Photodynamic therapy (PDT) is a promising therapeutic procedure for the management of a variety of solid tumors and non-malignant lesions. PDT is a two-step procedure that involves the administration of a photosensitizing agent [1-3], followed by activation of the drug with non-thermal light of a specific wavelength [4,5] (Figure 1). The anticancer effect of PDT is a consequence of a low-to-moderately selective degree of photosensitizer (PS) uptake by proliferating malignant cells, direct cytotoxicity of reactive oxygen species (ROS) and a severe vascular damage that impairs blood supply to the treated area [6,7]. Those biological effects of PDT are limited to the particular areas of tissues exposed to light. Additionally, PDT leads to activation of tumor directed, systemic immune responses [8-10].

After the absorption of photons the PS is transformed from its ground state to its triplet excited state via a short-lived excited singlet state (Figure 2) [11]. The triplet state can undergo two different reactions: (i) it can undergo electron or hydrogen atom transfer reactions with oxygen or a substrate producing free radicals and other reactive oxygen species; (ii) it can transfer its energy directly to ground-state triplet oxygen to form excited state singlet oxygen (Figure 3). The first process is called a type I reaction and the second a type II reaction [4]. The PDT effectiveness is therefore determined by oxygen supply and may be decreased in conditions where there is tissue hypoxia [12].

PDT as a treatment procedure has been accepted by the United States Food and Drug Administration for use in endo-bronchial and endo-esophageal cancer [13,14] and also as a treatment of premalignant and early malignant lesions of skin (actinic keratosis), bladder, breast, stomach and oral cavity [4,15].

The available body of literature suggests that there is clearly no single pathway that leads to cell death after PDT. In the present review we aim to describe and summarize different cell death pathways activated by PDT [16,17] and describe the key players controlling cell death related to PDT (Table 1).

Apoptosis is a very complex, multi-step, multi-pathway cell-death program that is genetically encoded in every cell of the body [18]. It can be initiated either through the activation of death receptors or the mitochondrial release of cytochrome c [19]. Both events eventually lead to activation of caspase cascades known as ‘executioner caspases’ such as caspase-3, -6 and -7 [20,21]. The active executioner caspases cleave cellular substrates, which leads to characteristic biochemical and morphological changes observed in dying cells [22]. Cleavage of nuclear lamins is followed by chromatin condensation and nuclear shrinkage; cleavage of the inhibitor of the DNase CAD (caspase activated deoxyribonuclease) causes DNA fragmentation. Cleavage of cytoskeletal proteins leads to cell fragmentation and formation of apoptotic bodies [23]. The apoptotic process is tightly controlled by various proteins [24,25]. It is well known that resistance of tumor cells to apoptosis might be an essential feature of cancer development thus, modulation of the key elements of apoptosis signaling may directly influence therapy-induced tumor-cell death [26,27].

It is generally recognized that mitochondria play a critical role in the apoptotic cascade by controlling the release of crucial factors involved in that process [67]. Among these factors cytochrome c plays an essential role. During the process of apoptosis, cytochrome c is released from mitochondria into the cytosol. Release of cytochrome c from mitochondria is controlled by proteins of the Bcl-2 family. In the cytosol, cytochrome c activates the caspases—a family of killer proteases—through formation of a complex with Apaf-1 (for apoptotic-protease activating factor-1), procaspase-9 and ATP or dATP. [68]. The opening of the mitochondrial membrane permeability transition pores, which results in the dissipation of the mitochondrial membrane potential (Δψ_m), has been proposed as the main mechanism for release of cytochrome c. Chiu and Olenick [69], showed that treatment of LY-R cells with Pc4 based PDT resulted in release of cytochrome c to the cytoplasm after 15 min, as estimated by an immunohistochemical method, and the loss of Δψ_m depended on PDT dose and the post treatment time. The observed loss of Δψ_m was only in those cells receiving the highest dose of PDT. It was concluded that the release of cytochrome c from mitochondria is independent of the loss of Δψ_m action after Pc4 mediated PDT.

Another work indicating an important role of cytochrome c in PDT-induced apoptosis was carried out by Vantieghem et al. [70]. In his approach, the overexpression of Bcl-2 in Pc60R1R2 cells revealed that mechanism of cytochrome c release after hypericin mediated PDT is caspase-dependent. The observed overexpression of Bcl-2 remarkably delayed cytochrome c release, but it did not protected cells from PDT induced death. These results showed that in cells overexpressing Bcl-2 addition of zVAD-fmk significantly suppressed cytochrome c release and apoptosis.

