International Journal of

Molecular Sciences
ArticleCell-Level Analysis Visualizing Photodynamic Therapy withPorphylipoprotein and Talaporphyrin Sodium
Mayuka Kamiyanagi 1,† , Atsushi Taninaka 1,2,† , Shunta Ugajin 1 , Yu Nagoshi 1 , Hiromi Kurokawa 3 ,Takahiro Ochiai 2 , Yusuke Arashida 1 , Osamu Takeuchi 1 , Hirofumi Matsui 3 and Hidemi Shigekawa 1, *
1
2
3

*
†

Citation: Kamiyanagi, M.; Taninaka,
A.; Ugajin, S.; Nagoshi, Y.; Kurokawa,
H.; Ochiai, T.; Arashida, Y.; Takeuchi,O.; Matsui, H.; Shigekawa, H.
Cell-Level Analysis Visualizing
Photodynamic Therapy with

Faculty of Pure and Applied Sciences, University of Tsukuba, 1-1-1 Tennodai, Tsukuba 305-8573, Japan
TAKANO Co., Ltd., Miyada-mura, Kamiina-gun, Nagano 399-4301, Japan
Fuculty of Medicine, University of Tsukuba, 1-1-1 Tennodai, Tsukuba 305-8575, Japan
Correspondence: hidemi@ims.tsukuba.ac.jp
These authors contributed equally to this work.

Abstract: We revealed the difference in the mechanism of photodynamic therapy (PDT) between
two photosensitizers: porphylipoprotein (PLP), which has recently attracted attention for its potential
to be highly effective in treating cancer, and talaporphyrin sodium (NPe6). (1) NPe6 accumulates inlysosomes, whereas PLP is incorporated into phagosomes formed by PLP injection. (2) PDT causes
NPe6 to generate reactive oxygen species, thereby producing actin filaments and stress fibers. In
the case of PLP, however, reactive oxygen species generated by PDT remain in the phagosomes
until the phagosomal membrane is destroyed, which delays the initiation of RhoA activation and
RhoA*/ROCK generation. (4) After the disruption of the phagosomal membrane, however, theoutflow of various reactive oxygen species accelerates the production of actin filaments and stressfibers, and blebbing occurs earlier than in the case of NPe6. (5) PLP increases the elastic modulus
of cells without RhoA activity in the early stage. This is because phagosomes are involved inpolymerizing actin filaments and pseudopodia formation. Considering the high selectivity anduptake of PLP into cancer cells, a larger effect with PDT can be expected by skillfully combining the
newly discovered characteristics, such as the appearance of a strong effect at an early stage.

Porphylipoprotein andTalaporphyrin Sodium. Int. J. Mol.
Sci. 2022, 23, 13140. https://doi.org/
10.3390/ijms232113140

Keywords: photodynamic therapy (PDT); talaporphyrin sodium (NPe6); porphylipoprotein (PLP);
phagosome; atomic force microscopy; elasticity

Academic Editors: João Paulo
C. Tomé, Augusto C. Tomé andLeandro M. O. Lourenço

1. Introduction

Received: 30 September 2022

Photodynamic therapy (PDT) treats diseases by injecting photosensitizers into thebody and irradiating them with light to generate reactive oxygen species (ROS) [1–4]. It is
noninvasive with few adverse effects and exhibits a high therapeutic effect on lesion areasthat are difficult to treat with surgical techniques. It is used not only for cancer therapy
but also for conditions such as age-related macular degeneration and skin diseases [5–7].
Recently, as a method to efficiently inactivate pathogens without inducing resistant bacteria, antimicrobial photodynamic therapy has been developed and applied to treatment
for periodontal disease [8–11]. Porphyrin has been mainly used as a photosensitizer. In
PDT, porphyrin ring is photoexcited to generate singlet oxygen, which is converted into
OH radicals and other ROS, such as H2 O2 , which destroy the cytoplasm [1,3,12,13]. The
performance of the photosensitizer determines the therapeutic effect of PDT, for example,
the degree of selective uptake into the affected area, permeability, and persistence. Photosensitizers accumulate in mitochondria, endoplasmic reticulum, and lysosomes at thecellular level. That changes the persistence of photosensitizers and the types of organelles
destroyed by ROS. For example, there is a difference in the effect of PDT with Porfimer
sodium and Talaporfin sodium (NPe6). Therefore, various photosensitizers suitable fortreatment have been developed so far [13].

Accepted: 26 October 2022
Published: 28 October 2022
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Int. J. Mol. Sci. 2022, 23, 13140. https://doi.org/10.3390/ijms232113140

