Irradiation- and Sensitizer-Dependent Changes in the Lifetime of

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Irradiation- and Sensitizer-Dependent Changes in the Lifetime of Intracellular Singlet Oxygen Produced in a Photosensitized Process Elsa F. F. da Silva,†,‡,§ Brian W. Pedersen,†,§ Thomas Breitenbach,† Rasmus Toftegaard,† Marina K. Kuimova,†,|| Luis G. Arnaut,‡ and Peter R. Ogilby*,† †

Center for Oxygen Microscopy and Imaging, Department of Chemistry, Aarhus University, DK-8000 Århus, Denmark Department of Chemistry, University of Coimbra, 3004 Coimbra, Portugal Department of Chemistry, Imperial College, SW7 2AZ London, United Kingdom

)

‡

ABSTRACT: Singlet oxygen, O2(a1Δg), was produced upon pulsed-laser irradiation of an intracellular photosensitizer and detected by its 1275 nm O2(a1Δg) f O2(X3Σg) phosphorescence in time-resolved experiments using (1) individual mammalian cells on the stage of a microscope and (2) suspensions of mammalian cells in a 1 cm cuvette. Data were recorded using hydrophilic and, independently, hydrophobic sensitizers. The microscope-based single cell results are consistent with a model in which the behavior of singlet oxygen reflects the environment in which it is produced; nevertheless, the data also indicate that a significant fraction of a given singlet oxygen population readily crosses barriers between phase-separated intracellular domains. The singlet oxygen phosphorescence signals reflect the effects of singlet-oxygen-mediated damage on cell components which, at the limit, mean that data were collected from dead cells and, in some cases, reflect contributions from both intracellular and extracellular populations of singlet oxygen. Despite the irradiation-induced changes in the environment to which singlet oxygen is exposed, the “inherent” intracellular lifetime of singlet oxygen does not appear to change appreciably as the cell progresses toward death. The results obtained from cell suspensions reflect key features that differentiate cell ensemble from single cell experiments (e.g., the ensemble experiment is more susceptible to the effects of sensitizer that has leaked out of the cell). Overall, the data clearly indicate that measuring the intracellular lifetime of singlet oxygen in a O2(a1Δg) f O2(X3Σg) phosphorescence experiment is a challenging endeavor that involves working with a dynamic system that is perturbed during the measurement. The most important aspect of this study is that it establishes a useful framework through which future singlet oxygen data from cells can be interpreted.

’ INTRODUCTION Singlet oxygen, O2(a1Δg), is the lowest excited electronic state of molecular oxygen. The photosensitized production and subsequent deactivation of singlet oxygen are ubiquitous processes in our oxygen- and light-containing world that have been of interest for over 50 years.14 Part of this interest reflects the fact that singlet oxygen has a characteristic chemistry that results in the oxygenation/oxidation of many organic functional groups.1,5 Among other things, this chemistry can result in the death of mammalian cells, either by initiating the cascade of events that characterize apoptosis or by inflicting sufficient local damage that necrotic death ensues.68 Singlet-oxygen-mediated cell death has been exploited in the medical procedure of photodynamic therapy wherein undesired cells (e.g., cancer cells) are destroyed upon the irradiation of a photosensitizer that is appropriately localized in the tissue to be treated.8,9 Over the years, an extensive effort has been expended to elucidate the mechanistic details of singlet-oxygen-mediated cell death.7,10 One important aspect of this work is the need to quantify the intracellular lifetime of singlet oxygen. Because singlet oxygen is a transient species, the dimensions of its localized “sphere of activity” around its point of photosensitized r 2011 American Chemical Society

production will depend both on its lifetime and on the diffusion coefficient of oxygen in that particular intracellular domain.3 Given the heterogeneous environment in the cell and the plethora of different targets for singlet oxygen reactions, the radius of this activity sphere can have an impact on the way in which singlet oxygen will influence cell function and, ultimately, cell death. Early estimates of the lifetime of intracellular singlet oxygen were obtained from photobleaching experiments11 or by extrapolating data from model solutions in which the 1275 nm O2(a1Δg) f O2(X3Σg) phosphorescence was recorded in time-resolved experiments.12 The perspective thus obtained was that the intracellular lifetime was very short (∼10300 ns), certainly with respect to the lifetime in neat H2O (∼3.5 μs) or neat hydrocarbons (∼1520 μs). More recently, O2(a1Δg) f O2(X3Σg) phosphorescence data have been recorded from steady-state imaging experiments on cells13 and pulsed-laserinitiated time-resolved experiments both on single cells1417 and Received: July 15, 2011 Revised: November 15, 2011 Published: November 27, 2011 445

dx.doi.org/10.1021/jp206739y | J. Phys. Chem. B 2012, 116, 445–461

The Journal of Physical Chemistry B on cell suspensions (i.e., ensembles).1823 A conclusion common to these latest studies is that the intracellular lifetime is longer than that obtained from the early indirect studies, with reported or estimated values ranging from ∼600 ns to ∼3 μs for an H2O-incubated cell. Although this latter range of lifetimes and the corresponding range in spheres of singlet oxygen activity are not large, the differences between respective reports have nevertheless been the source of an ongoing discussion for the past ∼5 years.3,7,24 A relevant point in this discussion is the observation of a H2O/D2O solvent isotope effect on both the intensity and lifetime of the 1275 nm phosphorescence signal obtained from cells.13,15,16,20,25 A key feature of the reasonably well-established mechanism of electronic-to-vibrational energy transfer that defines the effect of solvent on the rate of singlet oxygen deactivation is that the lifetime of singlet oxygen in D2O (∼67 μs) is ∼20 times longer than that in H2O (∼3.5 μs).2,26 Singlet oxygen phosphorescence data recorded from cells incubated with D2O, and independently H2O, likewise reveal a H2O/D2O solvent isotope effect, which indicates that the lifetime of intracellular singlet oxygen must be long enough such that, in the kinetic competition for the deactivation of intracellular singlet oxygen, solvent can effectively compete with reactive components of the cell (e.g., proteins). These kinetic data are also consistent with the observation that the effectivity of cell death mediated by singlet oxygen depends on the H2O/D2O ratio with which the cells had been incubated.2731 Perhaps the most significant recent observations in this field have been reports that the temporal profiles of the O2(a1Δg) f O2(X3Σg) phosphorescence signals evolve with the elapsed time of sensitizer irradiation. Specifically, data recorded from microscope-based single cell experiments17 as well as cuvettebased cell suspension experiments21,22 indicate that the rates of singlet oxygen production and removal both decrease with an increase in the elapsed irradiation time. These observations set the stage for our present study. For this work, we set out to perform both microscope-based single cell experiments and cuvette-based ensemble experiments to better elucidate irradiation-dependent phenomena that influence direct time-resolved measurements of the singlet oxygen lifetime in cells. One goal was to have both single cell and ensemble data performed in the same laboratory so as to avoid the propagation of misunderstandings associated with data recorded in separate laboratories. With the knowledge that the lifetime of singlet oxygen reflects the molecular composition of its immediate environment,2,3,26,32 a point that has also been demonstrated for intracellular singlet oxygen,16 another goal of our present work was to clearly identify factors that influence data recorded using these respective experimental approaches. We conclude that, irrespective of the experimental approach, measuring the “real” intracellular lifetime of singlet oxygen in a direct O2(a1Δg) f O2(X3Σg) phosphorescence experiment is a nontrivial exercise because it involves working with a dynamic system that is being perturbed during the measurement. Nevertheless, we are now able to provide definitive statements regarding irradiation-dependent changes in the temporal profile of O2(a1Δg) f O2(X3Σg) phosphorescence signals as measured in time-resolved photosensitized experiments from cell-containing systems. In turn, this information should provide a useful framework for future studies of the behavior of intracellular singlet oxygen.

