Paola A. Erba • Chiara Manfredi • H. William Strauss • Giuliano Mariani
The term “apoptosis” refers to the biologic process of programmed cell death, an active process requiring energy and regulated by the activation of specific genes, whose transcription and translation into active proteins determine death of the cell. The word “apoptosis” derives from two Greek roots, “apo” (which means from) and “ptosis” (which means falling). It was first coined by Kerr et al.1 over 40 years ago while investigating the rapid involution of hepatic parenchyma in response to acute surgical interruption of the portal venous blood supply to the liver.2 They demonstrated that cells within the ischemic hepatic lobes, interspersed among islands of normal tissue, transformed into small round vesicles of cytoplasm that often contained specks of condensed nuclear chromatin. These vesicles were then taken up and ingested by surrounding cells as well as by specialized (professional) mononuclear phagocytes. Although it was clear that these were dying cells, the process profoundly differed from necrosis because of the following: (i) the lack of adjacent inflammation; (ii) the mitochondria and ribosomes contained in the round vesicles remained intact throughout the process; and (iii) the aggregation of vesicles into clusters, which strongly suggested the “budding” or formation of these structures from the surface of the dying cell.
This new type of cell death contrasted with typical necrosis, in which dying cells lose membrane integrity, begin to swell up, then they spill their contents into surrounding tissues, thus causing local inflammation. Because the cells that Kerr observed in the liver actually decreased in size as they died, this type of cell death was also defined as “shrinkage necrosis.” Phenotypic appearance of apoptosis was soon reported to occur also within histologic specimens of basal cell carcinoma,3 within cancer specimens obtained from sites treated with radiotherapy, and within the adrenal cortex of rats treated with prednisolone.4 In addition, apoptosis associated with regression of experimental rat breast carcinoma after removal of the ovaries5 was also demonstrated.
However, the biologic mechanisms and pathways that drive apoptosis were at that time unknown. Continued investigation has shown that the histologic changes of apoptosis are preceded by an initiation stage called the “lag or trigger phase”6–9 and multiple triggers of apoptosis have now been identified, such as withdrawal of growth factors, DNA damage, immune reactions, ionizing radiation, chemotherapy, and ischemic injury.10–13 These and other triggers can start a cascade of events that lead to the morphologic changes of apoptosis. The lag time between exposure to the trigger(s) and the time when morphologic signs of apoptosis can be observed highly variable as it heavily depends on cell type, type of trigger(s), intensity and exposure, duration of trigger, and local environmental conditions.14
The apoptotic process can be divided into four different phases15,16: Initiation, execution, recognition/phagocytosis of the apoptotic bodies, and removal. All these phases, characterized by specific events, are morphologically defined by cell shrinkage and by a specific pattern of chromatin condensation, resulting in a dense crescentic mass close to the nuclear margin.17,18 Budding of the apoptotic bodies and the expression of new molecules on the cell surface serve as recognition signal for phagocytes to engulf the apoptotic cell, which thus dies without inflicting damage to viable neighboring cells.19–21 All these steps that allow the cell to die without inducing inflammatory response make apoptosis different from necrosis, which is characterized by the primary loss of cell membrane function and integrity, with consequent release into the environment of cytoplasm components causing local inflammation (Fig. 36.1).22
It was subsequently observed that apoptosis occurs physiologically in many circumstances, both during embryologic development23 and in adult life. In embryos, apoptosis is a mechanism that limits excessive proliferation (mitosis) during organogenesis. In adults, apoptosis regulates proliferation of immune cells and death because of viral infection, serving as a mechanism of defense from the outer injuries. For instance, neoplastic transformation can be considered as a condition characterized by a high rate of cell proliferation not counterbalanced by a corresponding higher of apoptosis level.
In physiologic conditions, the apoptotic machinery can be activated by different stimuli such as cytokines, severe DNA damage with no possibility of repair, decline of growth factors, low ATP supply, and various physical or chemical stimuli.24 On the other hand, causes of impaired activation of physiologic apoptosis can be found in specific mutations of cell cycle checkpoints where specific proteins are downregulated.
Apoptosis is a highly complex mechanism that can be activated by different signals that lead to the activation of specific apoptotic pathways but focus on the same end-point targets consisting of DNA fragmentation (with DNA laddering at 180- to 200-bp intervals), formation of the apoptotic body, and subsequent phagocytosis. The two different basic pathways activating apoptosis can be summarized as follows (Fig. 36.2):
• The extrinsic pathway represented by the tumor necrosis factor (TNF)-related apoptosis-inducing ligand (TRAIL) and its network of receptors.25
• The intrinsic pathway including the mitochondrial pathway and the endoplasmic reticulum pathway.26,27
A specific family of cysteine–aspartate-specific proteases called caspases plays a crucial role in the execution of apoptosis (Fig. 36.3). These proteases are activated by specific cleavage at an aspartic acid residue. Caspase family includes 14 different components that are present in virtually all cells.28 Caspases are activated by proteolytic cleavage of the zymogen through a reaction where the initiator caspases activate the executioner caspases. The executioner caspases are the effector molecules responsible for the phenotypic appearance of apoptotic cells. Such activation may occur in response to different stimuli, including developmental signals, as well as various forms of persistent cellular stress or injury.29
Initiator caspases, caspase-8, -9 and -10, activate the executioner caspases. Caspase-3, -6 and -7, the executioner caspases, have multiple intracellular targets, including inactivation of DNA repair enzymes and activation of DNA cleavage. Initiator caspases have long prodomains, whereas executioner caspases have short prodomains. The prodomain of the initiator caspases contains protein–protein interaction motifs for binding to upstream adaptor molecules.
Caspase regulation is under dual control by both activating factors (Apaf-1 and cytochrome c) and inhibiting factors (inhibitor of apoptosis proteins, IAPs), whose activity is in turn regulated by a complex network of upstream signaling pathways. Each caspase is also associated with a specific inhibitor, allowing the system to be strictly regulated by several positive and negative feedback mechanisms. The final enzyme activated within the cascade is caspase-3, which is activated both in the extrinsic pathway (by activation of the FAD CD95 receptor) and the intrinsic (mitochondrial) pathway and it is responsible for the activation of the target protein leading to DNA cleavage or disassembling of the cytoskeleton.
FIGURE 36.1. Schematic representation of the phenotypic cellular events leading to necrosis (upper pathway) or to apoptosis (lower pathway) following a variety of initiators. (From Galluzzi L, Vitale I, Abrams JM, et al. Molecular definitions of cell death subroutines: Recommendations of the Nomenclature Committee on Cell Death 2012. Cell Death Differ. 2012;19:107–120, with permission.)
