Due to its partial selectivity and minimally invasive character, photodynamic therapy (PDT) is often chosen for treating early stages of local, superficial cancers in hollow organs and other regions of the body accessible to light application.1,2 PDT is also considered as the treatment of choice for, among others, actinic keratosis (in dermatology), and wet age-related macular degeneration (in ophthalmology). To obtain the desired cytotoxic and healing effects, PDT relies on the presence in the target tissue of a photosensitizer (PS), molecular oxygen (), and administration of light at wavelengths absorbed by the PS. In this work, a time-tested method involving a biochemical precursor of the PS is used: topical or systemic administration of 5-aminolevulinic acid (ALA) leads, after a suitable drug-light interval, to biosynthesis and accumulation of photoactivable porphyrins (PAPs), including protoporphyrin IX (PpIX), the actual PS in the cells of the target tissue.3 Note that ALA-derivative-based PDT procedures are now approved for the treatment of premalignant and malignant conditions in dermatology.1
Despite encouraging results, some clinicians avoid using this therapy due to observed fluctuations in intra- and inter-patient therapeutic outcomes.4 These fluctuations, generally linked to uneven PS and/or tissular distribution, might in principle be counterbalanced by monitoring in real time the local PS and/or concentrations and then adjusting the light dosimetry.5,6 Measuring the local is also of interest for other types of treatments, such as radiotherapy, as it is well established that tissue oxygenation plays an important role in the outcome of such treatments.7 Note that previous studies8,9 have provided strong evidence that singlet oxygen () and other active oxygen species are the major active cytotoxic particles within the target tissues, which induce cell apoptosis. Therefore, accurate monitoring of , though quite difficult to do, might lead to a better prediction of the clinical outcome than other monitoring techniques. However, optimizing the treatment dosimetry obviously remains a challenge.10–13
Different approaches have been proposed to monitor PDT efficiently, with the goal of improving its clinical outcome.14,15 One of them is explicit dosimetry, in which the quantities of light, drugs, and oxygen are continuously and separately monitored during treatment. Alternatively, implicit dosimetry uses an indirect estimator of the biological damages, such as the degree of bleaching (photodegradation) of the photosensitizer, to predict the clinical outcome.5,16,17
This study uses the chicken embryo chorioallantoic membrane (CAM), an in vivo model well suited to characterize PDT’s vascular effects,18–20 to study the relationship between the evolution of tissular oxygen concentration during PDT and the resulting vascular damage. For this purpose, the quenching of PpIXs (from here on, the abbreviation PpIX encompasses both PpIX itself and the other photoactivable porphyrins) and delayed fluorescence (DF) by molecular oxygen21 are measured on the one hand by a fiber-based spectrometer22 to estimate , and on the other hand, on a simple, microphotographic-based evaluation index of the amount of vessel damage. The results of this study show that a robust, positive correlation exists between the reduction of tissular oxygen and PDT-created tissular damage.
Materials and Methods
PpIX Photodynamics and Diffusional Properties When Used as the PS
A summary of the photochemically induced reactions (type-II photochemical reaction pathway) can be represented by the following chemical scheme:23 to the long-lived triplet state , which has a lifetime ranging from a few microseconds up to a few milliseconds. This is why the tissular oxygen in the triplet ground state () has enough time to undergo a collision with the PS in the state, to form the highly reactive and cytotoxic singlet molecular oxygen . Once generated, undergoes deexcitation through several pathways, among which a low-probability radiative decay with emission of an infrared photon at 1270 nm. This luminescence is in fact characterized by a very weak signal, making its measurement not easily applicable in the clinical context (the luminescence emission of has a probability of and a characteristic lifetime of approx. 100 ns under in vivo conditions).10,24 These fast and weak signals from at 1270 nm are difficult to work with, especially under clinical conditions.
The idea was developed to measure the production of excited singlet oxygen via the “consumption” of the PpIX’s triplet state ( in the above chemical scheme), which is efficiently quenched by the molecular oxygen found in the tissue. Indeed, a fraction ranging from 55% to 80% of the PpIX population is quenched by , depending on the actual conditions.23 Therefore, measuring the photosensitizer’s luminescence lifetime offers an alternative to measuring the radiative decay of .25 Furthermore, in contrast to intensity measurement methods, lifetime measurement methods have the advantage of being inherently self-referential and more robust.26
PpIX’s phosphorescence signal, from the radiative path connecting to , may not be detected easily in vivo because the phosphorescence quantum yield is much too low and because this luminescence takes place at wavelengths too long to be easily detected.27 This difficulty can be overcome by measuring PpIX’s DF lifetime as a substitute for its phosphorescence lifetime.
