Molecular effectors of multiple cell death pathways initiated by photodynamic therapy

Biochimica et Biophysica Acta 1776 (2007) 86 – 107 www.elsevier.com/locate/bbacan

Review

Molecular effectors of multiple cell death pathways initiated by photodynamic therapy Esther Buytaert, Michael Dewaele, Patrizia Agostinis ⁎ Department of Molecular and Cell Biology, Faculty of Medicine, Catholic University of Leuven, Campus Gasthuisberg, Herestraat 49, B-3000 Leuven Belgium Received 1 May 2007; received in revised form 27 June 2007; accepted 1 July 2007 Available online 6 July 2007

Abstract Photodynamic therapy (PDT) is a recently developed anticancer modality utilizing the generation of singlet oxygen and other reactive oxygen species, through visible light irradiation of a photosensitive dye accumulated in the cancerous tissue. Multiple signaling cascades are concomitantly activated in cancer cells exposed to the photodynamic stress and depending on the subcellular localization of the damaging ROS, these signals are transduced into adaptive or cell death responses. Recent evidence indicates that PDT can kill cancer cells directly by the efficient induction of apoptotic as well as non-apoptotic cell death pathways. The identification of the molecular effectors regulating the cross-talk between apoptosis and other major cell death subroutines (e.g. necrosis, autophagic cell death) is an area of intense research in cancer therapy. Signaling molecules modulating the induction of different cell death pathways can become useful targets to induce or increase photokilling in cancer cells harboring defects in apoptotic pathways, which is a crucial step in carcinogenesis and therapy resistance. This review highlights recent developments aimed at deciphering the molecular interplay between cell death pathways as well as their possible therapeutic exploitation in photosensitized cells. © 2007 Elsevier B.V. All rights reserved.

Contents 1. 2.

Photodynamic therapy . . . . . . . . . . . . . . . . . . . . . . 1.1. Direct photokilling of cancer cells . . . . . . . . . . . . Molecular mechanisms of programmed cell death. . . . . . . . 2.1. Type I PCD: apoptosis . . . . . . . . . . . . . . . . . . 2.1.1. Mitochondrial apoptosis in photosensitized cells 2.1.2. Death receptor-mediated signaling . . . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

. . . . . .

87 88 88 89 91 95

Abbreviations: 1O2, singlet oxygen; Δψm, mitochondrial transmembrane potential; [Ca2+]cyt, cytosolic Ca2+-concentration; AlPc, aluminum phthalocyanine; ALA, 5-aminolevulinic acid; AIF, apoptosis inducing factor; ANT, adenine nucleotide translocator; Apaf-1, apoptosis protease activating factor 1; ASK1, apoptosis signalregulating kinase-1; ATF, activating transcription factor; Atg, autophagy-related gene; BA, bongkrekic acid; BH, Bcl-2 homology; CHO, Chinese hamster ovary; CHOP, C/EBP homologous protein; CsA, cyclosporine A; CypD, Cyclophilin D; DISC, death inducing signaling complex; DKO, double knockout; DR, death receptor; EndoG, endonuclease G; ER, endoplasmic reticulum; ERAD, ER-associated degradation; FADD, Fas-associated death domain; FRET, fluorescence resonance energy transfer; GADD34, growth arrest and DNA-inducible gene 34; GPx4, glutathione peroxidase 4; HA14-1, ethyl 2-amino-6-bromo-4-(1-cyano-2ethoxy-2-oxoethyl)-4H-chromene-3-carboxylate; HP, hematoporphyrin; Hsp, heat shock protein; IAP, inhibitor of apoptosis; IMM, inner mitochondrial membrane; IMS, intermitochondrial membrane space; IRE1, inositol-requiring enzyme 1; JNK, c-Jun N-terminal kinase; LC3, microtubule-associated light chain 3; MEF, murine embryonal fibroblast; MMP, mitochondrial membrane permeabilization; NAO, N-nonyl acridine orange; NPe6, N-aspartyl chlorin e6; Omi/HrtA2, Omi/High temperature requirement protein A2; OMM, outer mitochondrial membrane; PARP-1, poly(ADP-ribose) polymerase-1; PBR, peripheral benzodiazepine receptor; Pc 4, silicon phthalocyanine 4; PCD, programmed cell death; PDT, photodynamic therapy; PERK, pancreatic ER kinase (PKR)-like ER kinase; PI3K, phosphatidylinositol 3 kinase; PLA2, phospholipase A2; PpIX, protoporphyrin IX; PPME, pyropheophorbide-a methylester; PP1, protein phosphatase-1; PTP, permeability transition pore; RIP1, receptor interacting protein 1; ROS, reactive oxygen species; SERCA, sarco-endoplasmic reticulum Ca2+ ATPase; Smac/DIABLO, second mitochondria-derived activator of caspases/direct IAP binding protein with low pI; SnET2, tin etiopurpurin; TMP, 4-5′-8-trimethylpsolaren; TNF, tumor necrosis factor; TPA, 12-O-tetradecanoyl-phorbol-13-acetate; TRAF2, TNF receptor associated factor 2; TRAIL, TNF-related apoptosis-inducing ligand; UPR, unfolded protein response; VDAC, voltage-dependent anion channel; XBP1, X-box protein 1 ⁎ Corresponding author. Tel.: +32 16 34 57 15; fax: +32 16 34 59 95. E-mail address: [email protected] (P. Agostinis). 0304-419X/$ - see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.bbcan.2007.07.001

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

2.1.3. The role of ER stress in apoptotic 2.1.4. Caspase-independent apoptosis . . 2.2. Type II PCD: autophagic cell death . . . . 2.2.1. PDT as inducer of autophagy . . 2.3. Type III PCD: necrosis . . . . . . . . . . 2.3.1. Programmed necrosis in PDT? . . 3. Conclusions. . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . .

cell death after PDT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Photodynamic therapy Photodynamic therapy (PDT) is a minimally invasive therapeutic modality approved for treatment of cancer diseases and non-oncological disorders. This approach is based on the local or systemic administration and the selective accumulation and/or retention of a photosensitizing agent (photosensitizer) in tumor tissue followed by irradiation with visible light of a wavelength matching the absorption spectrum of the photosensitizer (therapeutic window: 600–800 nm) (reviewed in [1–4]). Following the absorption of photons the photosensitizer transforms from its ground singlet state into an excited singlet state, which can decay back to its ground state by emitting fluorescence. This fluorescent property can be used for diagnostic purposes with those dyes displaying a preferential uptake in cancerous tissues [2]. Importantly, a fraction of the excited singlet state molecules is transformed via intersystem crossing into the relatively long-lived (micro- to milliseconds) excited triplet state, which can either form free radicals or radical ions by hydrogen atom extraction or electron transfer to biological substrates (such as membrane lipids), solvent molecules or oxygen [1,2]. These radicals can interact with ground-state molecular oxygen to produce superoxide anion radicals,

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

87

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . .

95 98 99 99 100 101 101 102 102

hydrogen peroxides and hydroxyl radicals (Type I reaction). Alternatively, from the excited triplet state the compound can transfer its energy to ground-state molecular oxygen to form the non-radical but highly reactive singlet oxygen (1O2) (Type II reaction), as shown in Fig. 1. Both reactions can occur simultaneously and the ratio between them depends on the photosensitizer and the nature of the substrate molecules [1–4]. However, direct and indirect evidence supports a prevalent role for 1O2 in the molecular processes initiated by PDT [5]. The extent of photodamage and cytotoxicity after PDT in vivo is multifactorial and can depend on the photosensitizing molecule, its localization at the time of irradiation, the total administered dose, the total light exposure dose, light fluence rate, the time between administration of the photosensitizer and irradiation, the type of tumor and its level of oxygenation [1,3,4,6]. These factors modulate the role of three independent processes that contribute to efficient PDT-induced tumor destruction: direct cancer cell death, destruction of tumor vasculature causing tumor ischemia and activation of an immune response. As discussed in recent reviews on this subject [3] and [7], it is thought that long-term tumor control results from a combination of these processes. PDT in vivo has been shown to reduce the

Fig. 1. Action mechanism of photosensitizing agents in PDT. When accumulated in tumor tissue, a photosensitizer in its singlet ground state (PTS) is excited to higher energy levels (PTS∗) upon irradiation and absorption of a photon of the appropriate wavelength. Deactivation occurs either through the emission of fluorescence or via intersystem crossing leading to formation of the excited triplet state (3PTS). Subsequently, energy can be lost in the presence of biological substrates and molecular oxygen (3O2), via Type I and Type II reactions leading to the formation of free radicals and ROS (O2 −, HO , H2O2, and 1O2). These cytotoxic species cause cellular photodamage which can activate a repair mechanism or lead to cell death when the damage is beyond repair.

U U

88

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Table 1 Preferential subcellular localization and molecular targets of the photosensitizers described in this review Photosensitizer

Abbreviation/trade name Chemical class

Subcellular localization

Molecular target

Aluminium phthalocyanine Silicon phthalocyanine

AlPc Pc4

phthalocyanine phthalocyanine

Zinc(II)phthalocyanine

ZnPc

phthalocyanine

Bcl-2 [95] Cardiolipin [80] Bcl-2, Bcl-XL [92–94,100,128] ND

Monocationic porphyrin Protoporphyrin IX

MCP PpIX, Levulan® (5-ALA, HAL) ATX-s10

Porphyrin Porphyrin

Mitochondria [95] ER, Mitochondria [70,80,92–94,100,128] Golgi apparatus, Plasma membrane [206] Plasma membrane, Cytosol [12] Mitochondria, Cytosol, Cytosolic membranes [67,69,76,82,83,125] Mitochondria, Lysosomes [116]

ND ANT [66] SERCA [145] Bcl-2 [92] ND Bcl-2 [13,92] Complex I ? [208] SERCA [16] ND Bcl-2, Bcl-XL [92,97,102]

13,17-bis (1-carboxypropionyl) carbamoylethyl-8-etheny-2-hydroxy -3-hydroxyiminoethylidene-2,7,12,18 -tetraethyl porphyrin sodium Porifimer sodium Photofrin®

Porphyrin-oligomer

Verteporfin Tin etiopurpurin N-aspartylchlorin e6 Meta-tetrahydroxyphenylchlorin

Chlorin Chlorin Chlorin Chlorin

Golgi apparatus, Plasma membrane [205] Mitochondria, ER [66,145] Mitochondria, Lysosomes [92] Lysosomes, Endosomes [115] ER, Mitochondria[13,92]

Chloromethyl-X-Rosamine Phenantroperylenequinone Chlorophyll-a derivative Porphycene

Mitochondria [71,208] ER, Lysosomes [16] ER, Lysosomes [85] ER, Mitochondria [92,97,102]

CMXRos Hypericin Pyropheophorbide-a methylester 9-capronyloxytetrakis (methoxyethyl)porphycene

BPD-MA, Visudyne™ SnET2 NPe6 mTHPC, Foscan®, temoporfin MitoTracker Red Hyp PPME CPO

Porphyrin

Casp3, Casp9 [12] PBR [67,69], Cardiolipin [83] p53 [125] Bcl-2 [116]

ND: not determined.

number of clonogenic tumor cells through direct ROS-mediated photodamage [8] and evidence for both apoptotic and necrotic regions has been found in tumor biopsies after PDT [4]. The direct cytotoxic effect of PDT, which is the subject of this review, is the result of the incorporation of the sensitizer mainly into cellular membranes and the subsequent light-induced generation of ROS causing irreparable damage. 1.1. Direct photokilling of cancer cells At the molecular level direct tumor cell destruction by PDT is caused by the irreversible photodamage to vital subcellular targets, which include the plasma membrane and intracellular membranes of the mitochondria, lysosomes, Golgi apparatus and endoplasmic reticulum (ER). Since most dyes do not accumulate in cell nuclei, PDT has generally a much lower potential of causing DNA damage, mutations and carcinogenesis as compared to that induced by X-radiation at equitoxic fluences/doses [6]. It is generally accepted that the intracellular localization of the sensitizer coincides with the primary site of photodamage. This is because the photogenerated singlet oxygen has a very short life and very limited diffusion in biological systems (half-life: b 0.04 μs, radius of action: b0.02 μm), indicating that primary molecular targets of the photodynamic process must reside within few nanometers from the dye [9]. Some studies have reported on the relocalization of certain photosensitizers after irradiation [10–13], suggesting that besides the primary site, photodamage can be rapidly propagated to other subcellular locations. The molecular nature of the photo-oxidized targets has profound influence on the signaling pathways and mode of cell

death initiated following PDT. Generally, photoactive compounds localizing to the mitochondria or the ER promote apoptosis, within a certain threshold of oxidative stress, while PDT with photosensitizers targeting either the plasma membrane or lysosomes, can either delay or block the apoptotic program predisposing the cells to necrosis [14] and reviewed in [6]. Table 1 illustrates the preferential subcellular localization and the identified molecular targets of the photosensitizer's subset discussed in more details in this review. Recent evidence, moreover, indicates that autophagy may be induced by PDT into the attempt to repair and survive photoinjury to key organelles and be turned into a cell death signal if this initial response fails [15–17]. Hence, lethal mechanisms initiated by the photosensitization process appear to encompass the three major morphologies of programmed cell death, e.g. apoptotic, necrotic and autophagic cell death. In this review we discuss the major molecular players involved in the signal transduction of these cell death pathways. 2. Molecular mechanisms of programmed cell death Programmed cell death (PCD) is defined as a genetically encoded form of suicide occurring in a predictable place and time during embryonic development (reviewed in [18]). Three morphologically distinguished forms of PCD have been characterized during development [19]. Type I PCD is characterized by phenotypic changes involving nuclear condensation and general cellular shrinkage. Type II PCD is distinguished by a lysosomal-dependent digestion of the cell and the presence of autophagic vacuoles (autophagosomes) while Type III PCD is marked by cellular swelling and a rapid loss of

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

plasma membrane integrity. Nowadays these processes are defined as apoptosis, autophagic cell death and necrosis, respectively, and are cellular programs known to play a crucial role in normal development, tissue homeostasis and in eliminating abnormal and damaged cells (reviewed in [20,21]). 2.1. Type I PCD: apoptosis Apoptotic cell death is unquestionably the best-studied form of PCD and is considered to have the most widespread physiological, pathological and therapeutic role [22]. Defective regulation or execution of apoptosis disrupts the balance between cell proliferation and cell death, triggering a spectrum of diseases including cancer [23,24]. Apoptosis is an ATP-requiring process morphologically characterized by chromatin condensation, cleavage of chromosomal DNA into internucleosomal fragments, cell shrinkage, membrane blebbing, formation of apoptotic bodies without plasma membrane breakdown, exposure of phosphatidylserine in the outer leaflet of the plasma membrane, and phagocytosis by neighboring cells. At the biochemical level, apoptosis entails the

89

activation of caspases, a highly conserved family of cysteinedependent aspartate-specific proteases, of which 11 members have been identified in humans [25]. All caspases are synthesized as pro-enzymes or zymogens and are activated in response to an apoptotic signal by proximity-induced dimerization at a multimeric protein complex (initiator caspases) or by limited proteolysis by an upstream caspase (effector caspases) [26]. It is generally accepted that once activated, the effector caspases are responsible for most of the stereotypic morphological and biochemical changes observed during apoptosis by cleaving a restricted subset of vital substrates [27]. The apoptotic caspases act in two main converging pathways, called extrinsic and intrinsic, in which initiator caspases-8/-10 and -9 directly activate the effector procaspases-3 and -7 (reviewed in [26] and illustrated in Fig. 2). The extrinsic pathway is triggered by binding of death ligands (e.g. FasL, TNF-α, TRAIL) to their cell surface death receptors (DR) (e.g. Fas, TNF-RI, TRAIL receptor). This induces DR clustering and formation of the death-inducing signaling complex (DISC), the oligomeric platform which recruits the initiator procaspases-8 and-10 and results in their dimerization-induced activation.

Fig. 2. The two main converging pathways leading to the activation of effector caspases. The binding of a death ligand, belonging to the TNF-α superfamily, to its cognate receptor induces the formation of the DISC, the molecular platform mediating the activation of the initiator procaspase-8 in the extrinsic pathway of apoptosis. Caspase-8 subsequently leads to the proteolytic activation of the main effector caspases-3/7. The intrinsic or mitochondrial pathway is initiated by the release of apoptogenic factors from the intermembrane space of the mitochondria into the cytosol (see text). The release of cytochrome c in the cytosol leads to the subsequent formation of the apoptosome, the heptameric complex triggering the activation of the initiator caspase-9, which in turn processes and activates the effector caspases-3/7. The activation of the effector caspases results in the morphological and biochemical features of apoptotic cell death, including internucleosomal DNA fragmentation, membrane blebbing and cell shrinkage. The release of AIF from mitochondria and its subsequent nuclear translocation leads to large-scale DNA fragmentation and contributes to caspase-independent apoptosis. The caspase-8 mediated cleavage of the cytosolic pro-apoptotic Bcl-2 family member Bid provides a molecular link between the extrinsic and intrinsic apoptotic pathways.