After activation of caspase-3, the morphologic events of apoptosis quickly follow, resulting in the orderly breakdown of cellular proteins (including the cytoskeleton and nuclear matrix), and in the activation of poly-ADP-ribose polymerase (PARP), an enzyme that facilitates the degradation of nuclear DNA into 50- to 300-kb-sized pieces (DNA ladder formation). These morphologic events are collectively called the “execution phase.” The hallmark of the start of the execution phase is redistribution and exposure of phosphatidylserine (PS) on the outer layer of the cell membrane (Fig. 36.4).30 PS is a negatively charged aminophospholipid present in all cells that constitutes approximately 2% to 10% of total cellular lipid. Mammalian cells synthesize PS predominantly by converting phosphatidylcholine (PC) and phosphatidylethanolamine (PE) through a serine exchange reaction. The enzymes PS-synthase 1 and 2, present in the endoplasmic reticulum, catalyze the conversion having PC and PE as the substrates, respectively. PS appears to be crucial to the cell and, as such, is produced by different biosynthetic routes that can compensate each other to maintain a certain minimal level of PS in case one route fails.31,32 Several transport mechanisms including vesicular transport and lipid-transfer protein-mediated lipid-exchange between juxtapositioned bilayers33 allow PS to reach the cell membrane, where it is subject to the action of the aminophospholipid transporter (APLT), which translocates PS rapidly from the exoplasmic to the cytoplasmic leaflet if PS appears in the exoplasmic leaflet. APLT is a member of the family of P4-type ATPases, a class of ATPases that mediate in an ATP-dependent manner the transbilayer movement of phospholipids.34 APLT activity is present mostly in blood cells (including erythrocytes, leukocytes, platelets, and nucleated cells in the bone marrow),35 where they reside in the membrane and in trans-Golgi and Golgi-derived secretory vesicles. Once established, PS asymmetry between the inner and outer leaflets of the plasma membrane is stable and is maintained by “translocase,”36 whereas “floppase” maintains an asymmetric distribution of different phospholipids.37–39
APLT activity is required again if disturbances caused by, for example, membrane fusion processes during endo- and exocytosis occur. Inhibition of APLT activity only results in a slow rate of PS exposure,40 thus indicating that PS asymmetry of the plasma membrane is important for cell homeostasis. Certain conditions can cause translocation of PS to the outer leaflet of the plasma membrane, where it may initiate and participate in humoral and cellular processes such as blood coagulation and phagocytosis. In this process a third enzyme, “scramblase,”41 together with the simultaneous calcium ion–dependent deactivation of “translocase” and “floppase,” is needed to redistribute phospholipids on the plasma membrane. Scrambling (a rapid, ATP-independent and nonphospholipid-species–selective event) causes randomization of the phospholipids over the two membrane leaflets. Scramblase operates in erythrocytes,42 in activated platelets,43 and in apoptotic cells.44 Phospholipid scramblase 1 (PLSCR1) seems to be the protein more often involved in phospholipid scramble of the membrane.45 However, cells are able to scramble plasma membrane phospholipids in the absence of PLSCR1,46 and the level of PLSCR1 expression is not correlated with the capacity to externalize PS during apoptosis.47 No other candidate mechanisms have been hypothesized, thus indicating complexity of phospholipid scrambling and, possibly, diversity in scrambling mechanisms.48
FIGURE 36.2. A: Extrinsic apoptosis. Upon binding of the FAS ligand (FASL), the cytoplasmic tails of FAS (also known as CD95, a prototypic death receptor) trimers recruit several proteins, including FAS-associated protein with a death domain (FADD), cellular inhibitor of apoptosis proteins (cIAPs), c-FLIPs, and procaspase-8 (or -10). This event results in activation of caspse-8 and -10, according to a sequence that has been denominated “death-inducing signaling complex” (DISC). Nevertheless, within the DISC, there are some proteins that exert prosurvival functions (c-FLIPs and cIAPs). When lethal signals prevail, activated caspase-8 can directly trigger the caspase cascade by proteolytic maturation of caspase-3 (in type I cells) or can stimulate mitochondrial outer membrane permeabilization (MOMP) by cleaving the BH3-only protein BID (in type II cells). Dependence receptors such as DCC or UNC5B, which relay lethal signals in the absence of their ligand (netrin-1), can also initiate extrinsic apoptosis, by activating caspase-9, through the assembly of a DRAL- and TUCAN- (or NLRP1-) or by the dephosphorylation-mediated activation of death-associated protein kinase 1 (DAPK1) by UNC5B-bound protein phosphatase 2A (PP2A), respectively. DAPK1 can mediate the direct activation of executioner caspases or favor MOMP. B: Intrinsic apoptosis. Multiple intracellular stress conditions (e.g., DNA damage, cytosolic Ca2+ overload) generate prosurvival and prodeath signals and converge to a mitochondrion-centered control mechanism. If lethal signals prevail, MOMP leads to dissipation of the mitochondrial transmembrane potential (δψm), arrest of mitochondrial ATP synthesis, and δψm-dependent transport activities. Moreover, uncoupling of the respiratory chains leads to overgeneration of reactive oxygen species (ROS), and release into the cytosol of proteins that are normally confined within the mitochondrial intermembrane space (IMS). In particular, cytochrome c (CYTC) and the cytoplasmic adaptor proteins APAF1 and dATP drive the assembly of the apoptosome, a multiprotein complex that triggers the proteolytic caspase-9 → caspase-3 cascade. (From Galluzzi L, Vitale I, Abrams JM, et al. Molecular definitions of cell death subroutines: Recommendations of the Nomenclature Committee on Cell Death 2012. Cell Death Differ. 2012; 19:107–120, with permission.)
Translocation of PS to the membrane exoplasmic leaflet does not compromise the barrier function of the membrane. Once in the exoplasmic leaflet, PS may participate in a variety of processes depending on type and localization of the PS-exposing cell. Cell surface expression of PS has been found with aging erythrocytes, activated platelets, activated macrophages, endothelial cells of tumor blood vessels, apoptotic cells, apoptotic bodies, and cell-derived microparticles.
The exposure of PS functions as target for the so-called professional phagocytes, such as monocytes and macrophages or, at a slower pace, adjacent stromal cells such as fibroblasts and vascular smooth muscle cells that are responsible for ingesting the small membrane-bound packets called “apoptotic bodies.” Inhibition of PS exposure greatly impairs phagocytic capacity of the activated macrophages.49 First described for apoptotic lymphocytes,50 PS exposure is recognized as a ubiquitous phenomenon of apoptosis that is independent of cell type and of cell-death–inducing trigger51 and is phylogenetically conserved.52
In the early phases of apoptosis, before the start of autodigestion of DNA and the self-packaging of intracellular contents, cells can also be directly engulfed by phagocytes. It is currently unclear which of these modes of apoptotic cell removal dominates in vivo. The mode of removal may in fact depend on the actual local conditions surrounding unwanted cell(s).
Modulators of Apoptosis
Apoptosis can be regulated by different modulators such as ions, genes, proteins, and cellular organelles (mitochondria or endoplasmic reticulum). Calcium ions are constantly required during the apoptotic process and are necessary to activate endonucleases, transglutaminase-specific proteases, reorganization of the cytoskeleton, and scramblase.
Besides calcium ions, different proteins can determine whether apoptosis proceeds or stops. This is the case for the Blc-2 family, composed by proteins subdivided into two groups on the basis of their proapoptotic or antiapoptotic activity, respectively.
Similarly, as the Bcl-2 family components, p53 protein (so denoted because its molecular weight is 53 kDa and also called “the Genome Guardian”) plays a crucial role in the activation or inhibition of the apoptotic process. The p53 protein is normally involved in cell cycle progression, senescence, DNA metabolism, angiogenesis, and cellular differentiation. In particular, p53 plays a crucial role in blocking the cell cycle progression and/or in inducing apoptosis in response to cellular stress or DNA damage.53 In physiologic conditions, the transcription of p53 is downregulated by the transcription factor Mdm2 (murine double minute 2), which inhibits the activation and accelerates the degradation of p53 upon binding; however, if DNA is damaged, specific protein kinases (depending on the type and severity of damage) phosphorylate p53 and disrupt its binding with Mdm2, thus prolonging its biologic half-life. The activation of p53 determines either a mechanism repairing DNA damage, or (for more severe DNA damage) activates apoptosis.
FIGURE 36.3. Cross-talk between the inflammatory and apoptotic caspase network, illustrating the analogies and overlaps between formation of the inflammasome and apoptosome. Bold arrows are established links, and dashed arrows indicate possible interactions not yet fully established.
FIGURE 36.4. Schematic representation of phosphatidylserine (PS) redistribution on the cellular membrane and formation of blebs during apoptosis. PC, phosphatidylcholine; PE, phosphatidyl-ethanolamine. (From Blankenberg FG, Norfray JF. Multimodality molecular imaging of apoptosis in oncology. AJR Am J Roentgenol. 2011;197:308–317, with permission.)
Cytochrome c is also crucially important for apoptosis. Because it is normally located in the inner mitochondrial membrane, its release indicates that an irreversible stage of apoptosis has been reached. Cytochrome c binds to APAF1 and to caspase-9 forming a complex called “apoptosome” that, together with ATP, induces activation of the executioner caspase-3.
Mitochondria represent the core subcellular organelles in the execution of apoptosis. There are different proteins or factors that are normally located in the inner layer of the mitochondrial membrane. Among these, it is important to mention the apoptotic inducer factor (AIF), which is responsible for a caspase-independent apoptotic pathway. The mechanism by which AIF and cytochrome c are released from mitochondria is not well defined. AIF-dependent apoptosis seems to be activated in ATP deprivation conditions.54
Induction of Apoptosis
Apoptosis can be activated either by extracellular or by intracellular factors, depending on the specific signal, for example depletion of a growth factor. Extracellular factors correspond to specific ligands that activate a specific apoptotic pathway by binding to their specific receptor(s), which are therefore called “death receptors.” These receptors, normally located on the cell membrane, are members of the TNFR family, such as Fas APO-1 and Fas CD95, TNF receptor-1 (TNFR-1), DR-3 (TRAMP), DR-4 (TRAIL-R1), and DR-5 (TRAIL-R2).
When a “death ligand” (TNF or CD95L) binds to its specific receptor, the activated receptor recruits the adaptor proteins (FADD and TRADD) at their intracellular site, the death domain (DD). The complex is able to recruit procaspase-8 that, once activated, in turn activates procaspase-3, or cleavage of the proapoptotic protein Bid, that belongs to Bcl-2 family. Bid is activated by proteolysis and is able to penetrate into the mitochondria, thus determining release of cytochrome c in the cytosol.