https://www.mdpi.com/journal/ijms

Int. J. Mol. Sci. 2022, 23, 13140

2 of 12On the other hand, advances in technology have made drug delivery practical. Research is being actively pursued for further development. Although attached polyethylene
glycol polymer chains (PEG) to molecules or nanoparticles have been clinically approved,
there remain several concerns including immunogenicity, anti-PEG immune response,
biocompatibility and toxicity issues following chronic administration. Previous attempts tosolve the issues typically involve developing PEG linker-removable approaches [14,15].
Porphylipoprotein (PLP) nanoplatform integrates a variety of imaging and therapeutic functionalities including positron emission tomography (PET) imaging, activatablenear-infrared (NIR) fluorescence and photodynamic therapy (PDT) [14,15]. The tumorselective activation of NIR fluorescence facilitated the delineation of tumor, the selectively
activated photodynamic reactivity enabled potent PDT for tumor ablation and surgery bed
cleaning [14,15].
Heme Carrier Protein 1 which is a transporter involved in low-affinity heme-Fe or
folate (HCP1) is expressed more in cancer cells than in normal cells [16–18]. Therefore,
molecules with a porphyrin ring passing through HCP1 have been considered for use as
photosensitizers [18]. In fact, in vivo and in vitro, PLP was found to be selectively taken up
by cancer cells in large amounts and have a large therapeutic effect [18]. A high effect has
also been confirmed in NOZ (Human gallbladder carcinoma cell line) cells [18]. However,
the mechanism for PDT has not been clarified yet. Therefore, in this study, we investigatedthe properties of PLP in detail by comparing them with those of NPe6.
In a previous paper, we focused on PDT using talaporphyrin sodium (NPe6) andinvestigated how it behaves in cancer cells by combining conventional techniques such as
luminescence and Western blotting with atomic force microscopy (AFM) [19]. We examined
the mechanism of actin filament formation by analyzing the change in elasticity of living
cells. As a result, we succeeded in clarifying the relationship between stress fiber generation
and blood flow disturbance, which was difficult to analyze by optical methods [19]. We
used similar methods in this study. As a result, it became clear that the high function of
PLP is caused by a mechanism involving phagosomes known in the field of autophagy.
This function has the potential to enhance the effectiveness of PDT and is expected to play
an important role in the future.
2. Results and Discussions
2.1. Subcellular Distribution of Injected NPe6 and PLP
The cells used in this study were rat gastric mucosa-derived cancer-like mutant RGK1
cells [20]. NPe6 or PLP was added to RGK1 cells and confocal microscope observations werecarried out 24 h later (see Materials and Methods for more details). Figure 1 shows results:(a,g) bright field (BF) images, (b,h) endoplasmic reticulum fluorescence (ER) images, (c,i)
mitochondria fluorescence images, (d,j) magnified views of (e,k), and (f,l) merged imagesof the distributions of mitochondria and photosensitizer images. We first examined theintracellular distributions of NPe6 and PLP.
Figure 1e,k show that NPe6 and PLP accumulate in granular substances with a diameter of about 200 nm − 1 µm. Lysosomes and phagosomes are vesicles with a diameter of
200 nm − 1 µm among intracellular organelles. As can be seen from the merged figures, inthe case of NPe6, NPe6 are distributed even in areas where no mitochondria are distributed,
as indicated by the white arrows in Figure 1d. This is in good agreement with the factthat NPe6 is known to accumulate in lysosomes [21]. In addition, NPe6 shows uniform
fluorescence inside the granular material in the enlarged view (Figure 1e). Therefore, NPe6
accumulated in the lysosomes, and PLP is considered to accumulate in phagosomes. Since
water-soluble porphyrin derivatives accumulate in lysosomes [21,22], NPe6 is considered
to be dissolved in the fluid inside lysosomes.

Int. J. Mol. Sci. 2022, 23, 13140

3 of 12Figure 1. Confocal microscopy observation of RGK1 cells after NPe6 or PLP was added and allowed to stand for 24 h. (a,g) Bright field (BF) images, (b,h) endoplasmic reticulum (ER) fluorescence images, (c,i) mitochondria fluorescence images, (d,j) NPe6 and PLP fluorescence images and(e,k) their magnified views, and (f,l) merged images of (c,d), and (i,j), respectively. ER-Tracker™
Green (Invitrogen™) and MitoTracker™ Orange CMTMRos (Invitrogen™) were used for fluorescencestaining of ER and mitochondria, respectively. After adding these fluorescent probes together in asample with standing of 24 h after doping NPe6 or PLP, and leaving the mixture for 30 min, measurements were carried out while changing the excitation light and filters. Then, PDT was performed toobserve the distributions of NPe6 and PLP. As shown in Figure 1e,k, NPe6 and PLP accumulated in
granular substances with a diameter of about 200 nm–1 µm.

On the other hand, in the case of PLP, the granular substances aggregate near mitochondria, and many of them have a ring shape, as expected and indicated by the redarrows in Figure 1k. Phagosomes are formed at the ER and the mitochondria contact
sites and have a circular shape [23]. Therefore, PLP is considered to accumulate in thecircular membrane of phagosomes. Since phagosomes and lysosomes (indicated by theyellow arrows in Figure 1k) are known to fuse to form phagolysosomes [24–31], the brightspots with diameter of several hundred nanometers in Figure 1k may be lysosomes. The
relationship between PLP, phagosomes, and lysosomes will be examined later.
2.2. Time-Lapse Observation by Phase-Contrast Microscopy
In general, when the light intensity of PDT was high (300 mW/cm2 ), cells swelled andthen ruptured (necrosis), and when the light intensity was low (100 mW/cm2 ), blebbing
occurred in about 10 min [19]. Furthermore, when PDT was performed at a lower lightintensity as 34 mW/cm2 , the blebbing was delayed, and the cell membrane hardened,causing obstruction of the blood flow and destruction of cancer cells [19].
In a previous study using NPe6, irradiation with light of 34 mW/cm2 at a wavelengthof 655 nm for 1 min generated ROS, immediately after which actin filaments were generated,
and, myosin light chain formation subsequently occurred after 5 min, leading to theformation of stress fibers [19,32]. Therefore, to compare the mechanism of stress fiber
formation, RGK1 was first irradiated with light of 34 mW/cm2 at a wavelength of 655 nm
for 1 min under the same sample conditions, that is, RGK1 that had been allowed to standfor 24 h after the addition of NPe6 or PLP, and time-lapse observation was performed witha phase-contrast fluorescence microscope.
Figure 2a,b show phase-contrast observation images of NPe6 and PLP, respectively.
Most of the cells to which NPe6 was added did not start blebbing (undergo a “deathdance”) [33] even after 15 min, and some cells started to bleb from about 60 min. In contrast,
cells to which PLP was added began to contract 5 min after 1 min light irradiation, andblebbing occurred 10–15 min after irradiation. In addition, the time required for almost all

Int. J. Mol. Sci. 2022, 23, 13140

4 of 12

cells in the observed area to bleb was 300 min for NPe6 and 60 min for PLP. These results
suggest that PLP induces excessive myosin light chain activation [33], which triggers
blebbing, in a shorter time than NPe6.

