Photodynamic therapy (PDT) dosimetry is complex as many factors are involved and varied interdependently. Monitoring the biological consequence of PDT such as cell death is the most direct approach to assess treatment efficacy. In this study, we performed 5-aminolevlinic acid (ALA)-PDT in vitro to induce different cell death modes (i.e., slight cell cytotoxicity, apoptosis, and necrosis) by a fixed fluence rate of 10 mW/cm2 and varied fluences (1, 2, and 6 J/cm2). Time course measurements of cell viability, caspase-3 activity, and DNA fragmentation were conducted to determine the mode of cell death. We demonstrated that NADH fluorescence lifetime together with NADH fluorescence intensity permit us to detect apoptosis and differentiate it from necrosis. This feature will be unique in the use of optimizing apoptosis-favored treatments such as metronomic PDT.
©2011 Optical Society of America
Photodynamic therapy (PDT) is a cancer therapy that involves photosensitizer interacting with light and oxygen to generate reactive oxygen species (ROS) or singlet oxygen that cause cell death and tumor destruction. It has been approved for treatments of head and neck cancer and basal-cell carcinoma in the European Union, esophageal and endobronchial cancer in the United States, and cervical and gastric cancers in Japan . Clinical PDT dosimetry is complex because not only several treatment factors are involved (i.e., light dose, drug dose, light-drug duration, oxygen) but also these factors varied dynamically and interdependently. As a result, the biological consequence of PDT is not necessarily proportional to a single factor such as the irradiance dose rate in ionizing radiation therapy. Generally, four PDT dosimetry strategies are pursued: explicit, implicit, and direct dosimetry, and biological tissue response monitoring . Explicit dosimetry directly measures the three PDT components (light, drug, and oxygen) and sometimes incorporates a dose model to simulate the outcome . Implicit dosimetry intended to measure a single metric such as photosensitizer photobleaching that is predictive of the biological damage or outcome under certain circumstances. Direct dosimetry measures the ROS, particularly singlet oxygen, that is widely believed to dominate the causes of biological damage for most current photosensitizers and treatment doses used . With the availability of the new technology, direct dosimetry shows advantages over explicit and implicit dosimetry for its simplicity to measure single metric only without any dose model . However, studies have shown that singlet oxygen monitoring approach could fail to predict the tumor response . In situations like this, reporters of biological response to therapy would be necessary. At present, the biological response monitoring is limited to measure the tumor volume, tumor perfusion, and treatment-induced necrosis [2,6]. Advanced technologies such as MRI and optical spectroscopy allow in vivo monitoring tumor perfusion such as blood flow and hemoglobin oxygenation that are undergoing active evaluation by several groups for the prediction of long-term response to PDT treatment [7–11].
Outstanding long-term tumor control and tumor apoptosis were observed with metronomic PDT, which involves long time and the continuous slow delivery of photosensitizer and light [12,13]. Low fluence rate PDT also leads to more durable tumor responses and induces more evenly distributed apoptotic cells [14,15]. Direct and noninvasive monitoring cell apoptosis immediately after PDT may provide a good way to determine the biological consequence and thus treatment efficiency under these circumstances. In this study, we aimed to demonstrate the detection of cell death immediately after PDT using the intrinsic fluorescence of reduced nicotinamide adenine dinucleotide (NADH), whose intensity has been widely accepted as an optical probe of cellular metabolic state. NADH fluorescence lifetime has recently been studied for its use in monitoring cell metabolic activities [16–18] as well, and cell death [18–21]. In previous studies, we showed that NADH fluorescence lifetime increase at the early phase of mitochondrial and poly(ADP-ribose) polymerase-1 (PARP-1) mediated cell death induced by staurosporine (STS)  and N-methyl-N’-nitro-N-nitrosoguanidine (MNNG) , respectively. This lifetime increase occurred before the disruption of mitochondrial membrane potential (ΔΦ), one of the first stages in both apoptotic and necrotic pathways, and decrease of oxygen consumption rates [20,21]. Here, we performed 5-aminolevlinic acid (ALA)-PDT in non-small cell lung carcinoma H1299 cells with a fixed fluence rate at 10 mW/cm2 and manipulated light fluences at 1, 2, and 6 J/cm2 to induce slight cytotoxicity, primary apoptosis, and necrosis, respectively. We examined how NADH fluorescence responded to these modes of cytotoxicity by tracing time course of NADH fluorescence lifetime and intensity for each condition. We demonstrated that NADH fluorescence lifetime increased only when the cells primarily died through the apoptotic pathway, which was confirmed by DNA fragmentation analysis, cell morphology, and caspase-3 activation [22,23]. NADH fluorescence intensity shows significant decrease when primary cells died through necrosis and no change for slight cytotoxicity after PDT for 1 hour. Taken together, these results suggest that direct monitoring ALA-PDT induced cell death and differentiating different death modes by NADH fluorescence lifetime and intensity is possible.