90

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Caspase-8/-10 in turn cleaves and activates the effector procaspases-3 and -7 leading to apoptosis. In the intrinsic pathway of apoptosis mitochondria occupy a central role. Although mitochondria are life-essential organelles for the production of metabolic energy in the form of ATP, numerous intrinsic and extrinsic cell death stimuli activating different, but often overlapping signaling pathways, converge on mitochondria to induce the permeabilization of mitochondrial membranes (MMP). MMP results in the release into the cytosol of several apoptogenic proteins stored in the inter-mitochondrial membrane space (IMS). This event has been proposed to be the point-of-no-return in many cell death programs, with the notable exception of sympathetic neurons, which have been shown to be rescued from death downstream of the cytochrome c release [28,29]. IMS proteins identified so far with a prominent role in apoptosis include activators of caspases such as cytochrome c, Omi/HtrA2 (Omi stress-regulated endoprotease/High temperature requirement protein A 2) and Smac/DIABLO (second mitochondria-derived activator of caspase/direct IAP binding protein with a low pI), as well as apoptosis-inducing factor (AIF) and endonuclease G (EndoG), which act in a caspase-independent fashion (reviewed in [30]). Cytosolic cytochrome c, by binding to Apaf-1 (apoptotic protease-activating factor 1) and in the presence of ATP or dATP, leads to the recruitment and activation of procaspase-9 through the formation of an heptameric complex called the apoptosome. Once free in the cytosol, Omi/HtrA2 and Smac/Diablo antagonize the activity of endogenous inhibitors of caspases (IAPs) thereby promoting caspase activation, while AIF and EndoG translocate to the nucleus where they mediate chromatin condensation and large-scale DNA fragmentation, independently from caspase signaling. In spite of numerous proposed models, which are probably not mutually exclusive and might occur sequentially or concurrently in certain apoptotic settings, the molecular mechanism underlying MMP is still elusive (for reviews see [31–34]). A first model involves the mitochondrial permeability transition (PT) which is a sudden increase of the inner mitochondrial membrane (IMM) permeability to solutes with a molecular mass up to 1.5 kDa, allowing solutes and ions to enter the matrix. This process is ascribed to the opening of a high-conductance channel located in the IMM which is designated as the PT pore (PTP), which is strongly favored by mitochondrial Ca2+ uptake and by the exposure of mitochondria to damaging ROS. In this model a long lasting PTP opening results in the dissipation of the H+ gradient over the IMM with consequent loss of the mitochondrial transmembrane potential (Δψm) and osmotic matrix swelling which can lead to rupture of the outer mitochondrial membrane (OMM), possibly because the OMM has a smaller surface area compared to that of the IMM, resulting in the release of apoptogenic factors from the mitochondria. Moreover, progressive PTP opening and mitochondrial uncoupling, would lead to a drastic drop in the cytosolic ATP levels, and consequent loss of viability. It should be mentioned that while PTP opening always results in Δψm collapse, measuring Δψm drop does not indicate necessarily that IMM permeabilization has occurred, given that transient PTP opening can be triggered by diverse physiological stimuli unrelated to death signals.

Several molecules found in different mitochondrial compartments have been postulated to participate in the formation of a macromolecular complex generated at the contact sites of the mitochondrial membranes, which regulates the open and closed state of the PTP. For a detailed discussion on the role of PTP in cell death, its pathophysiological and therapeutic implications the readers are referred to recent exhaustive reviews on this subject [35] and [36]. The OMM voltage-dependent anion channel (VDAC), the IMM adenine-nucleotide translocator (ANT) and the matrix chaperone cyclophilin D (CypD) have been suggested to be central components of the PTP and the inhibition of cytochrome c release by the CypD-binding drug cyclosporin A (CsA) has been used in support of this model. However, the role of VDAC, ANT and CypD in PTP formation has recently been challenged as mitochondria derived from mice deficient in either VDAC1, ANT (both isoforms) or CypD still undergo PTP opening [37– 42] and the derived deficient cells can still die by apoptosis. This suggests that these proteins are not essential components of the PTP but likely fulfill a regulatory function in PTP opening [35]. Since CypD-deficient cells show resistance to necrotic cell death induced by ROS and Ca2+ overload and CypD-deficient mice display a high level of resistance to ischaemia/reperfusioninduced cardiac injury [38,41,42], it has been suggested that CypD plays a crucial role in necrotic cell death. In addition, the existence of a Ca2+-independent, CsAinsensitive unregulated PTP formed by cross-linking and misfolding of proteins at the mitochondrial membranes has also been proposed [43]. A second mechanism for MMP entails the formation of proteinaceous channels in the OMM through a process involving oligomerization of pro-apoptotic Bcl-2 members or interaction between Bcl-2 proteins and VDAC [44]. The Bcl-2 protein family comprises at least 20 members containing one of four Bcl-2 homology domains (BH1 to BH4). Anti-apoptotic Bcl-2 proteins (e.g. Bcl-2, Bcl-XL, Bcl-w) contain all four BH domains and reside at the cytoplasmic side of cellular membranes of mitochondria, ER and nucleus; proapoptotic Bcl-2 proteins can be subdivided into the “BH123 multidomain” (e.g. Bax, Bak) and “BH3-only” proteins (e.g. Bid, Bad) [23]. In healthy cells Bax is found as a cytosolic monomer that, upon apoptotic stimuli, changes conformation and integrates in the OMM where it oligomerizes [45], whereas Bak is associated with the OMM [45–47]. Although there is no doubt that anti-apoptotic and pro-apoptotic members of this family thwart each others function, the molecular mechanisms underlying pro-apoptotic Bcl-2 proteins action appears to be different. BH123 multidomain Bax and Bak proteins once activated undergo a conformational change (taking place in the cytosol for Bax, or at the mitochondrial membrane for Bak) that triggers their oligomerization/activation and the formation of supramolecular pores mediating MMP. The exact mechanism resulting in Bax and Bak activation remains still elusive, but it is thought be inhibited by the anti-apoptotic Bcl-2 members likely acting at the mitochondrial membranes [47,48]. When activated by different signaling pathways pro-apoptotic “BH3-only” proteins (e.g. Bad, Bim, Puma, Noxa) act predominantly, if not

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

solely, by binding to and neutralizing the activity of the antiapoptotic Bcl-2 proteins [49], thereby unleashing Bax and Bak from their negative control. Bax and Bak oligomers have been shown to be essential for MMP in several paradigms of apoptosis, given that fibroblasts deficient in both proteins (e.g. Bax/Bak double knockout) are particularly refractory to undergo MMP in response of a variety of death signals [50,51]. Molecular cross-talk between the extrinsic and intrinsic pathway is provided by the caspase-8-mediated cleavage of the cytosolic BH3-only protein Bid as truncated Bid (tBid) translocates to the mitochondria and facilitates the release of cytochrome c into the cytosol [52]. Although the mechanisms by which tBid promotes MMP are not fully elucidated, the ability of tBid to bind to the specific IMM phospholipid cardiolipin [53] and to insert into selected OMM lysolipids [54,55] has been suggested to improve interaction with Bax and Bak proteins. This argues that the ability of pro-apoptotic Bcl-2 proteins to induce MMP and promote the efflux of IMS proteins from mitochondria may require the coordinated interaction of lipids and proteins at the mitochondrial membranes [48]. Also, mitochondria cristae remodeling and mitochondria fragmentation observed in cells undergoing apoptosis [56,57] is likely to be regulated by changes in the lipid composition of mitochondrial membranes [58]. Mitochondria fragmentation or fission (see for a review [59]) has been proposed as an additional mechanism causative for MMP. However, this view has been recently challenged based on the observation that preventing the fragmentation of the mitochondrial network in HeLa and COS cells by inhibiting the mitochondrial fission machinery, results in a partial inhibition of the release of cytochrome c from the mitochondria but does not affect Bax/Bak-dependent apoptosis [60]. 2.1.1. Mitochondrial apoptosis in photosensitized cells The critical role of the mitochondrial pathway of caspase activation following PDT has been largely documented (reviewed in [6,61–63]). Typically following PDT, the release of mitochondrial cytochrome c into the cytosol is followed by the apoptosome-mediated caspase activation cascade, leading to the apoptotic morphotype. As observed in other apoptotic paradigms, during PDT-mediated apoptosis Δψm progressively collapses, intracellular ATP levels steadily drop and damage to mitochondria becomes irreversible ensuring cell death. This is especially true for those photosensitizing agents with a prevalent mitochondrial localization, including porphyrogenic sensitizers and phthalocyanine-related compounds, which upon irradiation swiftly mediate MMP [6,62]. Mitochondria are also critical executers of lethal pathways emanating from photodamage to other subcellular sites or organelles, although in this case the release of apoptogenic proteins from mitochondria is delayed. How mitochondrial membranes are permeabilized in response to PDT and whether cytochrome c release involves opening of the PTP or rather the formation of a specific channel is not completely understood and it is possible that these two mechanisms do not occur in a mutually exclusive fashion in

91

photosensitized cells. Moreover, the prevalent mechanism for MMP induced by photoactive compounds associated almost exclusively with, or produced by, the mitochondria or by dyes displaying a multiple subcellular distribution may be distinct and regulated by the activation of upstream signaling pathways utilizing mitochondria as ultimate death executers. The main molecular mechanism by which PDT-treated cells initiate MMP, e.g. whether it involves IMM or OMM permeabilization, is not merely a fundamental question but an issue of strategic therapeutic implication in PDT. It should be mentioned, however, that the exact mechanism of mitochondria breakdown during cell death is still intensively debated as discussed in recent reviews [35,36,64], thus far beyond the PDT paradigm of apoptosis induction. Intracellular Ca2+ overload, with consequent mitochondrial 2+ Ca -uptake, the increase in the cellular pro-oxidant state and the generation of free fatty acids, such as those produced by phospholipase A2 (PLA2), are known factors favoring PTP opening [35]. Since these apoptogenic signals are commonly produced by PDT with a variety of photoactive molecules [6,61– 63] and likely synergize each others cytotoxic effects, a causative role for PTP opening as a main molecular mechanism in photokilling appears obvious. The effect of the photosensitization process on mitochondria has been addressed in different studies using isolated mitochondria as well as intact cells. The major observations in support of a role of PTP in mitochondrial apoptosis entail (1) the extreme sensitivity of presumed PTP components to photo-oxidation [65,66]; (2) the affinity of certain photosensitizing agents for proteins of the mitochondrial membranes with an established or suggested role in the regulation of PT [66–69]; (3) the inhibition of cytochrome c release and photo-cytotoxicity by pharmacological PTP inhibitors (e.g. bongrekic acid, CsA, PK11195) [67– 70]; (4) measurements of mitochondrial-entrapped calcein showing occurrence of IMM permeabilization [70–72]. The extreme sensitivity of the ANT to thiol group oxidation at sites critical for nucleotide transport renders this IMM protein a putative target of the photogenerated 1O2, a ROS highly reactive with thiol groups [73]. Consistent with this hypothesis, light activation of the sensitizer verteporfin (e.g. benzoporphyrin derivative monoacid ring A) using isolated mitochondria caused a ROS-mediated and Bcl-2-inhibitable rapid Δψm loss, which was suppressed by the ANT-specific inhibitor bongrekic acid (BA) as well as by CsA [66]. Analysis of the effects of verteporfin-PDT upon reconstitution of purified ANT in proteoliposomes indicated that this photosensitizing agent caused permeabilization of proteoliposomes through a DTT- and catalase-inhibitable oxidation of the ANT, while it did not affect other proteins or lipids, such as the ROS-sensitive phospholipid cardiolipin [66]. Verteporfin-mediated photo-cytotoxicity was suppressed by Bcl-2, or by the ANT ligands ATP and ADP which inhibit PT in isolated mitochondria and artificial membranes [74], suggesting that PTP opening triggered by ANT oxidation is apoptogenic in this paradigm. Fluorescence studies using rat liver mitochondria irradiated either with hematoporphyrin (HP) or with the psoralenderivative 4,5′,8-trimethylpsolaren (TMP), indicate that while

92

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

both dyes photogenerate 1O2, HP inhibits PTP opening, likely by the photo-oxidation of critical histidine(s) of the pore functional domains, while TMP promotes it [75]. These studies establish that the specific mitochondrial microenvironment where 1O2 is photogenerated can have dramatically different effects on PTP opening [75]. However, conclusions on the molecular targets and effects of PDT based on experiments using isolated mitochondria should be confirmed in intact cells where the photosensitizing effects, either direct or indirect, on MMP can be kinetically coupled to apoptotic parameters such as cytochrome c release and caspase signaling, given that PTP opening can modulate different cell death modalities [36]. As additional caveat, incubation of isolated mitochondria with lipophilic photosensitizing agents would force their partitioning into mitochondrial membranes, which may not be the preferred intracellular locations exhibited by the dyes in intact cells. The high affinity of certain mitochondria-associated dyes for putative components of the PTP has been proposed to be mechanistically involved in causing intrinsic apoptosis following their light activation in cultured cells. The photosensitizer protoporphyrin IX (PpIX), which is synthesized in the mitochondrial matrix from its heme precursor 5-aminolevulinic acid (ALA) and thought to distribute upon formation within the cytosol and cytosolic membranes [76], is a well-known high affinity ligand of the peripheral benzodiazepine receptor (PBR). The PBR, a highly hydrophobic protein located in the OMM, has been suggested as a putative component of the PTP [64]. In different PDT paradigms utilizing PpIX as the photosensitizing agent, the PBR-specific ligand isoquinoline carboxiamide PK11195 proved efficient in reversing both photo-cytotoxicity and PBR binding affinity, thus establishing a correlation between these events in PDT-induced cell death [67–69]. The calcein–cobalt technique is a trustable imaging method to investigate the occurrence of IMM permeabilization in intact cells based on the cytosolic quenching of the fluorescence of calcein accumulated in mitochondria by the mitochondriaimpermeable Co2+ upon PT [77]. Using this technique, PDT of A431 cells with the silicon phthalocyanine 4 (Pc 4), a dye located at mitochondria and ER membranes, resulted in a 1O2mediated and CsA-inhibitable PTP opening followed by mitochondrial depolarization and cytochrome c release [70]. However, using mouse lymphoma LY-R cells Pc 4-PDT mediated the release of cytochrome c with light doses that were not sufficient to cause an immediate measurable change in transmembrane potential [78]. This suggests that the MMP mechanism utilized by the same dye can depend on the cellular context and probably reflects a differential distribution of the photosensitizer in intracellular membranes. Exposure of human osteosarcoma cells to laser irradiation by confocal laser scanning microscopy with Chloromethyl-XRosamine (CMXRos or MitoTracker Red), a dye generally used to image mitochondria in a variety of studies, resulted in IMM permeabilization and mitochondrial apoptosis by a mechanism which could not be inhibited by CsA [71]. In a further study by the same group, laser scanner beam irradiation of CMXRos within a subset of mitochondria of an individual

cell, was shown to induce two temporally distinguished signaling phases between irradiated and not irradiated mitochondria. Irradiated mitochondria underwent structural changes from a filamentous to a fragmented structure and lost their cytochrome c without major alteration in Δψm, while their non-irradiated counterparts underwent Ca2+-dependent but CsA- and caspaseinsensitive Δψm loss and only subsequently released cytochrome c [79]. These studies reveal the occurrence of interorganellar signaling and suggest that Ca2+-waves may be used to propagate a CsA-insensitive IMM permeabilization within mitochondria upon photosensitization [79]. Fluorescence resonance energy transfer (FRET) between the cardiolipin specific probe 10-N-nonyl acridine orange (NAO) and Pc 4 in the human prostate cancer PC-3 cells indicated that the dye resides near cardiolipin-containing sites, thus mainly on the IMM where this anionic phospholipid is prevalently, although not exclusively, found [80]. Although the consequence of the direct Pc 4-mediated cardiolipin photo-oxidation was not further explored, peroxidation of cardiolipin is known to disrupt the binding of cytochrome c to the IMM, thus facilitating the release of the soluble IMS pool of cytochrome c mediated by PT or by OMM permeabilization by pro-apoptotic Bcl-2 family proteins (reviewed in [81]). Further evidence in support of the role of 1O2-mediated photo-oxidation of unsaturated lipids is provided by the finding that PpIX-mediated cardiolipin photooxidation, cytochrome c release and apoptosis are inhibited by overexpressing the antioxidant enzyme GPx4, which efficiently detoxifies hydroperoxides from membranes, only when PpIX and GPx4 co-localize to the mitochondria [82,83]. ROS generation in mitochondria and subsequent lipid peroxidation can result in dramatic alterations of mitochondrial vital functions, such as Δψm maintenance, respiration and oxidative phosphorylation and mitochondrial Ca2+ buffering capacity, all factors which alone or in combination favor MMP (reviewed in [81]). With photosensitizing agents displaying a weaker association with mitochondrial membranes, such as the main ERlocalizing sensitizers hypericin, pyropheophorbide-a methylester (PPME) and meta-tetrahydroxyphenylchlorin (Foscan®), the essential role of PTP opening in MMP and apoptosis induction remains uncertain [16,72,84–86]. A photo-induced PT in hypericin-loaded murine 3T3 fibroblasts was detected by the calcein–cobalt quenching procedure [72]. However, in 3T3 cells, in rat/mouse T-cell hybridoma (PC60R1R2) and human Jurkat cells hypericin photo-induced Δψm loss was not inhibited by Bcl-2 overexpression, which delayed cytochrome c release and induction of apoptosis, nor by CsA or BA [72,84]. Similarly, no inhibitory effect of CsA on cytochrome c release and apoptosis was observed in PPME-photosensitized colon cancer cells [85]. Furthermore, albeit mitochondria isolated from murine embryonic fibroblasts (MEFs) doubly deficient for bax and bak can still undergo PT in vitro [87], pro-apoptotic Bax and Bak proteins are essential for MMP induction and apoptotic cell death following hypericin-mediated PDT [16]. These studies suggest that PT per se does not suffice to induce apoptosis, at least in certain PDT paradigms, and raise the possibility that photosensitized mitochondria can swell in a