Among the intracellular factors, p53 is activated by severe DNA damage, such as exposure to x-ray, chemotherapeutic agents, and various physical or chemical agents. For example, the rapid activation of p53 caused by ionizing radiation is mediated by the ataxia-telangiectasia–mutated (ATM) kinase. Upon activation, p53 and its downstream effectors (e.g., p21) regulate different responses, including cell cycle checkpoints, apoptosis, and stress-induced premature senescence (SIPS). In most human cell types, the primary response triggered by moderate doses of DNA-damaging agents is not apoptosis but a sustained proliferation block.55–58 The proliferation block triggered by ionizing radiation predominantly induces SIPS in p53-proficient cultures,59 whereas it induces the formation of multinucleated and polyploid giant cells in p53-deficient cultures.60 Evidence is emerging that such responses may represent cell survival mechanisms consequent to treatment with ionizing radiation. If DNA damage is severe and not repairable by the repair machinery, p53 is able to induce apoptosis by increasing the levels of Bax (a proapoptotic protein belonging to the Bcl-2 family), thus altering the Bcl2/Bax ratio. Upon activation, Bax moves from the cytoplasm onto the mitochondrial membrane, where it induces formation of a pore that causes the release of cytochrome c from the mitochondria. The process continues on with formation of the apoptosome and subsequent activation of procaspase-3.
The third apoptotic pathway is activated by damage/stress of the endoplasmic reticulum (ER). In fact, also on the ER surface there are various proteins involved in apoptosis. Caspase-12 (a murine caspase not yet identified in human tissues) is specifically associated with the ER stress, and its activation occurs as a response to ER stress either by the activation of calpain or by cleavage of caspase-7. An additional mechanism of induction can be linked to IRE-1, that can aggregate procaspase-12 at the ER membrane surface; recruiting of the cytosolic protein adaptor TRAF2 results in cleavage and activation of caspase-12, presumably via an induced proximity mechanism. Combination of caspase-12 with caspase-9 mediates a new intrinsic apoptosis pathway, apoptosome- and mitochondria-independent. Nevertheless, an ER-dependent apoptotic pathway can be activated also by a different protein, BAP31; this protein is normally located on the ER transmembrane space, where it normally acts by binding with nascent membrane proteins in transit between the ER and cis-Golgi network and exists in a complex with procaspase-8 and the antiapoptotic regulators Bcl-2 or Bcl-x. ER stress or other apoptotic signals lead to the cleavage of BAP31, with formation of the p20 fragment that initiates the following cascade: release of Ca2+ from the ER, upatake of Ca2+ into the mitochondria causing release of cytochrome c and initiation of apoptosis. Furthermore, caspase-8 can cleave BAP31 at the ER and thus stimulate Ca-dependent mitochondrial fission and enhances the release of cytochrome c. Thus, the caspase-derived fragment of BAP31 may coordinate cell death signals between the ER and mitochondria.
Regulation of Apoptosis by the Bcl-2 Family Proteins
The Bcl-2 family is composed of about 30 different members that result from the interaction of about 25 genes and can be classified on the basis of their proapoptotic or antiapoptotic activities. The presence of four conservative domains (BH1–BH4) permits to cluster these proteins into a single family, the denomination of which derives from the prototype protein, Bcl-2 (the 26-kDa product of the bcl-2 gene). In particular, the BH4 domain is present in all the antiapoptotic proteins such as Bcl-2, Bcl-xL, and Bcl-w, whereas the BH3 domain is typical of the proapoptotic proteins such as Bax, Bad, Bak, and Bok. Most of these proteins (that have at their C-terminus a group of hydrophobic amino acids permitting anchorage to membranes) are localized on the outer mitochondrial membrane and enable MOMP; alternative locations are in the endoplasmic or nuclear membrane.
In normal condition the Bcl-2 protein is mostly localized in the cytosol but, in response to an apoptotic stimulus, it moves onto the external mitochondrial membrane, where its overexpression enhances resistance to apoptosis61 by maintaining impermeability and the membrane potential of mitochondria. However, the activation of Bax or Bak disrupts mitochondrial permeability, with consequent release of cytochrome c and Smac/DIABLO from the mitochondria and formation of the apoptosome complex. Following an appropriate injury, Bax (which is normally confined in the cytosol) binds to the mitochondrial membrane by its hydrophobic anchorage and induces pore formation; thereafter, the voltage-dependent anion channel is responsible for selective translocation of AIF, cytochrome c, and SMAC/Diablo to the cytosol.
Bid, Bax, and Bad are also crucially important for perpetuating apoptosis. Bid is cleaved by caspase-8, and the fragment containing the BH3 domain activates release of cytochrome c. Bid functions as a proapoptotic factor acting in two ways: The first mimics the Bax pathway (pore formation), whereas the second one inhibits formation of the Bcl-XL-Apaf-1 complex. Bad activates the apoptotic pathway because its phosphorylated form binds to and inhibits the antiapoptotic Bcl-x and Bcl-2 proteins, thus allowing the continuation of the apoptotic cascade.
The Bcl-2 family includes members that exert their activity in the cytoskeleton too, where the final target is disassembling of the microtubules operated by the Bim protein, which is normally sequestered as a microtubule-associated dynein motor complex. By moving to the mitochondria during apoptosis, Bim enhances the proapoptotic activity and induces release of cytochrome c from the mitochondria.
Activation of Caspases as Apoptosis-Specific Proteases
Caspases are normally present in cells in their inactive form as zymogene. They are activated by specific cleavage by Apaf-1 which is, in turn, activated by the cytochrome c released from mitochondria, but can also be activated by the adaptor proteins at the cytoplasmic site of the death receptors described above.
Although activation of a particular caspase following an apoptotic stimulus is mostly tissue-specific, caspase-3 is ubiquitous. Caspase-3 is activated in the final phase of apoptosis, causing DNA cleavage.
Caspases can act on different substrates, mostly intracellular signal transporters. The most important and more extensively investigated substrate is PARP-1, a conserved multi-domain enzyme that is present in virtually all eukaryotes (except in yeast). PARP-1 is normally stored into the nucleus in an enzymatically inactive form bound to chromatin, with a density of 1 every 20 nucleosomes. When DNA damage occurs, PARP-1 catalyzes the cleavage of its substrate NAD into nicotinamide and ADP-ribose and polymerizes long ADP-ribose chains onto core histones, linker histone H1, and many other nuclear proteins (heteromodification), as well as onto itself (automodification). The final target of this chain is to activate the repair system in case of mild DNA damage; whereas, in severe DNA damage, a different mechanism leading to cellular death is activated. Other substrates for caspases are pRB, NuMA, DNA-PK. pRb is the product of the retinoblastoma gene determining cell cycle arrest; denomination of this gene derives from the childhood cancer retinoblastoma, because of somatic mutation of both alleles of the gene. Nuclear mitotic apparatus (NuMA) and DNA-PK, a critical component of the nonhomologous end-joining pathway, influence which pathway is ultimately utilized for repair.
APOPTOSIS AND DISEASE
Control of apoptosis is critical for normal tissue development, and tissue homeostasis, whereas inhibition of apoptosis contributes to the development and progression of cancer.62,63 Recognition and clearance of PS-expressing cells are extremely efficient in healthy tissues, which therefore contain, if any, a low steady-state level of PS-expressing cells. Different pathologic conditions can shift the balance between appearance and clearance of PS-expressing cells toward a sustained presence of PS-expressing cells as well as of cell remnants, such as apoptotic bodies and cell-derived microparticles in tissues. Many tissues can tolerate a surprisingly high level of cell death, because they compensate for the loss of cells through increased cell proliferation and regeneration.64 For example, a full-sized mammalian liver can regenerate after as much as 75% of the whole organ has been removed.65 Depending on the type of tissue damage, these regeneration processes involve several steps, including wound healing, the formation of proliferative blastema cells, differentiation, and patterning.66 As hypothesized as early as 1988,67 work in several model organisms has revealed that, unexpectedly, apoptosis may also drive cell proliferation during tissue regeneration.68–76 In particular, the proliferation component of regeneration, including the formation of blastema cells, is under the control of apoptotic cells through a series of events that has been termed “apoptosis-induced compensatory proliferation.”77
However, mitogenic signaling by apoptotic cells may contribute also to the development of neoplasms. The old concept that tumors resemble wounds that do not heal78,79 has more recently been supported by the observation that several pathways involved in tissue regeneration and stem cell self-renewal play prominent roles in human cancer as well.80–85 Therefore, even if wound healing, inflammation, tissue stress, or tissue damage on one hand promote a certain degree of apoptosis in cancer cells, on the other hand these dying cells may release mitogens that promote malignant growth. Even if the release of mitogens by apoptotic cells is not sufficient per se to promote the overall growth of a tumor, it may nevertheless stimulate proliferation of cancer stem cells86 or provide a supportive microenvironment facilitating tumor growth.87 This process could contribute to regrowth and relapse after favorable response of cancer to therapy, and also to distant seeding of metastases. This possibility deserves serious consideration for several reasons. First, virtually all cancer cells have acquired at least some degree of resistance toward apoptosis, and hence they have features of the so-called “undead” cells.88 Second, most existing anticancer therapies, including radiation and chemotherapy, kill cancer cells by apoptosis, and hence are expected to induce a “compensatory proliferation” response. Third, it has been demonstrated that compensatory growth is at the base of oncogenic cooperation between genetically distinct cells in a Drosophila tumor model.89 Therefore, the combination of apoptosis resistance and strong, therapy-induced cellular stress and damage may lead to the expansion of cancer stem cells and hence increase the likelihood of tumor regeneration and development of secondary tumors. Thus, development of a cancer (dysregulation leading to abnormal cell accumulation) and its successful treatment (iatrogenic modification stimulating tumor cell removal) may represent the two opposite sides of the apoptosis coin.