2. Materials and methods
2.1 Cell line and photodynamic therapy treatment
H1299 non-small cell lung carcinoma were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; Gibco, Invitrogrn Corp., Carlsbad, California, USA) supplemented with 5% fetal bovine serum, 100 units/ml penicillin and 50 mg/ml streptomycin at 37°C in 5% CO2 humidified atmosphere. Cells were seeded on 24 mm diameter glass coverslips (Paul Marienfeld GmbH & Co., Lauda- Konigshofen, Germany) at 2x105 cells 24 hours before ALA-PDT treatment. Before fluorescence lifetime imaging microscopy (FLIM), 5-ALA (A7793; Sigma, Buchs, Switzerland) was prepared by resuspending in the culture medium to a concentration of 1 mM. Cells were washed with the serum-free medium and incubated in serum-free medium  containing 1 mM 5-ALA for four hours. The drug dose (1 mM) used here was based on the cell viability study that there was no significant change over a range of drug dose from 0.5 to 2 mM. At the dose of 1 mM, PPIX fluorescence was detected maximal at 4 hr incubation time (data not shown). The cells were then washed once by phosphate buffer solution (PBS) and incubated in PBS. Subsequently the cells were irradiated by an 8 x 12 cm2 LED array with a wavelength peak at 633 nm. The light fluence rate at 10 mW/cm2 and three different light fluences at 1, 2, or 6 J/cm2 were used to treat the cells. Immediately after PDT treatment, the cells were either imaged for NADH fluorescence lifetime or incubated with fresh DMEM and 5% FBS at 37°C in 5% CO2 incubator for use in the analysis of cell viability, caspase-3 activity, and DNA fragmentation. Control cells were performed with light treatment only, drug treatment only, and no drug and light treatment. For fluorescence lifetime image, the cell-seeded coverslip was washed twice using PBS and then transferred into an imaging chamber filled with 1 ml DMEM with 5% FBS. All images were taken at 256 x 256 pixels resolution with the field of view (FOV) of 100 x 100 μm and the acquisition time of 20 minutes to obtain enough photons for reliable analysis of the NADH fluorescence lifetime. Time-lapsed NADH fluorescence lifetime images were obtained at the same site (same FOV) before, immediately after (0-20 minutes), and up to 2 hr after ALA-PDT.
2.2 Cell viability assay
Cell viability was determined by using the CellTiter-Blue® cell viability assay (Promega Corp., Madison, Wisconsin, USA). Cells at a concentration of 2x105 cells were seeded per 35 mm plate. After ALA-PDT treatment, we added the medium containing CellTiter-Blue® reagent into the cells, which were then incubated for 1 hour at 37°C in 5% CO2 humidified atmosphere. The viable cells were detected by a Victor2 1420 Multilabel counter (PerkinElmer, Foster, Massachusetts, USA) with an excitation wavelength of 560 nm and emission wavelength of 590 nm.