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Figure 1. Structures of the hydrophilic (TDFPPS) and lipophilic (TDFPPM) singlet oxygen sensitizers that, heretofore, have not been used in an intracellular τΔ study.

’ EXPERIMENTAL SECTION Instrumentation and Approach. Details of the femtosecond laser system used for sample irradiation have been described.33,34 The instrumentation and methods used to create images of the cell and monitor sample luminescence have likewise been described.17,3537 For the microscope-based experiments, the 420 nm exciting light was focused to yield a beam waist with a diameter of ∼1.4 μm at the image plane.14,34 In these studies, the cells were exposed to an oxygen-saturated medium while data were recorded. For the cuvette-based ensemble studies, the exciting light was a collimated beam ∼5 mm in diameter that propagated through the entire 1 cm path length of suspended cells in an air-saturated medium. Chemicals. The singlet oxygen sensitizers chosen for this work were (1) 5,10,15,20-tetrakis(N-methyl-4-pyridyl)-21H,23Hporphine (TMPyP, Sigma-Aldrich), (2) pyropheophorbide-a (PPa, Frontier Scientific), (3) 5,10,15,20-tetrakis(2,6-difluoro-3-sulfophenyl)porphine (TDFPPS), and (4) 5,10,15,20-tetrakis(2,6difluoro-3-(N-methylsulfamoyl)phenyl)porphine (TDFPPM). The hydrophilic molecule TMPyP and the lipophilic molecule PPa were used as received. TDFPPS and TDFPPM (Figure 1) were synthesized, isolated, and characterized using procedures that have been published.38,39 All of these dyes have an appreciable absorbance at the stated excitation wavelength of 420 nm. Note that, although our 840 nm fs pulses have a broad spectral output (∼1015 nm) as they exit the laser itself,33 the process of frequency-doubling to 420 nm narrows this spectral distribution appreciably. Bovine serum albumin (BSA, MW ∼ 65 kDa, Sigma-Aldrich), the BSA-fluorescein conjugate (Molecular Probes/Invitrogen, catalog # A23015), and D2O (99% D, EurisoTop) were used as received. Sodium azide, 1,2-dimyristoylglycero-3-phosphocholine, 446

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and Triton X-100 (all from Sigma-Aldrich) were likewise used as received. An established procedure was used to prepare the unilamellar vesicles from 1,2-dimyristoylglycero-3-phosphocholine.40 Cells. To maintain continuity with our previous work, HeLa cells were used for the microscope-based studies. The preparation of samples and the descriptions of our cultivating medium and so-called standard maintenance medium (SMM) have been described.25,36 For the present work, slight alterations in our general procedure were implemented. Briefly, the confluent HeLa cells were washed twice with a standard phosphate buffered saline solution, trypsinized with 0.25% trypsin/EDTA to facilitate detachment from the surface of the culture flask, and centrifuged at 1000 rpm for 2 min. The cells were subsequently seeded on polylysine-coated coverslips at a concentration of (46) 104 cells/mL. The cultivating medium used was Eagle’s minimal essential medium with Earl’s Salts supplied with 10% fetal calf serum, 1% L-glutamine, 100 U/mL penicillin, 100 μg/mL streptomycin, and 1% nonessential amino acids. After seeding, the cells were left for at least 24 h to settle and restart growth. Thereafter, the sensitizer to be incorporated was added to the medium, and the cells incubated for an additional 24 h. When the hydrophobic molecule TDFPPM was used, a maximum of 1% ethanol was added to the medium to facilitate dissolution of 12.5 μM of the dye. The exchange of intracellular H2O with D2O was achieved by changing the tonicity of the medium, as previously described,25 with the exception that, for some experiments, cells were only exposed to D2O-based SMM for a period that did not exceed ∼15 min and the laser experiment was performed immediately thereafter. Samples were prepared in the same way for the experiment in which A-549 epithelial cells were used. For the cuvette-based ensemble studies, it was more convenient to use nonadherent HL-60 cells (Human promyelocytic leukemia cells). These cells were maintained in 75 cm2 cultivation flasks in RPMI 1640 medium supplied with 10% fetal calf serum, 1% L-glutamine, 100 U/mL penicillin, and 100 μg/mL streptomycin at a concentration of ∼5 105 cells/mL. In a typical preparation, the cells were collected from the flask, centrifuged at 1000 rpm for 2 min, resuspended, and incubated with a 50 μM solution of TMPyP dissolved in the cultivating medium for 24 h at 37 °C. When the hydrophobic sensitizer TDFPPM was used, a maximum of 1% ethanol was added to the medium to facilitate dissolution of 12.5 μM of the dye. When PPa was used, a maximum of 1% DMSO was added to the medium containing 5 μM of the dye, and the system was incubated for 3 h (i.e., PPa incorporation into the cell is comparatively rapid). For selected experiments used to examine the effect of the medium, cell populations were also independently incubated with a solution of the sensitizer dissolved in our SMM. After incubation, the cell suspension was centrifuged (1000 rpm for 2 min) and the medium removed. Samples were then resuspended in D2Obased hypertonic SMM for 3 min to remove intracellular H2O via osmosis25 and then recentrifuged and suspended again in D2O-based SMM either with or without BSA, depending on the experiment to be performed. For a sample used in a given optical experiment, cells were present at a concentration of ∼106 cells/mL and the sample was continuously stirred to maintain the suspension.

phosphorescence experiments are reported, it is nevertheless useful to briefly review the model used when the kinetics of singlet oxygen production and decay in a photosensitized process are quantified. The sensitizers used in this study produce singlet oxygen in a collision-dependent process of energy transfer from the lowest energy triplet state, T1, to O2(X3Σg). The sensitizer fluorescent state, S1, is sufficiently short-lived to preclude quenching by O2(X3Σg) under our conditions. Thus, the sensitizer triplet state, produced via S1 f T1 intersystem crossing, is the sole precursor to singlet oxygen (eq 1). kform

kΔ

f O2 ða1 Δ g Þ f singlet oxygen removal T1 s 3 O2 ðX Σg Þ

ð1Þ

For our experiments, it is convenient to express the sum of all processes by which singlet oxygen can be removed (i.e., all chemical reaction and physical quenching channels) through the first-order rate constant kΔ.3 The reciprocal of this rate constant, 1/ kΔ, thus defines the lifetime of singlet oxygen, τΔ. Solving eq 1 for the concentration of singlet oxygen at a given time t after the rapid, pulsed-laser production of an initial population of sensitizer triplet states at t = 0, [T1]0, yields the expression shown in eq 2, where kT represents all channels for T1 removal, including processes that do not result in singlet oxygen production. As seen in eq 2, it is kT, not kform, that defines the rate of singlet oxygen production. The time-dependent intensity of the O2(a1Δg) f O2(X3Σg) phosphorescence signal is proportional to [O2(a1Δg)]t.