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

CsA-insensitive and reversible manner without the induction of cell death [71,88]. In conclusion, although the induction of PT by PDT has been shown in several models, whether opening of a CsA-sensitive or -insensitive PTP follows, depends crucially on the localization of the dye and its primary molecular targets, and may be causative for apoptosis in certain scenarios but not in all instances. 2.1.1.1. Pro-apoptotic Bcl-2 proteins in photokilling. The view that pro-apoptotic Bcl-2 family members constitute a critical gateway to permeabilize OMM in photosensitized cells is emerging from recent studies (for a recent review, see [62]). Consistently, relocation of Bax from the cytosol to the mitochondria has been reported in different PDT paradigms to occur with kinetics matching the release of cytochrome c [16,89,90]. A notable exception is provided by smooth muscle cells photosensitized with verteporfin, where Bax translocation to the mitochondria has been detected secondary to the release of cytochrome c [91]. The requirement of the multidomain Bcl-2 family member Bax (and Bak) for the induction of apoptotic cell death has been unambiguously proven by studying PDT cellular responses in cancer cells expressing undetectable Bax levels or in bax−/−/ bak−/− (DKO) MEFs. The apoptotic response of Pc 4-mediated PDT was studied in human breast cancer cells MCF-7c3 treated with Bax antisense oligonucleotides to suppress Bax expression, and in the human prostate cancer DU-145 cells, which do not express Bax. In these photosensitized cells the canonical hallmarks of apoptosis, including the release of cytochrome c from mitochondria, caspase activation and nuclear fragmentation were completely inhibited, while restoration of Bax expression in DU-145 cells re-sensitized cells to apoptotic cell death [89]. Furthermore, the MEF model revealed that the re-expression of mitochondriatargeted Bax is both necessary and sufficient to fully restore caspase activation and apoptotic cell death following hypericinPDT in apoptosis-deficient Bax−/−/Bak−/− cells [16]. The molecular mechanisms promoting Bax (and Bak) conformational change and activation during apoptosis are not completely understood (see for a recent review [47,48]) and for PDT, in particular, remain speculative. Since in healthy cells Bax is found as an inactive monomer mainly in cytosol or loosely bound to the mitochondria and ER membranes, the photo-generation of ROS at different subcellular sites must convey the molecular signal required for Bax translocation to and insertion in the OMM through the activation of specific, but ultimately converging, signaling pathways. A signal for Bax-mediated MMP may rely on the dramatic and specific photo-induced oxidation of the anti-apoptotic Bcl2 protein that results in its loss-of-function, by photosensitizing agents residing in membranes in its close vicinity [92–94]. Presumably this event lowers the Bcl-2 mitochondrial barrier function and acts as a permissive signal for Bax/Bak-mediated MMP resulting in rapid apoptosome-mediated caspase activation. In some cellular systems, such as human breast epithelial cell line MCF10A and in the human epidermoid carcinoma A431 cells, stable overexpression of Bcl-2 has been shown to

93

upregulate Bax levels as well, and the increased sensitivity to PDT of these cells has been attributed to the higher Bax/Bcl-2 ratio resulting from Bcl-2 photo-oxidation [95,96]. Interestingly, in the murine leukemia L1210 cells the effects of CPOmediated PDT on mitochondrial apoptosis were mimicked by the small molecule Bcl-2 ligand HA14-1 (ethyl 2-amino-6bromo-4-(1-cyano-2-ethoxy-2-oxoethyl)-4H-chromene-3-carboxylate) [97], which by antagonizing Bcl-2 binding to Bak and Bax can either induce mitochondrial apoptosis [98] or sensitize cells to a variety of apoptotic signals [99]. This supports the notion that ROS-mediated Bcl-2 damage suppresses its ability to prevent Bax oligomerization in the OMM [92,100], thereby lowering the threshold for MMP in the photosensitized cells. Furthermore, it is plausible that the generation of singlet oxygen and oxygen radicals by mitochondria-photosensitizing agents triggers either alone (e.g. by oxidation of neighboring proteins and lipids such as cardiolipin) or in combination with additional lethal signals, the activation of Bax and Bak and the consequent induction of MMP. Interestingly, HA14-1 induces the mitochondrial hallmarks of apoptosis in intact cells, but not in isolated mitochondria, suggesting that both mitochondrial and extramitochondrial signals are required [101]. Since photodamage involves also the ER-associated pool of Bcl-2 [80,102], the additional signal for Bax/Bak activation may be provided by changes in Ca2+ homeostasis following photo-induced injury to the ER, the main intracellular Ca2+ store. Given that Bcl-2 family members have been shown to regulate Ca2+ homeostasis besides mitochondrial functions in apoptosis [51,103], and that Ca2+ release as well as its subsequent uptake by mitochondria are key factors regulating PTP opening and cytochrome c release (reviewed in [104,105]), concomitant Bcl-2 photodamage at the mitochondria and ER sites may alter Ca2+ fluxes between these organelles in favor of MMP induction. Indeed, a decisive role for Bcl-2 at the ER in the control of cell death is supported by studies showing that Bcl-2 overexpression reduces resting ER Ca2+ concentration and the extent of capacitative Ca2+ entry [106,107]. In some models the photosensitization process has been shown to be accompanied by the intracellular accumulation of certain sphingolipids, such as ceramide and sphinganine [108– 110]. Because ceramide has been implicated as a pro-apoptotic signal in response to a diversity of cellular insults utilizing the mitochondrial pathway of apoptosis (reviewed in [111]), a role for this bioactive lipid in Bax-mediated MMP following PDT can be proposed. However, some uncertainty remains about the essential role of ceramide in PDT-mediated apoptosis. During Pc 4 photosensitization de novo ceramide is accumulated through the inhibition of ceramide conversion to complex sphingolipids through a direct photo-oxidation of the ER-associated serine palmitoyltransferase (SPT) and is pro-apoptotic [108]. Conversely, in PPME-PDT treated cells, ceramide accumulation involves the activation of acidic sphingomyelinase but it is not essential for apoptosis induction [109]. These studies suggest that the specific metabolic pathway regulating the intracellular accumulation of ceramide and other sphingolipids is of utmost importance in determining cell fate after PDT.

94

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Other potential contributors to Bax activation are fatty acids, such as arachidonic acid and lysophosphatidylcholine produced by the action of PLA2. These micelle-forming lipids can promote positive membrane curvature and have been shown to favor Bax pore forming activity [54,55]. Hence, activation of the cytosolic Ca2+-dependent cPLA2 downstream of the ER-Ca2+ release may lead to the generation of lipid molecules that, in concert with Bax, promote MMP and the consequent release of apoptogenic factors in response to ER photodamage. Consistent with this hypothesis, pharmacological inhibition of cPLA2 protects HeLa and T24 cells from hypericin-mediated photokilling and delays the kinetics of cytochrome c release and procaspase-3 activation [112]. In certain PDT scenarios the cleavage/activation of the BH3 only Bcl-2 family member Bid could represent the key signal for Bax and Bak activation. Bid can be cleaved not only by caspase8 but also by cathepsins released in the cytosol upon lysosomal membrane rupture [113,114], and this pathway may be relevant for those dyes localizing predominantly in these acidic organelles. Light activation of the photosensitizer N-aspartyl chlorin e6 (NPe6), a dye localizing predominantly in lysosomes and endosomes, in murine hepatoma 1c1c7 cells promoted Bid cleavage and mitochondrial apoptosis through the leakage of cathepsins from photo-oxidized lysosomes [115]. Pharmacological inhibitions of cathepsin B and D prevented mitochondrial apoptosis induction by the novel photosensitizing agent ATXs10 (13,17-bis (1-carboxypropionyl)carbamoylethyl-8-etheny2-hydroxy-3-hydroxyiminoethylidene-2,7,12,18-tetraethyl porphyrin sodium) [116]. Since the lysosomal compartment is highly susceptible to oxidation, the participation of lysosomal proteases in apoptosis induction following PDT could prove to have a more general role. However, permeabilization of lysosomal membranes does not precede MMP in photosensitized cells with hypericin [16], which displays a partial colocalization to lysosomes and, moreover, inhibitors of lysosomal proteases do not block cytochrome c release and apoptosis in these instances [16]. Hence, it could be that at the PDT doses used in the former studies, the amount of singlet oxygen generated at the lysosomes may not be sufficient to cause an immediate breakdown of the lysosomal membranes for which higher, possibly necrotic, PDT dose might be required. Alternatively, the temporal resistance of lysosomes to photodamage could be related to the rapid up-regulation of heat shock protein 70 (Hsp70) following photosensitization [117–120], since this molecular chaperone has been implicated in maintaining the integrity of lysosomal membranes in other apoptotic models [121]. Although mitochondrial apoptosis induced by PDT has been considered to be largely independent on the status of the tumor suppressor p53 as it is efficiently induced in p53-null cells [6,122,123] some recent studies indicate a partial dependence on p53-dependent pathways for photokilling [124–126]. In addition, apoptosis mediated by PpIX-PDT has been shown to occur, at least in part, through a p53-dependent pathway involving an up-regulation in the expression of the pro-apoptotic Bcl-2 family members PUMA and Bak [125]. Interestingly, PpIX appears to bind to p53 and to disrupt its interaction with its negative

regulator HDM2 in the HCT116 colon cancer cells [125], suggesting that this photosensitizer is endowed with the capability to regulate p53 transcriptional activity through a novel mechanism. These studies support the notion that multiple dye- and celltype specific mechanisms operate in photosensitized cells in concert to activate Bax, which constitutes an essential molecule in MMP and apoptosis of PDT-treated cells. 2.1.1.2. Anti-apoptotic Bcl-2 family members in PDT. A plethora of studies have provided conclusive evidence that elevation in Bcl-2 expression can cause resistance to chemotherapeutic drugs and radiation (for a recent review see [127]). In contrast, the role of Bcl-2 in PDT is not unambiguous. For instance, while partial protection from PDT-mediated apoptosis has been observed in cells overexpressing Bcl-2, paradoxically, in other instances, increased levels of Bcl-2 enhanced the efficiency of photokilling [6,62]. As mentioned before, light activation of a class of photosensitizers, including the tin etiopurpurin SnET2, the phthalocyanine Pc 4, the chlorin m-THPC Foscan®, the porphycene CPO, results in the targeting of the anti-apoptotic Bcl-2 protein for photodamage in several cell lines in vitro [13,92,95,102], and apparently also in vivo following Pc 4-PDT [80]. Reportedly, both the mitochondria- and ER-associated pools of the anti-apoptotic Bcl-2 proteins have been shown to be targeted by PDT with these photosensitizing agents [13,92,95,102]. Bcl-2/Bcl-xL photodamage by Pc 4-PDT results from an immediate, light- and ROS-dependent process causing crosslinking of the proteins, requires the insertion of Bcl-2/Bcl-xL into membranes and is independent from caspases or other proteolytic signals [94,128]. Interestingly, other OMM proteins such as Bak or VDAC, or cytosolic Bax, are undamaged by PDT indicating that the close vicinity of the target and photosensitizer is a prerequisite, as also demonstrated by FRET analysis [80]. ROS-induced Bcl-2 conformational change and PDT-mediated apoptosis are tightly associated processes. For instance, compounds such as the nontoxic bile acid ursodeoxycholic acid (UDCA) that increases the sensitivity of Bcl-2 to photodamage by promoting proximity to membrane-bound CPO, also enhances caspase-mediated photokilling [129]. Recently, a PDT dose-dependent Bcl-2 photodamage in MCF 7 cells, which lack caspase-3, with the ER-localizing dye Foscan® was suggested to involve specifically the ER-associated pool of Bcl2 [13]. Although no clear relationship between Bcl-2 photooxidation and effector caspase-6/-7 activation was found [13], it is still possible that this event contributes to photokilling by activating, as described further, caspase-independent lethal subroutines, a process that might not be readily appreciated due to the rapid induction of apoptotic cell death by caspase activation. Direct Bcl-2 modifications by hypericin- or PPME-photogenerated ROS have not been revealed [85,130]. Since these photosensitizing agents also localize to the ER in different cell lines, this indicates that ER-associated Bcl-2 is a selective target of a subclass of dyes displaying a high affinity for this protein. In hypericin-PDT, for example, a CDK1-mediated signaling pathway during a G2/M arrest preceding apoptosis is associated with

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

a Ser-70 phosphorylation of the mitochondria-bound Bcl-2, which increases its cytoprotective effects [130]. These studies clearly outline that Bcl-2 can undergo specific post-translational modifications following PDT; it can be either a direct target of the photogenerated ROS or a downstream effector of signal transduction pathways initiated by the primary photodamage to other molecules. Thus, it is not surprising that the changes in Bcl-2 function evoked by these processes are highly photosensitizer-dependent. 2.1.2. Death receptor-mediated signaling Exposure of different cancer cells to light activation using different dyes has been shown to result in an increased expression of mainly Fas and FasL, both in vitro (reviewed in [6,61,62]) and associated to TUNEL-positive cells in response to PDT in vivo [131,132], suggesting that the extrinsic pathway of caspase activation contributes to the PDT-mediated apoptotic response. Combination of PDT with Photofrin® and TNF-α resulted in an increased tumor killing in mice compared to either therapy alone [133] and addition of TNF-α to Photosan 3 photosensitized murine YAC-1 lymphoma cells decreased cell viability additively [134]. The apoptogenic effects of verteporfin on human Jurkat lymphoma cells at suboptimal levels of PDT [135] were increased in an additive fashion when recombinant TNF-α, FasL and/or TRAIL were combined. Addition of TNF-R1-Fc, TRAIL-R2-Fc, and Fas-Fc chimera proteins did not affect the extent of cytochrome c release and DNA fragmentation that occurred in response to PDT, thus suggesting the participation of two separate and eventually converging apoptotic pathways [135]. Consistent with this view, monitoring the effects of Photofrin®-PDT in single cell by fluorescent imaging indicated that caspase-3 was activated rapidly while caspase-8 remained inactive [136]. Furthermore, PDT-induced apoptosis has been shown to be unaltered in FADD deficient mouse embryonic fibroblasts [137] and in HeLa cells overexpressing CrmA, a cowpox specific inhibitor of caspase-8 and caspase-1 [138], indicating that engagement of the DR signaling pathway is dispensable for the induction of apoptosis by PDT. In favor of a more direct participation of the DR pathway of caspase activation in PDT are observations indicating that in certain PDT settings the multimerization and activation of Fas may occur in a ligand-independent fashion [139] and that activation of caspase-8 precedes mitochondrial cytochrome c release and apoptosis [140–143]. However, conflicting evidence on the hierarchical ordering of caspase activation following PDT remain since other studies indicated that caspase-8 processing was secondary to the activation of the mitochondrial caspase cascade, likely as part of a cytosolic amplification step [85,135,144]. In conclusion, these studies suggest that while a direct involvement of DR-signaling of caspase activation in PDTinduced apoptosis may be restricted to certain cell types, the extrinsic pathway may be propagated through autocrine/ paracrine signals resulting from the increased expression of death receptors and their ligands following PDT.