2.3 Analysis of sub-G1 contents
An aliquot of 1x106 cells were seeded on a 100-mm plate for 24 hr. After ALA-PDT treatment, the adherent and floating cells were collected and then centrifuged at 1500 rpm. The pellet was resuspended in 500 μl PBS to a concentration of 1x106 cells/ml. These resuspended cells were then fixed with 1 ml ice-cold methanol overnight at −20°C. The fixed cells were washed twice in 2 ml cold PBS. The cells were resuspended in 1 ml staining buffer containing RNase (50 g/ml) and propidium iodide (PI, 60 mg/ml) (Sigma, Buchs, Switzerland) in PBS. After incubation at 20°C for 30 minutes in the dark, the stained cells were analyzed in a fluorescence-activated cell sorter (FACSCalibur, Becton Dickinson, Franklin Lakes, New Jersey, USA). The percentage of cells in sub-G1 phase was analyzed using the WinMDI software (Becton Dickinson, Franklin Lakes, New Jersey, USA).
2.4 Caspase-3 activity assay
Caspase-3 activity was determined by fluorescent measurement of 7-amino-4 trifluomethylcoumarin (AFC) released from fluorogenic substrate Ac-DEVD-AFC (caspase-3 substrate) following ALA-PDT of H1299 cells. The cells were lysed in 50 μl lysis buffer (12.5 mM Tris-HCl, 1 mM dithiothreitol, 0.125 mM ethylenediaminetetraacetic acid (EDTA), 5% glycerol, and an aliquot of complete protease inhibitor mixture (Roche Applied Sciences, Mannheim, Germany) pH 7.0, and centrifuged at 12000 rpm and4 °C for 15 minutes. A 100 μg aliquot of protein was incubated with 20 μM fluorogenic substrate of caspase-3 (Calbiochem, San Diego, California, USA) in 500 μl assay buffer (50 mM Tris-HCl, 1 mM EDTA, and 10 mM EGTA (ethyleneglycol-bis- (β-aminoethylether)-N, N, N’, N’-tetraacetic acid), pH 7.0) at 37 °C for 30 min in the dark as described previously . The fluorescence intensity was determined by a spectrofluorometer (Hitachi F-3000, Tokyo, Japan) at an excitation wavelength of 408 nm and an emission wavelength of 505 nm.
2.5 NADH fluorescence lifetime imaging microscopy (FLIM)
Time-domain FLIM was performed using a 60X 1.45NA PlanApochromat oil objective lens (Olympus Corp., Tokyo, Japan) on a modified two-photon laser scanning microscope (FV300 with the IX71 inverted microscope, Olympus Corp., Japan) as described previously . In brief, NADH fluorescence was two-photon excited at 740 nm by a mode locked Ti:sapphire laser (Mira F-900, Coherent Inc., Santa Clara, California, USA) pumped by a solid-state continuous wave 532 nm Verdi laser (Coherent Inc., Santa Clara, California, USA). The emitted NADH fluorescent light was detected using a band pass filter of 450 ± 40 nm (Edmund Optics, Inc., Barrington, New Jersey, USA). Time-resolved detection was conducted by the single photon counting SPC-830 printed circuit board (Becker & Hickl GmbH, Berlin, Germany). Data were analyzed with the commercially available SPCImage v2.8 software (Becker & Hickl GmbH, Berlin, Germany) via a convolution of a double-exponential model function, and the instrument response function (IRF). The convoluted results were fitted to the experiment data to extract lifetime parameters τ1, τ2, a1, a2 and τm. τm is the mean lifetime defined as (a1τ1 + a2τ2)/(a1 + a2). IRF was measured using a second harmonic generated signal from a periodically poled lithium niobate crystal.
NADH fluorescence lifetime has recently been investigated as the biomarker of cell metabolic state [16,17] and cell death [19,20]. This “fluorescence lifetime” usually refers to parameters like free NADH fluorescence lifetime (τ1~400 to 500 ps), bound NADH fluorescence lifetime (τ2 ~2000 to 3000 ps), the corresponding amplitudes a1 and a2, mean lifetime (τm = a1τ1 + a2τ2), and/or the ratio of relative amplitudes of two lifetime components (a1/a2). In this study, we only report τm as the fluorescence lifetime.