½O2 ða1 Δ g Þt ¼

kform ½O2 ðX3 Σ g Þ½T1 0 kΔ kT

fexpðkT tÞ expðkΔ tÞg

ð2Þ It should be apparent from eq 2 that the singlet oxygen lifetime is most accurately quantified under conditions where kΔ < kT. This inequality generally occurs in weakly deactivating solvents (e.g., deuterated and/or hydrocarbon solvents), in the presence of low concentrations of reactive solutes and quenchers, and in media where the O2(X3Σg) concentration is comparatively high. Although a living cell is inhomogeneous and characterized by domains with different O2(X3Σg) concentrations and molecular composition, we are nevertheless generally faced with conditions that yield a situation where kΔ ≈ kT or kΔ > kT. For the latter case in particular, even if one can obtain kT in an independent experiment (as one should), the data that yields kΔ appear on the short rising portion of the time-resolved phosphorescence signal and are difficult to accurately quantify. Moreover, data recorded at times close to that at which the pulsed excitation light is delivered often show the effects of scattered light and luminescence from optics used,41 and this can preclude the accurate use of eq 2 in this time domain (even if a femtosecond laser is used as the excitation source15). Although this latter issue can sometimes be addressed through the use of expressions that are more extensive than eq 2,42,43 one is still left with a difficult problem. As outlined in the Introduction, there is a unique and characteristic solvent isotope effect on the lifetime of singlet oxygen (i.e., for the neat solvents D2O and H2O, kΔD2O = (67 μs)1 = 1.5 104 s1 and kΔH2O = (3.5 μs)1 = 2.9 105 s1). Most importantly, singlet oxygen experiments performed on cells likewise show a H2O/D2O isotope effect that is indicative of a solvent-dependent change in the magnitude of kΔ. Specifically,

’ RESULTS AND DISCUSSION Kinetic Model. Although it has been discussed at length in a plethora of papers in which time-resolved singlet oxygen 447



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kinetic profile of the O2(a1Δg) f O2(X3Σg) phosphorescence signal recorded. It is important to note, however, that the excitation light is scattered appreciably by the cell (Figure 2).36 Therefore, although a portion of the 1275 nm phosphorescence signal detected will come from a localized intracellular domain in or near the laser focal volume, an appreciable amount of the signal still comes from singlet oxygen produced in other parts of the cell simply because the sensitizer in these other areas readily absorbs the incident light that has been scattered. Because the scattered light is less intense, the O2(a1Δg) f O2(X3Σg) phosphorescence emitted from these latter spatial domains will likely contribute to the overall signal observed with a kinetic profile that is different from that in the laser focal volume. In short, pronounced oxygenation of reactive substrates within the localized spatial domain of the focused laser will not be the only thing that influences O2(a1Δg) f O2(X3Σg) phosphorescence data in our single cell experiments. [Based solely on controlling the excitation source, a true spatially localized singlet oxygen experiment can best be achieved using two-photon excitation.34,36 Unfortunately, the number of excited states typically produced in a two-photon experiment is sufficiently small as to make the O2(a1Δg) f O2(X3Σg) phosphorescence signal even more difficult to detect in a single cell experiment.34] Influence of Irradiation Power and Elapsed Irradiation Time. At present, to collect a time-resolved O2(a1Δg) f O2(X3Σg) trace from a cell in a medium containing 100% D2O, we typically irradiate TMPyP at 420 nm with a fluence of 7 kJ/cm2 and collect the 1275 nm emission for a period of 3 min using a multichannel scaler and a laser repetition rate of 1 kHz. Under these conditions, we generally observe morphological changes in the cell characteristic of necrotic death (Figure 3, left-hand column). Although cell death and/or changes in cell function can be assessed using a variety of assays, the use of bright field or phase contrast images to record morphology changes (e.g., membrane located vacuole formation, shrinkage, loss of adhesion to the plate, condensation of nuclear chromatin) is an established approach that combines accuracy with ease of implementation.31,35,44,45 Of course, such general morphological features are likely the result of changes in multiple cellular processes. With respect to the irradiation-dependent formation of vacuoles shown in the left-hand column of Figure 3, analogous vacuoles are likewise formed upon sensitizer irradiation in H2Oincubated cells. However, the time for vacuole appearance in the H2O-containing cells is longer than that in D2O-containing cells (i.e., evidence that singlet oxygen is more cytotoxic in D2O than in H2O).36 In contrast to the data shown on the left-hand side of Figure 3, irradiation of an intracellular sensitizer in a O2(a1Δg) f O2(X3Σg) phosphorescence experiment can, in some cases, cause no apparent change in cell morphology as assessed through a bright field or phase contrast image (Figure 3, right-hand column). In these cases, we find that the 1275 nm phosphorescence signals are generally more intense and, thus, more readily observed. The kinetics of these signals clearly evolve as a function of the elapsed irradiation time; both the rise and fall time constants obtained from a fit of eq 2 to the data (i.e., 1/kT and 1/kΔ) get longer with an increase in the elapsed irradiation time. The differences in the data shown in the left- and right-hand columns of Figure 3 principally reflect the fact that, in the lefthand column, the cells were alive at the start of the experiment, whereas, in the right-hand column, the cells were dying or already