Another support for the hypothesis that cytochrome c plays a vital role in PDT-induced apoptosis was provided by Varnes et al. [71]. In this study, an LD_99.9 dose of Pc4 PDT induced loss of cytochrome c from the mitochondria of LY-R cells, but also inhibited respiration and caused activation of caspase 3-like proteases. All events took place within 15 min of light exposure. This study provided clear evidence that cytochrome c leakage is dependent on activation of proteolytic proteases after PDT.

All death receptors belong to TNFR superfamily (tumor necrosis factor receptor) [72]. Most of them act as transmembrane signal transducers that respond to ligand binding. Some of them, however, do not transduce signals, but rather function as decoy receptors that compete for the interaction of cognate ligands with their signaling receptors. Each member of the TNFR superfamily has at least one specific ligand, but there are other ligands that bind to several receptors. The most complex example is APO2/TRAIL, which binds five different receptors [73].

TNFR signaling was discovered to be an important factor in immune response and such members like Fas and APO2/TRAIL induce apoptosis through a p-53 independent mechanism. The signaling members of TNFR superfamily can be divided into two main subgroups. One class contains cytoplasmic death domain, whereas the other class does not. Death domains mediate interaction of death receptors with death-domain–containing adaptor proteins and then act in the caspase-dependent pathway. Death domain containing receptors activate the apoptotic pathway by caspase-8 mediated cleavage of the pro-apoptotic Bcl-2 superfamily member BID [74]. This protein interacts with other molecules like Bax and Bak, which cause release of mitochondrial cytochrome c and SMAC/DIABLO, activating caspase-9 and eventually caspase-3. The other group use TNFR-associated factor to link these receptors to serine/threonine protein kinase cascades that regulate gene transcription [75].

Because treatment with factors that activate death receptor signaling in cancer cells may be an effective anticancer strategy it has been investigated whether PDT can affect this pro-apoptotic pathway.

The early evidence comes from the study designed by Ahmad et al. [76] where the involvement of the cell surface receptor Fas (CD95) pathway in A431 cells in Pc4 mediated PDT was investigated. It was observed that a significant time-dependent increase in the protein expression of Fas at 5, 15, 30 and 60 min post PDT occurred. In an immunoblot, Fas protein was observed as a doublet, which may indicate the insoluble and soluble forms of Fas. By using immunoprecipitation it was determined that Pc4-PDT resulted in a multimerization of Fas protein leading to its activation. To confirm that PDT induced apoptosis is due to Fas activation the involvement of Fas associated death domain level (FADD) was examined by using immunoblot analysis. It confirmed a time-dependent increase, for up to 1 h post-PDT in Fas protein levels. Pc4-PDT also caused activation of FADD-like interleukin-1 beta-converting enzyme (FLICE), which was evident from the appearance of cleaved products of pro-casapase-8.

To further evaluate the role of Fas in PDT mediated apoptosis, Ali et al. [77] used a model of Hypocrellin A (HA) and Hypocrellin B (HB) as photosensitizers in CNE2 and TW0-1 cells. This group observed elevated levels both Fas and FasL in both examined cell lines and these higher levels coincided with increase in phototoxicity of PDT. The enhanced expression of CD95/CD95L was detectable within 2 h after irradiation using HA and HB and was significantly increased at 3 h following light exposure. The specific involvement of Fas/FasL system in CNE2 and TW0-1 cell death induced by HA and HB was further supported by significant inhibition of apoptosis in the presence of either anti-CD95 or anti-CD95L antibodies.

There is also strong evidence that APO2/TRAIL is involved in PDT mediated apoptosis. Schempp et al. [78] used Hypericin mediated PDT in Jurkat cells and by using neutralizing antibodies against Fas, FasL, TRAIL and TNFR1 discovered that post PDT apoptosis can be inhibited by polyclonal anti-TRAIL antibody. The specificity of the inhibitory activity of this antibody was demonstrated by blocking TRAIL mediated killing of Jurkat cells. However, the mechanism of Hypericin-PDT mediated apoptosis based on TRAIL/TRAIL-receptors involvement is not clear and it may include increased shedding of TRAIL, stabilization of death inducing TRAIL receptors or down regulation of decoy receptors.