2.6 NADH fluorescence intensity measurement
NADH fluorescence intensity is measured by using a Victor2 1420 multilabel counter (PerkinElmer, Foster, Massachusetts, USA) at an excitation wavelength of 355 nm and an emission wavelength of 460 nm. Treated cells were resuspended in 200 μl PBS and transferred into a black OptiPlate-96F 96-well plate (Packed Bioscience, Perkin-Elmer, Foster, Massachusetts, USA) to measure the NADH fluorescence and NADH fluorescence intensity was normalized to cell numbers.
3.1 Characterization of cell death pathways induced by three different light doses of ALA-PDT
Slight cell death, primary apoptosis, and necrosis or non-apoptotic death was induced by treating cells at 1, 2, or 6 J/cm2, respectively, with a fixed fluence rate of 10 mW/cm2. In Fig. 1A , the cell viability of control cells was measured for cells without light and drug, with light only (6 J/cm2,10 mW/cm2), and with drug only treatments. No cell death was observed in all controls. The cell viability of PDT treated cells decreased slowly under the lowest fluence at 1 J/cm2 to remain 62% at 4 hr after PDT. It decreased rapidly within the first hour to be less than 30% with the highest dose at 6 J/cm2. At 2 J/cm2, cell viability decreased monotonically to 28% at 4 hr after PDT treatment. We then measured caspase-3 activity and DNA fragmentation to confirm the pathway of cell death as published reports . Figure 1B shows the time course caspase-3 activity immediately after PDT (0 hr), and at 1, 2, and 3 hr after PDT treatment. The maximal caspase-3 activity appeared at 2 and 1 hr after PDT for light fluence of 1 and 2 J/cm2, respectively. No caspase-3 activity was shown at 6 J/cm2. Figure 1C shows DNA fragmentation for three different controls and for PDT treated cells at 4 hr after PDT. The light fluence of 2 J/cm2 induced 35.89 ± 11.17% sub-G1 contents (Fig. 1C (e)). No obvious sub G1 content was observed for controls (Figs. 1C(a)-(c)) and PDT treated cells at 1 J/cm2 (Fig. 1C(d)). Cells treated with 6 J/cm2 did not show any cell cycle distribution (Fig. 1C(f)). Table 1 summarizes the statistic results for those experiments with a p-value less than 0.05 using student t-test by comparing with controls without drug and light. Taken together, we conclude that the light fluence of 2 J/cm2 induced cell death primarily by apoptosis. Cells treated with 6 J/cm2 underwent non-apoptotic pathway that neither their caspase-3 activity nor DNA fragmentation showed any sign of apoptosis. The expression of caspase-3 activity for cells treated with 1 J/cm2 suggested that partial cells died by apoptosis.
3.2 The increase of NADH fluorescent lifetime was only detectable in cells that mainly died by apoptosis
To determine whether PDT induced cell death is detectable by NADH fluorescence lifetime changes, we acquired time course micrographs of NADH fluorescent lifetime of H1299 cells within the same field of view (FOV) for three different controls (Fig. 2 ) and for PDT treated cells immediately after PDT for up to two hours under three different fluence conditions (Fig. 3 ). Controls without light and drug treatments were imaged at 0 to 20, 20 to 40, 60 to 80, and 100 to 120 minutes after the same preparation time for drug controls. Light controls (6 J/cm2, 10 mW/cm2) were imaged after light irradiation. Drug controls were imaged after 4 hr ALA incubation. PDT treated cells were imaged after 4 hr ALA incubation and light irradiation. Thus, time point at 0-20 minutes in Fig. 3 represents the time point immediately after PDT treatment. All micrographs are displayed using the same color bar of the time scale (200 to 3000 ps). We could examine cell morphology as well through these micrographs.
