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Bringing Cell Death AliveCardiovascular Toxicology (2003) 03 207–218 $25.00 (http://www.cardiotox.com) Humana Press © Copyright 2003 by Humana Press Inc. All rights of any nature whatsoever reserved. 1530-7905/01

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Bringing Cell Death Alive Ewald A. W. J. Dumont,1,* Chris P. M. Reutelingsperger,2 Guido Heidendal,3 and Leo Hofstra1 1

Department of Cardiology of the Cardiovascular Research Institute Maastricht; 2Department of Biochemistry of the Cardiovascular Research Institute Maastricht; and 3Department of Nuclear Medicine University Hospital Maastricht, University of Maastricht, Maastricht, The Netherlands

Abstract The role of apoptosis in ischemia and reperfusion of the heart has been widely debated. This controversy has been continued because of the lack of an apoptosis detection method that allowed obtaining detailed kinetic and quantitative information on apoptosis. Here we focus on recent findings that look into the detection of apoptosis following ischemia and reperfusion in the heart in animal models and in patients using Annexin-A5 based image technology. Following cardiac cell damage, one major characteristic finding is that apoptotic cells express phosphatidylserines (PS) on the outer leaflet of their cell membrane, serving as a “remove me” signal for the immune system. Annexin-A5, a native plasma protein with a high affinity for PS, can be used to measure this mode of cell death. Several Annexin-A5 based imaging systems have been developed to measure apoptosis from cell culture up to patients. In this review, implications, limitations, and clinical relevance of cell death imaging will be discussed. Key Words: Cell death; apoptosis; ischemia and reperfusion; imaging; fluorescence techniques; Annexin-A5; phosphatidylserine; SPECT imaging.

*Author to whom all correspondence and reprint requests should be addressed: Ewald A. W. J. Dumont, Department of Cardiology of the Cardiovascular Research Institute Maastricht, University of Maastricht, P.O. Box 616, 6200 MD Maastricht, The Netherlands. E-mail: Ewald.Dumont@ CARIM.unimaas.nl Received: 5/27/03 Accepted 5/27/03 Cardiovascular Toxicology, vol. 3, no. 3, 207–218, 2003

Cardiovascular Toxicology

In contrast to necrosis, apoptosis is a well-orchestrated biochemical cell death process that results in a typical morphology of the dead cell. Typically, apoptosis is characterized by cell shrinkage, chromatin condensation, membrane blebbing, and externalization of phosphatidylserines (PS) to the outer leaflet of the cell membrane. A central role in the biochemical execution of the cell is played by the caspases, a family of cysteine proteases that reside as inactive precursors in every cell. However, upon activation of the caspases these enzymes start to cleave many substrates, including deoxyribonucleases (DNases) and cytoskeletal proteins. This results in the classical appearance of the apoptotic cell and into cleavage of DNA. All apoptosis detection techniques rely on these biochemical and morphological alterations displayed by apoptotic cells. Most of these techniques, such as the widely used DNA electrophoresis and the terminal deoxynucleotidyl-mediated–dUTP nick-end labeling (TUNEL) assay require isolated cells or biopsies and are not suitable for in vivo detection of apoptosis. This limits the application of these techniques to obtain detailed kinetic and quantitative information on apoptosis. This information can be critical in the assessment of the efficacy of novel apoptosis

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modulating therapies, such as apoptosis inhibiting drugs in acute myocardial infarction or apoptosisinducing drugs in cancer. In this review, we discuss the development of the Annexin-A5 based imaging from bench to bedside and show examples of detection of cell death in experimental models and in patients with cardiac disease. In addition, we discuss the potential of Annexin-A5 based imaging technology to accelerate development of novel apoptosis modulating compounds (e.g., in cancer and in acute myocardial infarction). Finally, we explore the opportunities and limitations of the Annexin-A5 based imaging technology.

Acute Myocardial Infarction Acute myocardial infarction (AMI) is a clinical syndrome that results from injury of myocardial tissue caused by an imbalance between myocardial oxygen supply and demand. AMI usually occurs because of a sudden rupture of a vulnerable atherosclerotic plaque in the coronary artery leading to coronary occlusion. Each year there are about 1.5 million patients with AMI in the United States and there are approx 950,000 deaths (1). This is 40.6% of all deaths or one in every 2.5 death. Cardiovascular disease claims more lives each year than the next six leading causes of death combined (2). About every 29 s a person will suffer a coronary event, and about every minute someone will die from one (1). Therefore, the disease burden arising from cardiovascular disease remains high and continues to have a high emotional, social, and financial impact on our society. Treatment of AMI today is focused on the reopening of the occluded coronary artery, so-called reperfusion therapy. Large clinical trials have shown that reperfusion therapy with thrombolytic agents and/or invasive intervention (balloon and/or stent) results in smaller infarctions and better survival. However, despite these successes, up to 40% of patients with acute myocardial infarction develop heart failure, because of the loss of myocardial cells resulting from an AMI. Because the heart is not able to replace cardiomyocytes that were lost following myocardial infarction, it is important to find ways to protect the cardiac cells from dying during infarction. Therefore, treatments that are aimed to inhibit cell death during myocardial infarction deserve the center of