Further support for this hypothesis was provided by Granville et al. [79], who described a two-fold increase in the number of cells containing hypo diploid DNA after combined therapy PDT + TRAIL. They also described that combined therapy of PDT and Fas-ligand increased the number of apoptotic cells seven times. The final conclusion was that Jurkat cells undergoing PDT procedure become more sensitive to Fas-ligand and TRAIL induced killing.

Mitogen-activated protein kinases are proline-directed Ser/Thr protein kinases activated by dual phosphorylation on both tyrosine and threonine residues [87]. These enzymes are critical components of a complex cellular signaling network that ultimately regulates gene expression in response to variety extracellular stimuli. The three well known MAPK families are: the extracellular signal-regulated kinases (ERKs), the c-Jun N-terminal kinases/stress-activated protein kinases (JNKs/SAPKs), and the p38 MAPK [88]. Each of these enzymes is a target for phosphorylation cascades in which the sequential activation of three kinases constitutes a common signaling pathway. The best characterized MAPK pathway is the Ras/Raf/MEK cascade leading to the activation of ERK1/2 in response to growth factors [89]. JNK and p38 are key mediators of stress signals and inflammatory response [90].

A link between SAPK and p38 pathways and apoptosis has been suggested in several studies [91,92]. The evidence of the involvement of JNK1 and p38 was provided by Assefa et al. [93]. In hypericin-mediated PDT of A431, HaCaT, HeLa and L929 cell lines theye observed a rapid and persistent activation of JNK1 and severely inhibition of the basal levels of ERK2. It was also demonstrated that photo-activated hypericin blocked the EGF-mediated activation of ERK2, and the inhibition of ERK2 pathway was irreversible. They also observed that JNK1 activation occurred independently from caspase activities. Using simultaneous inhibition of both stress kinases they observed significant sensitization of cells to PDT-hypericin mediated apoptosis. They concluded that both JNK1 and p38 MAPK pathways played an important role in cellular resistance to PDT-induced apoptosis with hypericin.

The involvement of stress kinases in PDT mediated cell death was also investigated by Chan et al. [94]. They evaluated the role of JNK1 and involvement of singlet oxygen in triggering the JNK pathway. PDT triggered activation of stress kinase pathway in two steps: firstly the formation of singlet oxygen induced rapid activation of JNK, which led to activation of the caspase cascade and apoptosis. Secondly activated caspase-3 could then act on several apoptotic substrates including PAK2. Cleavage of PAK2 released its C-terminal catalytic fragment as a 36 kDa active kinase that can further perform a second step in JNK activation.

Xue et al. [95] investigated the involvement of Etk/Bmx kinase in PDT mediated apoptosis. Etk/Bmx is a newly discovered tyrosine kinase, commonly expressed in prostate epithelial and carcinoma cells. It might play an important role in the growth, development and anti-apoptotic action in epithelial cells. Using an overexpression model in LNcaP cells they found that Etk/Bmx is an effector of PI3-kinase and that the PI3-kinase/Etk pathway is involved in the protection of prostate carcinoma cells from apoptosis in response to PDT.

Ceramide is implicated in the cell-signaling pathway involved in apoptosis as it acts as a second messenger to permeabilize the outer mitochondrial membrane and facilitate the release of cytochrome c from the mitochondria [96]. Other modes of ceramide-mediated cell death involve the activation of stress-activated protein/JUN kinase (SAPK/JNK) [97], dephosphorylation of the retinoblastoma protein and upregulation of transcription factors. Evidence of ceramide involvement in PDT-mediated apoptosis was provided by Separovic et al. [98]. In phthalocyanine PDT model they demonstrated in the U937 cell line elevated levels of ceramide by 45, 37, 67, 118 and 134% at 1, 10, 30, 60 and 120 min, respectively, after PDT procedure. In CHO cells ceramide generation in response to PDT was rapid but did not reach such a high level. Ceramide accumulation was also observed in RIF1 cells. In this model ceramide levels were not different from control level for the first 10 min post-PDT but then reached a maximum of 118%, maintaining this level up to 2 h. The PDT dose produced a 99% loss of clonogenicity, however in RIF1 cell there was no evidence of apoptosis.