Principles and Experimental Setup for Determination Through the Measurement of the PS’s DF Lifetime
Tissular can be determined by measuring the lifetime of the PS’s triplet state, thanks to the Stern-Volmer relationship:Ref. 28, including the setup’s temporal resolution and level of background optical noise. The schematics of the experimental setup are shown in Fig. 1.
The plano-convex lens (3) is mounted on a 5-degrees-of-freedom holder, allowing for easy focusing of the laser beam on a predefined point (5) (for specific details about the setup or parts thereof, see Ref. 28). The luminescence emitted from the irradiated sample is collected by the distal end of the optical fiber (6) and returned to the enclosure, where a large fraction (probably more than 80%) of this light falls on the parabolic metal mirror (4) and is reflected and focused by it on a gateable photomultiplier (PMT) (7). The reflected laser light and unwanted optical background noise are filtered out by a band-pass filter in the filter wheel (8; center wavelength: 645 nm, full width at half maximum: 75 nm). The PpIX DF signal is recorded with a digital storage oscilloscope (DSO) connected to a PC (9), where the data processing and analysis takes place (see the next section for more detail). The signal delay generator (10) is used to synchronize the gating of the PMT with the laser pulses and to trigger the DSO with the DF signals. The advantages of this optical design include (a) the absence of spectral distortions because only reflections take place in the scrutinized part of the optical path; (b) no autoluminescence is induced in the parabolic mirror which, due to its metallic nature, does not interact with the excitation pulse; (c) the solid angle of acceptance of the parabolic mirror () is larger than the entrance solid angle of the optical fiber (), which leads to a high light recovery fraction; (d) the mirror’s parabolic shape eliminates the need of an additional lens to focus the luminescence emitted by the fiber tip (or the quartz cuvette) on the detector.
The time-resolved detection of the weak DF signal requires gating to avoid saturation of the detector by the much stronger prompt fluorescence (PF) signal. The photomultiplier tube is switched on about 3 μs after the laser pulse, allowing the intensity of any PF to decrease to the noise level.
Data Acquisition and Analysis
The PMT’s analog signal is supplied to the DSO for averaging (about 30 transients are used), and the averaged values are transferred to the PC for analysis. The luminescence decay is fitted to a sum (usually one or two components are retained) of exponential functions:25,29–31 using a self-written algorithm implementation. The calculated mathematical fits were graphically checked by plotting the residuals of each fit and the autocorrelation of the residuals,32,33 as shown in previous research.28
Handling of the CAMs and ALA Administration
The procurement and processing of the eggs to prepare the CAM has been described in previous research (such as Ref. 20). In brief, fertilized chicken eggs are transferred (blunt end up) into an automatic turn-incubator, set at 37°C, 65% relative humidity (RH), and normal atmospheric conditions (i.e., 21%). Given the geographical location (400 m above sea level) of the experimental setup, this corresponds to a partial pressure of approximately 150 mmHg of . On embryo development day 3 (EDD 3), a portion of the eggshell is removed at the pointed end of the egg, creating a hole of approximately 3 mm in diameter, which is covered with Parafilm cling film, and the pointed end is always kept pointing up. This procedure causes the embryo and its enveloping CAM to retract locally from this artificial opening, exposing at its surface a flattened portion of the membrane, and providing an access to observe and treat the CAM’s capillary plexus and vasculature. The eggs are then returned to the incubator (blunt end down) and kept in a static position. At EDD 11, the eggs are removed from the incubator for topical ALA administration: a zone of the CAM characterized by homogeneous vascularization is visually selected and a 20 μl droplet of a 0.15 M () ALA aqueous solution in 0.9% NaCl, adjusted to pH 6 with NaOH, is deposited in the center of this zone. The eggs are then returned to the incubator and kept static until further use. As a rule, any manipulation of the CAM after ALA application is performed in total darkness or under subdued light.
PpIX Buildup Kinetics After ALA Topical Administration
Limited experimental data have been published about the localization and buildup kinetics of PpIX in the CAM following topical administration of ALA. Moreover, most of the results have been obtained for tumors growing in this membrane.34–36 In view of optimizing DF signal quality, a first experiment using 10 eggs is performed, whereby each egg is examined during times ranging between 0 to 6 h following the topical administration of ALA.