Figure 2. Time-lapse phase-contrast images of RGK1, which had been left standing in an incubatorfor 24 h after adding NPe6 (a) or PLP (b), then irradiated with light of 34 mW/cm2 at a wavelengthof 655 nm for 1 min.

In Figure 2, the amount of light is the same the both NPe6 and PLP-added samples.
Thus, when the amount of light is increased, PLP is more effective in directly destroying
cancer cells through necrosis or necroptosis. The fact that PLP is taken up by cancer cells inlarge amounts with high selectivity also enhances the therapeutic effect of this method.
2.3. Fluorescence Observation of Actin Filament Formation Process
When PDT is performed with a low light intensity, it has a therapeutic effect not by theimmediate destruction of cancer cells but by the disturbance of blood flow [19,32,34,35]. The
ROS generated by PDT trigger the formation of stress fibers through the activation of RhoA
(Ras homolog family member A), producing actin filaments and myosin light chains [1,16,30].
In this case, the process of actin filament formation can be observed by fluorescence bystaining actin filaments with SPY-555 (Cytoskeleton Inc., Denver, CO, USA) [36,37]. This afluorescent probe enables the direct imaging of actin filaments in living cells [36,37].
Figure 3a,b, respectively show phase-contrast observation images and fluorescenceobservation images of actin filaments stained with SPY555 after PDT of RGK1 with NPe6
and PLP. For NPe6, as shown in Figure 2, the phase-contrast images showed almost no
structural changes in most of the cells even 14 min after exposure to light for 1 min.
The fluorescence intensity of actin filaments slightly increased immediately after lightirradiation and gradually increased until 14 min after light irradiation. For PLP, it can beseen from the phase-contrast observation images that the cells began to contract 4 min after
1 min of light irradiation, and many cells had contracted by 14 min after light irradiation.
The fluorescence intensity of actin filaments hardly increased immediately after lightirradiation but started to increase after 4 min.
Figure 4 shows the results of quantifying the fluorescence intensity from the images in
Figure 3. NPe6 showed an increase in fluorescence intensity after 1 min of irradiation. On
the other hand, in PLP, the fluorescence intensity hardly changed after 1 min of irradiation,
but the fluorescence intensity had increased 4 min after irradiation and reached the samelevel as that of NPe6. After 9 min, NPe6 continued to show a slight increase, while PLP
showed a sharp increase in fluorescence intensity. This indicates that the production ofactin filaments during 1 min of light irradiation is lower in PLP than in NPe6. The rapid
increase in fluorescence intensity at 9 min for PLP is considered to be due to the influx of
SPY-555 into cells due to damage to the cell membrane, as blebbing begins at 10–15 minaccording to Figure 2. This is because SPY-555 emits light when it enters the cells andinteracted with actin filaments. In the case of NPe6, as shown in the previous paper, an
increase similar to that of PLP was observed from about 30 min [19]. This is due to the time
lag in which the cell membrane is damaged.

Int. J. Mol. Sci. 2022, 23, 13140

5 of 12Figure 3. Results of time-lapse observation with a fluorescence microscopy. NPe6 (a) and PLP(b)-doped RGK1 samples were allowed to stand for 24 h in an incubator and then irradiated with alight of 34 mW/cm2 at a wavelength of 655 nm for 1 min.

Figure 4. Results of quantifying the fluorescence intensity from the fluorescence observation images
in Figure 3.

2.4. Analysis of RhoA Activation by Western Blotting
RhoA activation can be evaluated by Western blotting to assess RhoA*/ROCK binding
and subsequent actin filament formation, where RhoA* is activated RhoA. As revealed in
the previous NPe6 study, there is a threshold amount of ROS that is involved in the increasein actin filament production [19]. That is, there is an upper limit to the concentration of
RhoA activated and binds to ROCK. In the fluorescence observation, no increasing inluminescence of PLP was observed immediately 1 min of light irradiation. Therefore, todetermine whether RhoA activation and RhoA*/ROCK binding occurred in PLP-PDT as in
NPe6, changes in the amount of RhoA were quantified by Western blotting.

Int. J. Mol. Sci. 2022, 23, 13140

6 of 12Figure 5 shows the results. In the PDT with PLP, there was no change in the concentrationof RhoA after 1 min of irradiation, and the amount of RhoA decreased when the cells wereincubated for 4 min at 37 ◦ C in 5% CO2 after 1 min irradiation. The decrease in RhoA after
5 min and 15 min irradiation was almost the same as after 1 min of irradiation and 4 min ofstanding. Therefore, it is considered that the RhoA*/ROCK binding was saturated after 1 minof irradiation. In a previous experiment using NPe6, the concentration of RhoA decreased
after 1 min of irradiation and RhoA*/ROCK was saturated after 5 min of irradiation [19].
The difference between PDT using NPe6 and PLP is the difference in the decreasein RhoA concentration (activation of RhoA to RhoA*) due to 1 min of irradiation, and it
is considered that NPe6 forms RhoA*/ROCK more rapidly. However, the saturation of
RhoA*/ROCK and the initiation of blebbing due to the overactivation of myosin lightchains shown in Figure 2 are faster for PLP. This indicates that the timing at which ROS isgenerated and RhoA is activated upon irradiation with light is different for NPe6 and PLP.
That is, NPe6 gradually generates ROS that promotes RhoA activation immediately after
light irradiation and forms RhoA*/ROCK more slowly than PLP. However, although PLP
generates singlet oxygen (1 O2 ) immediately after light irradiation, it does not generate ROS
that promotes RhoA activation.