our attention. Despite many successes in experimental models of myocardial infarction, no benefit of pharmacological interventions has yet been shown in the patient with AMI. This may be due to lack of insight into the mechanisms and kinetics of cell death in the heart following ischemia and reperfusion. The use of Annexin-A5 based image technology may help to overcome these issues and may, therefore, result in the acceleration of drug development for prevention of cell death in AMI.

Apoptosis, How It all Started Based on the characteristic morphology when cells die, it was proposed in 1972 that death of normal but redundant cells, as well as some pathological ones, are suicides (2). That is, the cells activate an intracellular death program and kill themselves in a controlled way. From 1977, cell death genes have been studied extensively in the nematode worm Caenorhabditis elegans. From then on, DNA ladders were first observed and apoptosis genes identified (3). Apoptotic cells shrink and are then rapidly eaten by neighboring cells, before there is any leakage of their contents. The remaining apoptotic bodies are eaten and digested so quickly by adjacent cells and phagocytes, that usually few dead cells are seen, even when large numbers of cells have died. This could be the reason why apoptosis was neglected for so long. What we would today call truly apoptosis, or programmed cell death, was first recognized by Carl Vogt in 1842 who saw dying cells in the neuronal system of developing toad embryos. However, it is thought that the very first cells described, those from cork, by Hooke in 1665, were from corpses that had died through the apoptotic pathway (4). After studies by Walther Flemming in 1885 and others, cell death remained a subject of some interest in insect physiology. For instance, Ilya Mechnikov, won a Nobel prize in 1908 for his discovery of phagocytosis. In 1948 Saunders observed cell death in chick limbs and in 1949 Hamburger begins the exploration of the nerve growth factor. For the first time the morphological alterations in programmed cell death were described in 1964, in the review of Saunders named “shrinkage necrosis.” In 1972, Kerr first coined the term apoptosis to describe the process of programmed cell death.

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Signaling Pathways in Apoptotic Cell Death The search for targets to intervene in the apoptotic program has been the focus of many researchers in the last decade. Much pioneer work in apoptosis signaling research has been done in the worm C. elegans and the fruit fly Drosophila melanogaster. The first studies in C. elegans identified two genes, CED-3 and CED-4 (CED for cell death abnormal), required for apoptosis in the worm. If either gene is inactivated by mutation, the 131 cell deaths that normally happen during the development of the worm (which has 1090 cells when mature) fail to occur (5). Remarkably, the mutant worms with 131 extra cells have a normal life span. In contrast, more complex animals cannot survive without apoptosis: mutations that inhibit apoptosis in the fruit fly Drosophila melanogaster, for example, are lethal early in development, as are some mutations in mice (6,7). The protein encoded by the CED-3 gene was found to be very similar to the human protein called interleukin-1-converting enzyme (ICE) (8). The similarity between the CED-3 and ICE proteins was the first indication that the death program depends on protein cleavage (proteolysis). Soon new members of the CED-3/ICE family of proteases were identified, including more than ten in humans, many of which become activated in apoptosis. They are all cyscteine aspartic acid-specific proteases, and therefore called caspases (9). Each caspase is made as a large, inactive precursor or procaspase, which is itself activated, usually by another caspase (8). In apoptosis, caspases are activated in an amplifying proteolytic cascade, cleaving one another in sequence. Two distinct, but mutually not exclusive, pathways have been described in apoptotic cell death in mammalian cells, one occurring through death receptors (extrinsic pathway) and the other through the mitochondria (intrinsic pathway).

Cell Death Receptor-Mediated Programmed Cell Death (Extrinsic Pathway) In the death receptor pathway, soluble or cell surface death ligands, such as tumor necrosis factor-α


(TNF-α) and Fas ligand bind to their corresponding receptors inducing the recruitment and activation of caspase-8 or -10, thereby activating the cell death machinery (10). Adaptor proteins achieve activation of caspase precursors. Caspase-8 is activated when death effector domains (DEDs) bind to the C-terminal DED in the adaptor FADD (11). These caspases activate other downstream caspases, such as -3 -6, and -7, either directly (by proteolytic processing), or indirectly by cleaving Bid, a Bcl-2 family member. Bid activates also the mitochondrial pathway by permeabilizing the mitochondrial membrane (11). Once activated, the downstream or effector caspases fragment cellular proteins resulting in the orderly deconstruction of the cell. The effector caspase-3 cleaves the inhibitory subunit of the endonuclease ICAD, which leads to activation of CAD and fragmentation of nuclear DNA (12,13).