In Fig. 2, controls without drug and light treatments and light controls show no change on cell morphology and NADH fluorescence lifetime. Drug controls show shrinkage in cell morphology but no change in NADH fluorescence lifetime. In Fig. 3 (top row), cells treated with 1 J/cm2 did not show NADH fluorescence lifetime change. These cells tended to shrink immediately after PDT like drug controls shown in Fig. 2 (middle row). However, different from drug controls, these PDT treated cells returned back to their original size before PDT (as controls in Fig. 2, the top row) although the distribution of mitochondria (i.e., where NADH fluorescent) tended to be more punctuated and aggregated toward to nuclei. Cells treated with 2 J/cm2 (Fig. 3, middle row) became round shapes at 20-40 minutes and after. Their membranes showed blebbing structures (white arrow head 1) at 20-40 minute time point. At 1 hour after PDT, cells show ring-like signatures (white arrowhead 2), nuclei shrinkage, and mitochondria relocation toward to nuclei, which became more significant at 2 hr time point. The NADH fluorescence lifetime increased as early as immediately after PDT (blue color as indicated by white arrow head 3 at 0-20 minute micrograph), but the cell morphology at this time point remained the same as controls (Fig. 2, top row). Higher NADH fluoresce lifetime (blue dots as indicated by yellow arrow heads) appeared in those enlarged round cells. Finally under light treatment of 6 J/cm2 (Fig. 3, bottom row), cell morphology indicates that cells were not intact anymore, cell nucleus became small, and the nuclear structure was impaired. The NADH fluorescence lifetime shows no change that all cells appeared yellowish.
3.3 NADH fluorescent lifetime and intensity together differentiated between apoptosis, necrosis, and slight cytotoxicity and controls
Figure 4A shows the average results of NADH fluorescence lifetime on 6 samples (i.e., 6 independent replicated experiments) for each condition in control cells (without drug and light, light only, and drug only) and PDT treated cells at the fluence of 1, 2, or 6 J/cm2. Same as we observed in Figs. 2 and 3, NADH fluorescence lifetime only increased under light fluence of 2 J/cm2 in which cells underwent apoptotic pathway (Fig. 1B). Cells treated with 1 and 6 J/cm2 did not show NADH fluorescence lifetime change where cells underwent slight cell death and necrosis, respectively. The cell viability shown in Fig. 1A indicates that 1 J/cm2 treatment condition only induced slight cell death (<40% at 4 hr after PDT). Some of these cells might undergo apoptosis because of caspase-3 activation as shown in Fig. 1B although DNA fragmentation was not seen (Fig. 1C).
To demonstrate that our finding is clinically practical for use in PDT, NADH fluorescence lifetime alone is not enough for this purpose because PDT often induces both necrosis and apoptosis. The change of NADH fluorescence lifetime could only differentiate apoptosis from all other conditions including the controls, slight cytotoxicity, and necrosis (Figs. 3 and 4A), but it could not differentiate necrosis from controls and slight cytotoxicity. Based on many reports showing decreased NADH fluorescence intensity during/after cell death, we performed NADH fluorescence intensity measurement that has potential to differentiate necrosis from controls and slight cytotoxicity. Figure 4B shows that NADH fluorescence intensity decreased significantly (p-value = 0.036 by comparing with controls without light and drug) at light fluence of 6 J/cm2 but did not change at light fluence of 1 J/cm2 within 1 hr after PDT. Data at 0 hr after PDT treatment in Fig. 4B represents results immediately after PDT treatment. NADH fluorescence intensity did not show change between controls without drug and light, light and drug controls (data not shown). Our results suggest that primary apoptotic cells could be identified by measuring the signal of increased NADH fluorescence lifetime, and necrotic cells could be identified by detecting no NADH fluoresce lifetime change but significant decreased NADH fluorescence intensity.