p21/WAF1 involvement in PDT was investigated by Ahmad et al. [99]. They found that cells subjected to PDT resulted in a growth arrest particularly observed in G₀-G₁ phase. The treatment caused an accumulation of 47%, 51% and 69% growth-arrested cells at 3, 6 and 12 h post-PDT, respectively. They further assessed the influence of PDT on WAF1/p21 induction and using Western blot analysis revealed a significant induction of WAF1/p21 at 3, 6 and 12 h after treatment, when compared with basal levels. In addition, PDT resulted in a time-dependent decrease in cyclin D1, cyclin E, cdk2 and cdk6. The decrease of cyclin D1 and cdk6 was markedly more pronounced than that of cyclin E and cdk2. To confirm this data they investigated the effect of PDT on kinase activities associated with cdk. The radioactive kinase activity assay showed that PDT almost universally resulted in a time-dependent decrease in kinase activities associated with all the cdks and cyclin examined. They also examined the effect of PDT on the binding between WAF1/p21-cyklin and WAF1/p21-cdk and also cyclin-cdk and found that PDT resulted in increased binding of cyclin D1 and cdk6 toward WAF1/p21, whereas the binding of cyclin E and cdk2 did not change.

Necrotic cell death has been described as a violent and quick form of degeneration affecting large fractions of cell populations, characterized by cytoplasmic swelling, destruction of organelles and disruption of the plasma membrane, leading to the release of intracellular contents and consequent inflammation [100].

Necrosis has been referred to as accidental cell death, caused by physical or chemical damage and has generally been considered an unprogrammed process [101]. It is characterized by a pyknotic nucleus, cytoplasmic swelling, and progressive disintegration of cytoplasmic membranes, all of which lead to cellular fragmentation and release of material into the extracellular compartment. In necrosis, decomposition is principally mediated by proteolytic activity, but the precise identities of proteases and their substrates are poorly defined [102].

Studying the factors and parameters that cause cellular necrosis after PDT is not as easy as studying those factors which lead to apoptosis. The crucial factors in determining the type of cell death, e.g., apoptosis or necrosis following PDT are: the cell type, the presence of an intact set of apoptosis machinery, the subcellular localization of the PS, the light dose applied to activate it locally, and the oxygen partial pressure [103]. One factor that can be agreed upon by all commentators is that high dose PDT (either a high photosensitizer concentration or a high light fluence or both) tends to cause cell death by necrosis, while PDT administered at lower doses tend to predispose cells towards apoptotic cell death.

Nagata and colleagues [104] used the amphiphilic PS ATX-S10 (Na) and human malignant melanoma cells and found that light doses that led to less than 70% cytotoxicity induced mainly apoptosis; by contrast, most cells appeared necrotic with doses that induced 99% cytotoxicity. A common feature of the apoptotic program initiated by PDT is the rapid release of mitochondrial cytochrome c into the cytosol followed by activation of the apoptosome and procaspase-3. With PS localized in the plasma membrane the photosensitization process can rapidly switch the balance towards necrotic cell death likely due to loss of plasma membrane integrity and rapid depletion of intracellular ATP [105]. It is also possible that high doses of PDT can photochemically inactivate essential enzymes and other components of the apoptotic cascade such as caspases. For instance, Lavie and coworkers [106] used the perylenequinones (hypericin and dimethyl tetrahydroxyhelianthrone) and found high dose PDT inhibited apoptosis by interfering with lamin phosphorylation, or by photodynamic cross-linking of lamins.

Xue and Oleinick [107] compared Pc4-mediated PDT of MCF7 cells that lack caspase 3 with the same cell line with caspase 3 transfected back in. They found apoptotic indicators only in the caspase expressing cells which also showed more loss of viability by an assay involving reduction of a tetrazolium dye; however both cell lines showed an equal degree of cytotoxicity by a clonogenic assay. Dahle, Steele and Moan reported [108] that the mode of cell death induced by PDT depended on cell density. They seeded Madison Darby canine kidney II cells (MDCK II) at two different densities and incubated them with meso-tetra(4-sulfonatophenyl)porphine (TPPS4) for 18 h, washed and irradiated with blue light. Four hours later the cells were studied by fluorescence microscopy. With <55% total cell death the apoptotic fraction was significantly higher for cells in confluent monolayers than for cells growing in microcolonies at equitoxic doses. Confluent cells were 2.9 times more sensitive than cells in microcolonies partly due to a 1.5 times higher uptake of TPPS4 in monolayer cells. The difference in mode of cell death for the different cell densities was not related to any observable difference in subcellular localization pattern of TPPS4 at equitoxic doses of PDT.