Dysregulation of apoptosis can cause either the accumulation of unwanted cells (as it may happen in cancer) or the premature removal of useful and/or essential cells (as it may happen in Alzheimer’s disease or in some autoimmune rheumatologic disorders).
Externalization of PS on the outer leaflet of the cell membrane does not only occur in apoptosis, as other forms of cell death can also demonstrate this feature. PS externalization occurs in necrosis/oncosis, mitotic catastrophe, cell senescence, pyroptosis, PARP-1-mediated cell death, and autophagy.90 In particular, autophagy (type II cell death, also known as “self-eating”) exhibits considerable overlap with apoptosis.91 Autophagy is initiated by derepression of the mammalian-cell target of rapamycin (mTOR Ser/Thr kinase), followed by formation of the Beclin-1-class III phosphatidylinositol 3-kinase complex. This complex mediates the formation of isolation membranes that engulf cytoplasmic material, thus producing autophagosomes. Bcl-2 and Bcl-XL are regulators of beclin-1, and constitute therefore a link with apoptotic cell death.92 Autophagosomes fuse then with lysosomes to create autolysosomes. In the autolysosome, the inner membrane and the luminal contents of the autophagic vacuole are degraded without eliciting inflammation. The main role of autophagy is to remove unneeded, senescent, or damaged cytoplasmic contents and to allow a cell to survive periods of cellular famine through the autodigestion of intracellular DNA/RNA, proteins, and lipids into free nucleotides, amino acids, and fatty acids. Autophagic cell death can also represent an alternative to apoptosis, if this latter mechanism is damaged/inhibited.
A related form of cell death involves lysosomal stress followed by lysosomal membrane permeabilization and release of cathepsin that causes generalized proteolysis. Details of when and how this process occurs are still poorly understood, but it causes expression of PS on cell membranes to ligands.
Recently, reversible PS externalization in a process independent of cytochrome c release, caspase activation, or DNA fragmentation was demonstrated.93 However, it accounts for lower levels of PS exposure as compared with apoptosis and other forms of cell death.94,95 The relatively low levels of PS exposure observed with reversible PS externalization can readily be counteracted by removal of the physiologic stressor (nitric oxide, p53 activation, allergic mediators, or growth factor deprivation). If the stress remains uncorrected, the cell may undergo apoptosis. The ability to identify cells with low, potentially reversible levels of PS exposure may explain the uptake of PS-ligands outside regions of apoptosis (as seen histologically by TUNEL staining), for example, in patients with hypoxic-ischemic reperfusion injury of the heart96,97 or brain.98–100 In fact, even in regions with apoptosis/ischemic injury, more annexin V-positive cells after the administration of radiolabeled annexin V as compared to apoptotic nuclei at TUNEL staining have been demonstrated.97 These observations suggest that annexin V uptake after ischemic reperfusion injury may be also because of large numbers of stressed cells (not necessarily committed to apoptosis) with relatively low levels of PS expression in contrast to the relatively fewer cells with high levels of PS exposure that are irreversibly committed to apoptosis. The ability of radiolabeled annexin V to bind to “stressed” cells with relatively low levels of PS exposure also implies that annexin V imaging may be extremely sensitive and can thus be used to identify tissues or organs at risk for irreversible injury, such as seen in hypoxic-ischemic injury101 or chronic heart failure,102 or at sites of active disease that can be seen in infection,103,104 unstable atherosclerotic plaques,105allograft rejection,106 or autoimmune disorders.107 Therefore, imaging based on radiolabeled annexin V could be useful for the serial assessment of acute and chronic disorders of organs or tissues at risk for permanent damage in which prompt treatment may prevent irreversible cell injury and death.
Most chemotherapy agents cause tumor cell death by inducing apoptosis, whereas resistance to anticancer treatment is believed to involve mutations that lead to deregulated cell proliferation and suppression of the mechanisms that control apoptosis.108 Defects in the apoptotic pathways are also responsible for resistance to therapy, and new therapeutic approaches attempting to reactivate these pathways bypassing the block are currently being explored.109Analysis of timing, tissue localization, and extent of cell death induced by anticancer drugs is helpful for assessing the efficacy of therapy.110,111 In solid tumors, assessment of drug-induced apoptosis can be performed using biopsy material of paired tumor samples (before and after therapy) and several biopsy cores. However, levels of cell death observed in tissue sections are variable, both at baseline and following therapy. This variability is likely to reflect tumor heterogeneity and the presence in the tumor of different clones with different rates of spontaneous cell death and different sensitivities to the drug used. Pharmacokinetics of a chemotherapeutic drug as well as the inherent kinetics of cell death processes clearly also affect the results at any given time point. Imaging techniques are therefore potential alternatives to serial tumor biopsy, because they may integrate the overall response of a tumor and can therefore be less affected by tumor heterogeneity at the microscopic level. Modulating the apoptotic pathway offers special opportunities for targeted therapeutic interventions, as direct imaging of caspase activity or PS expression can be used for noninvasive monitoring of early drug responses to drugs.
Several approaches can be used to specifically stain cell suspensions and histologic specimens for detecting apoptosis. The most widely used are based on assays for terminal deoxynucleotidyltransferase (Tdt)-dUTP nick end labeling (TUNEL), originally introduced by Gavrieli et al. in 1992.6 TUNEL is based on the specific binding of TdT to the 3′-OH ends of fragmented DNA. After proteolytic treatment of histologic sections, TdT incorporates X-dUTP (X = biotin, DIG, or fluorescein) at sites of DNA breaks. Terminally modified nucleotide avidin peroxidase can then amplify the signal and allows the evaluation of labeled cells under light/fluorescent microscopy, flow cytometry, or via immunohistochemistry. Another DNA-based method is the detection of the internucleosomal fragmentation produced by endonucleases at expected intervals of 180 to 200 bp.112 Standard DNA extraction techniques are used to obtain the nucleic acids from either cells or homogenized tissue. DNA is then submitted to electrophoresis in an agarose gel to demonstrate the characteristic DNA ladder pattern.
Because PS externalization represents the hallmark of apoptosis, a variety of PS-binding compounds have been extensively investigated for imaging, including proteins,113 peptides,114–117 and small chemical entities.118 Peptides and small chemical compounds have the advantage of being quickly and efficiently cleared from the blood circulation. Therefore, in principle they exhibit optimal properties as imaging probes, because their signal-to-background ratio is favorably affected by this kinetic profile. However, the main disadvantage of these compounds is their low affinity for PS. On the other hand, proteins exhibit higher affinities for PS, but they are usually cleared from the blood circulation with slower kinetics because of their higher molecular weight.
FIGURE 36.5. Three-dimensional structure of annexin V. (From Extracellular Matrix Proteins, http://www.bbm1.ucm.es/public_html/res/prot/extmat.html; accessed January 2, 2013.)