Figure 2. (a) Brightfield image of a neuroblastoma cell containing the sensitizer protoporphyrin IX (PpIX). The white dot approximates the cross-sectional size of the focused laser beam used to irradiate the cell at 420 nm. (b) Image of the same cell based on the fluorescence of PpIX excited by the focused laser beam. The data clearly show that the incident laser light is scattered and creates PpIX excited states throughout the cell. These images are modified versions of previously published data.36

kΔ becomes smaller in D2O-incubated cells. One can exploit this solvent isotope effect to obtain more accurate values of kΔ through eq 2 simply because one moves further away from the undesired limiting condition of kΔ > kT. In these latter experiments, one must consider to what extent the cell responds adversely to the exchange of H2O for D2O. Data accumulated thus far indicate that, over the time course in which our experiments are performed, the presence of D2O by itself does not appear to have pronounced adverse effects on cell viability.25,31 Of course, by decreasing the magnitude of the solvent-dependent term in kΔ, one effectively increases the concentration of singlet oxygen available for reaction with cellular components which, in turn, can accelerate the appearance of morphological/chemical changes associated with cell death (i.e., singlet oxygen becomes more cytotoxic in D2O). As we will shortly establish, this in itself can influence the values of kΔ recorded in a time-resolved O2(a1Δg) f O2(X3Σg) phosphorescence experiment. Microscope-Based Single Cell Experiments. Influence of Spatial Localization and Scattered Light. In our single cell experiments, the excitation light is typically focused to a beam waist, at the sample, of ∼1 μm. The diameter of this spot is thus smaller than the diameter of a typical cell (∼20 30 μm).34,36 One can readily estimate that, in the laser focal volume, one could have appreciable singlet-oxygen-mediated changes in reactive substrates (e.g., protein oxidation).22,32 Such changes in the local environment could certainly have a significant effect on the 448



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Figure 3. Bright field images and 1275 nm singlet oxygen phosphorescence traces obtained from D2O-incubated cells containing the hydrophilic sensitizer TMPyP that were alive (left column) and that were dead or dying (right column) at the start of the experiment. These differences in cell viability reflect the influence of the incubation medium used in these experiments (i.e., use of our SMM yields a larger fraction of dead/dying cells). After moderate exposure of TMPyP to focused laser light (fluence of 7 kJ/cm2 for 3 min), vacuole formation indicative of cell necrosis was apparent for the cells that were initially alive, whereas essentially no morphological changes were observed upon irradiation of dead/dying cells (fluence of 7 kJ/cm2 for 9 min). The kinetic traces recorded after 9 min of elapsed light exposure (solid line in left column, O in right column) yield different rate constants for singlet oxygen formation and decay than those recorded after 3 min of elapsed irradiation (dotted line in left column, 9 in right column). In comparing the signal-to-noise levels in the respective kinetic traces, it is useful to note that the data in the left-hand column were recorded using 3 μs resolution (i.e., each data point reflects a larger time window), whereas the data in the right-hand column were recorded using 1 μs resolution; a difference that only further supports our contention that “the deader, the easier”.

dead when the experiment started. The lack of light-induced vacuole formation in this latter case is most likely a ramification of the fact that, in a dead or dying cell, the “machinery” that gives rise to the vacuole phenomenon may simply no longer work. On the basis of bright field images, certainly those shown in the left-hand column of Figure 3, it is reasonable to conclude that, over the typical time period required for acquisition of the 1275 nm phosphorescence signal, τΔ values obtained from single cells reflect an average of the effects of an evolving intracellular environment. This is substantiated by the fact that data recorded from a given cell in successive acquisition periods evolve in time; the rates of both signal appearance and disappearance decrease with an increase in the elapsed irradiation time. Under conditions

such as those shown in Figure 3, using the hydrophilic sensitizer TMPyP, we generally record values of τΔ that evolve from ∼15 to ∼40 μs for D2O-incubated cells (Table 1). These data are consistent with those we have obtained in previous single cell experiments.17 This phenomenon of irradiation-induced changes in τΔ has also been observed in ensemble experiments using suspensions of H2O-incubated cells.21,22 Specifically, Schlothauer et al. report that τΔ evolves from ∼0.5 to ∼1.0 μs upon prolonged irradiation of the hydrophobic sensitizer pheophorbide-a.21 In a second study by this same group,22 it was also noted that, after a O2(a1Δg) f O2(X3Σg) measurement on a given ensemble of cells, although the “outer shape of the cells was still normal” 449



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Table 1. Summary of τΔ Data from D2O-Incubated Mammalian Cells τΔ/μsa

commentsb

Single Cells hydrophilic sensitizers TMPyP TDFPPS

irradiation dependence

∼15 f ∼40

w/BSA

∼1520

intracellular τΔ for dead/dying cell


∼2540

reflects contribution of extracellular singlet oxygen

w/BSA

∼20

intracellular τΔ for dead/dying cell


f∼30


w/BSA

f∼17 f∼17

intracellular τΔ for dead/dying cell intracellular τΔ for dead/dying cell at the limit of low [chlorin]16


hydrophobic sensitizers TDFPPM chlorin

Cell Suspensions hydrophilic sensitizers TMPyP


∼40


w/BSA

∼4.5

expected for BSA quenching and indicates that

w/BSA

6(2

skin fibroblasts from Jimenez-Banzo et al.20

most singlet oxygen detected is extracellular hydrophobic sensitizers TDFPPM PPa


see text for discussion of problems

w/BSA

see text for discussion of problems


f∼27

data interpretation is not straightforward

w/BSA

kΔ in eq 2) and yield τΔ = 30 μs. The data shown here are representative in that the signal-to-noise ratio in the TDFPPM experiments was always smaller than that in the TDFPPS experiments.

influenced by the domain in which it is produced. We return to this point in a subsequent section. In any event, to further elucidate some of these points, there is a need to collect more data that addresses the issue of sensitizerdependent intracellular singlet oxygen lifetimes. To this end, we used two sensitizers that, heretofore, have not been exploited for intracellular τΔ experiments, TDFPPS and TDFPPM (Figure 1). Although the structural differences between TDFPPS and TDFPPM appear subtle, independent solubility experiments on these compounds are consistent with calculations of the 454



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Figure 7. Images of cells based on the fluorescence of lipophilic sensitizers that show dynamic light-induced changes. Data from TDFPPM-containing cells are shown in the left-hand column (a = short elapsed irradiation time, b = prolonged irradiation time). In this case, experiments were performed with A-549 human lung adenocarcinoma epithelial cells, and data from the same group of cells are shown in the respective panels. Data from chlorincontaining HeLa cells are shown in the right-hand column (c = short elapsed irradiation time in H2O-based medium, d = prolonged irradiation time in D2O-based medium). Here, data from two different groups of cells are shown, and panel c was recorded using a confocal microscope whereas panel d is a simple widefield image recorded using a CCD camera. In all cases, the cell nuclei appear as darker spots (white arrows). Panel d has previously been published.16 The images are shown in “false” colors assigned by the data-handling software.