Autophagy is a catabolic cellular mechanism that allows the cell to maintain a balance between the synthesis, degradation, and recycling of cellular products [109]. A variety of autophagic processes exist, all of which involve the lysosomal degradation of the cellular organelles and proteins. The most well-known mechanism proceeds in the following manner: A double membrane structure called autophagosome surrounds the target region, creating a vesicle that separates its contents from the rest of the cytoplasm. This vesicle is then transported and fused to the lysosome, forming a structure called the autophagolysosome, the contents of which are subsequently degraded by lysosomal hydrolases [110]. Besides facilitating the disposal of unwanted proteins, organelles, and invading microorganisms, autophagy also allows a cell to reallocate its nutrients from unnecessary processes to life-essential ones in times of starvation or stress.

The term “autophagy” originated in 1963 at the Ciba Foundation Symposium on Lysosomes, where it was used to describe the presence of membrane vesicles that contain disintegrating organelles and cytoplasm components. Evidence soon confirmed that autophagy is an adaptive, energy-generating process. Molecular control of autophagy came into focus in the late 1990s. Signaling pathways governing this process became better understood after the identification of the target of rapamycin kinase (TOR), which controls cell growth and protein synthesis [111]. In 1997, Ohsumi's group showed that yeast autophagy is similar to that of mammals, which allowed them to analyze the process in the genetically tractable yeast system. This study led to the discovery of the first autophagy-related gene, ATG1. Shortly thereafter, the first mammalian autophagy genes were identified [112].

One of the first diseases associated with autophagy was cancer [113]. It was discovered that Beclin1, an essential protein for autophagy, is also a tumor suppressor [114]. Further studies on the role of autophagy in cancer revealed some interesting properties of the disease. Initially, autophagy suppresses tumor growth through its production of Beclin1. This changes as the tumor becomes more advanced; autophagy begins to promote tumor progression by providing the mechanism for cells in the central, low nutrient, part of the tumor to obtain the energy that they need to stay alive. It was also found that autophagy blocks apoptotic pathways, thereby protecting cancer cells from treatment [114]. On the other hand, some cancer therapies induce autophagic cell death of tumor cells. This two-sided effect of autophagy on tumors can be exploited by anticancer therapy to provide better treatment for cancer patients.

It is still unclear exactly how autophagy affects the outcome of PDT [66,115,116]. In general, mammalian cells use autophagy as a defense against ROS mediated damage by clearing the cell of damaged organelles [117]. Depending on the type of ROS and degree of oxidative injury, PDT may stimulate autophagy [118] that either acts in a cytoprotective manner or induces autophagic cell death [119]. Autophagy may play a role in PDT induced apoptosis, but the two processes can also occur independently of one another [120]. A study done on murine leukemia L1210 cells found that a wave of autophagy occurs right before apoptosis [121]. It was also found that prevention of autophagy by silencing the Agt7 gene allowed photo-killing to occur at lower light doses. This observation is consistent with the theory that autophagy is a defense mechanism against PDT induced ROS [122]. The situation is different for tumor cells that do not have the ability to undergo apoptosis through a deficiency in Bax and Bac, which regulate the apoptotic pathway. In these cells, PDT induced autophagy stimulates a necrotic, caspase independent cell death [64]. Suppression of autophagy in apoptosis-deficient cells resulted in the inhibition of cell death during PDT. In general, the induction of autophagy in PDT treated cells occurs independently of an apoptotic outcome. While autophagy seems to play a pro-survival role in tumor cells that are capable of apoptosis, it has been shown to promote death in cells that are apoptosis-deficient.

In order to understand how PDT affects autophagy it is important to take note of the PDT affected proteins that are involved in this mechanism. Many proteins, some directly involved in the autophagic process, are damaged by PDT induced ROS. Some ER and mitochondrial photosensitizers cause damage to Bcl-2, to which Beclin1 (a pro-autophagic protein) binds. The IP3 protein, a regulator of the autophagic process associated with the ER [123,124] is affected by phthalocyanine photosensitizer Pc 4 [125]. The photosensitizer AlPcS causes damage to the mammalian target of rapamycin, mTOR, a cell growth regulator that takes part in the autophagic signaling pathway [126]. Other proteins, like Beclin1, Atg5, and Atg7, appear to be unaffected by PDT. Although many proteins involved in the autophagic process are photodamaged by PDT, it appears that those involved in the formation of autophagosomes remain active [120].