Figure 5. Results of estimating changes in RhoA concentration after PDT with PLP by Western
blotting. (a) Western blot photograph. (b) Graph quantifying the results of (a). The irradiation was
carried out at 0.0531 mW/cm2 .

2.5. Elastic Modulus Measurement by AFM
To examine the above results using changes in cell stiffness, we measured the local
elastic modulus of cells using AFM. Since the state of actin filament formation is related
to the elastic modulus of the cell, it is possible to determine the state of actin filament
formation by measuring the elastic modulus of the cell [19]. Figure 6 shows phase-contrast
images, topographic images, and elastic modulus maps of RGK1 with added NPe6 and PLP
after 1 min of light irradiation and 1 min irradiation and 4 min standing in an incubator at
37 ◦ C with 5% CO2 concentration.

Int. J. Mol. Sci. 2022, 23, 13140

7 of 12Figure 6. Phase-contrast images, topographic images, and elastic modulus maps of RGK1 with added
NPe6 (a–d) and PLP (e–h) after 1 min of light irradiation and 4 min of standing in an incubator
at 37 ◦ C with 5% CO2 concentration. Force curves were measured at 64 × 64 points in an area of
100 µm × 100 µm. The topo images were obtained by mapping the height when a force of 200 pN
was measured. The elastic modulus was evaluated using Hertz’s formula [38].

For individual cells, we estimated the average local elastic modulus around the cell
nucleus and the edge of the cell (referred to as the cell body). We also estimated the average
elastic modulus over the observed cells, and the results are shown in Figure 7.

Figure 7. Estimated average local elastic moduli around the cell nucleus and the edge of the cell(referred to cell body) over all observed cells. N = 44 (1 min) and 45 (1 min + 4 min) for NPe6 (a), and34 (1 min), and 26 (1 min + 4 min) for PLP (b), respectively.

Int. J. Mol. Sci. 2022, 23, 13140

8 of 12For NPe6, the elastic modulus showed little change after 1 min of light irradiation.
After 1 min of irradiation and 4 min of standing in an incubator, the elastic modulus
increased at the cell body and around the cell nucleus. This result corresponds well to theformation of stress fibers, as observed in a previous study [19]. In contrast, for PLP, theelastic modulus near the cell nucleus did not change after 1 min of light irradiation, but theelastic modulus at the cell body increased slightly. When the elastic modulus changes as aresult of a change in osmotic pressure due to cell contraction, the elastic modulus of thecell increases uniformly and independently of the cell nucleus periphery and the cell body.
Therefore, it is considered that this change in elastic modulus reflects some change in thecell structure. After 1 min of irradiation and 4 min of standing in an incubator, the elastic
modulus of the edge of the cell increased remarkably, approaching the state of NPe6. This
corresponds well to the results for actin filament generation shown in Figure 4. However,from the phase-contrast images in Figure 2, the cells contracted for PLP under both 1 minof light irradiation and 1 min of irradiation followed by 4 min of standing in a incubator,
whereas the contraction of cells was negligible for NPe6. These results suggest the existenceof a new mechanism for the case of PLP-PDT, which we will discuss in detail later.
For NPe6, the average elastic moduli around the cell nucleus and at the edge of thecell were estimated to be 4.97 ± 0.63 kPa and 13.8 ± 2.4 kPa before light irradiation and4.08 ± 0.48 kPa and 16.5 ± 2.3 kPa after 1 min irradiation, respectively. The average elastic
modulus around the cell nucleus did not change markedly, and the average elastic modulus
at the cell body increased by 19% (1.19-fold increase). The average elastic moduli around thecell nucleus and at the cell body were estimated to be 4.41 ± 0.54 kPa and 9.86 ± 2.03 kPa
before light irradiation, which increased to 6.17 ± 0.68 kPa and 18.2 ± 3.6 kPa after 1 minof light irradiation and 4 min of standing, respectively. The average elastic modulus of theedge of the cell increased by 85% (1.85-fold increase). The delay of 5 min for stress fiber
formation is related to the activation of myosin light chain after light irradiation [19,32].
For PLP, the average elastic moduli around the cell nucleus and at the edge of thecell were estimated to be 2.74 ± 0.27 kPa and 8.58 ± 1.49 kPa before light irradiation and2.67 ± 0.24 kPa and 11.8 ± 1.7 kPa after 1 min of light irradiation, respectively. The average
elastic modulus around the cell nucleus did not increase, but the average elastic modulus at
the cell body increased by 38% (1.38-fold increase), which is a larger increase than for NPe6.
The average moduli of elasticity around the cell nucleus and at cell body was estimatedto be 3.53 ± 0.35 kPa and 13.2 ± 1.7 kPa before light irradiation, which increased to4.33 ± 0.37 kPa and 22.3 ± 1.8 kPa after 1 min of light irradiation and 4 min of standing in
the incubator, respectively. The average moduli of elasticity around the cell nucleus andthe cell body increased. However, the rate of increase in the average elastic modulus of theedge of the cell was large, increasing by 69% (1.69-fold increase).
The standard error of the average elastic modulus after 1 min of irradiation and 4 minof standing was smaller for PLP than for NPe6. This suggests that PLP has a more uniform
effect on the whole cell, which may be consistent with the high therapeutic efficacy of PDT
observed in in vivo studies [18].
2.6. Mechanistic Differences between NPe6 and PLP
Based on the above results, we consider the reason for the difference in the effects
of NPe6 and PLP on PDT. NPe6 and PLP absorb the light around 655 nm. The molecular
structure that generates singlet oxygen is a porphyrin ring in NPe6 and a porphyrin ringand chlorine in PLP [13–15,18]. We discuss why the timing and amount of RhoA activationand switching of the RhoA*/ROCK generation mechanism are different.
Figure 8a,b, respectively show confocal fluorescence observation images of NPe6
and PLP added samples during 1 min of irradiation with light of 640 nm wavelength at
439 mW/cm2 . It has been suggested that NPe6 is taken up by lysosomes and PLP by phagosomes. For NPe6, no significant structural changes were observed in the lysosomes where
NPe6 is thought to accumulate. In contrast, phagosomes are deformed after irradiation,
and almost all of them are destroyed within 50 s to 1 min.