In Fig. 2(a), the resulting spatial distribution of PpIX in the CAM is shown, 4 h after ALA administration. The picture reveals a slightly more intense fluorescence (lighter areas) near the walls of the large vessels. As an illustrative example, the luminescence intensity in the administration zone is presented in Fig. 2(b). As shown in this figure and in agreement with Refs. 34–36 the maximum of PpIX accumulation is reached approximately 4 h after ALA administration.
PDT Irradiation and CAM Tissular Oxygen Depletion Measurements
The study design is guided by the following rationale (see below for description of experimental protocols): Protocol I is used to determine the existence of a robust relationship between PDT duration (up to 600 s) and , while using a minimum number of eggs: thus, each egg is submitted to a number of successive, cumulative irradiations and measurements.
Protocol II seeks to check the same relationship as protocol I using (a) a different batch of eggs, (b) intermediate durations for the incremental PDTs, and (c) a longer total treatment time (900 s), in order to confirm and improve the precision of the relation observed using protocol I, while still keeping the cumulated total measurement time under a certain maximum (6 times 3 s). In fact, in protocols I and II, measurements are performed once during each 3 s OFF period of the treatment laser (i.e., immediately after each incremental PDT irradiation period) and the difference with the initially determined baseline value is computed to be the tissular oxygen depletion assignable to the corresponding, cumulated, PDT treatment time.
Protocol III is used to prove that the parasitic effects of the multiple, intermediary, measurements used in protocols I and II are negligible (Fig. 3).
The normal, baseline value of the is determined for each egg before any PDT irradiation protocol is started. The treatment laser is then switched on and off and measurements performed according to one of the three experimental protocols described above. In protocol III, measurements are performed just once before and once after a continuous, predetermined irradiation period. The difference between those two values is computed to be the tissular oxygen depletion assignable to the corresponding, uninterrupted, PDT treatment time.
Details of the PDT irradiation and tissular oxygen measurement procedures are as follows: At EDD 11, 4 h after ALA administration, the egg is taken out of the incubator and placed under the combined PDT-irradiation/DF-measurement setup. The drug-light interval of 4 h allows time for the cells to produce sufficient tissular PpIX. In order to minimize re-oxygenation of the tissue during PDT by atmospheric oxygen, a round, 20-mm, 0.15-mm-thick microscope cover-glass (3) is gently floated on the CAM’s surface (1) after depositing two drops (2) of human physiologic serum ( of 0.9% NaCl; see Fig. 4). This liquid layer, acting as a cushion, is essential to avoid any adherence between the cover-glass and the CAM and possible subsequent injury to the CAM surface. Therefore, during PDT or tissular oxygen measurement, oxygen may diffuse only from the CAM vasculature and from the tissues adjacent to the treated zone, not from the shielded surface of the treated zone.
A PDT treatment zone, featuring blood vessels and capillaries with diameters ranging from 5 to 300 μm, is visually chosen, approximately in the middle of the zone under the cover-glass. The tip of the optical fiber (5) is aimed toward the center of this zone and held at a distance of above the cover-glass. It is kept in place by a metal fiber holder (6). The PDT light is provided by a CW solid-state diode laser (2) (Oxxius, 405 nm, 1.3 mW total power at the distal end of the fiber) and administered through a light distributor providing a homogenous spot (4) (MedLight Frontal light distributor, , core diameter 600 μm). This treatment fiber was chosen and its distance to the CAM adjusted so as to deliver a homogeneous PDT treatment spot (diameter 8 mm) on the selected zone of the CAM vasculature. Small irradiances () are preferred to avoid significant photothermal effects. The period during which the eggs are kept out of the incubator is limited to about 15 min. To keep the DF measurement durations small with respect to PDT exposure times, only 30 consecutive laser sweeps on the same area are used and averaged for the oxygen measurements. Thus, the total acquisition time for one measurement is 3 s, compared to a minimum PDT exposure time of 75 s, and the corresponding total light dose delivered by the probing laser is limited to , as compared to a minimum PDT light dose of , both at 405 nm. A light dose of is found to have a barely detectable PDT vascular effect. Therefore, the minimal PDT light dose is set to approximately double this value. At the end of the PDT treatment, the cover-glass is carefully removed from the treated zone. Only very small iatrogenic damages are identified after these mechanical operations. The eggs then are returned to the incubator while waiting for the vascular damage assessment.