NADH is an electron and proton donor in mitochondria to regulate the energy metabolism of the cell and to participate in many biological processes including DNA repair and transcription. Although NADH is not directly involved in any form of cell death processes, mitochondrial dysfunction is one key feature during all forms of cell death. Several studies in vitro and in vivo investigated the potential of using NADH fluorescence intensity as an intrinsic biomarker of cell death (either apoptosis or necrosis) monitoring . However, few have reported NADH fluorescence lifetime measurements during cell death. Pogue et al. observed no change of NADH fluorescence lifetime in vivo after BPD-PDT treatment . Their observation of no NADH lifetime change is likely because most of cells died through necrosis treated with a high fluence rate of 200 mW/cm2.We have previously observed in vitro an increase of NADH fluorescence lifetime during cell death mediated by mitochondria and PARP-1 activation, and no lifetime change during H2O2 induced necrosis [19,21]. The key finding of this study is that, using the same death inducer (i.e., ALA-PDT with different light doses to induce different death modes), NADH fluorescence lifetime only increased when primary cells underwent apoptosis and otherwise no lifetime changed. The results of this study are consistent with our previous findings by applying different death inducers.
In the in vivo situation, cells die after PDT through a mix of apoptosis and necrosis. The results of this study suggest that NADH fluorescence lifetime increases only when cell death was primarily apoptotic (e.g., cells treated with 2 J/cm2 showed less than 30% cell viability at 4 hr after PDT, caspase-3 activation and DNA fragmentation in Fig. 1, and increased lifetime in Figs. 3 and 4A), but not when less than 40% cells died 4 hr after PDT and part of them died by apoptosis (e.g., cells treated with 1 J/cm2 showed 62% cell viability at 4 hr after PDT, caspase-3 activation in Fig. 1B, but no lifetime change in Figs. 3 and 4A). We envision that monitoring NADH fluorescence lifetime will be unique but only useful for optimizing the treatment efficacy of apoptosis favored treatments such as metronomic PDT. When cells die pre-dominantly by necrosis, we envision that no NADH fluorescence lifetime but a significant decrease in NADH fluorescence intensity will be detected.
It is unclear what biological reaction takes place in those areas with higher NADH fluorescent lifetime values (blue color in the lifetime micrograph, Fig. 3) compared to the other areas of the cell. One thing we are sure is that this increased NADH fluorescence lifetime cannot be explained by increased cellular metabolism as shown in differentiated stem cells . We measured the substrate-supported and State 3 respiration rates immediately after and 1 hr after ALA-PDT for controls and all of three treatment conditions. Both substrate-supported and State 3 respiration rates show significant decrease for all PDT-treated cells immediately after light irradiation as compared to the control (data not shown). Similar results were found in our previous study during PARP-1 mediated cell death where both oxygen consumptions and ATP level decreased while NADH fluorescence lifetime increased . In the same study, we also excluded the possibility that increased NADH fluorescence lifetime was caused by increased NADH content. The increase of NADH fluoresce lifetime during cell death is likely due to some protein conformation changes, which in turn resulted in an increase of NADH interaction with surrounding proteins. Because many protein conformation changes could possibly occur altogether, a single cause that attributed to the change of NADH fluorescence lifetime during cell death may be difficult to comprehend.
It is generally accepted that cells have the characteristics of shrinkage, round shape and membrane blebbing during apoptotic process [27,28]. We did not observe cell shrinkage but round shape and membrane blebbing for PDT-treated cells with 2 J/cm2 in which caspase-3 activation and DNA fragmentation were elicited in the cell dead of apoptosis. We observed cell shrinkage for control cells treated with drug only and for PDT-cells treated with 1J/cm2, but these 2 groups of cells had different kinetic change in cell size within 2 hr FLIM imaging as mentioned in the Results: 1) PDT-treated cells shrank and then resumed to its control size within 2 hr after the treatment; 2) the mitochondria showed more punctuated pattern in PDT-treated cells than did drug control cells. Based on the cell shrinkage effect of low dose ALA-PDT treatment (1 J/cm2), we suspect that the shrinkage of drug control cells was due to the imaging laser light that introduced some PDT effect.