Annexin V, a 36-kDa vesicle-associated protein, is perhaps the most widely investigated PS-targeting moiety. It is a nonglycosylated single chain protein that belongs to the annexin supergene family. Its polypeptide is organized in an N-terminal tail with a C-terminal core containing four domains that form the annexin core, a slightly bent surface with a convex shape that interacts with the PS-containing phospholipid membrane (Fig. 36.5).119,120 Annexin V binds to PS in a Ca2+-dependent manner, with nanomolar affinity. Ca2+ ions bind to the annexin core surface at type II Ca2+-binding sites121 and form the prime contact by coordinating carbonyl and carboxyl groups of the protein and phosphoryl moieties of the glycerol backbone of membrane phospholipids (Fig. 36.6).120 The domains are composed mainly of α-helices and the Ca2+-binding sites protrude as loops. The overall binding affinity for PS arises from collaboration among the Ca2+-binding sites of the four domains, with a dominant role for domain 1.122 In solution annexin V is a monomer, but once bound to PS-expressing membrane three monomers build a trimer by protein–protein interaction; in turn, trimers assemble in a two-dimensional lattice covering the PS-expressing surface by trimer-trimer interactions (Fig. 36.7).123 Therefore, such two-dimensional protein network of annexin V at a PS-expressing cell surface determined internalization of annexin V124 by a unique pathway of pinocytosis. This pathway is initiated by disassembly of the cortical actin network underlying the PS-exposing membrane patch. Annexin V then binds to PS and crystallizes on the cell surface as closely packed trimers that cause the underlying membrane to bend inward. The invaginated membrane patch then closes on itself and is transported into the cytosol in a microtubule-dependent manner. This pathway does not seem to be related to clathrin- or caveolin-mediated endocytosis, as it is neither actin-driven nor preceded by membrane ruffling.125
Recombinant human (rh) annexin V exhibits PS-binding properties identical to those of annexin V purified from human tissue.126 Availability of recombinant annexin V spurred synthesis of a wide range of labeled forms of annexin V to image PS expression using optical, radionuclide, and magnetic resonance imaging.127 Initially, annexin V was coupled to fluorescent dye molecules and used as an apoptosis detection reagent for fluorescence microscopy and flow cytometry.14 Subsequently, annexin V was coupled to a radionuclide (99mTc) or iron oxide nanoparticles128 to detect apoptosis noninvasively in animals129 and in patients130,131 using radionuclide imaging techniques or magnetic resonance, respectively.
FIGURE 36.6. Binding of annexin V to phosphatidylserine (PS) expressed on the outer layer of cell membrane. Binding to PS occurs only after formation of a trimer structure of annexin V. (Reproduced from “Annexins: Molecular Structure to Cellular Function.” Barbara AS, ed. Austin, TX: Landes Bioscience, 1996, with permission.)
Photonic imaging methods have also been used to image annexin V labeled with fluorochromes132 or near-infrared (NIR) fluorochromes.133 Furthermore, Schellenberger et al.134 described labeling of annexin V with superparamagnetic iron oxide (SPIO) nanoparticles, for MR detection. Gd-containing annexin V-coated liposomes for positive or bimodal MR contrast have also been developed135,136 as well as multimodal contrast agents for combined MRI and fluorescent imaging.136
Most preclinical noninvasive imaging of PS expression has been carried out with radionuclide and magnetic resonance imaging. In fact, low tissue penetration of photons and autofluorescence of extracellular matrix components have so far hampered development of noninvasive optical imaging of PS expression. Recently, near-infrared fluorescent (NIRF) probes and fluorescence-mediated tomography (FMT) have been developed, thus making noninvasive optical imaging feasible.137 Second-generation annexin A5 has been coupled to the NIRF probe Vivo-750 via thiol chemistry and employed successfully to quantify the anticancer effect of cytotoxic compounds in a mouse cancer model using noninvasive FMT.
Radionuclide Labeling of Annexin V
Radiolabeling of rh-annexin V with 99mTc-pertechnetate can be performed by the penthioate radioligand (N2S2) method.138 This method starts with the chelation of 99mTc in the presence of stannous gluconate to yield 99mTc-gluconate, which is then reacted with acidified penthioate ligand (tetrafluorophenyl 4,5-bis-(S-1-ethoxy-ethylmercaptoacetamido) pentanoate, TFP) under heating to form a stable 99mTc-N2S2 complex. The technetium diamide dimercaptide N2S2 ester complex is then randomly conjugated to the N-H groups of lysine of the protein at basic pH. However, the N2S2 labeling method is cumbersome, has low labeling efficiency (30% to 40%), and entails a high nonspecific excretion of radioactivity into the bowel via biliary excretion. For these reasons, an improved labeling method using the bifunctional agent hydrazinonicotinamide (HYNIC) was selected for clinical trials.139 In this procedure, HYNIC (succinimidyl [6-hydrazinopyridine-3-carboxylic acid]), also known as (succinimidyl [6-hydrazinonicotinic acid]), is used to randomly modify the accessible N-terminal groups in the lysine residues of rh-annexin V. The resultant compound can then be lyophilized and stored indefinitely for labeling with 99mTc. Labeling of reconstituted HYNIC-annexin V is performed simply by reacting the conjugate with 99mTc-pertechnetate in the presence of stannous tricine for 5 to 10 minutes at room temperature. Similarly as the penthioate radioligand (N2S2 method), 99mTc-HYNIC-annexin V shows the greatest accumulation in the kidneys, liver, and urinary bladder. However, 99mTc-HYNIC-annexin does not undergo any bowel excretion, thus resulting in excellent imaging profile for the abdominal region. Unfortunately, 99mTc-HYNIC-annexin V exhibits high accumulation in the renal cortex, thus limiting visualization of pararenal structures.138
FIGURE 36.7. Structural modifications of annexin V, enabling its binding to phosphatidylserine expressed on the outer layer of the cell membrane. Upper panels: Trimer of the annexin V-calcium complex represented in the solid state from the top (upper left) and from the side (upper right) of the three-fold axis. The protein backbone is in blue, calcium ions are in magenta, and sulfate ions are rendered as balls and sticks. Subunits within the trimer are arranged so that the ion channels within each subunit are parallel to the three-fold axis. Lower panels: Annexin trimer in presence of zinc ions rather than calcium ions, represented in the solid state from the top (lower left) and from the side (lower right) of the three-fold axis. In this case the three subunits arrange to form a propeller-like structure, with more globular trimer–trimer arrangement, is more globular; orientation of the ion channels is not parallel, and thus cannot interact with the phospholipid bilayer simultaneously. (Source: http://www.chem.sc.edu/faculty/lebioda/projects/anxV.htm; accessed January 2, 2013).
However, because the N-H groups of lysine are also present on the surface of the annexin V core, radiolabeling may compromise its binding to PS.140 Therefore, to avoid reduction of binding affinity, annexin V variants have been generated for site-directed labeling at the concave side of the molecule using thiol chemistry. Thiol-linkage sites have been obtained either in extensions of the N-terminus141,142 or between the N-terminal tail and the concave side of annexin V, and successfully coupled with a number of compounds without adversely affecting its binding affinity for PS.128,143–146 These proteins have an endogenous site for 99mTc chelation consisting of six amino acid tags added at the N-terminus, followed by the 1 to 320 amino acid sequence of wild-type annexin V, with only the amino acid cys-316 mutated to serine. 99mTc chelation is thought to occur via formation of an N3S structure involving the N-terminal cysteine and the immediately adjacent amino acids. The purified protein is then reduced and stored for later labeling with 99mTc using glucoheptonate as an exchange reagent. Both the V-117 and V-128 variants have major advantages over HYNIC-annexin V, including a 50% to 75% lower renal accumulation and a markedly improved in vivo localization to sites of apoptosis in animal models.140 Using related annexin mutants, it has been found that all four calcium-binding sites are needed for full in vitro and in vivo binding of annexin V. Mutation (loss of function) of any one of the four calcium-binding sites decreases apoptosis-related in vivo localization of the tracer by 25%, and any two site mutations result in a 50% decline of uptake. The adverse effects of the random modification of annexin V have also been observed with 111In-DTPA-PEG-annexin V.147 Further work has also established that random modification of the lysine residues of annexin V with HYNIC, mercaptoacetyltriglycine (MAG3), or fluorescein isothiocyanate decreases by 50% nonspecific liver uptake of such form of annexin V compared with self-chelating (site-specific) protein, as also does conjugation with biotin. Despite its prolonged circulation time, the uptake of 111In-DTPA-PEG-annexin V in a mammary carcinoma was not increased relative to a nonspecific control protein pegylated in the same manner. Therefore, any specificity for PS in the tumor had been lost, probably because the annexin V was heavily modified on amino groups to attach enough PEG. Annexin V has also been proven to be quite heat-labile, as it loses most of its activity with heating at 56°C for 10 minutes148 (whereas being quite stable at 37°C), thus precluding the use of many different types of labeling chemistries.