95

2.1.3. The role of ER stress in apoptotic cell death after PDT 2.1.3.1. Photodamage to the endoplasmic reticulum. Oxidative damage to the ER following PDT can result in dramatic changes in ER homeostasis, which can be further propagated to the mitochondrial cell death machinery [6,61,62]. Different ERassociated dyes have been shown to stimulate a rapid increase in the intracellular calcium concentrations ([Ca2+]cyt) immediately after photoactivation. Based on the use of the cellpermeable Ca2+ chelators (e.g. BAPTA- or EGTA-AM) to buffer the [Ca2+]cyt increase or inhibitors of mitochondrial Ca2+ uptake (e.g. ruthenium red) after PDT, either no clear cut causative role [16,85,102,145], an apoptogenic [146–151] or even a protective role [152–154] for the increase in intracellular Ca2+ has been proposed. These different functional outcomes could be related to the source of Ca2+ mediating the rise in the [Ca2+]cyt, as both influx of extracellular Ca2+ and the release of Ca2+ from different intracellular stores (e.g. ER, mitochondria) have been observed following photosensitization (reviewed in [62]). In some PDT paradigms the increase in [Ca2+]cyt is associated to an immediate 1O2-mediated damage to the sarco/endoplasmic reticulum Ca2+-ATPase-2 (SERCA2) pump [16,145], which by coupling ATP hydrolysis to Ca2+ transport from the cytosol across the ER membrane, maintains the level of resting [Ca2+]ER three to four orders of magnitude higher than the [Ca2+]cyt [155]. The dramatic perturbations in Ca2+ homeostasis caused by the incapability of the cells to refill ER Ca2+ pools following SERCA 1O2 attack, rather than the increase in [Ca2+]cyt per se, were functionally linked to photokilling as antioxidants prevented SERCA loss, re-established Ca2+ homeostasis and conferred resistance to cell death [16]. Interestingly, rapid Ca2+ overload per se was not sufficient for the induction of mitochondrial apoptosis in these PDT paradigms [16,145]. This notion is further supported by the observation that in Bax/ Bak deficient cells in which the lower level of resting [Ca2+]ER has been corrected by the overexpression of SERCA, which restores sensitivity to PT and apoptosis in response to a subset of cellular stresses in the DKO cells [51], hypericin-PDT does not induce apoptotic cell death, for which the re-expression of mitochondria Bax is strictly required [16]. However, depletion of the ER Ca2+ pool committed the cells to death, as Bax/Bak deficient cells were equally well photokilled by hypericin, through a caspase-independent subroutine of cell death [16]. This is also supported by studies demonstrating that Pc 4induced photokilling of Bax-deficient MCF7 or HCT116 cells still proceeds, although in a delayed form, in the absence of cytochrome c release, drop of Δψm and caspase activation [89]. Thus, in spite of the essential role of Bax (and Bak) for MMP and apoptosis, these reports indicate that the commitment event in cell death occurs upstream to and independent of Bax and Bak. Indeed, recent findings indicate that interfering with the sequestration of Ca2+ into the ER is sufficient to trigger cell death as part of a stress response [156–158]. The signaling mechanisms transducing the primary photodamage to the ER into cell death pathways in PDT-treated cells are not completely solved, but a role for the activation of

96

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

phospholipases, calpains and Bap31 has been proposed in recent studies. Rapid release of Ca2+ from intracellular pools following photosensitization of murine lymphoma L5178Y cells with aluminum phthalocyanine (AlPc) was found to be associated with a rapid activation of phospholipase C (PLC) and the breakdown of membrane phosphoinositides. In this system, the PLC inhibitor, U73122, blocked both the rapid and transient increase in inositol-1,4,5-trisphosphate and intracellular Ca2+ levels as well as PDT-induced apoptosis [159]. PKC inhibition or its down-regulation by prolonged incubation with 12-Otetradecanoyl-phorbol-13-acetate (TPA) protected Chinese hamster ovary (CHO) cells from AlPc-mediated PDT cytotoxicity [160] suggesting that Ca2+ -dependent signaling pathways mediated by PKC activation can contribute to photokilling in this system. Furthermore, inhibitors of cPLA2 have been shown to impede apoptotic cell death in some PDT paradigms [112,159], likely by blocking the breakdown of membrane phospholipids and the resulting release of arachidonic acid, which is a known apoptotic mediator [112,161]. The Ca2+-dependent cysteine proteases calpains have been recently shown to be activated either in the absence of caspase signaling or in parallel to the canonical caspase-dependent pathways using bisulfonated aluminum phthalocyanine (AlPcS2) [162] or 5-ALA [163] as the photosensitizers. Calpains can contribute to apoptosis by cleaving Bcl-2 family members, including Bax and Bid or by modulating caspase activity (e.g. activation of caspase-12, or inhibition of effectors caspases), with which they share common substrates [104]. However, cleavage of Bax has not been reported in PDT-treated cells [162] and inhibition of calpains is not sufficient to block apoptotic cell death in some PDT contexts either [16,162,163]. These observations suggest that the activation of calpains is not an essential step in the initiation of apoptosis after PDT, but likely contributes to the final elimination of the dying cells and/or participates in non-apoptotic cell death subroutines. The ER-mitochondria cross-talk in HeLa cells following PDT with verteporfin was proposed to involve the integral ER membrane protein Bap31 [144]. Bap31 has been reported to undergo cleavage by caspase-8 and to mediate a Ca2+-dependent pathway leading to mitochondrial fission which enhances the release of cytochrome c and apoptosis [164]. However, caspase8 mediated cleavage of Bap31 occurred secondary to the initial cytochrome c release in verteporfin-photosensitized cell [144], suggesting that in this paradigm the ER-mitochondrial cross talk is engaged to amplify the mitochondrial caspase activation cascade. The major molecular signals evoking MMP in photosensitized cells are illustrated in Fig. 3. 2.1.3.2. Activation of the UPR in response to PDT-mediated ER stress. Apart from Ca2+ storage and signaling, the main function of the ER is folding, modifying and sorting of newly synthesized proteins. Disturbances in any of these functions can lead to ER stress. Ca2+ overload and depletion of the ER Ca2+ pool are insults leading to ER stress, changes in protein folding

and subsequent activation of the unfolded protein response (UPR). The UPR is a primarily pro-survival response activated to reduce the accumulation and aggregation of unfolded or misfolded proteins and to restore normal ER functioning by the induction of molecular chaperones, translational attenuation and ER-associated degradation (ERAD) (reviewed in [165]). However, if ER stress persists and cannot be repaired, the UPR can activate a cell death program, which generally converges into the caspase-activation cascade. The UPR is initiated through dissociation of the ER chaperone GRP78/Bip upon accumulation of unfolded proteins from three ER transmembrane stress receptors: the pancreatic ER kinase (PKR)-like ER kinase (PERK), activating transcription factor 6 (ATF6) and inositolrequiring enzyme 1 (IRE1), allowing their sequential activation by homodimerization and autophosphorylation. PERK activation results in eIF2α-phosphorylation causing inhibition of general protein translation, which alleviates protein loading on the ER. However, this event also leads to the translation of certain genes, such as that encoding for activating transcription factor 4 (ATF4). ATF4 promotes cell survival through the induction of genes involved in restoring ER homeostasis, but it can also favor apoptosis through the induction of the transcription factor C/EBP homologous protein (CHOP). CHOP induction is thought to tip the balance towards apoptosis under conditions of persistent ER stress [166,167]. CHOP positively regulates the expression of the protein phosphatase-1 (PP1) interacting protein growth arrest and DNA damage-inducible gene 34 (GADD34), which causes PP1 to dephosphorylate eIF2α thereby resulting in translational recovery, while it downregulates the expression of Bcl-2, thereby increasing the sensitivity of the cell to undergo apoptosis [168]. ATF6 leads to the induction of ER chaperone proteins such as GRP78/Bip, GRP94, protein disulphide isomerase (PDI), and converges also in the regulation of the transcription factors CHOP and XBP1 (X-box binding protein 1). A second branch of the UPR involves the IRE1-XBP1 pathway. IRE1 leads to cleavage of XBP1 generating the active transcription factor XBP1s, which induces a negative feedback loop relieving the PERK-mediated translational block. This event might represent termination of the UPR, returning the ER to normal functioning when the UPR has been successful, or, when the stress persists to cell death by proapoptotic protein synthesis [167]. Additionally, via the adaptor molecule TNF receptor associated factor 2 (TRAF2) and apoptosis signal-regulating kinase-1 (ASK1), IRE1 may lead to activation of JNK (c-Jun N-terminal kinase) and p38MAPK . This provides a link with the apoptotic pathway through regulation of both anti- and pro-apoptotic Bcl-2 family proteins by JNK [46] and the p38MAPK -mediated phosphorylation/ activation of CHOP [169]. The execution of the UPR-mediated cell death involves the activation of caspases. Both murine caspase-12 and human caspase-4 have been reported to associate with the ER, and in rodents the ER-associated caspase-12 has been proposed to be the initiator caspase in ER stress pathways to apoptosis. In this pathway, it has been proposed that calpain (m-calpains) is a central player in the conversion of Ca+2 signals from the stressed ER to the caspase-12 activation [170], which can then lead to caspase-9/caspase-3/7 activation and apoptosis

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

97

Fig. 3. Signaling pathways leading to MMP in photosensitized cells. Depending on the nature of the photosensitizer (PTS) and its intracellular localization, the initial photodamage can involve different molecules with the consequent activation of specific death pathways converging into MMP. Mitochondria-localized PTS can cause an immediate and light-dependent photodamage to mitochondria gatekeepers, such as the anti-apoptotic Bcl-2 and Bcl-xL proteins, or to putative PTP components, prompting the release of caspase-activating molecules, such as cytochrome c, in the cytosol. PTS accumulating prevalently in the lysosomes induce lysosomal membrane permeabilization leading to the release of lysosomal hydrolases and cleavage of the pro-apoptotic BH3-only protein Bid, which can culminate in Baxmediated MMP and caspase activation. With ER localizing PTS, extensive photodamage to the SERCA2 pump leads to ER Ca2+ depletion, which initiates Baxdependent mitochondrial apoptosis. Increase in intracellular Ca2+ concentration can result in the activation of cPLA2 and generation of arachidonic acid (AA). Mitochondrial Ca2+ uptake, ROS and AA can synergize to induce PTP resulting in MMP. Pathways leading to the intracellular accumulation of ceramide (Cer) in photosensitized cells can be propagated from ER-associated or lysosomal (aSmase) ceramide metabolizing enzymes. In most cases pro-apoptotic effectors Bax/Bak are essential effectors of MMP and consequent caspase activation, molecular events which rapidly precipitate apoptotic cell death.

in a cytochrome c and Apaf-1-independent manner [171,172]. However, the general role of caspase-12 and caspase-4 in ER stress is still debatable [167] as most humans express a truncated prodomain-only form of caspase-12 and both caspases belong to the subclass of inflammatory caspases, with a proposed role in inflammation and innate immunity rather than in ER stress. The main pathways activated by ER stress are illustrated in Fig. 4. Recent data indicate that following PDT different heat shock proteins as well as the ER chaperones, GRP78/Bip, Grp94, calreticulin, PDI, calnexin are induced in a time dependent manner [13,119,149,163]. In addition, induction of CHOP and activation of the ER stress-mediated caspase-12 are taking place in photosensitized cells ([119,149], Buytaert E and Dewaele M, unpublished results). Given that PDT-mediated apoptosis with a mitochondrial and ER-localizing porphyrin has been reported to be attenuated in chop deficient cells [118], it seems reasonable to assume that sustained UPR leading to the

induction of CHOP contributes to apoptosis induction in photosensitized cells. A recent genome-wide analysis in hypericin-PDT exposed bladder cancer cells (T24 cells) revealed that proximal molecular sensors and effectors of the UPR are induced in a coordinated manner [119]. Consistent with the observation that in this PDT model the most apical lethal signal is a fast ER Ca2+ depletion induced by photo-oxidation of the SERCA pump [16], these studies further uncover that perturbations in the ER caused by accumulation of photo-oxidated/misfolded proteins can persistently activate the UPR pathway. The nature of the pathways activated by ER stress and cell fate following UPR induction are not fully understood, but increasing evidence indicates that this pathway is used to induce both adaptive and cell death responses and could affect tumor growth, promote dormancy and influence therapy outcome [173]. Hence the functional impact of the UPR on the modulation and efficiency of PDT-induced cell death in cancers should be further evaluated both in vitro as well as in vivo.

98

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Fig. 4. Molecular interplay between the ER and the mitochondria during cellular stress. Cellular stress impinging on the ER, results in the activation of mitochondriadependent and mitochondria-independent apoptotic pathways. The uptake of excessive amounts of Ca2+ by the mitochondria, upon Ca2+ overload, results in the mitochondrial release of cytochrome c and induction of the apoptosome leading to the caspase activation cascade. In addition, the release of Ca2+ from the ER may cause the activation of calpains, which can process and activate caspase-12 (the human homologue is caspase-4). Caspase-12 can directly activate caspase-9, thereby contributing to apoptosis via a mitochondria-independent pathway (see text). Under conditions of ER stress, the accumulation of unfolded proteins in the ER triggers the activation of proximal effectors of the unfolded protein response (UPR). In this pathway the induction of CHOP via ATF-4 and XBP1s by the coordinated action of PERK, ATF-6 and IRE1, leads to the down-regulation of the expression of Bcl-2, which favors cytochrome c efflux and mitochondrial apoptosis. In a second branch of the UPR, IRE1 recruits TRAF-2 in a complex to activate caspase-12. The IRE1-TRAF-2 complex also recruits the MAPKKK, ASK1 which in turn results in the activation of the stress activated p38 and JNK MAPKs involved in the modulation of apoptosis induction.

2.1.4. Caspase-independent apoptosis Caspases appear to constitute a major but not the sole determinant for the manifestation of apoptotic morphology. Indeed, caspase inhibition has been consistently shown to protect cells only transiently or not efficiently, against apoptotic cell death induced by a variety of stress signals including PDT (reviewed in [6]). This is because once MMP has occurred cell death can be propagated, albeit with delayed kinetics, regardless of caspase activity, through caspase-independent cell death pathways. Known mitochondrial inducers of caspase-independent cell death are the flavoprotein AIF and the endonuclease Endo G. These IMS pro-apoptotic proteins are released in the cytosol following MMP and are thought to translocate to the nucleus where they are involved in DNA fragmentation and chromatin condensation in a caspase-independent fashion [30]. Recent studies have implicated the mitochondrial release of AIF as key mediator of caspase-independent apoptosis following PDT [69,91,174]. In verteporfin-mediated PDT of smooth muscle cells, AIF has been shown to be released from mitochondria together with the caspase activators cytochrome c and Smac/DIABLO [91]. In HAL-loaded Reh cells [69] and Jurkat cells [174], the release of mitochondrial AIF in the cytosol was followed by its nuclear translocation and by z-VAD-fmkindependent large scale DNA degradation. In the Reh cell model, AIF release was accompanied with Δψm loss and was

blocked by BA or by PK11195, evoking the involvement of PT as mechanism for MMP, while cytochrome c was retained in the mitochondria [69]. To explain this observation it was suggested that PpIX-mediated mitochondrion-photosensitization leads to a site-specific PT modulation which would allow the specific AIF release [69]. Discrepancy regarding as to whether AIF release precedes, is concomitant to, or follows the release of others IMS proteins from mitochondria as revealed by these studies, is not restricted to PDT scenarios but reflects a general and still unresolved issue. In some instances it has been shown that cytochrome c release precedes the release of other IMS proteins and may promote AIF and Smac/Diablo release through a caspase-dependent mechanism [175], whereas in other paradigms cytochrome c follows the caspase-independent efflux of AIF, Smac/Diablo and Endo G [176,177]. In the latter case it is thought that the major pool of cytochrome c remains tightly associated to the IMM and requires the cooperation of additional signals (e.g. cardiolipin oxidation, Bax/Bak, mitochondria fission) in order to be released in the cytosol [177]. In conclusion, irrespective of the mechanism of MMP, these data indicate that the induction of apoptotic cell death after PDT underlie the activation of caspase-dependent and-independent pathways, which can act in parallel to ensure the death of the doomed cells. Additionally, as mentioned before, caspase signaling is not required for the induction of necrosis and likely also

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

for autophagic cell death, two cell death modalities, which have been described to occur in response to PDT. 2.2. Type II PCD: autophagic cell death Macroautophagy, hereafter called autophagy (literally “selfeating” in Greek) is a major catabolic process initiated in eukaryotic cells to generate the intracellular building blocks through the recycling of cytoplasmic components [178]. In yeast, where this process has been best studied, autophagy functions as a survival or adaptive mechanism allowing the maintenance of vital functions during nutrient-limiting conditions. In mammalian cells autophagy may also promote cell survival by removing damaged organelles, toxic metabolites or intracellular pathogens. However, autophagy may as well promote cell death through excessive self-digestion and degradation of essential cellular constituents (for recent reviews on autophagic cell death the readers are referred to [179–181]). Although many unanswered questions remain concerning the molecular players of the “autophagic cell death”, this cellular program is thought to proceed in the absence of caspase-signaling or even to be activated under conditions of caspase inhibition [182,183]. The initial step in autophagy is the formation of a doublemembrane structure that sequesters cytoplasmic components as well as organelles and shapes the autophagic vacuoles or the socalled autophagosomes [178]. Eventually, autophagosomes fuse with lysosomes and their cytoplasmic material is degraded by lysosomal hydrolases. A family of autophagy-related genes (Atg) discovered in yeast and almost integrally conserved in all eukaryotic phyla, controls the formation of the autophagosome. Autophagy is regulated by class I and class III phosphatidylinositol 3 kinase (PI3K) signaling pathways, which have been reported to inhibit and stimulate autophagy, respectively [184]. Formation and completion of the sequestering vesicle is a complex mechanism which is under positive control of the class III PI3K complex and involves several autophagy regulators including Atg6/Beclin 1 and Atg8/LC 3 (for an extensive review on the molecular machinery of autophagy, see [178]). The functional contribution of this catabolic process in cell death is still uncertain as it is currently unclear whether autophagy directly contributes to death or is a failed effort to preserve cell viability. However, recent studies suggest that autophagy may regulate cancer development and progression as well as response to cytotoxic therapy (reviewed in [181,185]. Several anticancer agents including tamoxifen, rapamycin, arsenic trioxide, temozolomide, histone deacetylase inhibitors, ionizing radiation, vitamin D analogue and etoposide, have been reported to kill cancer cells and to concurrently induce autophagy as part of a stress response [185]. However, the molecular mechanisms utilized by these drugs to trigger autophagy in the cancer cells and the direct implication of autophagic cell death following anticancer treatment are still unresolved questions. 2.2.1. PDT as inducer of autophagy Recent studies have shown that PDT may induce nonapoptotic cell death associated with the induction of autophagy [16,17].