generally appears that BSA has no effect on the appearance rate of singlet oxygen phosphorescence in the TDFPPM experiment. These observations could reflect, in part, the well-established fact that, with similar sensitizer triplet-state lifetimes, singlet oxygen production is slower in aqueous solvents simply because the concentration of oxygen is roughly 10 times smaller than that in hydrocarbon solvents (i.e., from eq 2, kT(aqueous) < kT(hydrocarbon)). We return to related aspects of this issue in our discussion below of data recorded from cell suspensions. In any event, the data in Figure 6 provide further evidence that the kinetic behavior of singlet oxygen reflects its local environment. Influence of Adding Singlet Oxygen Quenchers: H2O. The lifetime of singlet oxygen in neat D2O (∼67 μs)53 is significantly longer than that in neat H2O (∼3.5 μs).63 Thus, increasing the percentage of H2O in a D2O-based system is tantamount to adding aliquots of a singlet oxygen quencher. We previously reported that singlet oxygen lifetimes obtained from TMPyPsensitized single cell experiments indeed get shorter as the amount of H2O added to the system is increased.25 The rate constant for singlet oxygen quenching obtained from these early experiments, (3.0 ( 0.7) 103 s1 M1,25 is consistent with that expected for H2O [i.e., (3.5 μs 55 M)1 ∼ 5 103 s1 M1]. For the present study, in light of our evidence for an extracellular population of singlet oxygen, we performed new H2O quenching

experiments using TMPyP and, independently, TDFPPM as sensitizers under conditions where BSA was added to the medium. We can draw several important conclusions from these H2O/ D2O data. Let us start with the data recorded using the hydrophilic sensitizer TMPyP (Figure 8). First, the plot of kΔ against the concentration of added H2O is reasonably linear and yields kq = (2.5 ( 0.2) 103 s1 M1, which is consistent with our earlier results obtained in the absence of BSA and for the general quenching of singlet oxygen by H2O (vide supra). Second, if we accept that singlet oxygen is more cytotoxic in D2O than H2O, then, upon irradiation of a system with more D2O and the production of a cell that is “more dead”, components of the cell should appear as a less effective quencher of singlet oxygen in the kinetic competition with solvent-mediated deactivation. With the latter in mind, one might expect to see curvature in a plot of kΔ against the concentration of added H2O. It is clear from the data in Figure 8 that, within the errors of our experiment, we obtain a linear plot of kΔ against the concentration of added water. Thus, for these cells that were already dead/dying at the start of the experiment (i.e., they were incubated for 24 h in our maintenance medium), most of the pertinent intracellular damage was already done and/or this damage is irrelevant to the intracellular domains from which a significant fraction of our 455



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domain of a bimolecular process. On the other hand, the rate constants for quenching by NaN3 are comparatively large [(1 ( 0.1) 108 and (8 ( 1) 108 s1 M1 for TMPyP- and chlorinsensitized sensitized intracellular processes, respectively]16 and approach the so-called diffusion-controlled limit of a bimolecular process, certainly in the more viscous intracellular domains.16,17 With these points in mind, recall now that reactions that occur at or near the diffusion-controlled limit are more sensitive to changes/differences in local viscosity than those that occur at the pre-equilibrium limit; a point that has been explicitly made with respect to the quenching of singlet oxygen.6567 Thus, it is reasonable to see differences in the rate constant for NaN3mediated removal of singlet oxygen sensitized by a hydrophilic and independently, hydrophobic sensitizer, particularly in a cell where local changes in viscosity can be pronounced during photoinduced cell death.17 The fact that kq for the TMPyPsensitized process, (1 ( 0.1) 108 s1 M1, is appreciably less than the kq in bulk water, (5.1 ( 0.1) 108 s1 M1, indicates that this quenching experiment was not adversely influenced to a great extent by an extracellular singlet oxygen population; the magnitude of the rate constant observed is determined by the intracellular viscosity.16 A more subtle point that also needs clarification is the fact that the rate constant for quenching of singlet oxygen by NaN3 in the chlorin-sensitized process [(8 ( 1) 108 s1 M1] is greater than that for the quenching of singlet oxygen in a bulk nonviscous aqueous solution [(5.1 ( 0.1) 108 s1 M1].16 The most likely explanation for this phenomenon is that, for a singlet oxygen population produced in a lipophilic membrane-localized domain, quenching by the nominally hydrophilic NaN3 occurs principally at the membrane interface under conditions in which NaN3 penetrates an appreciable distance into the membrane. With respect to the latter, there is indeed precedence for the penetration of a small charged solute into a liposomal bilayer.68 Thus, the comparatively large value of kq in the chlorin-sensitized process reflects a solvent effect on the quenching rate constant. These data again support the thesis that, even though singlet oxygen readily crosses barriers between phase separated domains, the overall behavior of singlet oxygen reflects the intracellular domain in which it is produced. Cuvette-Based Cell Suspension Experiments. To complement our microscope-based single cell studies, we carried out a series of independent experiments using cell suspensions. For these studies, we chose to use HL-60 cells specifically because they are nonadherent. Even though it is possible to detect time-resolved O2(a1Δg) f O2(X3Σg) phosphorescence signals from an H2O-incubated suspension of cells, one is still faced with the difficult task of decoupling the rate constant for singlet oxygen removal, kΔ, from the rate constant that determines the rate of singlet oxygen formation, kT (eq 2). Simply put, because kT ∼ kΔ in an H2Oincubated cell, and because the time constants involved (i.e., 1/kT and 1/kΔ) are comparatively short, it is difficult to accurately quantify kΔ, even with the appropriate control experiments. Thus, we opted only to demonstrate some key points using D2O-incubated cells. Experiments were performed under a variety of conditions using TMPyP, PPa or TDFPPM as a sensitizer. Experiments with TMPyP as the Sensitizer. Representative O2(a1Δg) f O2(X3Σg) phosphorescence traces recorded using TMPyP as the sensitizer are shown in Figure 9. The following

Figure 8. Plot of kΔ against the concentration of H2O in a D2O-based medium containing 0.75 mM BSA. The hydrophilic molecule TMPyP was used as the sensitizer. Each point represents the average of data from at least 5 cells, and the slope yields kq = (2.5 ( 0.2) 103 s1 M1. Because these cells were incubated for 24 h in our “maintenance” medium prior to the experiment, it is clear that we started with cells that were already dying/dead.