SUMMARY OF THE MAIN STUDIES ON THE USE OF RADIOPHARMACEUTICALS TO EVALUATE APOPTOSIS EITHER IN VITRO OR IN VIVO
Several approaches to label annexin V with fluorine-18 (18F)149 for PET imaging have been developed. One method has used N-succinimidyl 4-fluorobenzoate to synthesize F-annexin V. The fluorine-labeled agent has lower uptake in the liver, spleen, and kidney than HYNIC-annexin V. Another method involves site-specific derivatization with an 18F-maleimide-labeled compound of mutant annexin V-117 or annexin V-128.142 However, both methods need more preclinical investigations before further development as imaging agents for apoptosis in patients.
Other radionuclide derivatives of annexin V have been developed, including annexin V-labeled 123I150 or 124I151 and 64Cu-labeled streptavidin for PET imaging after pretargeting of PS with biotinylated annexin V.152
Pilot Clinical Applications of Imaging Apoptosis with Radiolabeled Annexin V
Using SPECT imaging with 99mTc-rh-annexin V, Belhocine et al.130 investigated 15 patients with either primary or metastatic lung cancer lesions before and after chemotherapy. In this pilot study, a negative scan after therapy (i.e., no change in tumor uptake versus the pretreatment baseline scan) correlated well with a lack of treatment response in six of eight patients; the remaining two patients (with metastatic breast cancer) actually had a clinically significant response to Taxol-based chemotherapy despite no change in 99mTc-rh-annexin V uptake. All seven patients with increased tumor uptake versus baseline (i.e., positive annexin V study) had an objective response to chemotherapy (tumor shrinkage of tumor). Such increase in annexin V uptake was observed 40 to 48 hours after chemotherapy in five of these seven patients (1 NHL, 1 HL, 1 SCLC, and 2 NSCLC), whereas it was observed at 20 to 24 hours post treatment in the remaining two patients (1 NSCLC and 1 SCLC). These findings suggest some variability in the optimal timing for imaging with annexin V following antitumor treatment, thus emphasizing the need to define the best timing for administration of radiolabeled annexin V when designing imaging trials for assessing with this agent tumor response to therapy.153 Several studies in different animal models confirm a wide variability in the time course of radiolabeled annexin V uptake following a single administration of chemotherapeutic agents (Table 36.1).154–160
99mTc-HYNIC-annexin V is currently being investigated in Phase II/III trials,161 as several studies have confirmed the potential clinical utility of 99mTc-HYNIC-annexin V in determining the efficacy of chemotherapy.131,162–164
In this regard, even the rate of spontaneous apoptosis assessed by a pretreatment 99mTc-HYNIC annexin-V scan may be useful to predict subsequent response to chemo- and radiotherapy.165 In fact, although the baseline tumor uptake of 99mTc-HYNIC annexin V differs significantly from patient to patient, in general patients displaying tumor progression or stable disease exhibited a higher than background uptake in their tumor tissue, whereas patients responding to treatment display a comparable or lower than the background uptake in their tumor.
Kartachova et al.166 found that the degree of tumor response to platinum-based chemotherapy in NSCLC patients correlated with the percentage increase in tumor uptake of 99mTc-HYNIC-annexin V versus baseline evaluated 48 hours after the first injection of cisplatin. Patients achieving only disease stabilization exhibited a slightly increased, unchanged, or even a slightly decreased annexin V tumor uptake, whereas in patients with progressive disease a marked decrease of annexin V tumor uptake was observed. The same group evaluated patients with primary untreated head and neck squamous cell carcinoma with 99mTc-HYNIC annexin V SPECT before treatment.167 On univariate as well as multivariate analysis, only the 99mTc-HYNIC-annexin V tumor-to-background ratio of the primary tumor distinguished using a cutoff value of 2 was predictive of recurrence-free survival and of overall survival. When lymph node status was considered (N0 versus N1-N2-N3 disease) 99mTc-HYNIC-annexin V tumor-to-background ratios were included in the multivariate model, both N status and the 99mTc-HYNIC-annexin V tumor-to-background ratios of the primary tumor showed an independent association with overall survival.
Treatment-induced changes of 99mTc-HYNIC-rh-annexin V uptake in patients with head and neck squamous cell carcinoma evaluated before and within 48 hours after the first course of cisplatin-based chemoradiation (with a radiation dose to the tumor of 6 to 8 Gy) was determined in primary tumor, lymph node metastases, and normal tissue included in the irradiation field. Patients with primary tumors with a high 99mTc-HYNIC-rh-annexin V uptake also had high 99mTc-HYNIC-rh-annexin V uptake in lymph node metastases. Treatment-induced increment in uptake of 99mTc-HYNIC-rh-annexin V in primary tumors and in lymph node metastases showed large interpatient differences. A high correlation was observed on a patient-to-patient basis between the maximum δ-uptake induced by treatment both in the primary tumor and in the lymph node metastases. However, in this small series of patients the treatment-induced increment in 99mTc-HYNIC-rh-annexin V uptake in primary tumor did not predict outcome: No correlation between 99mTc-HYNIC-rh-annexin V uptake and early response was observed. In addition, 99mTc-HYNIC-rh-annexin V uptake also did not predict the locoregional control rates within the first 2 years of follow-up, possibly because of advanced stages of head and neck squamous cell carcinoma with large and necrotic tumors. Treatment-induced 99mTc-HYNIC-rh-annexin V uptake in relation to radiation dose in the parotid glands was observed, indicating early treatment-related apoptosis in a normal tissue at mean radiation doses as low as 3 to 8 Gy and one course of cisplatin.168 The uptake of 99mTc-HYNIC-rh-annexin V in the parotid glands showed a radiation dose-response relationship: Glands that had received higher doses of radiation demonstrated increased 99mTc-HYNIC-rh-annexin V uptake. Already after a low dose (6 to 8 Gy), parotid glands may be affected, thus suggesting early start of loss of parotid gland function. This is in agreement with observations in experimental rodent and monkey models in which apoptosis was induced early, after doses of up to 5 Gy.169,170
It has also been recently reported that the uptake of annexin V in normal tissues, such as the spleen and bone marrow (that are susceptible to drug-induced injury), is not significantly changed following chemotherapy or prior administration of radiolabeled annexin V within a 48-hour period.171 Furthermore, the biodistribution of radiolabeled annexin V in the kidneys, liver, and whole body remained unchanged after chemotherapy or prior administration of the same tracer.
Kartachova et al.172 systematically explored how to best assess chemotherapy-induced increases in annexin V uptake. SPECT studies in 38 patients with lymphoma (n = 31), NSCLC (n = 4), and head and neck squamous cell carcinoma (n = 3) were analyzed for maximal counts per pixel in the tumor volume (Cmax) for every target lesion in addition to grading on a visual four-grade score (i.e., Cmax/expressed as percentages of baseline values: Grade-1, decrease >25%; grade 0, 1% to 25% decrease; grade +1, 1% to 25% increase; grade +2, >25% increase; visual analysis: 0 = absent, 1 = weak, 2 = moderate, 3 = intense). Both the quantitative and visual assessments of increments in annexin V uptake after treatment correlated well with therapeutic outcome as evaluated by RECIST criteria. Excellent intra- and interobserver reproducibility was found, thus suggesting that chemotherapy-induced increases in annexin V uptake evaluated by SPECT can be employed for the early noninvasive assessment of anticancer treatment efficacy (24 to 48 hours after initiation of therapy), weeks before actual tumor shrinkage can be detected by conventional imaging. Examples of clinical imaging in lymphoma patients exhibiting different responses to radiation therapy are shown in Figures 36.8–36.10, obtained when imaging apoptosis with 99mTc-HYNIC-annexin V.
Semiquantitative 99mTc-HYNIC-annexin V tumor uptake (expressed as percent fraction of injected activity) in visible tumor lesions divided by the tumor volume derived from SPECT imaging has been found to correlate linearly with the histologic score of Fas ligand expression, whereas no correlation was found with the histologic score of matrix metalloproteinase-9, as well as with the number of tumor-infiltrating lymphocytes (CD45 staining) or microvessel density.173
Annexin V imaging can be problematic for detecting modest responses to therapy, such as those often seen clinically in patients with solid tumors. In fact, several factors complicate imaging of tumor cell death, such as nonspecific tracer uptake in tumors because of enhanced permeability and retention (caused by leaky vessels and poor lymphatic drainage in tumors) that may mask small variations in specific annexin V uptake because of cell death.174 Thus, enhanced permeability and retention175 together with the effects of anticancer drugs on the vasculature might result in altered tracer accumulation. In fact, using a 11C-labeled Sel-tagged annexinV ([11C]-Anx5-ST)176,177 in a SCID mice implanted with human squamous carcinoma cells, a trend to reduced [11C]-AnxA5-ST uptake 72 hours after treatment with doxorubicin has been observed.178 In this regard, doxorubicin has been reported to affect endothelial cell function179 and the actual imaging readout for assessing the specific uptake of annexin V should therefore be estimated as the difference between [11C]-Anx5-ST uptake and uptake of a size-matched control ligand ([11C]-mTrx-GFP-ST) after treatment, rather than simply the uptake of [11C]-Anx5-ST post-doxorubicin versus baseline. Whether changes in passive, nonspecific tracer accumulation have influenced the adequacy of other studies with annexin V ligands to reveal therapy-induced cell death cannot be excluded.