99

Due to the high reactivity of photogenerated ROS it is not surprising that autophagy is initiated in an attempt to remove oxidatively damaged organelles or to degrade large aggregates of cross-linked proteins, produced by photochemical reactions, which cannot be removed by the ubiquitin–proteasome system or by the ERAD. Since autophagy is a self-limiting process, it is possible that its persistence results in a metabolic and bioenergetic collapse, which is causative for cell death [15]. Alternatively, the function of autophagy could be orchestrated by dedicated signaling molecules and switched from a survival to a lethal pathway in certain instances. So far two studies reported on the induction of autophagy in PDT-treated cells and on the relationship between autophagy and cell death [16,17]. In a first study the commitment event and the subsequent cell death modalities in hypericin-mediated PDT were examined in wild type MEFs and in apoptosis-deficient Bax−/−Bak−/− MEFs. In these cells all the biochemical hallmarks of apoptosis are prevented while photokilling continues unaltered through the induction of a non-apoptotic cell death pathway, which is associated to the ultrastructural and biochemical features of autophagy [16]. The pharmacological blockade of autophagy by the phosphatidylinositol 3-kinase (PI3K) class III inhibitor wortmannin, in apoptosis-deficient DKO cells results in a significant reduction of cell death, which evidences that PDT can simulate an autophagic cell death pathway, at least under conditions of apoptosis inhibition [16], as shown in Fig. 5. Further studies have shown that mitochondrial apoptosis and autophagy are concurrently promoted downstream of ER damage in PDT-treated cells ([16,17] and Dewaele M, unpublished results). The simultaneous induction of autophagy and apoptosis has been reported in murine leukemia L1210 cells after PDT with the porphycene CPO [17,186]. Interestingly, protection of CPO-mediated Bcl-2 photodamage reduced not only caspase activation but also the conversion of LC-3 I, e.g. the mammalian homologue of Atg8, into the lipidated LC-3 II form, a biochemical hallmark of autophagy [187]. This led to the suggestion that loss of native Bcl-2 may regulate both apoptotic and autophagic pathways following CPO-mediated PDT [186]. This hypothesis is based on the observation that Bcl-2 and/or Bcl-xL can inhibit not only apoptosis but also Beclin 1-dependent autophagy and Beclin 1-dependent autophagic cell death [188], through a direct interaction, which has been recently shown to require the BH3 domain in Beclin 1 [189]. It is important to mention, however, that the concomitant engagement of both apoptosis and autophagy occurs also in HeLa cells following hypericin-mediated PDT [Dewaele et al., unpublished results] regardless of a reduction in the expression levels of anti-apoptotic Bcl-2 proteins [130], thus suggesting that other molecular determinants can play a key role in this process. The specific ROS-damaged subcellular site or organelle and the cargo of the autophagic vacuole are factors which could potentially influence the outcome of the autophagic process in PDT-treated cells. For instance, depending on the type and degree of organellar dysfunction, a degradative pathway targeting preferentially the photo-oxidized organelle or the cross-

100

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Fig. 5. Autophagy induction by photo-oxidative damage to intracellular organelles/targets. Photodamage to key organelles like the ER, mitochondria and lysosomes following PDT with different PS could trigger autophagy as part of a survival pathway into the attempt to remove the dysfunctional organelle. Photo-oxidative damage to the ER or specifically to the anti-apoptotic Bcl-2/Bcl-xL proteins, which may alter Bcl-2/Bcl-xL interaction with Beclin 1, favor the induction of autophagy (see text). In apoptosis-deficient cells, as seen in cells lacking the pro-apoptotic multidomain Bax and Bak proteins and/or caspase signaling, an autophagic cell death pathway is revealed.

linked protein aggregates for autophagy could be specifically initiated. Examples of specific degradation pathways for damaged mitochondria, a process called mitophagy, have been documented in cultured hepatocytes [190] and in yeast [191]. In mammalian cells mitophagy often precedes apoptosis and requires caspase inhibition in order to be fully appreciated (discussed in [192]). It still remains to be clarified though whether mitophagy delays cell death, by preventing leaking mitochondria from spilling out their pro-apoptotic proteins and from generating ROS, or accelerates cell demise, by degrading the major contributors of cellular ATP production [192]. In situations where the ER is the main photo-damaged organelle and mitochondria are spared from major alterations as reported in Bax-deficient cells [16,89,193] autophagy could target the ER for extensive engulfment and degradation resulting in the activation of a cell death pathway, through a molecular mechanism which still needs to be clarified [16]. In apoptosis-competent cells where mitochondria are undergoing rapidly MMP and caspases are active in response to PDT, the function of the autophagic pathway(s) initiated by ER photodamage should be further evaluated. Recent studies have established that there are instances where the ER is preferentially targeted for autophagic degradation to support survival in yeast [194,195], and auto-

phagy is initiated following ER stress in mammalian cells [196–199]. In conclusion, more studies are required to identify the molecular mediators which may turn autophagy into a bona fide cell death pathway and to better understand the cross-talk between autophagy and the apoptotic machinery in PDT-exposed cancer cells. Defining the threshold of PDT-induced autophagy compatible with survival of the cancer cells could also provide important insights in this matter, as after a sublethal PDT dose the induction of autophagy may contribute to cancer cell survival rather than death. Last but not least it is important to assess whether autophagy is an in vivo tumor response evoked by PDT and whether the use of pharmacological modulators of this catabolic process in combination with PDT will prove beneficial to improve its therapeutic efficacy. 2.3. Type III PCD: necrosis Necrosis is morphologically characterized by vacuolization of the cytoplasm, swelling and breakdown of the plasma membrane resulting in an inflammatory reaction due to release of cellular contents and pro-inflammatory molecules. Necrosis is thought to be the end result of a bio-energetic catastrophe

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

resulting from ATP depletion to a level incompatible with cell survival (reviewed in [20]). However, although necrosis has long been described as a passive, unorganized way to die, recent evidence suggests that necrotic cell death can be actively propagated as part of a signal transduction pathway. For instance, the engagement of TNF- and Fas receptors in certain cell lines trigger necrosis through the activation of RIP1 (receptor interacting protein 1) kinase, under conditions of caspase inhibition [200,201]. Moreover, RIP1 has been suggested to be a key element of the JNK-mediated necrotic cascade activated by poly(ADP-ribose) polymerase-1 (PARP-1) following oxidative stress injury [202]. Thus these observations indicate that in certain circumstances necrotic cell death may rely on the activation of an intracellular program in response to specific cues (for a recent review see [203,204]). 2.3.1. Programmed necrosis in PDT? A shift from apoptotic to necrotic cell death using a particular dye can be usually instigated by increasing the intensity of the PDT dose (e.g. by increasing the light dose or concentration of the dye). This generates a massive induction of ROS leading to an immediate bioenergetic catastrophe, drastic drop in ATP levels and general metabolic inhibition. Necrosis is the major cell death morphology induced by PDT with compounds localized to the plasma membrane (reviewed in [6,61,62]). This is likely due to a rapid loss of plasma membrane integrity, incapability to maintain ion fluxes across the plasma membrane and fast depletion of intracellular ATP, following photosensitization as shown in studies using Photofrin® [205] or zinc(II) phthalocyanine [206]. In certain PDT paradigms necrosis, and not secondary necrosis consequent to apoptotic cell death, appears to be the preferential mode of cell death also for photosensitizers originally found in or subsequently relocalized to other subcellular compartments. This suggests that signaling pathways that orchestrate necrosis rather than apoptosis may exist. Although a biochemical pathway mediating necrosis following PDT has not been identified yet, certain factors, such as Ca2+ overload, the origin and type of generated ROS, may be decisive to promote a necrotic cell death pathway. A central role of Ca2+ in photo-oxidative initiation of necrotic cell death of neuronal and glial cells by photosens, a mixture of different sulfonated aluminum phthalocyanines AlPcSn, has been reported to involve calmodulin and CaMKII signaling [207]. In an epithelial breast tumor cell line [83] the mode of cell death was found site-specific with respect to the distribution of a fixed level of PpIX. Apoptosis was the preferred mode of photokilling when PpIX was associated to the mitochondria, while this pathway was preempted by necrosis, occurring in the absence of cytochrome c release and caspase signaling, when most of the porphyrin diffused to other cellular targets [83]. Some cationic porphyrins, which have been shown to relocalize from plasma membrane to the cytosol during irradiation, have been reported to cause photoinactivation of procaspase-9 and procaspase-3 [12]. As inhibition of caspases has been documented to be a signal shifting the mode of cell death in favor of necrosis, this photosensitizer-

101

mediated caspase inactivation mechanism in the cytosol could result in the activation and propagation of a necrotic cell death pathway. Using specific ROS quenchers in PPME-photosensitized cells it was suggested that 1O2 produced at the ER/Golgi membranes mediates necrosis, whereas ROS other than 1O2 and likely produced by the mitochondria would activate the intrinsic apoptosis pathway [85]. In HeLa cells loaded with CMXRos (Mitotracker Red) intense light activation of the dye caused the photogeneration of both 1O2 and superoxide anion (O2·−) followed by necrotic cell death [208]. In this model a post-PDT massive intramitochondrial ROS accumulation was blocked by diphenyleneiodonium, and inhibitor of flavin-containing enzymes, suggesting the involvement of mitochondria Complex I in this process. Indeed, increasing evidence suggest that Complex I is an important, if not the major, source of O2·− generated by the mitochondria [81]. In this model necrosis was blocked by the mitochondriatargeted antioxidant (MitoQ) while pharmacological PT- (e.g. CsA and BA) and caspase inhibitors were not protective. Conversely, CMXRos-mediated apoptosis was not affected by MitoQ, but it was counteracted by Bcl-2 and caspase inhibition [208]. These PDT models indicate that necrotic or apoptotic cell death following PDT may be differentially regulated by the type and site of ROS produced within the cells, and point to O2·− generated by Complex I as a pro-necrotic signal following the initial photodamage. This would be consistent with studies indicating a relevant role for mitochondria-produced ROS in TNFα-induced necrotic killing and with the observations of the cytoprotective effect of the Complex I inhibitor rotenone (for a review see [203,204]). 3. Conclusions The raising interest in PDT as promising anticancer treatment is witnessed by the increasing literature on the cell death mechanism elicited in cancer cells by the photoactivation of a diverse group of second generation photosensitizers. Recent evidence indicate that PDT can evoke the main cell death morphologies which have been described; apoptotic, necrotic and autophagic cell death, which explains why in some systems the specific inhibition of one death signal is not sufficient to block PDT-mediated cell death. Recent studies point out that while apoptosis is probably the preferred path to cell death, is not the unique one. If the apoptotic route is compromised, the photodamaged cell fated to die will utilize the autophagic or necrotic programs for its demise. Understanding the molecular differences and identifying the cross-talk between these cell death programs will certainly prove crucial to the development of new therapeutic modalities in PDT aimed at increasing the killing efficacy of the cancer cells, in which one or several of these pathways might have been inactivated. Moreover, a better knowledge of the way cancer cells die following PDT will also contribute to a better understanding of the impact that different cell death modalities have on the innate and adaptive immune responses and on therapeutic outcome.

102

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

Acknowledgements Research in the author's laboratory is supported by the Onderzoekstoelage (OT) from the K.U.Leuven, the Interuniversitaire Attractiepolen (IAP VI/P18) of the Federal Belgian Government and by F.W.O grant G.0104.02. We apologize to those colleagues whose work has not, or only partially, been cited in this review due to space limitation. References [1] B.W. Henderson, T.J. Dougherty, How does photodynamic therapy work? Photochem. Photobiol. 55 (1992) 145–157. [2] K. Berg, P.K. Selbo, A. Weyergang, A. Dietze, L. Prasmickaite, A. Bonsted, B.O. Engesaeter, E. ngell-Petersen, T. Warloe, N. Frandsen, A. Hogset, Porphyrin-related photosensitizers for cancer imaging and therapeutic applications, J. Microsc. 218 (2005) 133–147. [3] D.E. Dolmans, D. Fukumura, R.K. Jain, Photodynamic therapy for cancer, Nat. Rev., Cancer 3 (2003) 380–387. [4] T.J. Dougherty, C.J. Gomer, B.W. Henderson, G. Jori, D. Kessel, M. Korbelik, J. Moan, Q. Peng, Photodynamic therapy, J. Natl. Cancer Inst. 90 (1998) 889–905. [5] M. Niedre, M.S. Patterson, B.C. Wilson, Direct near-infrared luminescence detection of singlet oxygen generated by photodynamic therapy in cells in vitro and tissues in vivo, Photochem. Photobiol. 75 (2002) 382–391. [6] N.L. Oleinick, R.L. Morris, I. Belichenko, The role of apoptosis in response to photodynamic therapy: what, where, why, and how, Photochem. Photobiol. Sci. 1 (2002) 1–21. [7] A.P. Castano, P. Mroz, M.R. Hamblin, Photodynamic therapy and antitumour immunity, Nat. Rev., Cancer 6 (2006) 535–545. [8] B.W. Henderson, S.M. Waldow, T.S. Mang, W.R. Potter, P.B. Malone, T.J. Dougherty, Tumor destruction and kinetics of tumor cell death in two experimental mouse tumors following photodynamic therapy, Cancer Res. 45 (1985) 572–576. [9] J. Moan, K. Berg, The photodegradation of porphyrins in cells can be used to estimate the lifetime of singlet oxygen, Photochem. Photobiol. 53 (1991) 549–553. [10] K. Berg, K. Madslien, J.C. Bommer, R. Oftebro, J.W. Winkelman, J. Moan, Light induced relocalization of sulfonated meso-tetraphenylporphines in NHIK 3025 cells and effects of dose fractionation, Photochem. Photobiol. 53 (1991) 203–210. [11] D. Kessel, M. Conley, M.G. Vicente, J.J. Reiners, Studies on the subcellular localization of the porphycene CPO, Photochem. Photobiol. 81 (2005) 569–572. [12] D. Kessel, Relocalization of cationic porphyrins during photodynamic therapy, Photochem. Photobiol. Sci. 1 (2002) 837–840. [13] S. Marchal, A. Francois, D. Dumas, F. Guillemin, L. Bezdetnaya, Relationship between subcellular localisation of Foscan and caspase activation in photosensitised MCF-7 cells, Br. J. Cancer 96 (2007) 944–951. [14] D. Kessel, Y. Luo, Y. Deng, C.K. Chang, The role of subcellular localization in initiation of apoptosis by photodynamic therapy, Photochem. Photobiol. 65 (1997) 422–426. [15] E. Buytaert, G. Callewaert, J.R. Vandenheede, P. Agostinis, Deficiency in apoptotic effectors Bax and Bak reveals an autophagic cell death pathway initiated by photodamage to the endoplasmic reticulum, Autophagy 2 (2006) 238–240. [16] E. Buytaert, G. Callewaert, N. Hendrickx, L. Scorrano, D. Hartmann, L. Missiaen, J.R. Vandenheede, I. Heirman, J. Grooten, P. Agostinis, Role of endoplasmic reticulum depletion and multidomain proapoptotic BAX and BAK proteins in shaping cell death after hypericin-mediated photodynamic therapy, FASEB J. 20 (2006) 756–758. [17] D. Kessel, M.G. Vicente, J.J. Reiners Jr., Initiation of apoptosis and autophagy by photodynamic therapy, Lasers Surg. Med. 38 (2006) 482–488.