Mitochondrial-Mediated Programmed Cell Death (Intrinsic Pathway) In the mitochondrial pathway, which is thought to be the most important in ischemia and reperfusion damage of the heart, the central event is the translocation of cytochrome c, AIF, and endonuclease G from mitochondria to the cytoplasm (10). This pathway is initiated by death stimuli that include next to ischemia and reperfusion also oxidative stress, genotoxic stress, ultraviolet radiation, chemotherapeutic agents and calcium excess. Once in the cytoplasm, cytochrome c binds Apaf-1 stimulating its oligomerization and the subsequent recruitment of procaspase-9 into a large complex termed the apoptosome. Following its activation, caspase-9 activates downstream effector caspases. The death receptor pathway and mitochondrial pathway are linked by Bid, which gets cleaved by caspase-8 following death receptor activation. Subsequently, the carboxy fragment of Bid, termed tBid, translocates to the mitochondria where it inserts into the outer mitochondrial membrane and stimulates cytochrome c release through its interactions with Bax and/or Bak, which are pro-apoptotic proteins. A decision to die should not be taken lightly, and so it is not surprising that the death program is regu-

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lated in complex ways, both from inside and outside the cell. A major class of intracellular regulators is the Bcl-2 family of proteins, which, like the caspase family, has been conserved in evolution from worms to humans (14). Bcl-2 and Bcl-xL are antiapoptotic proteins, which are a component of the outer mitochondrial membrane; inhibit cell death indirectly by competing with Bax and Bak for tBid and possibly through their direct interactions with Bax and Bak. The human Bcl-2 gene, which was initially identified in a common form of lymphocyte cancer, has a chromosome abnormality, which causes excessive production of the Bcl-2 protein. The high levels of Bcl-2 promote cancer by inhibiting apoptosis, thereby prolonging cell survival. Remarkably, the human Bcl-2 gene is so similar to CED-9 that it can suppress apoptosis in C. elegans when artificially introduced into the worm (15). So far, 15 Bcl-2 family members have been identified so far in mammals. Some, such as Bcl-2 and Bcl-X, suppress apoptosis; others such as Bax, Bad, and Bid, promote it. The ratio of suppressors to promoters helps determine a cell’s susceptibility to apoptosis. The Bcl-2 family proteins regulate apoptosis in several ways (14). None of the known mammalian Bcl-2 family apoptosis suppressors binds directly to caspases, although they can inhibit caspase activation indirectly (14). Some block the ability to activate procaspase-9; some inhibit the release of cytochrome c from mitochondria, and some bind to the apoptosis promotors and inhibit their function. Many proteins of the Bcl-2 family have a hydrophobic tail that enables them to bind to the outside surface of mitochondria, the endoplasmic reticulum and the nucleus, and they are thought to function mainly on these organelles. Bax, for example, acts as an apoptosis promotor, in part at least, by binding to mitochondria, where it can interfere with mitochondrial function and release cytochrome c (16). The activity of Bcl-2 family members can be regulated by an increase or decrease of the expression of the gene that encodes them or by modification of the proteins themselves, by either the addition of phosphate (phosphorylation) or cleavage by proteases. The cleavage of Bcl-2 by caspases, for instance, inactivates Bcl-2’s inhibitory function, whereas the cleavage of the apoptosis promotor Bid by caspases releases a fragment that triggers the release of cytochrome c


from mitochondria; both events accelerate the death process (17,18). Other inhibitors include ARC or FLIP (inhibitor of caspase-8 activation), and XIAP or related proteins (inhibitors of caspases-3 and -9). When apoptosis is induced, in addition to cytochrome c, SMAC/Diablo is also released from the mitochondria that bind to XIAP precluding its inhibition of caspases.

How Annexin-A5 Works The detection of apoptotic cells using the AnnexinA5 assay is based on the high affinity of Annexin-A5 for PS, a membrane phospholipid that is exteriorized by cells during apoptosis. PS externalization on the cell membrane is one of the essential changes on the apototic cell, making it a target for phagocytes (19). In apoptotic cell death, the PS that in normal cells is only present on the inner leaflet of the cell membrane facing the cytosol is translocated to the outer leaflet of the cell membrane (20,21). Phagocytes have a PS receptor, which specifically binds to PS, engulfing apoptotic cells (22). Our group started investigating Annexin-A5 in 1992 as a phospholipid binding protein. AnnexinA5 is a member of a super family of proteins, the Annexins, which all bind to phospholipid surfaces (23). Annexin-A5 binds with high affinity to PS in the presence of Ca2+ ions, which change the confirmation of its binding site (24,25). How the mechanism of PS externalization is linked to the apoptotic machinery is yet unknown. After the discovery that apoptotic cells specifically express PS, the Annexin-A5 imaging assay was developed to measure apoptosis in vitro, e.g., for the fluorescence detection in flow- and laser scanning cytometry (26–28). Now, the Annexin-A5 assay is one of the most popular apoptosis detection systems for in vitro use.