In summary, we demonstrated that the mode of cell death in response to ALA-PDT (i.e., apoptosis or non-apoptosis) is dependent on the light dose applied. The mode of cell death can be differentiable by changes in cellular NADH fluorescent lifetime and intensity: increase in the NADH fluorescent lifetime could be related to the execution of the apoptotic program. Moreover, there was no change in cellular NADH fluorescent lifetime but a decrease in NADH fluorescence intensity indicating non-apoptotic cell death. Our findings suggest that the change of NADH fluorescent lifetime and intensity may be a noninvasive indicator of the mode of PDT-induced cell death
We acknowledge the use of the imaging core facility of National Yang-Ming University in carrying out part of the experiments reported in this study. This work was supported by National Science Council of Taiwan grants NSC94-2321-B-010-004-YC, NSC98-2112-010-002, and NSC97-2320-B-010-013-MY3 and by the Ministry of Education of Taiwan grant 97QC021 multidisciplinary training program for talented college students.
References and links
4. M. T. Jarvi, M. J. Niedre, M. S. Patterson, and B. C. Wilson, “Singlet oxygen luminescence dosimetry (SOLD) for photodynamic therapy: current status, challenges and future prospects,” Photochem. Photobiol. 82(5), 1198–1210 (2006). [CrossRef] [PubMed]
5. M. J. Niedre, A. J. Secord, M. S. Patterson, and B. C. Wilson, “In vitro tests of the validity of singlet oxygen luminescence measurements as a dose metric in photodynamic therapy,” Cancer Res. 63(22), 7986–7994 (2003). [PubMed]
6. K. K. Wang, S. Mitra, and T. H. Foster, “Photodynamic dose does not correlate with long-term tumor response to mTHPC-PDT performed at several drug-light intervals,” Med. Phys. 35(8), 3518–3526 (2008). [CrossRef] [PubMed]
8. J. C. Finlay and T. H. Foster, “Hemoglobin oxygen saturations in phantoms and in vivo from measurements of steady-state diffuse reflectance at a single, short source-detector separation,” Med. Phys. 31(7), 1949–1959 (2004). [CrossRef] [PubMed]
10. G. Yu, T. Durduran, C. Zhou, H. W. Wang, M. E. Putt, H. M. Saunders, C. M. Sehgal, E. Glatstein, A. G. Yodh, and T. M. Busch, “Noninvasive monitoring of murine tumor blood flow during and after photodynamic therapy provides early assessment of therapeutic efficacy,” Clin. Cancer Res. 11(9), 3543–3552 (2005). [CrossRef] [PubMed]
11. B. Chen, B. W. Pogue, I. A. Goodwin, J. A. O’Hara, C. M. Wilmot, J. E. Hutchins, P. J. Hoopes, and T. Hasan, “Blood flow dynamics after photodynamic therapy with verteporfin in the RIF-1 tumor,” Radiat. Res. 160(4), 452–459 (2003). [CrossRef] [PubMed]
12. S. K. Bisland, L. Lilge, A. Lin, R. Rusnov, and B. C. Wilson, “Metronomic photodynamic therapy as a new paradigm for photodynamic therapy: rationale and preclinical evaluation of technical feasibility for treating malignant brain tumors,” Photochem. Photobiol. 80(1), 22–30 (2004). [CrossRef] [PubMed]
13. A. Bogaards, A. Varma, K. Zhang, D. Zach, S. K. Bisland, E. H. Moriyama, L. Lilge, P. J. Muller, and B. C. Wilson, “Fluorescence image-guided brain tumour resection with adjuvant metronomic photodynamic therapy: pre-clinical model and technology development,” Photochem. Photobiol. Sci. 4(5), 438–442 (2005). [CrossRef] [PubMed]