Vascular Damage Assessment
The treated eggs are taken out of the incubator 16 to 18 h after termination of the PDT irradiation protocol. This standard time period has been in use in our laboratory for several years, partly for practical reasons, as the measurements are performed overnight.37 Although it was not methodically optimized, it corresponds to a state of the CAM where PDT damage has had the time to develop further while the counteracting processes of vessel repair and neo-angiogenesis have not yet interfered significantly with the PDT results.37
An intravenous (IV) injection of 20 μl of a 150 μM solution of fluorescein isothiocyanate-dextran (FITC-dextran, 150 kDa, in NaCl 0.9%, Sigma Aldrich) is administered into one of the large CAM veins, while 0.1 ml of India ink (Parker-Quink) is injected under the CAM into the extraembryonic cavity to increase contrast and screen out the continuously changing background fluorescence.38 Vascular damage due to PDT is then examined and recorded photographically with an epifluorescence microscope (Nikon E600FN, excitation wavelength between 450 and 490 nm, emission wavelength ). The damage extent is classified on the basis of blood vessel closure: The diameter of the largest blood vessels closed by the PDT treatment is used as an arbitrary index of vascular damage. Figure 5 shows examples of actual micrographs.
A drug-light interval of 4 h was chosen for the PDT procedure and used throughout this study [see Fig. 3(a) and 3(b) for the experimental determination of this time interval]. The zone where ALA is topically administered then is selected to perform DF measurements, as this results in an adequate signal-to-noise ratio. The treatment laser delivers at 405 nm.
In Vivo Measurement of Tissular as a Function of PDT Irradiation Time
In a first series of experiments, DF-based measurements were performed on a number of egg batches (each one comprising 12 to 20 eggs), subjected to increasing cumulated PDT irradiation times. Three slightly different illumination protocols were used (see Fig. 4 and PMGQ3PMGQ and Sec. 2 of this paper).
The left vertical scale of Fig. 6 shows the measured reciprocal lifetime of PpIX’s DF plotted against the total illumination time (horizontal axis). The right vertical scale of the figure gives the corresponding (computed) tissular , using the Stern-Volmer equation, in which was derived from previous experiments.22
Clearly, the tissular oxygen concentration decreases during the first 500 s of cumulated PDT time, for a cumulated light dose of . At longer treatment times, it reaches an asymptotic residual value, close to zero. The relatively large error bars () probably relate to inhomogeneities in the distribution of the PS and/or local differences in the aptitude of CAM tissues to renew the consumed by the PDT process, depending on their level of metabolism and degree of vascularization. It can be seen that differences ascribable to illumination protocol, if any, are negligible in this experimental context.
Tissular Oxygen Depletion as an Index of PDT Efficiency
A second experiment was then performed on five lots of 20 eggs, submitted to different PDT optical doses, using illumination Protocol III (see above for details of egg/CAM handling).
The baseline value of tissular was first determined for each untreated egg before starting the chosen PDT irradiation protocol, using the described PpIX DF reciprocal lifetime determination. The eggs were then submitted to PDT illumination under a constant irradiance of at 405 nm for different durations (150, 300, 450, 600, 900 s), corresponding to optical light doses of 0.38, 0.75, 1.14, 1.50, and , respectively.
Immediately after treatment, each egg’s tissular concentration was again measured with the same method. The difference between the initial and final DF reciprocal lifetimes was used as a measure of tissular depletion ascribable to the PDT treatment. Figure 7 illustrates this increasing oxygen depletion as a function of PDT illumination time.
The eggs were then returned to the incubator for 16 to 18 h. Finally, the extent of vascular damage caused by the PDT was determined using a simple arbitrary damage index described above.
In Fig. 8(a), the vascular damage index is plotted as a function of the PDT treatment duration. As expected, the data shows a clear, positive correlation between those two parameters. Although, as already noted, the tissular does not decrease significantly anymore for PDT illumination periods longer than about 500 s (Fig. 6), the observed tissular damage does continue to increase with increasing light doses, at least until the irradiation time of 900 s. This suggests that the PDT mechanism is still operating at very low oxygen concentrations.