In Vivo Imaging of Apoptosis in Animal Models The possibility of detection of cell death in vitro with Annexin-A5 raised the question whether labeled Annexin-A5 would be suitable for the in vivo detection of cell death in the complex environment of the whole organism.

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A first effort to study PS expression in the complex environment of the whole organism was addressed by Van den Eijnde et al. After intracardiac injection of living mouse embryos with biotinylated Annexin-A5, they showed after fixation on light microscopic level Annexin-A5 binding to apoptotic cells independent of germ layer (29). They found Annexin-A5 binding to interdigital webspaces at E13. Injection of a mutant Annexin-A5 for its four phospholipid binding sites by site-directed mutagenesis (M1234) did not bind to these cells in vivo (30), indicating that wild-type Annexin-A5 binding was specific and related to activation of the cell death program. Together, these data demonstrated that Annexin-A5 was able to detect programmed cell death during development in the complex in vivo environment. A next question we aimed to address was whether the Annexin-A5 protocol could be used to visualize programmed cell death in disease models in experimental models in vivo. For this purpose we developed an ischemia and reperfusion (I/R) model in the living mouse heart in situ. After intubation and ventilation of the mouse in a temperature-controlled environment, 30 min of ischemia and 90 min of reperfusion was applied. Before the end of the reperfusion phase the biotinylated Annexin-A5 was injected through a cannula into the jugular vein. Light-microscopic analysis of the injured area revealed binding of Annexin-A5 to cardiomyocytes undergoing programmed cell death as shown in Fig. 1. Further analysis of the Annexin-A5 positive cells showed caspase-3 activation and DNA laddering. Again in this model the mutant Annexin-A5 (M1234) did not bind any cardiomyocyte in the affected area. The data in this I/R model in the adult mouse show that PS externalization on injured cardiomyocytes can be visualized with intravenous-administered Annexin-A5 into the jugular vein (31). A disadvantage of the biotin labeled Annexin-A5 method is that biopsies and histologic processing are still needed. To overcome this problem, we injected fluorescent-labeled Annexin-A5 and modified our operation microscope with a fluorescence module to visualize binding of fluorescent Annexin-A5 during ischemia and reperfusion in the beating murine heart in situ on a cellular level (Fig. 2A) (32). With the same imaging protocol of 30 min of ischemia and 90 min of reperfusion but injection of fluorescent-labeled


Annexin-A5, revealed increased binding to injured cardiomyocytes starting directly after reperfusion reaching maximum fluorescence intensity after 20 min. Annexin-A5 binding was quantitated as the AnnexinA5 positive area as percentage of the area at risk. Inhibition of the apoptotic program with zVAD-fmk, a pancaspase inhibitor, reduced Annexin-A5 positive cells in the affected area from 20.2% to 10% (pretreatment) (32) (Fig. 2). A big advantage is that the kinetics of intracellular processes can be studied on a cellular level in the living organism, but the heart must be exposed which is not feasible in routine clinical practice. For the purpose of closed chest cell death imaging we turned to radiolabeled Annexin-A5 and subsequent nuclear imaging as first performed by Blankenberg et al. (33). They used technetium99m-labeled Annexin-A5 (Tc-Anx-A5) in several animal models and scintigraphy to visualize apoptosis noninvasively in a variety of animal models. After successful application of this imaging method in our I/R model in mice showing enhanced uptake of Tc-Anx-A5 in the area at risk (Fig. 3), we developed this method to be used in patients.

In Vivo Imaging of Cell Death in Patients After Acute Myocardial Infarction Today’s treatment of patients after an acute myocardial infarction (AMI) is aimed on restoring blood flow of the obliterated coronary artery, with either primary angioplasty (PTCA) or thrombolytic therapy. Animal studies from our and many other groups show that following reperfusion, programmed cell death is substantially enhanced and greatly contributes to the total damage (31). These studies also confirmed that inhibition of caspases decrease reperfusion-induced programmed cell death and reduced infarct size (34,35). From these studies, we concluded that it was most likely that after AMI and reperfusion therapy in patients apoptosis was contributing to cardiomyocyte cell death. To evaluate the possibility to detect programmed cell death in patients with AMI, we injected systemically Tc-Anx-A5 in seven patients with an AMI and successful reperfusion therapy. Single-photon emis-

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Fig. 1. Immunohistochemical staining of the heart of a mouse. The heart was subjected to 30 min of ischemia and 90 min of reperfusion. Thirty minutes before the end of the reperfusion time period, biotinylated Annexin-A5 was injected systemically into the jugular vein. At the end of the reperfusion period, the heart was routinely fixed and developed with DAB (brown staining of Annexin-A5, apoptotic cells) and counterstained with hematoxylin (blue nuclear staining). At the transverse section in A, the affected area is delineated and on high magnification (B) binding of Annexin-A5 to the cell membrane can be seen.