FIGURE 36.8. Patient with non-Hodgkin lymphoma in the left side of the neck. Axial fused SPECT/CT images clearly show increase in 99mTc-annexin V uptake from weak prior to radiotherapy (A) to intense (B) early after starting the first course of radiotherapy. On baseline CT (C) and on CT obtained 4 weeks after radiotherapy (D) a complete response is seen (circle). (Image courtesy of Dr. Renato A. Valdés Olmos, Nuclear Medicine Division, Diagnostic Oncology, The Netherlands Cancer Institute–Antoni van Leeuwenhoek Hospital, Amsterdam, The Netherlands.)
The relationship between annexin V SPECT imaging and [18F]FDG uptake has not been systematically compared in clinical trials. Studies in tumor models, however, have demonstrated that an enhanced apoptotic reaction (increased uptake of radiolabeled annexin V) is correlated with decreased [18F]FDG uptake 48 hours after the start of cytotoxic chemotherapy.180 In a clinical trial of 45 patients with breast cancer receiving three administrations of neoadjuvant chemotherapy, a significant reduction in [18F]FDG uptake was associated with a marked increase in the fraction of TUNEL-positive (apoptotic) tumor cells.181
Because apoptosis is an energy-dependent process (at least initially), glucose demand may actually increase temporarily following effective antitumor treatment.182 One example of such occurrence can be the “metabolic flare” often observed on [18F]FDG PET imaging following hormonal therapy for estrogen receptor–positive human breast cancer.183
FIGURE 36.9. Patient with non-Hodgkin lymphoma in the left axilla. Axial fused SPECT/CT images show slight increase in 99mTc-annexin V uptake from moderate prior to radiotherapy (A) to moderately intense (B) early after starting the first course of radiotherapy. On baseline CT (C) and CT obtained 4 weeks after radiotherapy (D) a partial response was observed (circle). (Image courtesy of Dr. Renato A. Valdés Olmos, Nuclear Medicine Division, Diagnostic Oncology, The Netherlands Cancer Institute–Antoni van Leeuwenhoek Hospital, Amsterdam, The Netherlands.)
In conclusion, several issues limit clinical use of radiolabeled annexin V probes. The main concern is suboptimal pharmacokinetics of radiolabeled annexin V, resulting in high background activity in the abdominal region. Another concern is the partial inability of annexin V probes to distinguish apoptosis from necrosis, because PS is exposed also during necrosis because of disruption of plasma membrane integrity. Finally, the optimal time window for detecting treatment-induced apoptosis in cancer patients has yet to be defined.184 In this regard, it should be emphasized that evaluating the time course of 99mTc-annexin V uptake is critical for any given clinical condition, because treatment-induced apoptosis may differ from tumor type to tumor type and possibly also from one chemotherapy agent to another agent. The clinical setting is also quite different from the experimental animal models, where the various parameters determining detection of treatment-induced apoptosis can be strictly controlled, including possible coadministration of different pro- or antiapoptotic stimuli.185
Imaging Apoptosis with Phosphatidylserine-Binding Peptides and Small Molecules
Compared to protein probes, peptides and small molecules have some favorable characteristics, including fast clearance from the circulation, efficient tissue penetration, and high target-to-nontarget ratios. Consequently, several peptide sequences have been screened for targeting PS exposure in apoptotic cells, using bacteriophage display technology. Optical imaging after the systemic administration of the fluorescein-labeled CLSYYPSYC peptide to tumor-bearing nude mice (H460-cell xenograft model) treated with a single dose of camptothecin indicated satisfactory homing of the peptide in the tumor.117
ApoSense molecules are small nonpeptidic fluorescent compounds developed on the basis of the γ-carboxyglutamic acid (Gla) structure. In response to apoptosis, these molecules exhibit selective membrane binding, transmembrane transport, and accumulation in the cytoplasm of apoptotic cells, being instead excluded from viable cells.186,187 Besides the fluorescence feature, some of these ApoSense molecules contain a fluorine atom, which facilitates radiolabeling with 18F for PET imaging. Recently, synthetic zinc(II)-dipicolylamine also showed some potential to image apoptotic and necrotic cells in vivo.188 However, further characterization of these apoptosis imaging probes is required to fully explore the potential of these low–molecular-weight alternatives to annexin V.
Besides annexin V, other proteins also bind to PS with high affinity. For example, the C2A domain (14.2 kDa) of another vesicle-associated protein, synaptotagmin I, binds with nanomolar affinity to negatively charged phospholipids in membranes, including PS, in a calcium-dependent manner.
Molecular Imaging of PS with Synaptotagmin I
As a whole molecule, synaptotagmin I is not suitable for imaging because of its transmembrane domain. Using recombinant technology, the soluble PS-binding C2A domain was expressed by Escherichia colias a fusion protein with Gluthation-S-transferase (GST). Although the affinity for binding PS is higher for C2A (Kd = 20 to 40 nM) than for the fusion protein C2A-GST (Kd ≅ 115 nM), C2A-GST was developed as a ligand because labeling of C2A reduces its PS-binding affinity.189 Labeling of GST-C2A likely occurred predominantly at the GST moiety. C2A-GST conjugated with fluorochromes, radionuclides, and superparamagnetic iron oxide particles using random chemical linkage retains its PS-binding properties.190,191 Whether site-directed chemical linkage will yield a superior PS imaging ligand has not been clarified so far.
FIGURE 36.10. Patient with non-Hodgkin lymphoma in the left groin. Axial fused SPECT/CT images show no changes in 99mTc-annexin V uptake prior to radiotherapy (A) and early after starting the first course of radiotherapy (B). On baseline CT (C) and CT obtained 4 weeks after radiotherapy (D) no response was observed (circle). (Image courtesy of Dr. Renato A. Valdés Olmos, Nuclear Medicine Division, Diagnostic Oncology, The Netherlands Cancer Institute–Antoni van Leeuwenhoek Hospital, Amsterdam, The Netherlands.)
On the other hand, C2A has been labeled with 99mTc for SPECT imaging of apoptosis in NSCLC patients treated with paclitaxel, showing significantly increased tumor uptake post therapy.192 99mTc-C2A-GST has also been employed in a reperfused acute myocardial infarction rat model. Ex vivo and in vivo data indicate that both specific binding and passive leakage contribute to the accumulation of the radiotracer in the area at risk.191 Finally, a recent in vitro study suggested that the C2A derivative binds more specifically to apoptotic and necrotic cells than does annexin V.193
Molecular Imaging of PS with Lactadherin
PS imaging with lactadherin purified from bovine milk has so far been limited to in vitro studies only.194 In particular, lactadherin has been coupled with fluorescein isothiocyanate via random chemical linkage.195 In principle, lactadherin has several advantages as a PS imaging agent over annexin V and synaptotagmin I, as its binding to membranes is directly proportional to PS content and independent from both phosphatidylethanolamine and Ca2+.195The drawback of lactadherin is its posttranslational modification, which precludes expression of functional lactadherin by recombinant technology in an E. colisystem.
Caspase peptide substrates containing either a radionuclide (e.g., an 18F-labeled caspase-inhibiting analog196), or a bioluminescence label197 or a far-red198 or near-infrared optical fluorochrome199 have been developed to detect apoptosis. Using firefly luciferase-based bioluminescence imaging in nude mice, Laxman et al.197 developed a caspase-cleavable reporter probe able to detect tumor apoptosis following chemotherapy. Although not relevant clinically, optical imaging techniques may become particularly useful in the screening of targeted drugs in preclinical studies.