singlet oxygen signal originates. The latter conclusion is likely to be the most pertinent given our independent observation that the intracellular value of τΔ recorded in the presence of BSA does not depend appreciably on the elapsed time of sensitizer irradiation (i.e., τΔ ∼ 1520 μs, vide supra). As with other experiments performed using a lipophilic sensitizer, it was difficult to obtain systematic results that could be used to obtain an accurate value of kq for H2O quenching of intracellular singlet oxygen generated using TDFPPM. Nevertheless, we consistently observed decreases in τΔ for TDFPPMsensitized singlet oxygen as the concentration of H2O in the medium was increased. This observation is consistent with those obtained from the TMPyP experiment and our independently published H2O quenching experiments in which chlorin was used as the sensitizer.16 The data thus indicate that, for singlet oxygen generated in lipophilic intracellular domains, H2O is still an effective a quencher. This result is entirely in keeping with our model in which singlet oxygen readily crosses the interface between hydrophobic and hydrophilic domains. Pre-equilibrium vs Diffusion-Controlled Quenching. At this juncture, it is useful to revisit our previously published data on the quenching of intracellular singlet oxygen by NaN3.16 Although NaN3 is known to be cytotoxic over long periods of exposure (∼>6 h),64 we have independently ascertained using several viability assays that our cells do not show an adverse response to NaN3 over the time course of our experiments (∼1 h).16 Indeed, at comparatively high concentrations of NaN3 (∼10 mM) added to the medium, one can protect against the adverse effects of singlet oxygen produced in a photosensitized reaction and maintain a D2O-incubated cell viable for at least 1 h upon irradiation.31 For the present discussion, we need to address the fact that the rate constants for quenching by NaN3 depended significantly on whether a hydrophilic or a hydrophobic sensitizer was used. One must first recognize that the absolute magnitude of the rate constant for quenching by H2O, for example, is small (∼3 103 s1 M1) and falls in the so-called pre-equilibrium 456



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decreased appreciably and our data yielded τΔ ∼ 0.6 μs with no change in the time constant for singlet oxygen formation. This is consistent with the expectation given the rate constant of 5 108 s1 M1 for the quenching of singlet oxygen by NaN3 in water.16 (3) For cells in the presence of 0.75 mM BSA, the intensity of our O2(a1Δg) f O2(X3Σg) phosphorescence signal likewise decreased appreciably and we recorded τΔ ∼ 4.5 μs (Figure 9b). Our observation of such a pronounced BSA-dependent effect on the time-resolved O2(a1Δg) f O2(X3Σg) signal is consistent with (a) TMPyP-sensitized data recorded from suspensions of skin fibroblasts by Jimenez-Banzo, et al.20 and (b) data sensitized by a hydrophilic aluminum phthalocyanine recorded from suspensions of leukemia cells by Niedre et al.18 Independent control experiments performed using the BSA-fluorescein conjugate indicate that, as with our HeLa cells, BSA does not enter our HL-60 cells during our experiments. These observations indicate that, in contrast to our microscope-based single cell experiments, an appreciable amount of the singlet oxygen phosphorescence signal detected in our TMPyP-sensitized cell suspension experiments and, by inference, in the suspension experiments of Jimenez-Banzo et al.20 and Niedre et al.,18 comes from an extracellular population of singlet oxygen. One thing common to all three of these studies is the use of a water-soluble sensitizer. One possible explanation for the data is that, after sensitizer incorporation, the procedures used to wash the cells with sensitizer-free medium are ineffective and do not completely remove residual extracellular sensitizer. However, a more likely possibility is that once the cells are washed and resuspended in a sensitizer-free medium, the intracellular sensitizer diffuses out to the extracellular domain along the newly established concentration gradient. This process will be enhanced upon light-induced perturbation of the cells and the associated changes that make the cell membrane more permeable to the sensitizer (vide supra, Figure 5). It is important to recognize that the effect of sensitizer diffusion into the extracellular medium will be more pronounced in the O2(a1Δg) f O2(X3Σg) phosphorescence measurement performed on a suspension of cells because the excitation light propagates through a 1 cm thick sample that is dominated more by the medium than by the cells. On the other hand, in our microscope-based single cell studies, the incident light is focused into the cell and the extracellular medium is irradiated only as a consequence of scattered light (i.e., the fraction of extracellular excitation is much smaller in the microscope-based study). The τΔ value of ∼4.5 μs obtained from these cell suspensions with BSA is consistent with what is expected for the quenching of singlet oxygen by BSA in a homogeneous solution.14,48 [i.e., at 0.75 mM BSA, a τΔ value of 4.5 μs points to the expected value of ∼3 108 s1 M1 for the quenching of singlet oxygen by BSA.] As an addendum to this discussion on the differences between our microscope-based experiments and the cell suspension experiments, we must also consider that, in the suspension experiments, collision- and shear-dependent perturbations associated with the process of stirring the cells may also exacerbate the permeability of an already leaky membrane associated with a light-perturbed cell (the sample is continuously stirred to maintain the suspension of cells during the measurement). Indeed, the phenomenon of a stirring-dependent perturbation has been documented in independent experiments on HL-60 cells.69

Figure 9. Time-resolved O2(a1Δg) f O2(X3Σg) phosphorescence traces obtained from suspensions of HL-60 cells in a D2O-based medium. The sensitizer used for these particular experiments, TMPyP, was irradiated at 420 nm using an average power of 1 mW (laser operated at 1 kHz, therefore 1 μJ/pulse). Data were acquired over a period of 1 min. A fit of eq 2 to the data is shown as a solid line in each trace. (a) Trace recorded after an elapsed irradiation period of 3 min, yielding τΔ = 40 μs. (b) Data recorded under the same conditions as that in panel a, only with 0.75 mM BSA present in the medium; τΔ ∼ 4.5 μs.

observations were repeatedly made from independent experiments using D2O-incubated cells: (1) In the absence of the added quenchers BSA and NaN3, we recorded singlet oxygen lifetimes in the range of ∼3942 μs, with time constants for singlet oxygen formation of ∼2.04.5 μs (Figure 9a). The latter were consistent with independent measurements with these cells of TMPyP phosphorescence performed at 900 nm, a wavelength where there is no emission from singlet oxygen. Although an irradiation-dependent increase in τΔ was observed under our conditions, this increase was subtle and occurred only over the range ∼3942 μs. Identical lifetimes were recorded from cells that had been incubated in our SMM instead of the cultivating medium. (2) For cells in the presence of 3.2 mM NaN3, the intensity of the O2(a1Δg) f O2(X3Σg) phosphorescence signal 457