15. B. W. Henderson, S. O. Gollnick, J. W. Snyder, T. M. Busch, P. C. Kousis, R. T. Cheney, and J. Morgan, “Choice of oxygen-conserving treatment regimen determines the inflammatory response and outcome of photodynamic therapy of tumors,” Cancer Res. 64(6), 2120–2126 (2004). [CrossRef] [PubMed]
16. D. K. Bird, L. Yan, K. M. Vrotsos, K. W. Eliceiri, E. M. Vaughan, P. J. Keely, J. G. White, and N. Ramanujam, “Metabolic mapping of MCF10A human breast cells via multiphoton fluorescence lifetime imaging of the coenzyme NADH,” Cancer Res. 65(19), 8766–8773 (2005). [CrossRef] [PubMed]
17. M. C. Skala, K. M. Riching, A. Gendron-Fitzpatrick, J. Eickhoff, K. W. Eliceiri, J. G. White, and N. Ramanujam, “In vivo multiphoton microscopy of NADH and FAD redox states, fluorescence lifetimes, and cellular morphology in precancerous epithelia,” Proc. Natl. Acad. Sci. U.S.A. 104(49), 19494–19499 (2007). [CrossRef] [PubMed]
18. H. W. Wang, Y. H. Wei, and H. W. Guo, “Reduced nicotinamide adenine dinucleotide (NADH) fluorescence for the detection of cell death,” Anticancer. Agents Med. Chem. 9(9), 1012–1017 (2009). [PubMed]
19. H. W. Wang, V. Gukassyan, C. T. Chen, Y. H. Wei, H. W. Guo, J. S. Yu, and F. J. Kao, “Differentiation of apoptosis from necrosis by dynamic changes of reduced nicotinamide adenine dinucleotide fluorescence lifetime in live cells,” J. Biomed. Opt. 13(5), 054011 (2008). [CrossRef] [PubMed]
20. J. S. Yu, H. W. Guo, C. H. Wang, Y. H. Wei, and H. W. Wang, “Increase of reduced nicotinamide adenine dinucleotide fluorescence lifetime precedes mitochondrial dysfunction in staurosporine-induced apoptosis of HeLa cells,” J. Biomed. Opt. 16(3), 036008 (2011). [CrossRef] [PubMed]
21. H. W. Guo, Y. H. Wei, and H. W. Wang, “Reduced nicotinamide adenine dinucleotide fluorescence lifetime detected poly(adenosine-5′-diphosphate-ribose) polymerase-1-mediated cell death and therapeutic effect of pyruvate,” J. Biomed. Opt. 16(6), 068001 (2011). [CrossRef] [PubMed]
22. D. Grebeňová, K. Kuželová, K. Smetana, M. Pluskalová, H. Cajthamlová, I. Marinov, O. Fuchs, J. Souček, P. Jarolím, and 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(2), 71–85 (2003). [CrossRef] [PubMed]
23. N. L. Oleinick, R. L. Morris, and I. Belichenko, “The role of apoptosis in response to photodynamic therapy: what, where, why, and how,” Photochem. Photobiol. Sci. 1(1), 1–21 (2002). [CrossRef] [PubMed]
24. F. Giuntini, L. Bourré, A. J. MacRobert, M. Wilson, and I. M. Eggleston, “Improved peptide prodrugs of 5-ALA for PDT: rationalization of cellular accumulation and protoporphyrin IX production by direct determination of cellular prodrug uptake and prodrug metabolization,” J. Med. Chem. 52(13), 4026–4037 (2009). [CrossRef] [PubMed]
25. B. W. Pogue, J. D. Pitts, M. A. Mycek, R. D. Sloboda, C. M. Wilmot, J. F. Brandsema, and J. A. O’Hara, “In vivo NADH fluorescence monitoring as an assay for cellular damage in photodynamic therapy,” Photochem. Photobiol. 74(6), 817–824 (2001). [CrossRef] [PubMed]
26. H. W. Guo, C. T. Chen, Y. H. Wei, O. K. Lee, V. Gukassyan, F. J. Kao, and H. W. Wang, “Reduced nicotinamide adenine dinucleotide fluorescence lifetime separates human mesenchymal stem cells from differentiated progenies,” J. Biomed. Opt. 13(5), 050505 (2008). [CrossRef] [PubMed]