By high-throughput screening and further structural optimization, several isatin (1H-indole-2,3-dione) sulfonamide analogs were identified to have nanomolar potency in inhibiting the executioner caspases, caspase-3, and caspase-7.200 For example, small-animal PET using 1-[4-(2-18F-fluoroethoxy)-benzyl]-5-(2-phenoxymethyl-pyrrolidine-1-sulfonyl)-1H-indole-2,3-dione (18F-WC-II-89) revealed high uptake of the radiotracer in the liver of a cycloheximide-treated rat, relative to the untreated control.196 Because caspases are intracellular targets, the radiolabeled imaging probes are usually lipophilic or contain a cell-penetrating moiety to achieve target access, which dampens the enthusiasm for these probes.201 Another problem for caspase-based imaging is that caspase inhibitors usually lack selectivity, as they are also effective inhibitors of various cathepsins. Recently, Bogyo’s group identified irreversible inhibitors and active site probes of the caspases (activity-based probes) that showed both broad and narrow selectivity within this family of proteases. Further optimization identified sequences with lower legumain reactivity and complete lack of reactivity toward the cathepsins.202 Optical imaging probes have been developed by conjugating such activity-based probes with near-infrared fluorescent tags and with a cell-permeable peptide sequence.
Activatable Probes for Caspases
The high background activity of caspase imaging with directly labeled probes can be overcome by activatable probes, which typically consist of three functional components. For example, a caspase-activatable probe, TcapQ(647), was synthesized so to include a Tat-peptide–based permeation peptide sequence, an effector caspase recognition sequence (Asp-Glu-Val-Asp, DEVD), and a flanking optically activatable pair comprising a far-red quencher (QSY 21) and a fluorophore (Alexa Fluor 647). Under baseline conditions, high quenching efficiencies resulting in low background fluorescence were observed. On exposure to executioner caspases, TcapQ(647) was specifically cleaved, thereby releasing the fluorophore from the quencher and enabling imaging of apoptosis. In vivo experiments demonstrated the ability of TcapQ(647) to detect parasite-induced apoptosis in human colon xenograft and liver abscess mouse models.203 Cell-permeable polymeric nanoparticles have also been prepared for apoptosis imaging. The close spatial proximity of the near-infrared fluorochromes in polymeric nanoparticles resulted in an autoquenched state and strong near-infrared fluorescence signal emitted in apoptotic cells.204 Theranostic agents that combine therapeutic components and a caspase-activatable imaging moiety have also been reported.205 Most activatable probes use the substrate DEVD to achieve caspase-3 cleavage after being internalized into lysosomes. Legumain is a lysosomal cysteine protease that plays a pivotal role in the endosomal/lysosomal degradation system. Cathepsins also become activated at the low pHs found in lysosomes. Thus, both cathepsins and legumain will digest DEVD and restore the optical signal even in nonapoptotic cells.202 Such cross-reaction may result in nonspecific activation of the activatable probes and thus increase the background activity of in vivo imaging. Key issues for future development of activatable probes include how to improve internalization and how to reduce noncaspase cleavage.
Reporter Gene Imaging of Caspase Activity
Bioluminescence imaging has been employed to image apoptosis with several seminal apoptosis-responsive reporter gene constructs.206,207 One reporter gene contains firefly luciferase gene flanked by ER (residues 281–599 of the modified mouse estrogen receptor sequence) with DEVD linker. A caspase-3–specific cleavage of the recombinant product results in the restoration of luciferase activity in cells undergoing apoptosis.207 Ray et al. developed a fusion protein construct by combining three different reporter proteins (red fluorescent protein, firefly luciferase, and HSV1-sr39 truncated thymidine kinase), linked through a caspase-3–recognizable polypeptide linker. Upon apoptosis induction with 8-μM staurosporine, the fusion protein showed a significant increase in firefly luciferase and red fluorescent protein activity in 293 T cells.208
A DEVD-containing cyclic luciferase to detect caspase activation has also been reported.209 Two fragments of DnaE intein were fused to neighboring ends of firefly luciferase connected with a DEVD sequence. After translation into a single polypeptide in living cells, the amino and carboxy terminals of luciferase were conjugated by protein splicing, which resulted in a closed circular polypeptide chain. When the substrate sequence was digested by caspases, luciferase was changed into an active form and restored its activity. Rather than modifying the reporter gene to respond to caspases, luciferase reporter gene substrate can also be modified to reflect caspase activity. One such example is a proluminescent, caspase-activated DEVD-aminoluciferin reagent (Caspase-Glo 3/7; Promega).210,211
Imaging Mitochondrial Membrane Potential
Another approach to detecting apoptotic cell death is to target the collapse of mitochondrial membrane potential (δψm), a hallmark of the initiating phase of apoptosis. Phosphonium cations are sufficiently lipophilic to permeate the membrane lipid bilayer and accumulate in cells as a function of the transmembrane voltage gradient. Because of the high mitochondrial membrane potential, most of the phosphonium cations accumulate within mitochondria. A PET agent, 18F-fluorobenzyl triphenylphosphonium, demonstrated good uptake in H345 cells. Selective collapse of δψm caused a substantial decrease in cellular uptake of 18F-fluorobenzyl triphenylphosphonium, compared with controls.212 In an orthotopic prostate tumor model, docetaxel caused a marked decrease (52.4%) of 18F-fluorobenzyl triphenylphosphonium tumor uptake within 48 hours, whereas [18F]FDG uptake was much less affected (12%).213Compared with the transient nature of PS externalization, loss of δψm is an ongoing process not limited to a time window, which may allow time-independent detection of apoptosis. However, “negative” contrast because of decreased accumulation of the imaging probes in apoptotic cells and cellular efflux mediated by the multidrug-resistance proteins may limit its clinical application.214
Despite almost two decades of intensive investigations, there is still no fully validated method for radionuclide imaging of apoptosis in humans. Although several radiopharmaceuticals have been proposed for this purpose, none of them have been licensed for human use. 99mTc-annexin V is perhaps the radiotracer that has been more extensively used in Phase II trials (for instance, in patients with NSCLC), but its validation as a prognostic imaging agent to predict response to antitumor treatment has not been completed. This delay in the clinical applications of an approach that holds such a high promise based on the results obtained both in vitro and in animal tumor models is caused by several factors; in particular, the lack of GMP-grade annexin V kits for clinical imaging trials after Theseus Imaging Corporation (Cambridge, MA) has closed its operation. It had limited clinical use of the tracer. This problem, together with the suboptimal biodistribution profile of 99mTc-HYNIC-annexin V, has stimulated several attempts to synthesize new radiolabeled moieties to image apoptosis and perform preclinical characterization. However, biologic characterization of the new molecules has not progressed sufficiently to pass the preclinical investigation phase. No new radiopharmaceutical has reached the clinical phase. Among all the radiolabeled agents, 99mTc-annexin V-128 is currently under development for human use. Expectation is high also for the 18F-labeled ligand of Caspase-3, [18F]fluoroethylazide (an isatin sulfonamide).
This scenario is further complicated by the complex clinical settings, where apoptosis imaging would be helpful. There is much wider variability in the clinical settings than the experimental settings, where most of the factors can be strictly controlled; for instance, to monitor the time course of the various events leading to apoptosis after exposure to antitumor treatment(s). In fact, it is now well established that each tumor exhibits great variability in the apoptotic response to each specific chemotherapeutic agent, or to the combination of such agents with the medications commonly used to reduce their side effects, particularly steroids. All of these effects have not been explored with reference to apoptosis imaging. Furthermore, no data are available on the effect of biologic agents on the pattern of apoptosis imaging agents, neither when used alone nor when used in combination with other antiproliferative drugs. Similarly, the effects of combined modalities on apoptosis (for instance, chemotherapy plus radiation therapy) are still to be explored.
A major crucial factor for the design of clinical trials aimed at imaging apoptosis as a prognostic indicator of tumor response to treatment is how to select the optimal timing to image the patient after treatment with the antitumor agent. In fact, the 24-hour time point proposed in the first clinical trial in NSCLC patients was arbitrarily chosen, despite the fact that several studies in animal models (e.g., at 3 hours) have shown that earlier time points might be preferable, at least following treatment with taxanes. Such choice, however, should be based on results of preclinical data in vitro and in vivo experimental models, adjusted for the pharmacokinetic profile in humans and on pilot investigations exploring the pattern of expression of the apoptosis-associated key molecules following the proapoptotic stimulus in patients.
An additional interesting issue is raised by the notion that PS expression on cell membranes following a proapoptotic injury does not always initiate an irreversible process that invariably leading to death. In fact, in some cases the PS-annexin V complex is internalized by the cell, thus indicating that the cell will survive the proapoptotic injury. This occurrence might open the road to a novel approach to additional therapy, for instance by conjugating annexin V with a cytotoxic agent (i.e., a β-emitting radionuclide or other toxins).
Based on all the evidence summarized in this chapter, the promise for efficient radionuclide imaging of apoptosis remains high. It is reasonable to expect that, as soon as GMP-grade radiopharmaceuticals will become available, full clinical validation of radionuclide imaging of apoptosis in oncology will be within reached.
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