Figure 8(b) shows the vascular damage index as a function of the reciprocal lifetime difference, a measure of tissular oxygen reduction during the treatment. Here again, a strong, positive correlation between the two parameters is clearly visible.
This paper reports a first study aiming at measuring, in vivo and in real time, the level of molecular oxygen contained in tissue, using the DF of a PS, in order to optimize PDT. The results demonstrate a clear correlation between the PDT-induced vascular damage and the reduction in tissular oxygen concentration, as assessed by the change in DF lifetime. This agrees with previous works8,9,39,40 regarding the role played by oxygen in PDT-induced tissue damage. Thus, adjusting the PDT light dose based on a real-time estimate of tissular monitored through the PS’s DF appears feasible. This approach then might enable controlling the efficacy of PDT, and reducing intra- and inter-patient fluctuations in this treatment’s clinical outcome.
The method presented here does not require the use of exogenous oxygen sensors since it is based on detecting the DF of the PS itself. This feature is important because, besides ALA, several other precursors of PpIX are now approved by the medical authorities of a number of countries. This is an implicit light dosimetry method because the value (or its reduction, both being related to the production of singlet oxygen) is measured close to the PS (about 1 μm). Such a method is likely to be much more directly related to PDT’s tissular effects than approaches relying on explicit light dosimetry (light dose, PS global concentration, etc.).5,16,41–43 In addition, most of these other methods for quantifying the outcome of PDT suffer from the fact that the is not measured at a subcellular or tissular scale.
In similar research, Mik et al. report in vitro and in vivo experiments using PpIX’s DF lifetime signals, to assess the amount of oxygen in tissues.27,44 However, as compared to other oxygen-sensing techniques, DF measurements have the advantage that they record the oxygen concentration in close proximity to the photosensitizer. In fact, given the diffusivity of molecular oxygen () in tissue (),45,46 and using PpIX’s triplet state maximum lifetime in vivo (), the distance reachable by an molecule to interact with excited PpIX molecules is calculated to be on the order of (.47 If the same computation is done for singlet oxygen (), characterized by much shorter lifetime of approximately 250 ns, the reachable radius is less than 40 nm,24 indicating that the photo-damages are induced at the close vicinity of the photosensitizing molecule. On the other hand, the critical oxygen diffusion distance, usually defined as the distance between a hypoxic tissue (i.e., a tissue in a situation of critically low oxygen level) and the nearest vessel was measured by Grote et al. to be on the order of 100 μm in a tumoral tissue.45 It is therefore clear that this DF-based oxygen sensing method is able to spatially discriminate between hypoxic zones and normally oxygenated tissue, providing that the excitation and/or emission light are administered and/or detected with the appropriate resolution. It is also worth noting that during the several hours of incubation, ALA is absorbed by the vasculature, where it circulates, and then diffuses out of the vasculature into almost all egg tissues. This is why PpIX are formed and accumulated apparently with different concentrations in most of the tissues of the chicken embryo.
Since in this study both the light used to excite the DF and the light used for PDT have the same wavelength, the DF excitation might itself induce some level of reduction, PpIX photobleaching, and vascular damage, thus leading to artifacts. However, the DF excitation light dose used here is much smaller than the one used to induce the smallest detectable PDT effect, and in a previous study,22 it was demonstrated that, in the absence of PDT irradiation, the DF excitation light does not induce any detectable vascular effect in similar conditions. Finally, the DF lifetimes measured with the three different illumination protocols confirm that the contribution to vessel closure of this excitation light (which differs according to the protocol used) is negligible (see Fig. 6).
A drug-light interval of 4 h was used for the present study. It should be noted that other incubation periods may lead to other values of the DF lifetimes, even if all other conditions are identical, since it is well known that the PpIX buildup, which takes place in specific tissue/cellular compartments, is followed by a diffusion of the PS to other compartments that present other microenvironments.48 The changed microenvironments then may lead to changed local and hence change in DF lifetime, as well as changed vascular occlusion efficiency. As a consequence, it could be that the results reported in Figs. 6 and 8 may differ if longer or shorter drug-light intervals are used.