Fig. 2. (A) In vivo real time fluorescence molecular imaging system. Real time imaging of cell death in the heart of the living mouse using fluorescence imaging. A shows the experimental setup of the equipment. The mouse chest was opened and fluorescent Annexin-A5 was administered systemically 10 min before ischemia. Reperfusion was established after 30 min of ischemia.

sion computed tomography (SPECT) was performed 15 h after injection. As illustrated in Fig. 4, SPECT imaging showed enhanced binding of the radiotracer in six of seven patients (36). Important was that different infarct locations, ranging from small inferior Cardiovascular Toxicology

wall infarctions to large anterior infarctions could be visualized. After the initial Tc-Anx-A5 SPECT we also performed a Tc-99m sestamibi scan 3 d after the initial AMI, a well-established method to delineate perfusion defects in the heart after an AMI. Annexin-

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Fig. 2. (B) At various time-points, images were taken with a high-sensitive cooled CCD camera. Binding of fluorescent Annexin-A5 can be noticed on the cell membrane of the single cell in the beating murine heart in vivo. I/R: ischemia time in min/reperfusion time in min.

A5 binding perfectly coincided with the sestamibi defect (37). These data show that programmed cell death is a part of reperfusion-induced cell death after an acute infarction followed by reperfusion therapy. In addition, these data indicate that imaging of apoptosis with Annexin-A5 may help to assess the efficacy of possible strategies to modulate programmed cell death (38). Cardiovascular Toxicology

In Vivo Imaging of Cell Death in Patients with an Intracardiac Tumor To demonstrate the potential of cell death detection in a variety of cardiac diseases, we will discuss the detection of cell death in patients with an unknown intracardiac mass. Choosing the right therapeutic approach in treating cancer is to know the exact type

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Fig. 3. Imaging of cell death in the heart of the living mouse using nuclear imaging. The mouse chest was closed after 30 min of ischemia and imaging was performed 16 h after reperfusion. The affected area of the heart at risk (arrow) shows enhanced technetium-99m Annexin-A5 uptake. Excretion of the Annexin-A5 by kidney and liver is imminent.

Fig. 4. Single-photon emission computed tomographic (SPECT) analyses of the heart of an AMI patient. After an AMI, the patient underwent primary percutaneous transluminal coronary angioplasty (PTCA) of the infarct-related blood vessel. Twenty hours after intravenous injection of Tc-99m Annexin V, SPECT analysis was performed. To evaluate ultimate infarct size by SPECT, after 3 d, the patient received Tc-99m sestamibi. The perfusion defect on the Sestamibi SPECT is co-localized in concordance with the Annexin-A5 uptake as is shown by short axis (upper panel) and vertical long axis (lower panel) reconstructions. Cardiovascular Toxicology

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of tumor present. Therefore, biopsies are needed, although tumors sometimes appear on hard to reach spots, such as in the brain or in the heart. Sometimes taking biopsies can be a risk for the patient (e.g., tumor spread or bleeding disorders). Malignant tumors have been shown to have a high proliferation and high apoptotic rate (39). Therefore, noninvasive detection of apoptotic cells in a tumor could predict possible malignancy. Although cardiac tumors are uncommon, they almost invariably interfere with normal blood flow and may become life threatening. Because of the localization in the heart and possible spread of malignant cells, biopsies are difficult. Therefore, we studied the possibility to use the Tc-Anx-A5 protocol for noninvasive prediction of the possible malignancy of intracardiac tumors. Our first patient with an echographically proven large tumor in the left ventricle with almost complete obstruction of the mitral valve was systemically injected with Tc-AnxA5 (Fig. 5). SPECT imaging of the cardiac region showed enhanced uptake of the Tc-Anx-A5 in the heart, resembling the contour of the echographically detected tumor. To obtain information of the tumor location within the left ventricle, dual isotope imaging with thallium was performed, showing the tumor exactly within the left ventricle (see Fig. 5). These in vivo imaging data obtained with Annexin-A5 suggested a high rate of cell death in the intracardiac tumor, which may indicate a malignant nature of the tumor. After surgical removal of the tumor, immunohistochemistry showed a high-grade malignant sarcoma, Annexin-A5 binding to the cell membranes and cytoplasmic caspase-3 activation of apoptotic tumor cells. Using the same protocol for a second patient with a benign myxoma in the atrium of the heart showed no enhanced Tc-Anx-A5 uptake. This study suggests that Annexin-A5 can be used for noninvasive detection of malignant tumors and can be of value for clinical decision making.