[18] R.A. Lockshin, Z. Zakeri, Programmed cell death and apoptosis: origins of the theory, Nat. Rev., Mol. Cell Biol. 2 (2001) 545–550. [19] J.U. Schweichel, H.J. Merker, The morphology of various types of cell death in prenatal tissues, Teratology 7 (1973) 253–266. [20] A.L. Edinger, C.B. Thompson, Death by design: apoptosis, necrosis and autophagy, Curr. Opin. Cell Biol. 16 (2004) 663–669. [21] R.A. Lockshin, Z. Zakeri, Apoptosis, autophagy, and more, Int. J. Biochem. Cell Biol. 36 (2004) 2405–2419. [22] N.N. Danial, S.J. Korsmeyer, Cell death: critical control points, Cell 116 (2004) 205–219. [23] D. Hanahan, R.A. Weinberg, The hallmarks of cancer, Cell 100 (2000) 57–70. [24] B. Zhivotovsky, G. Kroemer, Apoptosis and genomic instability, Nat. Rev., Mol. Cell Biol. 5 (2004) 752–762. [25] M. Lamkanfi, W. Declercq, M. Kalai, X. Saelens, P. Vandenabeele, Alice in caspase land. A phylogenetic analysis of caspases from worm to man, Cell Death Differ. 9 (2002) 358–361. [26] K.M. Boatright, G.S. Salvesen, Mechanisms of caspase activation, Curr. Opin. Cell Biol. 15 (2003) 725–731. [27] A.U. Luthi, S.J. Martin, The CASBAH: a searchable database of caspase substrates, Cell Death Differ. 14 (2007) 641–650. [28] M. Deshmukh, K. Kuida, E.M. Johnson Jr., Caspase inhibition extends the commitment to neuronal death beyond cytochrome c release to the point of mitochondrial depolarization, J. Cell Biol. 150 (2000) 131–143. [29] I. Martinou, S. Desagher, R. Eskes, B. Antonsson, E. Andre, S. Fakan, J.C. Martinou, The release of cytochrome c from mitochondria during apoptosis of NGF-deprived sympathetic neurons is a reversible event, J. Cell Biol. 144 (1999) 883–889. [30] G. van Loo, X. Saelens, M. van Gurp, M. MacFarlane, S.J. Martin, P. Vandenabeele, The role of mitochondrial factors in apoptosis: a Russian roulette with more than one bullet, Cell Death Differ. 9 (2002) 1031–1042. [31] M. Crompton, E. Barksby, N. Johnson, M. Capano, Mitochondrial intermembrane junctional complexes and their involvement in cell death, Biochimie 84 (2002) 143–152. [32] J.C. Martinou, D.R. Green, Breaking the mitochondrial barrier, Nat. Rev., Mol. Cell Biol. 2 (2001) 63–67. [33] N.J. Waterhouse, J.E. Ricci, D.R. Green, And all of a sudden it's over: mitochondrial outer-membrane permeabilization in apoptosis, Biochimie 84 (2002) 113–121. [34] D.R. Green, G. Kroemer, The pathophysiology of mitochondrial cell death, Science 305 (2004) 626–629. [35] A. Rasola, P. Bernardi, The mitochondrial permeability transition pore and its involvement in cell death and in disease pathogenesis, Apoptosis 12 (2007) 815–833. [36] G. Kroemer, L. Galluzzi, C. Brenner, Mitochondrial membrane permeabilization in cell death, Physiol. Rev. 87 (2007) 99–163. [37] C.P. Baines, R.A. Kaiser, T. Sheiko, W.J. Craigen, J.D. Molkentin, Voltage-dependent anion channels are dispensable for mitochondrialdependent cell death, Nat. Cell Biol. 9 (2007) 550–555. [38] C.P. Baines, R.A. Kaiser, N.H. Purcell, N.S. Blair, H. Osinska, M.A. Hambleton, E.W. Brunskill, M.R. Sayen, R.A. Gottlieb, G.W. Dorn, J. Robbins, J.D. Molkentin, Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death, Nature 434 (2005) 658–662. [39] J.E. Kokoszka, K.G. Waymire, S.E. Levy, J.E. Sligh, J. Cai, D.P. Jones, G.R. MacGregor, D.C. Wallace, The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore, Nature 427 (2004) 461–465. [40] E. Basso, L. Fante, J. Fowlkes, V. Petronilli, M.A. Forte, P. Bernardi, Properties of the permeability transition pore in mitochondria devoid of Cyclophilin D, J. Biol. Chem. 280 (2005) 18558–18561. [41] T. Nakagawa, S. Shimizu, T. Watanabe, O. Yamaguchi, K. Otsu, H. Yamagata, H. Inohara, T. Kubo, Y. Tsujimoto, Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death, Nature 434 (2005) 652–658. [42] A.C. Schinzel, O. Takeuchi, Z. Huang, J.K. Fisher, Z. Zhou, J. Rubens, C. Hetz, N.N. Danial, M.A. Moskowitz, S.J. Korsmeyer, Cyclophilin D is a

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

[43]

[44] [45]

[46]

[47]

[48] [49]

[50]

[51]

[52]

[53]

[54]

[55]

[56]

[57]

[58]

[59] [60]

[61]

component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia, Proc. Natl. Acad. Sci. U. S. A. 102 (2005) 12005–12010. L. He, J.J. Lemasters, Regulated and unregulated mitochondrial permeability transition pores: a new paradigm of pore structure and function? FEBS Lett. 512 (2002) 1–7. Y. Tsujimoto, S. Shimizu, VDAC regulation by the Bcl-2 family of proteins, Cell Death Differ. 7 (2000) 1174–1181. A. Gross, J. Jockel, M.C. Wei, S.J. Korsmeyer, Enforced dimerization of BAX results in its translocation, mitochondrial dysfunction and apoptosis, EMBO J. 17 (1998) 3878–3885. G.J. Griffiths, L. Dubrez, C.P. Morgan, N.A. Jones, J. Whitehouse, B.M. Corfe, C. Dive, J.A. Hickman, Cell damage-induced conformational changes of the pro-apoptotic protein Bak in vivo precede the onset of apoptosis, J. Cell Biol. 144 (1999) 903–914. A. Antignani, R.J. Youle, How do Bax and Bak lead to permeabilization of the outer mitochondrial membrane? Curr. Opin. Cell Biol. 18 (2006) 685–689. S. Lucken-Ardjomande, J.C. Martinou, Newcomers in the process of mitochondrial permeabilization, J. Cell Sci. 118 (2005) 473–483. S.N. Willis, J.I. Fletcher, T. Kaufmann, M.F. van Delft, L. Chen, P.E. Czabotar, H. Ierino, E.F. Lee, W.D. Fairlie, P. Bouillet, A. Strasser, R.M. Kluck, J.M. Adams, D.C. Huang, Apoptosis initiated when BH3 ligands engage multiple Bcl-2 homologs, not Bax or Bak, Science 315 (2007) 856–859. M.C. Wei, W.X. Zong, E.H. Cheng, T. Lindsten, V. Panoutsakopoulou, A.J. Ross, K.A. Roth, G.R. MacGregor, C.B. Thompson, S.J. Korsmeyer, Proapoptotic BAX and BAK: a requisite gateway to mitochondrial dysfunction and death, Science 292 (2001) 727–730. L. Scorrano, S.A. Oakes, J.T. Opferman, E.H. Cheng, M.D. Sorcinelli, T. Pozzan, S.J. Korsmeyer, BAX and BAK regulation of endoplasmic reticulum Ca2+: a control point for apoptosis, Science 300 (2003) 135–139. X. Luo, I. Budihardjo, H. Zou, C. Slaughter, X. Wang, Bid, a Bcl2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors, Cell 94 (1998) 481–490. F. Gonzalvez, F. Pariselli, P. Dupaigne, I. Budihardjo, M. Lutter, B. Antonsson, P. Diolez, S. Manon, J.C. Martinou, M. Goubern, X. Wang, S. Bernard, P.X. Petit, tBid interaction with cardiolipin primarily orchestrates mitochondrial dysfunctions and subsequently activates Bax and Bak, Cell Death Differ. 12 (2005) 614–626. G. Basanez, J.C. Sharpe, J. Galanis, T.B. Brandt, J.M. Hardwick, J. Zimmerberg, Bax-type apoptotic proteins porate pure lipid bilayers through a mechanism sensitive to intrinsic monolayer curvature, J. Biol. Chem. 277 (2002) 49360–49365. A. Goonesinghe, E.S. Mundy, M. Smith, R. Khosravi-Far, J.C. Martinou, M.D. Esposti, Pro-apoptotic Bid induces membrane perturbation by inserting selected lysolipids into the bilayer, Biochem. J. 387 (2005) 109–118. L. Scorrano, M. Ashiya, K. Buttle, S. Weiler, S.A. Oakes, C.A. Mannella, S.J. Korsmeyer, A distinct pathway remodels mitochondrial cristae and mobilizes cytochrome c during apoptosis, Dev. Cell 2 (2002) 55–67. M. Germain, J.P. Mathai, H.M. McBride, G.C. Shore, Endoplasmic reticulum BIK initiates DRP1-regulated remodelling of mitochondrial cristae during apoptosis, EMBO J. 24 (2005) 1546–1556. T. Garofalo, A.M. Giammarioli, R. Misasi, A. Tinari, V. Manganelli, L. Gambardella, A. Pavan, W. Malorni, M. Sorice, Lipid microdomains contribute to apoptosis-associated modifications of mitochondria in T cells, Cell Death Differ. 12 (2005) 1378–1389. G.M. Cereghetti, L. Scorrano, The many shapes of mitochondrial death, Oncogene 25 (2006) 4717–4724. P.A. Parone, D.I. James, C.S. Da, Y. Mattenberger, O. Donze, F. Barja, J.C. Martinou, Inhibiting the mitochondrial fission machinery does not prevent Bax/Bak-dependent apoptosis, Mol. Cell. Biol. 26 (2006) 7397–7408. P. Agostinis, E. Buytaert, H. Breyssens, N. Hendrickx, Regulatory pathways in photodynamic therapy induced apoptosis, Photochem. Photobiol. Sci. 3 (2004) 721–729.

103

[62] R.D. Almeida, B.J. Manadas, A.P. Carvalho, C.B. Duarte, Intracellular signaling mechanisms in photodynamic therapy, Biochim. Biophys. Acta 1704 (2004) 59–86. [63] A.C. Moor, Signaling pathways in cell death and survival after photodynamic therapy, J. Photochem. Photobiol., B 57 (2000) 1–13. [64] P. Bernardi, A. Krauskopf, E. Basso, V. Petronilli, E. Blachly-Dyson, L.F. Di, M.A. Forte, The mitochondrial permeability transition from in vitro artifact to disease target, FEBS J. 273 (2006) 2077–2099. [65] P. Costantini, A.S. Belzacq, H.L. Vieira, N. Larochette, M.A. de Pablo, N. Zamzami, S.A. Susin, C. Brenner, G. Kroemer, Oxidation of a critical thiol residue of the adenine nucleotide translocator enforces Bcl-2-independent permeability transition pore opening and apoptosis, Oncogene 19 (2000) 307–314. [66] A.S. Belzacq, E. Jacotot, H.L. Vieira, D. Mistro, D.J. Granville, Z. Xie, J.C. Reed, G. Kroemer, C. Brenner, Apoptosis induction by the photosensitizer verteporfin: identification of mitochondrial adenine nucleotide translocator as a critical target, Cancer Res. 61 (2001) 1260–1264. [67] D. Kessel, M. Antolovich, K.M. Smith, The role of the peripheral benzodiazepine receptor in the apoptotic response to photodynamic therapy, Photochem. Photobiol. 74 (2001) 346–349. [68] T.J. Dougherty, A.B. Sumlin, W.R. Greco, K.R. Weishaupt, L.A. Vaughan, R.K. Pandey, The role of the peripheral benzodiazepine receptor in photodynamic activity of certain pyropheophorbide ether photosensitizers: albumin site II as a surrogate marker for activity, Photochem. Photobiol. 76 (2002) 91–97. [69] I.E. Furre, S. Shahzidi, Z. Luksiene, M.T. Moller, E. Borgen, J. Morgan, K. Tkacz-Stachowska, J.M. Nesland, Q. Peng, Targeting PBR by hexaminolevulinate-mediated photodynamic therapy induces apoptosis through translocation of apoptosis-inducing factor in human leukemia cells, Cancer Res. 65 (2005) 11051–11060. [70] M. Lam, N.L. Oleinick, A.L. Nieminen, Photodynamic therapy-induced apoptosis in epidermoid carcinoma cells. Reactive oxygen species and mitochondrial inner membrane permeabilization, J. Biol. Chem. 276 (2001) 47379–47386. [71] T. Minamikawa, A. Sriratana, D.A. Williams, D.N. Bowser, J.S. Hill, P. Nagley, Chloromethyl-X-rosamine (MitoTracker Red) photosensitises mitochondria and induces apoptosis in intact human cells, J. Cell Sci. 112 (Pt 14) (1999) 2419–2430. [72] R. Chaloupka, P.X. Petit, N. Israel, F. Sureau, Over-expression of Bcl-2 does not protect cells from hypericin photo-induced mitochondrial membrane depolarization, but delays subsequent events in the apoptotic pathway, FEBS Lett. 462 (1999) 295–301. [73] C. Salet, G. Moreno, F. Ricchelli, P. Bernardi, Singlet oxygen produced by photodynamic action causes inactivation of the mitochondrial permeability transition pore, J. Biol. Chem. 272 (1997) 21938–21943. [74] A.P. Halestrap, K.Y. Woodfield, C.P. Connern, Oxidative stress, thiol reagents, and membrane potential modulate the mitochondrial permeability transition by affecting nucleotide binding to the adenine nucleotide translocase, J. Biol. Chem. 272 (1997) 3346–3354. [75] G. Moreno, K. Poussin, F. Ricchelli, C. Salet, The effects of singlet oxygen produced by photodynamic action on the mitochondrial permeability transition differ in accordance with the localization of the sensitizer, Arch. Biochem. Biophys. 386 (2001) 243–250. [76] Z. Ji, G. Yang, V. Vasovic, B. Cunderlikova, Z. Suo, J.M. Nesland, Q. Peng, Subcellular localization pattern of protoporphyrin IX is an important determinant for its photodynamic efficiency of human carcinoma and normal cell lines, J. Photochem. Photobiol., B 84 (2006) 213–220. [77] V. Petronilli, G. Miotto, M. Canton, M. Brini, R. Colonna, P. Bernardi, L.F. Di, Transient and long-lasting openings of the mitochondrial permeability transition pore can be monitored directly in intact cells by changes in mitochondrial calcein fluorescence, Biophys. J. 76 (1999) 725–734. [78] S.M. Chiu, N.L. Oleinick, Dissociation of mitochondrial depolarization from cytochrome c release during apoptosis induced by photodynamic therapy, Br. J. Cancer 84 (2001) 1099–1106. [79] M.G. Lum, P. Nagley, Two phases of signalling between mitochondria during apoptosis leading to early depolarisation and delayed cytochrome c release, J. Cell Sci. 116 (2003) 1437–1447.