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9 of 12Figure 8c shows a schematic diagram of PLP uptake by phagosomes and the destruction of the phagosome membrane by PDT. PLP induces phagosome formation and isincorporated therein. PLP is considered to accumulate in the phagosome membrane, andthereby a ring-like structure was observed in the fluorescence image shown in Figure 1.
Phagosomes and lysosomes are known to fuse to form phagolysosomes [23,26–31]. As
shown in Figure 1, phagosomes are fused with granular substances with a diameter ofseveral hundred nanometers that emit strong fluorescence, which could be lysosomes.
Phagolysosomes are known to produce non-mitochondrial generation of ROS [39]. Therefore, in addition to ROS generated from the porphyrin ring and chlorine of PLP, other ROS
are also generated by the oxidation of substances flowing from lysosomes. It is consideredthat large amounts of ROS are released when the phagosome membrane is destroyed bylight irradiation of the PLP accumulated there. The reason why RhoA is not immediately
activated by light irradiation in PLP, unlike NPe6, is well explained by the fact that it takes
time for the phagolysosomal membrane to be destroyed.

Figure 8. Confocal fluorescence observation images of (a) NPe6 and (b) PLP during 1 min of
irradiation with light of 640 nm wavelength and 439 mW/cm2 . (c) Schematic diagram of PLP uptake
and destruction of phagosome membrane. In (a), the contrast has been adjusted.

As clarified in previous NPe6 research, there is a threshold amount of ROS involvedin the increase in actin filament formation [19]. That is, there is an upper limit to theconcentration of RhoA that is activated and binds to ROCK. When an amount of ROS
that greatly exceeds this threshold is generated in a short time, the surplus ROS destroy
cells regardless of the increase in actin filament formation [19]. When the amount of ROS
generated exceeds the threshold but does not cause cell necrosis, the concentration of RhoA,
which binds to ROCK, reaches the upper limit and the elastic modulus stops increasing.
This is because the amount of ROCK is almost fixed. After 1 min of irradiation and 4 minof standing in an incubator, the amount of singlet oxygen generated is greater in PLP
than in NPe6. However, as shown in Figure 7, both PLP and NPe6 resulted in similar
increases in the average elastic modulus of the cell edges. This is probably because bothphotosensitizers reached the upper limit of the concentration of RhoA* that binds to ROCK.
Finally, we consider the reason why the elastic modulus of the cell edge slightly
increased immediately after irradiation in the case of PLP. As shown in the schematic
diagram of Figure 8c, the phagosome membrane is broken, and the release of ROS andhydrolytic enzymes causes cell contraction, increasing the tension at the cell edges and the
local elastic modulus. However, this observation is inconsistent with the absence of RhoA
activation as shown in Figure 5. On the other hand, it is known that actin polymerization