Clinical Relevance of Cell Death Imaging After an acute myocardial infarction followed by reperfusion, we have previously shown that reperfusion of the infarct area of the mouse heart induces programmed cell death, which can be visualized noninvasively with Tc-99m–labeled Annexin-A5. These findings triggered us to extend our studies to Cardiovascular Toxicology

humans using the Annexin-A5 imaging protocol. The results presented show the feasibility of noninvasive monitoring of the dynamics of cell death in the heart. Now, the lack of a potent inhibitor of apoptotic cell death keeps researchers and cardiologists from studying the beneficial effects of these cell-death inhibitors using the Annexin-A5 imaging protocol. In the field of cardiac transplant research, Blankenberg et al. and our group have shown in animal studies detection of cell death in vivo with Tc-99m– labelled Annexin-A5 and nuclear imaging. Increased uptake of Annexin-A5 was seen in transplanted hearts, which correlated with the in vitro detection of DNA fragmentation by terminal deoxynucleotidyl-mediated- dUTP nick-end labeling (TUNEL). Recently, Narula et al. used the Annexin-A5 protocol to detect apoptosis in human cardiac allograft transplant recipients. Normally, endomyocardial biopsies are performed frequently to detect allograft rejection, which are not without any risk. In this study of 18 recipients, 13 patients had negative and five had positive myocardial uptake of Annexin-A5. These latter five demonstrated at least moderate transplant rejection and caspase-3 staining, suggesting apoptosis in their biopsy specimens. Immunosuppressive therapy with cyclosporine alleviated histologic evidence of apoptosis and rendered Annexin-A5 scans negative. This study shows the clinical feasibility and safety of Annexin-A5 imaging for noninvasive detection of transplant rejection by targeting cell membrane phospholipid alterations that are commonly associated with the process of apoptosis (40). In the imaging of neoplasia in oncology, follow-up of antitumor treatment is based on anatomical changes, which take days to weeks to be detected with computed tomography (CT) or magnetic resonance imaging (MRI). Using the Annexin-A5 protocol, molecular changes during cell death can be monitored directly after the first treatment and changes in therapeutic intervention can be made in a much faster way. As discussed earlier in this review, assessment of the potential malignancy of a tumor, contraindicated for taking biopsies, seems feasible. Furthermore, this technique allows measurement of the efficacy of therapies targeting cell death, such as antitumor therapies, on an individual basis. Hence, the Annexin-A5 imaging protocol might contribute to the growing propensity to base clinical decisions on the biological properties of the individual rather than on the statistics of a population.

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Limitations of Nucleotide Imaging A major disadvantage of current nuclear imaging techniques is that the resolution is approx 10 mm. Small numbers of cell undergoing programmed cell death cannot be detected, as well as small metastasis in tumor imaging. One solution may be the conjugation of Annexin-A5 with a marker, which can be used for MRI and/or PET, which have a higher spatial resolution than nuclear imaging. Such image modalities could be helpful in patients with heart failure, patients with unstable atherosclerotic plaques or patients with small (metastasizing) tumors. Next to the resolution difficulties, the timing of imaging is dependent on the blood pool activity after injection of the Tc-AnxA5. Until now, first images only could be made after 10–15 h after injection, making this protocol unsuitable for acute imaging for instance in acute coronary syndromes. Furthermore, Annexin-A5 is excreted through liver, kidney and bladder and this leads to a high background signal within these organs (see Fig. 3). Cell death detection in these organs therefore, for instance seen in kidney transplant rejection or hepatitis cannot be imaged with this protocol. The aforementioned organs overshadow the gut, making it difficult to use the protocol for intra-abdominal malignancies. To overcome this problem, our group is now testing different mutations of Annexin-A5, which show different biodistribution and excretion patterns.

Future Perspectives Until now, imaging of Annexin-A5 in patients has only been performed with SPECT imaging with a relaFig. 5. (opposite page) Annexin-A5 imaging and immunohistochemical analysis of an intracardiac tumor in a patient. The tumor mass in the heart of a patient was diagnosed by echocardiography (A, within white circumference). The patient then received thallium and Tc-99m Annexin-A5 intravenously for dual isotope SPECT analyses. B shows the uptake of thallium in the heart by short axis reconstructions. C shows the uptake of Tc-99m Annexin-A5 (within white circumference), which was predominantly located in the ventricular lumen. The tumor was surgically removed, routinely fixed and sectioned. The sections were stained using anti-Annexin-A5 antibodies (D) and antiactivated caspase-3 antibodies (E) either alone or in combination (F). Cardiovascular Toxicology