The relatively large error bars affecting the DF lifetimes presented in Figs. 6Fig. 7–8 may well be due to local differences in tissue oxygen renewal during PDT. These local differences could, for example, be due to spatial and/or temporal differences in the metabolic activity, blood perfusion, inhomogeneous distribution of the PS, etc. Therefore, one approach to reducing these fluctuations would consist of probing a larger area (typically 1–2 cm in diameter) so that the measurement is averaged over a larger surface. These observations furthermore suggest that it might be of interest to image the PS’s DF lifetime at “high” spatial resolution in the hope of getting information regarding the changes of tissue oxygenation at a tissular and even a cellular level. Recording such images at different times after the beginning of PDT and at different drug-light intervals would certainly help to identify treatment strategies to target specific tissue structures (vessels versus stroma, for instance).
Finally, it should be noted that due to the instrumentation and the shot noise, the fluctuations affecting the DF lifetime measurements are negligible compared to the other factors mentioned above. Indeed, repeated measurements performed in identical conditions, at the same location and on the same sample, confirm this. Nevertheless, if necessary, the shot noise can be improved further by reducing the time during which the detector is blinded to reject the prompt fluorescence. The resulting increase of DF radiant energy detected will decrease the shot noise, but it also will increase the risk of detecting (a) the emission of other endogenous luminophores presenting “long” (i.e., between 100 ns and 1 μs) luminescence lifetimes; and (b) luminescence emitted by the instrumentation (laser spontaneous emission, luminescence from the optics in the setup). This is why, in this study, a time delay of 3 μs was selected to “blind” the detector, which corresponds to an optimal tradeoff between this delay and the observed signal-to-noise ratio.
The error bars affecting the measurements of the diameter of the largest occluded blood vessels, which are shown in Fig. 8 are likely to be due to various factors, including the following:
(a) The selection of the area treated by PDT. Indeed, somewhat different results may be obtained if the relative density of large and small vessels, or of veins versus arteries, differs from place to place. Although the relative importance of this source of fluctuations is difficult to assess quantitatively, it probably plays a minor role, as described by Lange et al.20 who used a similar metric to quantify the vascular effects induced by PDT in the CAM model.
(c) Intrinsic errors while measuring the vessels diameter. The fluctuations due to this last element could be reduced using quantitative image analysis programs to characterize the vascular network, as proposed by Ref. 51. One may conclude, considering the abovementioned factors, which are possibly responsible for the rather large fluctuations affecting measurements of the DF lifetime and blood vessel closure efficacy, that probing of larger well-selected areas, as indicated above, most likely will help to reduce these fluctuations.
The results presented in Fig. 8(b) indicate that the vascular occlusion effects of PDT and the differences in reciprocal DF lifetime, which is a metric for the tissular oxygen reduction, are correlated. For illumination periods longer than 500 s (), it is noteworthy that the vascular damage continues to increase, despite the fact that the measured residual does not decrease significantly anymore after 500 s (it is then already very close to 0%, as shown in Fig. 6). This indicates that some PDT-related mechanism continues to be effective at very low . One possible explanation is that all the oxygen provided by the bloodstream is consumed immediately during the PDT illumination. This hypothesis is realistic since the blood vessels affected by PDT, depending on their size among others, will close to a large extent, after the end of the illumination,52 at the applied conditions. Consequently, in such conditions, the PDT effect would be “oxygen supply dependent,” as reported elsewhere.53–55 Other phototoxic mechanisms that do not involve molecular oxygen, as suggested by Plaetzer et al.,56 could also cause a PDT effect in the absence of a significant . It is possible that experiments similar to those presented in this paper, but performed at different fluence rates, may help to reveal the nature of these effects.
Finally, note that the approach presented here is of interest, not only for PpIX-based PDT, but also when other PSs with a detectable DF or a difficult-to-detect phosphorescence is used. One may even suggest that the technique presented in this paper be used to monitor the during the treatment of age-related macular degeneration (AMD) with Visudyne®. Indeed, application of the DF measurements to the retina may be helped by the unique optical properties of the eye, and using such measurements may help compensate, to some extent, for the inter-patient fluctuations observed during AMD-PDT.57
In conclusion, the present work has demonstrated in vivo that the DF of a photosensitizer can be used to determine the in tissues and help to predict PDT’s tissular effects. Applications of this approach to other systems are numerous, and this technology may help improve understanding of the mechanisms involved in the oxygenation and photosensitization of various materials, including biological tissues.
This work was supported by the Swiss National Science Foundation (Grant No. 205320-130518) and funded in part by the J. Jacobi Trust. We gratefully acknowledge editorial support by Matthieu Zellweger.