Int. J. Mol. Sci. 2022, 23, 13140

10 of 12and pseudopodia formation are induced during phagosome formation [29]. Therefore, theincrease in the elastic modulus of the cell edges at the early stage of PDT with PLP may
be caused by such a mechanism, which is different from the mechanism activating RhoA
with ROS discussed in the case of NPe6. Nevertheless, it was revealed that PLP induces
RhoA activation through a mechanism different from that of NPe6 and exhibits different
dynamical effects of PDT.
3. Materials and Methods
NPe6 is a photosensitizer with a porphyrin ring and has been widely used in PDT. PLP
is a photosensitizer that contains a hydrophobic drug-loadable core enveloped in a porphyrin–
lipid monolayer and constrained by ApoA-1 mimetic R4F (Ac-FAEKFKEAVKDYFAKFWD)
peptide networks [14,15]. The cells used in this study were rat gastric mucosa-derived
cancer-like mutant RGK1 cells [20]. RGK1, a chemically induced oncogenic cancer-likemutant of RGM1 (RGM1, a rat gastric epithelial cell line, was purchased from RIKEN CELLBANK (Ibaraki, Japan)), was established [20]. RGK1 cells were cultured in DMEM/F12
without L-glutamine (Sigma-Aldrich Japan K.K., Tokyo, Japan). NPe6 (final concentration
78 µM) or PLP (final concentration 19 µM) added to samples, which were allowed tostand for 24 h in an incubator at 37 ◦ C with a CO2 concentration of 5%. These values are
not toxic [18]. In this study, it was carried out for 24 h so that the amount of sensitizer
entering the cells was saturated. There is an experiment in which the incubation time wasvaried [18]. It has been confirmed that it does not saturate in 3 h but saturates in 12 h. We
used 24 h to saturate the uptake. Phase-contrast and fluorescence observations and theirtime-lapse observations were performed with a IX83 microscope system (Olympus Corp.)
equipped with a stage top incubator (Tokai Hit Co., Ltd.). A super-resolution observation
unit (Yokogawa Electric Corp., CSU-W1 SoRa) was installed in the IX83 system and used toobserve the accumulation sites of NPe6 and PLP. For the light irradiation of PDT, a whitelight source of the IX83 system for fluorescence observation equipped with a 655 nm filter
and a 640 nm semiconductor laser in the CSU-W1 SoRa system was used.
NPe6 or PLP was added to RGK1 cells cultured in a dish of 60 mm diameter andallowed to stand for 24 h in an incubator at 37 ◦ C with a CO2 concentration of 5%. These cells
were also used for Western blotting to quantify the amount of RhoA. AFM observation was
performed by setting the AFM system (Oxford Instruments MFP-3D-BIO) on a microscope(Olympus Corp. IX71) so that the elastic modulus of the same cells could be measured
before and after light irradiation. After AFM observation using the IX71 system, the sample
was moved to the stage top incubator placed on an IX83, irradiated with light, and thenreturned to the IX71 system. The same cells were found by phase-contrast observation, andAFM observation was performed on them.
4. Conclusions
We investigated the difference in the mechanism of PDT between two photosensitizers, PLP, which has recently attracted attention for its potential to be highly effective intreating cancer, and NPe6, which has been widely used. It was revealed that PLP induces
RhoA activation through a mechanism different from that of NPe6 and exhibits different
dynamical effects of PDT: (1) NPe6 accumulates in lysosomes, whereas PLP is incorporated
into phagosomes, whose formation is induced by PLP injection into cells. (2) PDT causes
NPe6 to generate ROS, thereby producing actin filaments and stress fibers. (3) In the caseof PLP, ROS generated by PDT remain in the phagosomes until the phagosomal membrane
is destroyed, which delays the initiation of RhoA activation and RhoA*/ROCK generation.
(4) However, after the disruption of the phagosome membrane, various ROS outflow, accelerating the production of actin filaments and stress fibers, and blebbing occurs earlier thanin the case of NPe6. (5) In the initial stage of PDT, PLP increased the elastic modulus of cells
without RhoA activity. The increase in the cell edge’s elastic modulus of the cell edges iscaused by a mechanism different from that considered so far for NPe6. That may be because
phagosomes are involved in polymerizing actin filaments and pseudopodia formation.

Int. J. Mol. Sci. 2022, 23, 13140

11 of 12Considering the high selectivity and uptake of PLP into cancer cells, a greater effectwith PDT can be expected by skillfully combining the newly discovered characteristics,
such as the appearance of a strong effect at an early stage. For example, as mentioned in
the introduction, it has been confirmed that PLP is highly effective against NOZ cells [18].
Considering that the mechanism is common, this result is expected to work as a similar
mechanism for other cells. Since PLP accumulates in phagosomes, which can be visualized, it is also expected to play an important role in studying the details of phagosome(autophagosome) dynamics in living cells.
Author Contributions: M.K. took data and analyzed them with S.U., Y.N. and A.T. Besides, A.T.
developed the system. H.K., T.O., Y.A., H.M. and O.T. provided analytical and technical advices.
H.S. organized and supervised the project and edited the paper with A.T. All authors have read andagreed to the published version of the manuscript.
Funding: We acknowledge the financial supports of a Grant-in-Aid for Scientific Research (17H06088,
JP19H02591, JP19K16854) from Japan Society for the Promotion of Science and Japan Core Research
Evolutional Science and Technology (CREST) (JPMJCR1875).
Institutional Review Board Statement: Not applicable.
Informed Consent Statement: Not applicable.
Data Availability Statement: Not applicable.
Acknowledgments: We thank Yokogawa Electric Corporation for the super resolution observations.
Conflicts of Interest: The authors declare no conflict of interest.

References
1.
2.
3.
4.
5.

6.

7.
8.
9.
10.

11.

12.
13.
14.