PDT can also affect autophagy by damaging organelles that are directly involved in the process. Several photosensitizers target autophagy-related organelles such as lysosomes and endosomes. When tetra(4-sulphonatophenyl)-porphine (TPPS4) photosensitizer is used photo-oxidation of the organelle matrix occurs [127]. In this type of PDT, lysosomal enzymes are inactivated before the membrane ruptures, which allows for specific targeting of this organelle without causing damage to the rest of the cell. This treatment can be used to selectively enrich autophagosomes [128]. On the other hand, NPe6 and TPPS2 photosensitizers bind to the lysosomal membrane, causing it to rupture upon irradiation. The released, but not neutralized proteases can cause induction of apoptosis by cathepsin-mediated cleavage of Bid [129]. The lysosomes, autophagosomes, endosomes, and autolysosomes of the cells treated with this amphiphilic photosensitizer will have it in their membranes. Upon irradiation, the membranes will break and release their contents into the cytoplasm. Based on these results, it can be concluded that the effect of PDT on the autophagy of a tumor cell depends on the type of photosensitizer used.

The effects of a damaged lysosome on the autophagic process were explored in an autophagic flux study of murine hepatoma 1c1c7 cultures. These cultures were sensitized with NPe6 and irradiated with LD90 light dose. Acridine orange (AO) staining of acidic organelles was done in order to observe the effect of PDT on the lysosome. No lysosomal damage was observed when cells were exposed to either light or photosensitizer alone. In those cells that received PDT, AO staining was completely lost within an hour, and within two hours autophagosome accumulation was observed [130]. This indicates that autophagosomes can be formed in the absence of lysosomes, but autophagy cannot be completed due to the lack of these organelles.

Based on studies with various cancer lines and photosensitizers it can be concluded that PDT directly induces autophagy. This is independent of photosensitizer target, as autophagy was observed with photosensitizers that localize in the ER, mitochondria, lysosomes, and endosomes. A second conclusion that can be drawn is that apoptosis often occurs in cells that are already undergoing autophagy and is also a result of PDT. The rates of autophagy and apoptosis depend on the cancer cell type, photosensitizer, and light dosage. Depending on cell type, autophagy will either promote or prevent cell death from PDT. In cells that are able to undergo apoptosis autophagy alleviates the destructive effects of PDT by recycling damaged organelles [131]. This opens up a possibility that PDT of these cancer cells can be enhanced by suppressing pro-autophagic proteins. Autophagy has the opposite effect on cells that are apoptosis-deficient, promoting cell death through necrosis. The last conclusion that can be made is that autophagic processes are compromised, but not prevented, in PDT protocols that employ lysosome targeting photosensitizers. In these procedures, autophagy is initiated by formation of autophagosomes but cannot be completed by fusing with the lysosome. This is because the proteins involved in autophagosome formation (Beclin1, Atg5) are not photodamaged by PDT.

PDT can lead to all three forms of cell death, namely apoptosis, necrosis and autophagy (Figure 4). The response to PDT may vary not only with the cell type or its genetic or metabolic potential but also with the experimental model, total fluence delivered, different types of photosensitizers and their intracellular localization. The initial site of PDT-related damage may determine which cell death pathway is initially activated. The extent of PDT related damage may also regulate how the PDT treated cells respond. It is possible that the autophagy process is activated as an initial rescue mechanism, when the PDT damaged cells try to contain and removed damaged proteins. Only when the PDT damage is sufficiently robust and the cells are damaged beyond repair does apoptosis occur. PDT, at its highest dose may also lead to necrosis, as the proteins that participate in both autophagy and apoptosis may be immediately destroyed and the cellular integrity may be broken.

It is also important to bear in mind that most of the reported in vitro data are also difficult to translate into the in vivo situation where unequal light distribution or inhomogeneous photosensitizer accumulation may lead to a variable cellular response within PDT treated tumors. Additionally, the shutdown of tumor vessels may lead to local depletion of nutrients and oxygen and therefore trigger secondary PDT related necrosis. Lastly, as PDT leads to activation of tumor directed immune response, some cancer cells are killed via apoptosis by cytotoxic T cells.

The understanding of the cascade of events playing a role in PDT-mediated apoptosis is far from complete. Defining these events will hopefully result in designing better PDT protocols, which could have wider applications as a cancer therapeutic modality.