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Thus, we performed experiments using TDFPPM. Unfortunately, as in our single cell studies, we found it difficult to obtain consistent results with this sensitizer in samples of suspended cells. Part of the problem may reflect the difficulties associated with using a hydrophobic dye that tends to aggregate in aqueous media. Although we were able to observe a rapidly decaying 1275 nm emission signal (τ < ∼4 μs) from D2O-based suspensions of HL60 cells containing TDFPPM upon irradiation at 420 nm, this signal does not have an appreciable component of singlet oxygen phosphorescence, as ascertained in a NaN3 quenching study. In light of our results using PPa (vide infra), this observation may reflect the way in which TDFPPM is incorporated and/or localized in HL-60 cells (i.e., (1) TDFPPM tends to aggregate and, as such, may not be efficiently incorporated and/or (2) a high membrane-localized concentration may not result in appreciable singlet oxygen production and/or may efficiently quench the singlet oxygen that is made). Experiments with PPa as the Sensitizer. Given our inability to record a singlet oxygen phosphorescence signal from a suspension of TDFPPM-containing HL-60 cells, we set out to see if acceptable data could be recorded from a cell suspension using a different lipophilic sensitizer. Pyropheophorbide-a, PPa, is readily incorporated into a variety of cells and localizes in the plasma membrane and other membranes found in the cytoplasm.35,51,70,71 Moreover, despite facile irradiation-induced bleaching of this dye, we have been able to record singlet oxygen phosphorescence signals from these cells in microscope-based experiments.35 Thus, it seemed reasonable to try to record time-resolved singlet oxygen phosphorescence data from a suspension of HL-60 cells containing PPa. The following observations were made upon 420 nm irradiation of PPa in suspensions of HL-60 cells (Figure 10): (1) An appreciable amount of the 1275 nm emission signal observed can be attributed to singlet oxygen phosphorescence on the basis of a NaN3 quenching experiment. (2) The rates of signal appearance and disappearance clearly decrease with an increase in the elapsed irradiation time. (3) The addition of BSA to the surrounding medium removes a subtle long-lived component on the signal decay (i.e., removes the 18 and 27 μs components on the data sets of early and prolonged irradiation, respectively). (4) The addition of BSA to the surrounding medium also causes the short-lived component of signal decay to increase (i.e., τ becomes longer) by ∼30%. Unfortunately, the interpretation of these data is not as straightforward as it might otherwise seem. In the least, from the BSA experiment, it appears that the bulk of the signal that we assign to singlet oxygen indeed originates from inside the cell. Moreover, even though we are working with D2O-incubated cells, the lifetime of singlet oxygen recorded from this cell suspension certainly appears to be shorter than that recorded in our single cell experiments. One might argue that the shorter values of τΔ recorded from this suspension of PPa-containing cells could be a manifestation of a higher concentration of effective intracellular singlet oxygen quenchers (e.g., proteins) that have yet to be fully oxygenated/ oxidized upon elapsed irradiation (i.e., the irradiation fluence required to record a singlet oxygen signal from cell suspensions is less than that used in our single cell experiments, vide supra). However, this interpretation contradicts the conclusions obtained through a variety of independent single cell experiments

Figure 10. Time-resolved 1275 nm emission traces recorded upon 420 nm irradiation of PPa in suspensions of HL-60 cells in a D2O-based medium. (a) Data recorded in the absence of added BSA. The trace recorded after an elapsed irradiation period of 1 min (O) is best fit as the sum of two exponentially decaying functions with τ1 = 2.9 ( 0.1 μs and τ2 = 18 ( 1 μs (line shown). The trace recorded after an elapsed irradiation period of 4 min (9) is best fit as the difference of exponential functions (e.g., eq 2) with τrise = 0.8 ( 0.1 μs and τdecay1 = 4.1 ( 0.1 μs and τdecay2 = 27 ( 3 μs (line shown). The trace recorded after an elapsed irradiation period of 4 min (1), but with added NaN3 at 50 mM, is best fit with a single exponentially decaying function with τ = 3.2 ( 0.1 μs (line shown). (b) Data recorded in the presence of 0.75 mM BSA. The trace recorded after an elapsed irradiation period of 1 min (O) is best fit with a single exponentially decaying function with τdecay = 3.7 ( 0.1 μs (line shown). The trace recorded after an elapsed irradiation period of 4 min (9) is best fit as the difference of two exponential functions with τrise = 1.0 ( 0.1 μs and τdecay = 5.4 ( 0.1 μs (line shown). The trace recorded after an elapsed irradiation period of 4 min (2), but with added NaN3 at 50 mM, is best fit with a single exponentially decaying function with τ = 3.0 ( 0.1 μs (line shown).

On the basis of this albeit limited data set, we nevertheless conclude that quantifying intracellular values of τΔ can be difficult, if not impossible, when a hydrophilic sensitizer is used in a cell suspension experiment. Experiments with TDFPPM as the Sensitizer. Having questioned the credibility of using a hydrophilic sensitizer in a cell suspension experiment, the inference is that more accurate data could be obtained through the use of a hydrophobic sensitizer. 458


The Journal of Physical Chemistry B (i.e., we have ascertained that a τΔ value of ∼15 20 μs likely represents the “inherent” intracellular lifetime in a D2Oincubated cell, irrespective of the extent of irradiation). A more likely explanation for these PPa data is that, in the membrane-localized domains where singlet oxygen is produced, quenching by high local concentrations of PPa would yield a shorter value of τΔ. The most convincing support for this interpretation comes from independent PPa experiments we performed on (1) unilamellar vesicles (i.e., liposomes) prepared from 1,2-dimyristoyl-glycero-3-phosphocholine and (2) single HeLa and neuronal cells. In the first case, liposomes were prepared and then exposed to an aqueous solution of PPa (with 1% added DMSO to facilitate PPa dissolution). Both the absorption and fluorescence spectra of these PPa-containing liposomes were characteristic of PPa aggregation (i.e., spectrally broadened absorption bands, weak fluorescence intensity). Most importantly, we were unable to detect a PPa-sensitized singlet oxygen phosphorescence signal from these liposomes. The latter could be due to efficient quenching by high local concentrations of PPa and/or by the inability of PPa aggregates to make appreciable amounts of singlet oxygen. However, upon the addition of the surfactant Triton X-100 to the liposome suspension, we were able to see an appreciable singlet oxygen signal that correlated to fragmentation of the liposomes into smaller units in which the PPa was presumably less-tightly packed. Analogous observations were made from HeLa cells and neurons into which PPa had been incorporated; addition of Triton X-100 led to changes in the PPa absorption and fluorescence spectra characteristic of a decrease in the extent of PPa aggregation and this, in turn, was accompanied by the appearance of a singlet oxygen phosphorescence signal. This interpretation based on high membrane-localized concentrations of PPa is also consistent with our earlier chlorin experiments where τΔ decreased with an increase in the time to which the cell was exposed to sensitizer-containing medium.16 Unfortunately, corresponding experiments performed thus far with our HL-60 cells have not yielded the expected changes; changing either the PPa concentration in the medium or the time of incubation does not result in corresponding changes in the kinetics of the singlet oxygen signal. These latter data may indicate that, irrespective of the PPa concentration in the extracellular medium, local concentrations in the HL-60 membranes may not appreciably change. If our suggestion that PPa quenches singlet oxygen is indeed correct, it is likely that the pheophorbide-a-sensitized singlet oxygen data reported by R€oder’s group21,22 may be subject to the same phenomenon (i.e., their reported values of τΔ would likewise be shorter than what would be expected solely for the ambient intracellular environment). Of course, data interpretation in these suspension experiments is further compounded by the fact that not only are we in a domain where kT ∼ kΔ but also it appears that both kT and kΔ change during the course of the experiment. Among other things, the clear irradiation-dependent change in kT could reflect changes in (1) the extent to which PPa binds to proteins as the proteins are oxidized by singlet oxygen and/or (2) the viscosity of the environment around the sensitizer, as discussed earlier. Unraveling the complexities of these particular PPa-sensitized cell suspension data will certainly require a plethora of difficult experiments that exceed the scope of our present report.