Sharman, W.M.; Allen, C.M.; van Lier, J.E. [35] Role of Activated Oxygen Species in Photodynamic Therapy. In Methods in
Enzymology; Singlet Oxygen, UV-A, and Ozone; Academic Press: Cambridge, MA, USA, 2000; Volume 319, pp. 376–400.
Stables, G.I.; Ash, D.V. Photodynamic Therapy. Cancer Treat. Rev. 1995, 21, 311–323. [CrossRef]
Pass, H.I. Photodynamic Therapy in Oncology: Mechanisms and Clinical Use. JNCI J. Natl. Cancer Inst. 1993, 85, 443–456.
[CrossRef]
Dougherty, T.J.; Kaufman, J.E.; Goldfarb, A.; Weishaupt, K.R.; Boyle, D.; Mittleman, A. Photoradiation Therapy for the Treatmentof Malignant Tumors. Cancer Res. 1978, 38, 2628–2635. [PubMed]
Treatment of Age-Related Macular Degeneration with Photodynamic Therapy(TAP) Study Group. Photodynamic Therapyof Subfoveal Choroidal Neovascularization in Age-Related Macular Degeneration with Verteporfin: Two-Year Results of2 Randomized Clinical Trials—TAP Report 2. Arch. Ophthalmol. 2001, 119, 198–207.
Blinder, K.J.; Bradley, S.; Bressler, N.M.; Bressler, S.B.; Donati, G.; Hao, Y.; Ma, C.; Menchini, U.; Miller, J.; Virgili, G.; et al. Effectof Lesion Size, Visual Acuity, and Lesion Composition on Visual Acuity Change with and without Verteporfin Therapy for
Choroidal Neovascularization Secondary to Age-Related Macular Degeneration: TAP and VIP Report No. 1. Am. J. Ophthalmol.
2003, 136, 407–418. [CrossRef] [PubMed]
Kennedy, J.C.; Pottier, R.H.; Pross, D.C. Photodynamic Therapy with Endogenous Protoporphyrin: IX: Basic Principles andPresent Clinical Experience. J. Photochem. Photobiol. B Biol. 1990, 6, 143–148. [CrossRef]
Cieplik, F.; Deng, D.; Crielaard, W.; Buchalla, W.; Hellwig, E.; Al-Ahmad, A.; Maisch, T. Antimicrobial Photodynamic Therapy—
What We Know and What We Don’t. Crit. Rev. Microbiol. 2018, 44, 571–589. [CrossRef]
Afrasiabi, S.; Partoazar, A.; Chiniforush, N.; Goudarzi, R. The Potential Application of Natural Photosensitizers Used in
Antimicrobial Photodynamic Therapy against Oral Infections. Pharmaceuticals 2022, 15, 767. [CrossRef]
Ichinose-Tsuno, A.; Aoki, A.; Takeuchi, Y.; Kirikae, T.; Shimbo, T.; Lee, M.-C.; Yoshino, F.; Maruoka, Y.; Itoh, T.; Ishikawa, I.;et al. Antimicrobial Photodynamic Therapy Suppresses Dental Plaque Formation in Healthy Adults: A Randomized Controlled
Clinical Trial. BMC Oral Health 2014, 14, 152. [CrossRef]
Pérez-Laguna, V.; Gilaberte, Y.; Millán-Lou, M.I.; Agut, M.; Nonell, S.; Rezusta, A.; Hamblin, M.R. A Combination of Photodynamic Therapy and Antimicrobial Compounds to Treat Skin and Mucosal Infections: A Systematic Review. Photochem. Photobiol.
Sci. 2019, 18, 1020–1029. [CrossRef]
Foote, C.S. Definition of Type I and Type II Photosensitized Oxidation. Photochem. Photobiol. 1991, 54, 659. [CrossRef] [PubMed]
Ethirajan, M.; Chen, Y.; Joshi, P.; Pandey, R.K. The Role of Porphyrin Chemistry in Tumor Imaging and Photodynamic Therapy.
Chem. Soc. Rev. 2010, 40, 340–362. [CrossRef] [PubMed]
Lovell, J.F.; Jin, C.S.; Huynh, E.; Jin, H.; Kim, C.; Rubinstein, J.L.; Chan, W.C.W.; Cao, W.; Wang, L.V.; Zheng, G. Porphysome
Nanovesicles Generated by Porphyrin Bilayers for Use as Multimodal Biophotonic Contrast Agents. Nat. Mater. 2011, 10, 324–332.
[CrossRef] [PubMed]

Int. J. Mol. Sci. 2022, 23, 13140

15.
16.

17.
18.
19.
20.

21.
22.
23.
24.

25.
26.

27.
28.
29.
30.
31.

32.
33.

34.
35.

36.
37.

38.
39.