tively low resolution of 10 mm. Enhancement of resolution of Annexin-A5 imaging in patients is needed and other imaging modalities, such as MRI and PET must be explored. In an animal model of non-Hodgkin lymphoma increased uptake of Annexin-A5 is measured during successful treatment. These tumor-imaging programs are now extended into clinical practice. In the United States, the Annexin-A5 protocol is used for the follow up of the efficacy of chemotherapeutic treatment of breast carcinoma. Our group is working on methods to quantify the amount of Annexin-A5 uptake in the imaging studies. This allows us to quantify Annexin-A5 uptake in relation to the infarcted area in patients treated after suffering an acute myocardial infarction. This approach can also be of use in oncology in monitoring treatment efficacy of tumors. At last but not least, the lack of a potent cell death inhibitor, which can be used in humans, keeps us from proving whether the AnnexinA5 protocol is a reliable endpoint to test apoptosis blocking therapy in patients suffering an acute myocardial infarction. Until this moment, it is not known how long PS is expressed on the cell membrane during the cell death program. We have started a study in patients with a sub-acute infarction. Preliminary data showed a trend that PS could be detected up to 24–48 h after the initial event, which indicates that PS detection could be used as an “ischemic memory,” which could be clinically relevant for stating an acute event that has been going on in the previous period of 24 h, although no ECG alterations are present.

Acknowledgments Part of this work was supported by grants of the Dutch Scientific Organization (NWO-902.26.184) and the Dutch Heart foundation (NHS98.195).

References 1. American Heart Association. Heart and Stroke Update, 2001. 2. Kerr, J.F., Wyllie, A.H., and Currie, A.R. (1972). Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br. J. Cancer 26:239–257. 3. Lockshin, R.A. and Zakeri, Z. (2001). Programmed cell death and apoptosis: origins of the theory. Nat. Rev. Mol. Cell. Biol. 2:545–550. 4. Vaux, D. and Korsmeyer, S.J. (1999). Cell death in development. Cell 96:245–254.

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5. Ellis, R. (1991). Mechanisms and functions of cell death. Annu. Rev. Cell Biol. 7:663–698. 6. White, K., Grether, M.E., Abrams, J.M., Young, L., Farrell, K., and Steller, H. (1994). Genetic control of cell death in Drosophila. Science 264:677–683. 7. Kuida, K, Zheng, T.S., Na, S., et al. (1996). Decreased apoptosis in the brain and premature lethality in CPP32deficient mice. Nature 384:368–372. 8. Nicholson, D.W. and Thornberry, N.A. (1997). Caspases: killer proteases. Trends Biochem. Sci. 22:299–306. 9. Alnemri, E.S., Livingston, D.J., Nicholson, D.W., et al. (1996). Human ICE/CED-3 protease nomenclature. Cell 87:171. 10. Lassus, P. (2002). Requirement for caspase-2 in stressinduced apoptosis before mitochondrial permeabilization. Science 297:1352–1354 11. Muzio, M. Chinnaiyan, A.M., Kischkel, F.C., et al. (1996). FLICE, a novel FADD-homologous ICE/CED-3 like protease, is recruited to the CD95 (Fas/APOP-1) death-inducing signalling complex. Cell 85:817–827. 12. Liu, X., Zou, H., Slaughter, C., and Wang, X. (1997). DDF, a heterodimeric protein that functions downstream of caspase-3 to trigger DNA fragmentation during apoptosis. Cell 89:175–184. 13. Enari, M., Sakahira, H., Yokoyama, H., et al. (1998). Caspase-activated DNase that degrades DNA during apoptosis, and its inhibitor ICAD. Nature 391:43–50. 14. Adams, J.M. and Cory, S. (1998). The Bcl-2 protein family: Arbiters of cell survival. Science 281:1322–1326. 15. Vaux, D.L., Weissman, I.L., and Kim, S.K. (1992). Prevention of programmed cell death in Caenorhabditis elegans by human bcl-2. Science 258:1955–1957. 16. Green, D.R. and Reed, J.C. (1998). Mitochondria and apoptosis. Science 281:1309–1312. 17. Luo, X., Budihardjo, I., Zou, H., Slaughter, C., and Wang, X. (1998). Bid, a Bcl-2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors. Cell 94:481–490. 18. Li, H., Zhu, H., Xu, C., and Yuan, J. (1998). Cleavage of Bid by caspase-8 mediates the mitochondrial damage in the Fas pathway of apoptosis. Cell 94:491–501. 19. Fadok, V.A., Voelker, D.R., Campbell, P.A., Cohen, J.J., Bratton, D.L., and Henson, P.M. (1992). Exposure of phosphatidylserine on the surface of apoptotic lymphocytes triggers specific recognition and removal by macrophages. J. Immunol. 148:2207. 20. Zwaal, R.F. and Schroit, A.J. (1997). Pathophysiologic implications of membrane phospholipid asymmetry in blood cells. Blood 89:1121. 21. Verhoven, B., Schlegel, R.A., and Williamson, P. (1995). Mechanisms of phosphatidylserine exposure, a phagocyte recognition signal, on apoptotic T lymphocytes. J. Exp. Med. 182:1597. 22. Blankenberg, F.G., Naumovski L., Tait, J.F., Post, A.M., and Strauss, H.W. (2001). Imaging cyclophosphamideinduced intramedullary apoptosis in rats using Tc-99mradiolabeled annexin V. J. Nucl. Med. 42:309. 23. Reutelingsperger, C.P. (2001). Annexins: key regulators of haemastasis, thrombosis, and apoptosis. Thromb. Haemost. 86:413–419. Cardiovascular Toxicology