104

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107

[80] R.L. Morris, K. Azizuddin, M. Lam, J. Berlin, A.L. Nieminen, M.E. Kenney, A.C. Samia, C. Burda, N.L. Oleinick, Fluorescence resonance energy transfer reveals a binding site of a photosensitizer for photodynamic therapy, Cancer Res. 63 (2003) 5194–5197. [81] S. Orrenius, V. Gogvadze, B. Zhivotovsky, Mitochondrial oxidative stress: implications for cell death, Annu. Rev. Pharmacol. Toxicol. 47 (2007) 143–183. [82] T. Kriska, W. Korytowski, A.W. Girotti, Hyperresistance to photosensitized lipid peroxidation and apoptotic killing in 5-aminolevulinate-treated tumor cells overexpressing mitochondrial GPX4, Free Radical Biol. Med. 33 (2002) 1389–1402. [83] T. Kriska, W. Korytowski, A.W. Girotti, Role of mitochondrial cardiolipin peroxidation in apoptotic photokilling of 5-aminolevulinatetreated tumor cells, Arch. Biochem. Biophys. 433 (2005) 435–446. [84] A. Vantieghem, Y. Xu, W. Declercq, P. Vandenabeele, G. Denecker, J.R. Vandenheede, W. Merlevede, P.A. de Witte, P. Agostinis, Different pathways mediate cytochrome c release after photodynamic therapy with hypericin, Photochem. Photobiol. 74 (2001) 133–142. [85] J.Y. Matroule, C.M. Carthy, D.J. Granville, O. Jolois, D.W. Hunt, J. Piette, Mechanism of colon cancer cell apoptosis mediated by pyropheophorbide—A methylester photosensitization, Oncogene 20 (2001) 4070–4084. [86] M.H. Teiten, S. Marchal, M.A. D'Hallewin, F. Guillemin, L. Bezdetnaya, Primary photodamage sites and mitochondrial events after Foscan photosensitization of MCF-7 human breast cancer cells, Photochem. Photobiol. 78 (2003) 9–14. [87] U. De Marchi, S. Campello, I. Szabo, F. Tombola, J.C. Martinou, M. Zoratti, Bax does not directly participate in the Ca(2+)-induced permeability transition of isolated mitochondria, J. Biol. Chem. 279 (2004) 37415–37422. [88] T. Minamikawa, D.A. Williams, D.N. Bowser, P. Nagley, Mitochondrial permeability transition and swelling can occur reversibly without inducing cell death in intact human cells, Exp. Cell Res. 246 (1999) 26–37. [89] S.M. Chiu, L.Y. Xue, J. Usuda, K. Azizuddin, N.L. Oleinick, Bax is essential for mitochondrion-mediated apoptosis but not for cell death caused by photodynamic therapy, Br. J. Cancer 89 (2003) 1590–1597. [90] D.J. Granville, J.R. Shaw, S. Leong, C.M. Carthy, P. Margaron, D.W. Hunt, B.M. McManus, Release of cytochrome c, Bax migration, Bid cleavage, and activation of caspases 2, 3, 6, 7, 8, and 9 during endothelial cell apoptosis, Am. J. Pathol. 155 (1999) 1021–1025. [91] D.J. Granville, B.A. Cassidy, D.O. Ruehlmann, J.C. Choy, C. Brenner, G. Kroemer, B.C. van, P. Margaron, D.W. Hunt, B.M. McManus, Mitochondrial release of apoptosis-inducing factor and cytochrome c during smooth muscle cell apoptosis, Am. J. Pathol. 159 (2001) 305–311. [92] D. Kessel, M. Castelli, Evidence that bcl-2 is the target of three photosensitizers that induce a rapid apoptotic response, Photochem. Photobiol. 74 (2001) 318–322. [93] L.Y. Xue, S.M. Chiu, N.L. Oleinick, Photochemical destruction of the Bcl-2 oncoprotein during photodynamic therapy with the phthalocyanine photosensitizer Pc 4, Oncogene 20 (2001) 3420–3427. [94] J. Usuda, S.M. Chiu, E.S. Murphy, M. Lam, A.L. Nieminen, N.L. Oleinick, Domain-dependent photodamage to Bcl-2. A membrane anchorage region is needed to form the target of phthalocyanine photosensitization, J. Biol. Chem. 278 (2003) 2021–2029. [95] H.R. Kim, Y. Luo, G. Li, D. Kessel, Enhanced apoptotic response to photodynamic therapy after bcl-2 transfection, Cancer Res. 59 (1999) 3429–3432. [96] M. Srivastava, N. Ahmad, S. Gupta, H. Mukhtar, Involvement of Bcl-2 and Bax in photodynamic therapy-mediated apoptosis. Antisense Bcl-2 oligonucleotide sensitizes RIF 1 cells to photodynamic therapy apoptosis, J. Biol. Chem. 276 (2001) 15481–15488. [97] D. Kessel, M. Castelli, J.J. Reiners Jr., Apoptotic response to photodynamic therapy versus the Bcl-2 antagonist HA14-1, Photochem. Photobiol. 76 (2002) 314–319. [98] J.L. Wang, D. Liu, Z.J. Zhang, S. Shan, X. Han, S.M. Srinivasula, C.M. Croce, E.S. Alnemri, Z. Huang, Structure-based discovery of an organic compound that binds Bcl-2 protein and induces apoptosis of tumor cells, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 7124–7129.

[99] F. Manero, F. Gautier, T. Gallenne, N. Cauquil, D. Gree, P.F. Cartron, O. Geneste, R. Gree, F.M. Vallette, P. Juin, The small organic compound HA14-1 prevents Bcl-2 interaction with Bax to sensitize malignant glioma cells to induction of cell death, Cancer Res. 66 (2006) 2757–2764. [100] J. Usuda, K. Azizuddin, S.M. Chiu, N.L. Oleinick, Association between the photodynamic loss of Bcl-2 and the sensitivity to apoptosis caused by phthalocyanine photodynamic therapy, Photochem. Photobiol. 78 (2003) 1–8. [101] J. An, Y. Chen, Z. Huang, Critical upstream signals of cytochrome C release induced by a novel Bcl-2 inhibitor, J. Biol. Chem. 279 (2004) 19133–19140. [102] D. Kessel, M. Castelli, J.J. Reiners, Ruthenium red-mediated suppression of Bcl-2 loss and Ca(2+) release initiated by photodamage to the endoplasmic reticulum: scavenging of reactive oxygen species, Cell Death Differ. 12 (2005) 502–511. [103] P. Pinton, R. Rizzuto, Bcl-2 and Ca2+ homeostasis in the endoplasmic reticulum, Cell Death Differ. 13 (2006) 1409–1418. [104] S. Orrenius, B. Zhivotovsky, P. Nicotera, Regulation of cell death: the calcium-apoptosis link, Nat. Rev., Mol. Cell Biol. 4 (2003) 552–565. [105] M.J. Berridge, P. Lipp, M.D. Bootman, The versatility and universality of calcium signalling, Nat. Rev., Mol. Cell Biol. 1 (2000) 11–21. [106] P. Pinton, D. Ferrari, P. Magalhaes, K. Schulze-Osthoff, V.F. Di, T. Pozzan, R. Rizzuto, Reduced loading of intracellular Ca(2+) stores and downregulation of capacitative Ca(2+) influx in Bcl-2-overexpressing cells, J. Cell Biol. 148 (2000) 857–862. [107] R. Foyouzi-Youssefi, S. Arnaudeau, C. Borner, W.L. Kelley, J. Tschopp, D.P. Lew, N. Demaurex, K.H. Krause, Bcl-2 decreases the free Ca2+ concentration within the endoplasmic reticulum, Proc. Natl. Acad. Sci. U. S. A. 97 (2000) 5723–5728. [108] V. Dolgachev, M.S. Farooqui, O.I. Kulaeva, M.A. Tainsky, B. Nagy, K. Hanada, D. Separovic, De novo ceramide accumulation due to inhibition of its conversion to complex sphingolipids in apoptotic photosensitized cells, J. Biol. Chem. 279 (2004) 23238–23249. [109] J.Y. Matroule, G. Bonizzi, P. Morliere, N. Paillous, R. Santus, V. Bours, J. Piette, Pyropheophorbide-a methyl ester-mediated photosensitization activates transcription factor NF-kappaB through the interleukin-1 receptor-dependent signaling pathway, J. Biol. Chem. 274 (1999) 2988–3000. [110] B. Wispriyono, E. chmelz, H. Pelayo, K. Hanada, D. Separovic, A role for the de novo sphingolipids in apoptosis of photosensitized cells, Exp. Cell Res. 279 (2002) 153–165. [111] T.A. Taha, T.D. Mullen, L.M. Obeid, A house divided: ceramide, sphingosine, and sphingosine-1-phosphate in programmed cell death, Biochim. Biophys. Acta 1758 (2006) 2027–2036. [112] N. Hendrickx, M. Dewaele, E. Buytaert, G. Marsboom, S. Janssens, B.M. Van, J.R. Vandenheede, P. de Witte, P. Agostinis, Targeted inhibition of p38alpha MAPK suppresses tumor-associated endothelial cell migration in response to hypericin-based photodynamic therapy, Biochem. Biophys. Res. Commun. 337 (2005) 928–935. [113] T. Cirman, K. Oresic, G.D. Mazovec, V. Turk, J.C. Reed, R.M. Myers, G.S. Salvesen, B. Turk, Selective disruption of lysosomes in HeLa cells triggers apoptosis mediated by cleavage of Bid by multiple papain-like lysosomal cathepsins, J. Biol. Chem. 279 (2004) 3578–3587. [114] V. Stoka, B. Turk, S.L. Schendel, T.H. Kim, T. Cirman, S.J. Snipas, L.M. Ellerby, D. Bredesen, H. Freeze, M. Abrahamson, D. Bromme, S. Krajewski, J.C. Reed, X.M. Yin, V. Turk, G.S. Salvesen, Lysosomal protease pathways to apoptosis. Cleavage of bid, not pro-caspases, is the most likely route, J. Biol. Chem. 276 (2001) 3149–3157. [115] J.J. Reiners Jr., J.A. Caruso, P. Mathieu, B. Chelladurai, X.M. Yin, D. Kessel, Release of cytochrome c and activation of pro-caspase-9 following lysosomal photodamage involves Bid cleavage, Cell Death Differ. 9 (2002) 934–944. [116] S. Ichinose, J. Usuda, T. Hirata, T. Inoue, K. Ohtani, S. Maehara, M. Kubota, K. Imai, Y. Tsunoda, Y. Kuroiwa, K. Yamada, H. Tsutsui, K. Furukawa, T. Okunaka, N.L. Oleinick, H. Kato, Lysosomal cathepsin initiates apoptosis, which is regulated by photodamage to Bcl-2 at mitochondria in photodynamic therapy using a novel photosensitizer, ATX-s10 (Na), Int. J. Oncol. 29 (2006) 349–355.

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107 [117] M. Nonaka, H. Ikeda, T. Inokuchi, Inhibitory effect of heat shock protein 70 on apoptosis induced by photodynamic therapy in vitro, Photochem. Photobiol. 79 (2004) 94–98. [118] S. Wong, M. Luna, A. Ferrario, C.J. Gomer, CHOP activation by photodynamic therapy increases treatment induced photosensitization, Lasers Surg. Med. 35 (2004) 336–341. [119] E. Buytaert, J.Y. Matroule, S. Durinck, P. Close, S. Kocanova, J.R. Vandenheede, P.A. de Witte, J. Piette, and P. Agostinis, Molecular effectors and modulators of hypericin-mediated cell death in bladder cancer cells, (submitted for publication). [120] S. Mitra, E.M. Goren, J.G. Frelinger, T.H. Foster, Activation of heat shock protein 70 promoter with meso-tetrahydroxyphenyl chlorin photodynamic therapy reported by green fluorescent protein in vitro and in vivo, Photochem. Photobiol. 78 (2003) 615–622. [121] J. Nylandsted, M. Gyrd-Hansen, A. Danielewicz, N. Fehrenbacher, U. Lademann, M. Hoyer-Hansen, E. Weber, G. Multhoff, M. Rohde, M. Jaattela, Heat shock protein 70 promotes cell survival by inhibiting lysosomal membrane permeabilization, J. Exp. Med. 200 (2004) 425–435. [122] H.B. Lee, A.S. Ho, S.H. Teo, p53 Status does not affect photodynamic cell killing induced by hypericin, Cancer Chemother. Pharmacol. 58 (2006) 91–98. [123] M. Weller, M. Trepel, C. Grimmel, M. Schabet, D. Bremen, S. Krajewski, J.C. Reed, Hypericin-induced apoptosis of human malignant glioma cells is light-dependent, independent of bcl-2 expression, and does not require wild-type p53, Neurol. Res. 19 (1997) 459–470. [124] D.S. Lim, S.M. Bae, S.Y. Kwak, E.K. Park, J.K. Kim, S.J. Han, C.H. Oh, C.H. Lee, W.Y. Lee, W.S. Ahn, Adenovirus-mediated p53 treatment enhances photodynamic antitumor response, Hum. Gene Ther. 17 (2006) 347–352. [125] J. Zawacka-Pankau, N. Issaeva, S. Hossain, A. Pramanik, G. Selivanova, A.J. Podhajska, Protoporphyrin IX interacts with wild-type p53 protein in vitro and induces cell death of human colon cancer cells in a p53dependent and-independent manner, J. Biol. Chem. 282 (2007) 2466–2472. [126] M. Mitsunaga, A. Tsubota, K. Nariai, Y. Namiki, M. Sumi, T. Yoshikawa, K. Fujise, Early apoptosis and cell death induced by ATX-S10Na (II)mediated photodynamic therapy are Bax- and p53-dependent in human colon cancer cells, World J. Gastroenterol. 13 (2007) 692–698. [127] J.M. Adams, S. Cory, The Bcl-2 apoptotic switch in cancer development and therapy, Oncogene 26 (2007) 1324–1337. [128] L.Y. Xue, S.M. Chiu, A. Fiebig, D.W. Andrews, N.L. Oleinick, Photodamage to multiple Bcl-xL isoforms by photodynamic therapy with the phthalocyanine photosensitizer Pc 4, Oncogene 22 (2003) 9197–9204. [129] M. Castelli, J.J. Reiners, D. Kessel, A mechanism for the proapoptotic activity of ursodeoxycholic acid: effects on Bcl-2 conformation, Cell Death Differ. 11 (2004) 906–914. [130] A. Vantieghem, Y. Xu, Z. Assefa, J. Piette, J.R. Vandenheede, W. Merlevede, P.A. de Witte, P. Agostinis, Phosphorylation of Bcl-2 in G2/M phase-arrested cells following photodynamic therapy with hypericin involves a CDK1-mediated signal and delays the onset of apoptosis, J. Biol. Chem. 277 (2002) 37718–37731. [131] T. Yokota, H. Ikeda, T. Inokuchi, K. Sano, T. Koji, Enhanced cell death in NR-S1 tumor by photodynamic therapy: possible involvement of Fas and Fas ligand system, Lasers Surg. Med. 26 (2000) 449–460. [132] B. Chen, T. Roskams, Y. Xu, P. Agostinis, P.A. de Witte, Photodynamic therapy with hypericin induces vascular damage and apoptosis in the RIF-1 mouse tumor model, Int. J. Cancer 98 (2002) 284–290. [133] D.A. Bellnier, Potentiation of photodynamic therapy in mice with recombinant human tumor necrosis factor-alpha, J. Photochem. Photobiol., B 8 (1991) 203–210. [134] I. Reiter, G. Schwamberger, B. Krammer, Effect of photodynamic pretreatment on the susceptibility of murine tumor cells to macrophage antitumor mechanisms, Photochem. Photobiol. 66 (1997) 384–388. [135] D.J. Granville, H. Jiang, B.M. McManus, D.W. Hunt, Fas ligand and TRAIL augment the effect of photodynamic therapy on the induction of apoptosis in JURKAT cells, Int. Immunopharmacol. 1 (2001) 1831–1840.

105

[136] Y. Wu, D. Xing, W.R. Chen, Single cell FRET imaging for determination of pathway of tumor cell apoptosis induced by photofrin-PDT, Cell Cycle 5 (2006) 729–734. [137] B. Nagy, W.C. Yeh, T.W. Mak, S.M. Chiu, D. Separovic, FADD null mouse embryonic fibroblasts undergo apoptosis after photosensitization with the silicon phthalocyanine Pc 4, Arch. Biochem. Biophys. 385 (2001) 194–202. [138] Z. Assefa, A. Vantieghem, W. Declercq, P. Vandenabeele, J.R. Vandenheede, W. Merlevede, P. de Witte, P. Agostinis, The activation of the c-Jun N-terminal kinase and p38 mitogen-activated protein kinase signaling pathways protects HeLa cells from apoptosis following photodynamic therapy with hypericin, J. Biol. Chem. 274 (1999) 8788–8796. [139] H. Takahashi, Y. Itoh, Y. Miyauchi, S. Nakajima, I. Sakata, A. IshidaYamamoto, H. Iizuka, Activation of two caspase cascades, caspase 8/3/6 and caspase 9/3/6, during photodynamic therapy using a novel photosensitizer, ATX-S10(Na), in normal human keratinocytes, Arch. Dermatol. Res. 295 (2003) 242–248. [140] N. Ahmad, S. Gupta, D.K. Feyes, H. Mukhtar, Involvement of Fas (APO1/CD-95) during photodynamic-therapy-mediated apoptosis in human epidermoid carcinoma A431 cells, J. Invest. Dermatol. 115 (2000) 1041–1046. [141] S. Zhuang, M.C. Lynch, I.E. Kochevar, Caspase-8 mediates caspase-3 activation and cytochrome c release during singlet oxygen-induced apoptosis of HL-60 cells, Exp. Cell Res. 250 (1999) 203–212. [142] S.M. Ali, S.K. Chee, G.Y. Yuen, M. Olivo, Hypericin induced death receptor-mediated apoptosis in photoactivated tumor cells, Int. J. Mol. Med. 9 (2002) 601–616. [143] S.M. Ali, S.K. Chee, G.Y. Yuen, M. Olivo, Photodynamic therapy induced Fas-mediated apoptosis in human carcinoma cells, Int. J. Mol. Med. 9 (2002) 257–270. [144] D.J. Granville, C.M. Carthy, H. Jiang, G.C. Shore, B.M. McManus, D.W. Hunt, Rapid cytochrome c release, activation of caspases 3, 6, 7 and 8 followed by Bap31 cleavage in HeLa cells treated with photodynamic therapy, FEBS Lett. 437 (1998) 5–10. [145] D.J. Granville, D.O. Ruehlmann, J.C. Choy, B.A. Cassidy, D.W. Hunt, B.C. van, B.M. McManus, Bcl-2 increases emptying of endoplasmic reticulum Ca2+ stores during photodynamic therapy-induced apoptosis, Cell Calcium 30 (2001) 343–350. [146] O. Inanami, A. Yoshito, K. Takahashi, W. Hiraoka, M. Kuwabara, Effects of BAPTA-AM and forskolin on apoptosis and cytochrome c release in photosensitized Chinese hamster V79 cells, Photochem. Photobiol. 70 (1999) 650–655. [147] X. Ding, Q. Xu, F. Liu, P. Zhou, Y. Gu, J. Zeng, J. An, W. Dai, X. Li, Hematoporphyrin monomethyl ether photodynamic damage on HeLa cells by means of reactive oxygen species production and cytosolic free calcium concentration elevation, Cancer Lett. 216 (2004) 43–54. [148] M. Ogata, O. Inanami, M. Nakajima, T. Nakajima, W. Hiraoka, M. Kuwabara, Ca(2+)-dependent and caspase-3-independent apoptosis caused by damage in Golgi apparatus due to 2,4,5,7-tetrabromorhodamine 123 bromide-induced photodynamic effects, Photochem. Photobiol. 78 (2003) 241–247. [149] N.K. Mak, K.M. Li, W.N. Leung, R.N. Wong, D.P. Huang, M.L. Lung, Y.K. Lau, C.K. Chang, Involvement of both endoplasmic reticulum and mitochondria in photokilling of nasopharyngeal carcinoma cells by the photosensitizer Zn-BC-AM, Biochem. Pharmacol. 68 (2004) 2387–2396. [150] M. Hermes-Lima, How do Ca2+ and 5-aminolevulinic acid-derived oxyradicals promote injury to isolated mitochondria? Free Radical Biol. Med. 19 (1995) 381–390. [151] Z. Zhou, H. Yang, Z. Zhang, Role of calcium in phototoxicity of 2-butylamino-2-demethoxy-hypocrellin A to human gastric cancer MGC803 cells, Biochim. Biophys. Acta 1593 (2003) 191–200. [152] L.C. Penning, M.H. Rasch, E. Ben-Hur, T.M. Dubbelman, A.C. Havelaar, Z.J. Van der, S.J. Van, A role for the transient increase of cytoplasmic free calcium in cell rescue after photodynamic treatment, Biochim. Biophys. Acta 1107 (1992) 255–260. [153] L.C. Penning, M.J. Keirse, J. VanSteveninck, T.M. Dubbelman, Ca(2+)mediated prostaglandin E2 induction reduces haematoporphyrin-