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’ CONCLUSIONS It is well-established that the most accurate and desirable approach to quantify the lifetime of singlet oxygen, τΔ, in a given system is to record the O2(a1Δg) f O2(X3Σg) phosphorescence in a laser-based time-resolved experiment. Although performing these experiments can be difficult even with a relatively “simple” system (e.g., homogeneous liquid sample), the experiments are particularly challenging when the goal is to monitor τΔ in a live cell. Much of this latter challenge arises from the fact that singlet oxygen is cytotoxic. Thus, the system on which measurements are made is being perturbed during the measurement. In turn, this is reflected in the O2(a1Δg) f O2(X3Σg) phosphorescence data obtained. From this perspective, it is not surprising that the temporal profile of the O2(a1Δg) f O2(X3Σg) phosphorescence signal obtained in photosensitized experiments depends on the accumulated number of photons delivered to the cell-containing system. Perhaps the easiest way, although not the only way, to describe this phenomenon in terms of molecular events is to recognize that chemical reactions of singlet oxygen with proteins, for example, can readily change the local intracellular environment such that conditions that influence the production and removal of singlet oxygen can likewise change. However, despite these expected environmental changes, we have demonstrated that, once we account for the effects of certain irradiationdependent phenomena (e.g., sensitizer leaking out of the cell), the “inherent” intracellular lifetime of singlet oxygen does not appear to change dramatically as the cell progresses toward death. Given the heterogeneity of a cell, one might also expect that the kinetics of singlet oxygen production and decay/removal would depend on whether a hydrophilic or a hydrophobic sensitizer is used. In experiments performed to address this latter point, the data obtained are consistent with a model in which the comparatively long-lived intracellular singlet oxygen readily crosses between phase-separated domains. Nevertheless, key aspects of singlet oxygen’s behavior still depend on the local environment in which it is produced. Examples of the latter include the rate of singlet oxygen formation and the rate with which selected solutes remove/deactivate singlet oxygen in diffusion-dependent processes. We have also confirmed that hydrophilic sensitizers can readily diffuse out of the cell during a O2(a1Δg) f O2(X3Σg) phosphorescence measurement that, in turn, gives rise to the production of a singlet oxygen population in the extracellular medium. If the O2(a1Δg) f O2(X3Σg) phosphorescence measurement is performed on a suspension of cells, this timedependent ratio of intracellular:extracellular singlet oxygen can influence the magnitude of the measured value of τΔ differently than when the O2(a1Δg) f O2(X3Σg) measurement is performed in a microscope-based single cell experiment simply because a larger fraction of the extracellular volume is probed in the experiments performed on a cell suspension. In all cases, if the overall rate of singlet oxygen removal (i.e., kΔ or 1/τΔ) is roughly equivalent to the rate of singlet oxygen formation (i.e., kT), quantifying the data can be a challenging and errorfilled endeavor. In comparing data obtained in single cell experiments to data obtained from cell suspensions, we demonstrated that differences in cell lines and cell handling procedures can influence the O2(a1Δg) f O2(X3Σg) phosphorescence signal. At the limit, 459



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a phosphorescence signal that is readily detected in one experiment (e.g., TDFPPM in single HeLa cells) may not be observed in another experiment with the same sensitizer (TDFPPM in a suspension of HL-60 cells). These observations may reflect, in part, cell-dependent and cell-handling-dependent ways in which the sensitizer is localized in the cell (e.g., high local concentrations that may quench any singlet oxygen that is made and/or aggregation that may preclude the formation of singlet oxygen in the first place). In conclusion, the results reported herein begin to establish a useful framework with which to interpret and discuss measured values of τΔ from such a complicated system. In itself, we believe this is an important step in mechanistic studies designed to better elucidate the roles played by singlet oxygen in cell signaling and, ultimately, cell death.

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’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Author Contributions §

These authors contributed equally to this work.

’ ACKNOWLEDGMENT This work was supported by the Danish National Research Foundation. E.F.F.d.S. thanks the Portuguese Foundation for Science and Technology for a fellowship (grant BD/46658/ 2008) and M.K.K. is thankful for an EPSRC Career Acceleration Fellowship (UK). ’ REFERENCES (1) Foote, C. S. Acc. Chem. Res. 1968, 1, 104–110. (2) Schweitzer, C.; Schmidt, R. Chem. Rev. 2003, 103, 1685–1757. (3) Ogilby, P. R. Chem. Soc. Rev. 2010, 39, 3181–3209. (4) Paterson, M. J.; Christiansen, O.; Jensen, F.; Ogilby, P. R. Photochem. Photobiol. 2006, 82, 1136–1160. (5) Clennan, E. L.; Pace, A. Tetrahedron 2005, 61, 6665–6691. (6) Oleinick, N. L.; Morris, R. L.; Belichenko, I. Photochem. Photobiol. Sci. 2002, 1, 1–21. (7) Redmond, R. W.; Kochevar, I. E. Photochem. Photobiol. 2006, 82, 1178–1186. (8) Bonnett, R. Chemical Aspects of Photodynamic Therapy; Gordon and Breach Science Publishers: Amsterdam, 2000. (9) Phillips, D. Photochem. Photobiol. Sci. 2010, 9, 1589–1596. (10) Klotz, L.-O.; Kr€oncke, K.-D.; Sies, H. Photochem. Photobiol. Sci. 2003, 2, 88–94. (11) Moan, J.; Berg, K. Photochem. Photobiol. 1991, 53, 549–553. (12) Baker, A.; Kanofsky, J. R. Photochem. Photobiol. 1992, 55, 523–528. (13) Zebger, I.; Snyder, J. W.; Andersen, L. K.; Poulsen, L.; Gao, Z.; Lambert, J. D. C.; Kristiansen, U.; Ogilby, P. R. Photochem. Photobiol. 2004, 79, 319–322. (14) Skovsen, E.; Snyder, J. W.; Lambert, J. D. C.; Ogilby, P. R. J. Phys. Chem. B 2005, 109, 8570–8573. (15) Snyder, J. W.; Skovsen, E.; Lambert, J. D. C.; Poulsen, L.; Ogilby, P. R. Phys. Chem. Chem. Phys. 2006, 8, 4280–4293. (16) Kuimova, M. K.; Yahioglu, G.; Ogilby, P. R. J. Am. Chem. Soc. 2009, 131, 332–340. (17) Kuimova, M. K.; Botchway, S. W.; Parker, A. W.; Balaz, M.; Collins, H. A.; Anderson, H. L.; Suhling, K.; Ogilby, P. R. Nat. Chem. 2009, 1, 69–73. (18) Niedre, M.; Patterson, M. S.; Wilson, B. C. Photochem. Photobiol. 2002, 75, 382–391. 460



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