24. Swairjo M. A., Concha, N.O., Kaetzel, M.A., Dedman, J.R., and Seaton, B.A. (1995). Ca2+-bridging mechanism and phospholipid head group recognition in the membrane-binding protein annexin V. Nat. Struct. Biol. 2:968. 25. Andree, H.A., Reutelingsperger, C.P., Hauptmann, R., Hemker, H C., Hermens, W. T., and Willems, G.M. (1990). Binding of vascular anticoagulant alpha (VAC alpha) to planar phospholipid bilayers. J. Biol. Chem. 265:4923. 26. Blankenberg, F.G. and Strauss, H.W. (2001). Will imaging of apoptosis play a role in clinical care? A tale of mice and men. Apoptosis 6:117. 27. Martin, S.J., Reutelingsperger, C.P., McGahon, A.J., et al. (1995). Early redistribution of plasma membrane phosphatidylserine is a general feature of apoptosis regardless of the initiating stimulus: inhibition by overexpression of Bcl2 and Abl. J. Exp. Med. 182:1545. 28. Bedner, E., Li, X., Gorczyca, W., Melamed, M.R., and Darzynkiewicz, Z. (1999). Analysis of apoptosis by laser scanning cytometry. Cytometry 35:181. 29. Van den Eijnde, S.M., Luijsterburg, A.J, Boshart, L., et al. (1997). In situ detection of apoptosis during embryogenesis with annexin V: from whole mount to ultrastructure. Cytometry 29:313–320. 30. Mira, J.P., Dubois, T., Oudinet, J.P., et al. (1997). Inhibition of cytosolic phospholipase A2 by annexin V in differentiated permeabilized HL-60 cells. Evidence of crucial importance of domain I type II Ca2+-binding site in the mechanism of inhibition. J. Biol. Chem. 272:10474. 31. Dumont, E., Hofstra, L., van Heerde, W.L., et al. (2000). Cardiomyocyte death induced by myocardial ischemia and reperfusion measurement with recombinant human annexin-V in a mouse model. Circulation 102:1564. 32. Dumont, E.A., Reutelingsperger, C.P., Smits, J.F., et al. (2001). Real-time imaging of apoptotic cell-membrane changes at the single-cell level in the beating murine heart. Nat. Med. 7:1352. 33. Blankenberg, F.G., Katsikis, P.D., Tait, J.F., et al. (1998). In vivo detection and imaging of phosphatidylserine expression during programmed cell death. Proc. Natl. Acad. Sci. USA 95:6349. 34. Holly, T., Drincic, A., Byun, Y., et al. (1999). Caspase inhibition reduces myocyte cell death induced by myocardial ischemia and reperfusion in vivo. Mol. Cell Cardiol. 31:1709. 35. Yaoita, H., Ogawa, K., Maehara, K., and Maruyama, Y. (1998). Attenuation of ischemia/reperfusion injury in rats by a caspase inhibitor. Circulation 97:276. 36. Hofstra, L., Liem, I.H., Dumont, E.A., et al. (2000). Visualisation of cell death in vivo in patients with acute myocardial infarction. Lancet 356:209. 37. Piwnica-Worms, D., Kronauge, J., and Chiu, M. (1990). Uptake and retention of hexakis (2-methoxyisobutyl isonitrile) technetium(I) in cultured chick myocardial cells. Mitochondrial and plasma membrane potential dependence. Circulation 82:1826. 38. Reutelingsperger, C.P. and Hofstra, L. (2000). Visualisation of cell death. Lancet 356:2014. 39. Holmgren, L., O’Reilly, M., and Folkman, J. (1995). Dormancy of micrometastases: balanced proliferation and apoptosis in the presence of angiogenesis suppression. Nat. Med. 1:149–153.

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Volume 3, 2003