analysis of the structural data obtained by MRI of mouse heart. M‐mode echocardiography (ECHO) is one of the most widely used methods for functional analysis of heart. In Chapter 3, ECHO was compared to CMRI for mouse cardiac functional assessment. It was observed that ECHO overestimated the end‐
diastolic, end‐systolic and stroke volumes compared to CMRI. Although the absolute 99
values of the functional parameters obtained from ECHO and MRI were different, both the modalities exhibited similar trends in healthy and failing mouse hearts. Bland‐
Altman analysis showed that there was general agreement in the two modalities for cardiac functional analysis. First application of the developed CMRI method was performed in permanent occlusion model of myocardial infarction (MI) in mice. CMRI showed the decline in the cardiac function in mouse after myocardial infarction. Significant increase in the end‐
diastolic volume and decrease in the ejection fraction was seen in the MI hearts. It was also observed that after myocardial infarction, the infarcted hearts can be grouped in two categories based on the extent of remodeling, especially hypertrophy. CMRI was also used for assessment of cardiac hypertrophy in a transgenic mouse model of hypertension. The hypothesis was that chronic hypertension would lead to maladaptive cardiac hypertrophy. CMRI was able to show the decrease in the cardiac function and the increase in left‐ventricular width in the transgenic mice, signifying cardiac hypertrophy. It was observed that the increase in LV wall thickness of transgenic mice was not seen in the ECHO measurements. This established CMRI, with its higher spatial resolution, to be a more effective tool in the assessment of cardiac hypertrophy. The next novel application of CMRI was to monitor labeled mouse mesenchymal stem cells (MSCs) in mouse hearts. sFor this purpose, mouse MSCs were labeled using iron‐oxide based MRI contrast agent. CMRI method for stem‐cell imaging was developed and tested using in vitro and ex vivo imaging. Labeled stem cells, injected in the peri‐infarct region of the mouse heart, were monitored for 4 weeks. The presence of labeled‐MSCs in mouse hearts was observed using CMRI and confirmed using Prussian blue staining at the end of the study period. In conclusion, an MR‐based imaging and functional analysis method for mouse hearts was developed and optimized to give high‐resolution images at 11.7 T. The 100
method was successfully applied to assess myocardial infraction and cardiac hypertrophy, and to monitor stem‐cell therapy in mouse heart. 101
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