106

[154]

[155]

[156]

[157]

[158]

[159]

[160]

[161]

[162]

[163]

[164]

[165] [166]

[167]

[168]

[169]

[170]

[171]

[172]

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107 derivative-induced cytotoxicity of T24 human bladder transitional carcinoma cells in vitro, Biochem. J. 292 (Pt 1) (1993) 237–240. A. Hubmer, A. Hermann, K. Uberriegler, B. Krammer, Role of calcium in photodynamically induced cell damage of human fibroblasts, Photochem. Photobiol. 64 (1996) 211–215. M.J. Berridge, M.D. Bootman, H.L. Roderick, Calcium signalling: dynamics, homeostasis and remodelling, Nat. Rev., Mol. Cell Biol. 4 (2003) 517–529. H.J. Chae, S.W. Chae, K.H. Weon, J.S. Kang, H.R. Kim, Signal transduction of thapsigargin-induced apoptosis in osteoblast, Bone 25 (1999) 453–458. Z. Pan, D. Damron, A.L. Nieminen, M.B. Bhat, J. Ma, Depletion of intracellular Ca2+ by caffeine and ryanodine induces apoptosis of Chinese hamster ovary cells transfected with ryanodine receptor, J. Biol. Chem. 275 (2000) 19978–19984. Z. Pan, M.B. Bhat, A.L. Nieminen, J. Ma, Synergistic movements of Ca(2+) and Bax in cells undergoing apoptosis, J. Biol. Chem. 276 (2001) 32257–32263. M.L. Agarwal, H.E. Larkin, S.I. Zaidi, H. Mukhtar, N.L. Oleinick, Phospholipase activation triggers apoptosis in photosensitized mouse lymphoma cells, Cancer Res. 53 (1993) 5897–5902. M.H. Rasch, K. Tijssen, J.W. Lagerberg, W.E. Corver, J. VanSteveninck, T.M. Dubbelman, The role of protein kinase C activity in the killing of Chinese hamster ovary cells by ionizing radiation and photodynamic treatment, Photochem. Photobiol. 66 (1997) 209–213. A. Gugliucci, L. Ranzato, L. Scorrano, R. Colonna, V. Petronilli, C. Cusan, M. Prato, M. Mancini, F. Pagano, P. Bernardi, Mitochondria are direct targets of the lipoxygenase inhibitor MK886. A strategy for cell killing by combined treatment with MK886 and cyclooxygenase inhibitors, J. Biol. Chem. 277 (2002) 31789–31795. R.D. Almeida, E.R. Gomes, A.P. Carvalho, C.B. Duarte, Calpains are activated by photodynamic therapy but do not contribute to apoptotic tumor cell death, Cancer Lett. 216 (2004) 183–189. D. Grebenova, K. Kuzelova, K. Smetana, M. Pluskalova, H. Cajthamlova, I. Marinov, O. Fuchs, J. Soucek, P. Jarolim, Z. Hrkal, Mitochondrial and endoplasmic reticulum stress-induced apoptotic pathways are activated by 5-aminolevulinic acid-based photodynamic therapy in HL60 leukemia cells, J. Photochem. Photobiol., B 69 (2003) 71–85. D.G. Breckenridge, M. Stojanovic, R.C. Marcellus, G.C. Shore, Caspase cleavage product of BAP31 induces mitochondrial fission through endoplasmic reticulum calcium signals, enhancing cytochrome c release to the cytosol, J. Cell Biol. 160 (2003) 1115–1127. M. Schroder, R.J. Kaufman, ER stress and the unfolded protein response, Mutat. Res. 569 (2005) 29–63. H. Zinszner, M. Kuroda, X. Wang, N. Batchvarova, R.T. Lightfoot, H. Remotti, J.L. Stevens, D. Ron, CHOP is implicated in programmed cell death in response to impaired function of the endoplasmic reticulum, Genes Dev. 12 (1998) 982–995. E. Szegezdi, S.E. Logue, A.M. Gorman, A. Samali, Mediators of endoplasmic reticulum stress-induced apoptosis, EMBO Rep. 7 (2006) 880–885. K.D. McCullough, J.L. Martindale, L.O. Klotz, T.Y. Aw, N.J. Holbrook, Gadd153 sensitizes cells to endoplasmic reticulum stress by downregulating Bcl2 and perturbing the cellular redox state, Mol. Cell. Biol. 21 (2001) 1249–1259. X.Z. Wang, D. Ron, Stress-induced phosphorylation and activation of the transcription factor CHOP (GADD153) by p38 MAP Kinase, Science 272 (1996) 1347–1349. Y. Tan, N. Dourdin, C. Wu, V.T. De, J.S. Elce, P.A. Greer, Ubiquitous calpains promote caspase-12 and JNK activation during endoplasmic reticulum stress-induced apoptosis, J. Biol. Chem. 281 (2006) 16016–16024. N. Morishima, K. Nakanishi, H. Takenouchi, T. Shibata, Y. Yasuhiko, An endoplasmic reticulum stress-specific caspase cascade in apoptosis. Cytochrome c-independent activation of caspase-9 by caspase-12, J. Biol. Chem. 277 (2002) 34287–34294. R.V. Rao, S. Castro-Obregon, H. Frankowski, M. Schuler, V. Stoka, R.G. del, D.E. Bredesen, H.M. Ellerby, Coupling endoplasmic reticulum stress

[173] [174]

[175]

[176]

[177]

[178]

[179] [180] [181] [182]

[183]

[184]

[185]

[186] [187] [188]

[189]

[190]

[191]

[192]

[193]

[194]

to the cell death program. An Apaf-1-independent intrinsic pathway, J. Biol. Chem. 277 (2002) 21836–21842. Y. Ma, L.M. Hendershot, The role of the unfolded protein response in tumour development: friend or foe? Nat. Rev., Cancer 4 (2004) 966–977. I.E. Furre, M.T. Moller, S. Shahzidi, J.M. Nesland, Q. Peng, Involvement of both caspase-dependent and-independent pathways in apoptotic induction by hexaminolevulinate-mediated photodynamic therapy in human lymphoma cells, Apoptosis 11 (2006) 2031–2042. D. Arnoult, B. Gaume, M. Karbowski, J.C. Sharpe, F. Cecconi, R.J. Youle, Mitochondrial release of AIF and EndoG requires caspase activation downstream of Bax/Bak-mediated permeabilization, EMBO J. 22 (2003) 4385–4399. S.W. Yu, H. Wang, M.F. Poitras, C. Coombs, W.J. Bowers, H.J. Federoff, G.G. Poirier, T.M. Dawson, V.L. Dawson, Mediation of poly(ADPribose) polymerase-1-dependent cell death by apoptosis-inducing factor, Science 297 (2002) 259–263. R.T. Uren, G. Dewson, C. Bonzon, T. Lithgow, D.D. Newmeyer, R.M. Kluck, Mitochondrial release of pro-apoptotic proteins: electrostatic interactions can hold cytochrome c but not Smac/DIABLO to mitochondrial membranes, J. Biol. Chem. 280 (2005) 2266–2274. B. Levine, D.J. Klionsky, Development by self-digestion: molecular mechanisms and biological functions of autophagy, Dev. Cell 6 (2004) 463–477. Y. Tsujimoto, S. Shimizu, Another way to die: autophagic programmed cell death, Cell Death. Differ. 12 (Suppl. 2) (2005) 1528–1534. J. Debnath, E.H. Baehrecke, G. Kroemer, Does autophagy contribute to cell death? Autophagy 1 (2005) 66–74. D. Gozuacik, A. Kimchi, Autophagy as a cell death and tumor suppressor mechanism, Oncogene 23 (2004) 2891–2906. S. Shimizu, T. Kanaseki, N. Mizushima, T. Mizuta, S. rakawa-Kobayashi, C.B. Thompson, Y. Tsujimoto, Role of Bcl-2 family proteins in a nonapoptotic programmed cell death dependent on autophagy genes, Nat. Cell Biol. 6 (2004) 1221–1228. L. Yu, A. Alva, H. Su, P. Dutt, E. Freundt, S. Welsh, E.H. Baehrecke, M.J. Lenardo, Regulation of an ATG7-beclin 1 program of autophagic cell death by caspase-8, Science 304 (2004) 1500–1502. A. Petiot, E. Ogier-Denis, E.F. Blommaart, A.J. Meijer, P. Codogno, Distinct classes of phosphatidylinositol 3′-kinases are involved in signaling pathways that control macroautophagy in HT-29 cells, J. Biol. Chem. 275 (2000) 992–998. Y. Kondo, T. Kanzawa, R. Sawaya, S. Kondo, The role of autophagy in cancer development and response to therapy, Nat. Rev., Cancer 5 (2005) 726–734. D.Kessel, Protection of Bcl-2 by salubrinal, Biochem. Biophys. Res. Commun. 346 (2006) 1320–1323. N. Mizushima, Methods for monitoring autophagy, Int. J. Biochem. Cell Biol. 36 (2004) 2491–2502. S. Pattingre, A. Tassa, X. Qu, R. Garuti, X.H. Liang, N. Mizushima, M. Packer, M.D. Schneider, B. Levine, Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy, Cell 122 (2005) 927–939. A. Oberstein, P.D. Jeffrey, Y. Shi, Crystal Structure of the Bcl-XL-Beclin 1 Peptide Complex: BECLIN 1 IS A NOVEL BH3-ONLY PROTEIN, J. Biol. Chem. 282 (2007) 13123–13132. S.P. Elmore, T. Qian, S.F. Grissom, J.J. Lemasters, The mitochondrial permeability transition initiates autophagy in rat hepatocytes, FASEB J. 15 (2001) 2286–2287. I. Kissova, M. Deffieu, S. Manon, N. Camougrand, Uth1p is involved in the autophagic degradation of mitochondria, J. Biol. Chem. 279 (2004) 39068–39074. M. Kundu, C.B. Thompson, Macroautophagy versus mitochondrial autophagy: a question of fate? Cell Death Differ. 12 (Suppl. 2) (2005) 1484–1489. S.M. Chiu, L.Y. Xue, K. Azizuddin, N.L. Oleinick, Photodynamic therapy-induced death of HCT 116 cells: Apoptosis with or without Bax expression, Apoptosis 10 (2005) 1357–1368. S. Bernales, K.L. McDonald, P. Walter, Autophagy counterbalances endoplasmic reticulum expansion during the unfolded protein response, PLoS Biol. 4 (2006) e423.

E. Buytaert et al. / Biochimica et Biophysica Acta 1776 (2007) 86–107 [195] T. Yorimitsu, U. Nair, Z. ang, D.J. Klionsky, Endoplasmic reticulum stress triggers autophagy, J. Biol. Chem. 281 (2006) 30299–30304. [196] M. Ogata, S. Hino, A. Saito, K. Morikawa, S. Kondo, S. Kanemoto, T. Murakami, M. Taniguchi, I. Tanii, K. Yoshinaga, S. Shiosaka, J.A. Hammarback, F. Urano, K. Imaizumi, Autophagy is activated for cell survival after endoplasmic reticulum stress, Mol. Cell. Biol. 26 (2006) 9220–9231. [197] Y. Kouroku, E. Fujita, I. Tanida, T. Ueno, A. Isoai, H. Kumagai, S. Ogawa, R.J. Kaufman, E. Kominami, T. Momoi, ER stress (PERK/ eIF2alpha phosphorylation) mediates the polyglutamine-induced LC3 conversion, an essential step for autophagy formation, Cell Death Differ. 14 (2007) 230–239. [198] A. Criollo, M.C. Maiuri, E. Tasdemir, I. Vitale, A.A. Fiebig, D. Andrews, J. Molgo, J. Diaz, S. Lavandero, F. Harper, G. Pierron, S.D. di, R. Rizzuto, G. Szabadkai, G. Kroemer, Regulation of autophagy by the inositol trisphosphate receptor, Cell Death Differ. 14 (2007) 1029–1039. [199] M. Hoyer-Hansen, L. Bastholm, P. Szyniarowski, M. Campanella, G. Szabadkai, T. Farkas, K. Bianchi, N. Fehrenbacher, F. Elling, R. Rizzuto, I.S. Mathiasen, M. Jaattela, Control of macroautophagy by calcium, calmodulin-dependent kinase kinase-beta, and Bcl-2, Mol. Cell 25 (2007) 193–205. [200] N. Holler, R. Zaru, O. Micheau, M. Thome, A. Attinger, S. Valitutti, J.L. Bodmer, P. Schneider, B. Seed, J. Tschopp, Fas triggers an alternative, caspase-8-independent cell death pathway using the kinase RIP as effector molecule, Nat. Immunol. 1 (2000) 489–495. [201] T. Vanden Berghe, M. Kalai, L.G. van, W. Declercq, P. Vandenabeele,

[202]

[203]

[204] [205]

[206]

[207]

[208]

107

Disruption of HSP90 function reverts tumor necrosis factor-induced necrosis to apoptosis, J. Biol. Chem. 278 (2003) 5622–5629. Y. Xu, S. Huang, Z.G. Liu, J. Han, Poly(ADP-ribose) polymerase-1 signaling to mitochondria in necrotic cell death requires RIP1/TRAF2mediated JNK1 activation, J. Biol. Chem. 281 (2006) 8788–8795. N. Festjens, T. Vanden Berghe, P. Vandenabeele, Necrosis, a wellorchestrated form of cell demise: signalling cascades, important mediators and concomitant immune response, Biochim. Biophys. Acta 1757 (2006) 1371–1387. W.X. Zong, C.B. Thompson, Necrotic death as a cell fate, Genes Dev. 20 (2006) 1–15. Y.J. Hsieh, C.C. Wu, C.J. Chang, J.S. Yu, Subcellular localization of Photofrin determines the death phenotype of human epidermoid carcinoma A431 cells triggered by photodynamic therapy: when plasma membranes are the main targets, J. Cell. Physiol. 194 (2003) 363–375. C. Fabris, G. Valduga, G. Miotto, L. Borsetto, G. Jori, S. Garbisa, E. Reddi, Photosensitization with zinc (II) phthalocyanine as a switch in the decision between apoptosis and necrosis, Cancer Res. 61 (2001) 7495–7500. A. Uzdensky, A. Lobanov, M. Bibov, Y. Petin, Involvement of Ca2+- and cyclic adenosine monophosphate-mediated signaling pathways in photodynamic injury of isolated crayfish neuron and satellite glial cells, J. Neurosci. Res. 85 (2007) 860–870. B.V. Chernyak, D.S. Izyumov, K.G. Lyamzaev, A.A. Pashkovskaya, O.Y. Pletjushkina, Y.N. Antonenko, D.V. Sakharov, K.W. Wirtz, V.P. Skulachev, Production of reactive oxygen species in mitochondria of HeLa cells under oxidative stress, Biochim. Biophys. Acta 1757 (2006) 525–534.