12 of 12Cui, L.; Lin, Q.; Jin, C.S.; Jiang, W.; Huang, H.; Ding, L.; Muhanna, N.; Irish, J.C.; Wang, F.; Chen, J.; et al. A PEGylation-Free
Biomimetic Porphyrin Nanoplatform for Personalized Cancer Theranostics. ACS Nano 2015, 9, 4484–4495. [CrossRef] [PubMed]
Nakai, Y.; Inoue, K.; Abe, N.; Hatakeyama, M.; Ohta, K.; Otagiri, M.; Hayashi, Y.; Yuasa, H. Functional Characterization of
Human Proton-Coupled Folate Transporter/Heme Carrier Protein 1 Heterologously Expressed in Mammalian Cells as a Folate
Transporter. J. Pharm. Exp. 2007, 322, 469–476. [CrossRef]
Le Blanc, S.; Garrick, M.D.; Arredondo, M. Heme Carrier Protein 1 Transports Heme and Is Involved in Heme-Fe Metabolism.
Am. J. Physiol.-Cell Physiol. 2012, 302, C1780–C1785. [CrossRef]
Kurokawa, H.; Ito, H.; Matsui, H. Porphylipoprotein Accumulation and Porphylipoprotein Photodynamic Therapy Effects
Involving Cancer Cell-Specific Cytotoxicity. Int. J. Mol. Sci. 2021, 22, 7306. [CrossRef]
Taninaka, A.; Ugajin, S.; Kurokawa, H.; Nagoshi, Y.; Kamiyanagi, M.; Takeuchi, O.; Matsui, H.; Shigekawa, H. Direct Analysis of
the Actin-Filament Formation Effect in Photodynamic Therapy. RSC Adv. 2022, 12, 5878–5889. [CrossRef]
Shimokawa, O.; Matsui, H.; Nagano, Y.; Kaneko, T.; Shibahara, T.; Nakahara, A.; Hyodo, I.; Yanaka, A.; Majima, H.J.; Nakamura, Y.;
et al. Neoplastic Transformation and Induction of H+,K+-Adenosine Triphosphatase by N-Methyl-N0 -Nitro-N-Nitrosoguanidine
in the Gastric Epithelial RGM-1 Cell Line. In Vitro Cell. Dev. Biol. Anim. 2008, 44, 26–30. [CrossRef]
Henderson, B.W.; Dougherty, T.J. How Does Photodynamic Therapy Work? Photochem. Photobiol. 1992, 55, 145–157. [CrossRef]
Nishida, K.; Tojo, T.; Kondo, T.; Yuasa, M. Evaluation of the Correlation between Porphyrin Accumulation in Cancer Cells andFunctional Positions for Application as a Drug Carrier. Sci. Rep. 2021, 11, 2046. [CrossRef]
Hamasaki, M.; Furuta, N.; Matsuda, A.; Nezu, A.; Yamamoto, A.; Fujita, N.; Oomori, H.; Noda, T.; Haraguchi, T.; Hiraoka, Y.; et al.
Autophagosomes Form at ER–Mitochondria Contact Sites. Nature 2013, 495, 389–393. [CrossRef]
Duclos, S.; Diez, R.; Garin, J.; Papadopoulou, B.; Descoteaux, A.; Stenmark, H.; Desjardins, M. Rab5 Regulates the Kiss and
Run Fusion between Phagosomes and Endosomes and the Acquisition of Phagosome Leishmanicidal Properties in RAW 264.7
Macrophages. J. Cell Sci. 2000, 113, 3531–3541. [CrossRef]
Goren, M.B.; D’Arcy Hart, P.; Young, M.R.; Armstrong, J.A. Prevention of Phagosome-Lysosome Fusion in Cultured Macrophages
by Sulfatides of Mycobacterium Tuberculosis. Proc. Natl. Acad. Sci. USA 1976, 73, 2510–2514. [CrossRef]
Mizushima, N.; Yamamoto, A.; Hatano, M.; Kobayashi, Y.; Kabeya, Y.; Suzuki, K.; Tokuhisa, T.; Ohsumi, Y.; Yoshimori, T.
Dissection of Autophagosome Formation Using Apg5-Deficient Mouse Embryonic Stem Cells. J. Cell Biol. 2001, 152, 657–668.
[CrossRef]
Peña-Ramos, O.; Chiao, L.; Liu, X.; Yu, X.; Yao, T.; He, H.; Zhou, Z. Autophagosomes Fuse to Phagosomes and Facilitate theDegradation of Apoptotic Cells in Caenorhabditis Elegans. eLife 2022, 11, e72466. [CrossRef]
Becken, U.; Jeschke, A.; Veltman, K.; Haas, A. Cell-Free Fusion of Bacteria-Containing Phagosomes with Endocytic Compartments.
Proc. Natl. Acad. Sci. USA 2010, 107, 20726–20731. [CrossRef]
Lee, H.-J.; Woo, Y.; Hahn, T.-W.; Jung, Y.M.; Jung, Y.-J. Formation and Maturation of the Phagosome: A Key Mechanism in Innate
Immunity against Intracellular Bacterial Infection. Microorganisms 2020, 8, 1298. [CrossRef]
Scott, C.C.; Botelho, R.J.; Grinstein, S. Phagosome Maturation: A Few Bugs in the System. J. Membr. Biol. 2003, 193, 137–152.
[CrossRef]
Lancaster, C.E.; Fountain, A.; Dayam, R.M.; Somerville, E.; Sheth, J.; Jacobelli, V.; Somerville, A.; Terebiznik, M.R.; Botelho, R.J.
Phagosome Resolution Regenerates Lysosomes and Maintains the Degradative Capacity in Phagocytes. J. Cell Biol. 2021, 220,e202005072. [CrossRef]
Suzuki, T.; Tanaka, M.; Sasaki, M.; Ichikawa, H.; Nishie, H.; Kataoka, H. Vascular Shutdown by Photodynamic Therapy Using
Talaporfin Sodium. Cancers 2020, 12, 2369. [CrossRef] [PubMed]
Ohgushi, M.; Matsumura, M.; Eiraku, M.; Murakami, K.; Aramaki, T.; Nishiyama, A.; Muguruma, K.; Nakano, T.; Suga, H.; Ueno,
M.; et al. Molecular Pathway and Cell State Responsible for Dissociation-Induced Apoptosis in Human Pluripotent Stem Cells.
Cell Stem Cell 2010, 7, 225–239. [CrossRef] [PubMed]
Nowak-Sliwinska, P.; van Beijnum, J.R.; van Berkel, M.; van den Bergh, H.; Griffioen, A.W. Vascular Regrowth Following
Photodynamic Therapy in the Chicken Embryo Chorioallantoic Membrane. Angiogenesis 2010, 13, 281–292. [CrossRef] [PubMed]
Cavin, S.; Riedel, T.; Rosskopfova, P.; Gonzalez, M.; Baldini, G.; Zellweger, M.; Wagnières, G.; Dyson, P.J.; Ris, H.-B.; Krueger, T.;et al. Vascular-Targeted Low Dose Photodynamic Therapy Stabilizes Tumor Vessels by Modulating Pericyte Contractility. Lasers
Surg. Med. 2019, 51, 550–561. [CrossRef]
Lukinavičius, G.; Reymond, L.; D’Este, E.; Masharina, A.; Göttfert, F.; Ta, H.; Güther, A.; Fournier, M.; Rizzo, S.; Waldmann, H.;
et al. Fluorogenic Probes for Live-Cell Imaging of the Cytoskeleton. Nat. Methods 2014, 11, 731–733. [CrossRef]
Lukinavičius, G.; Umezawa, K.; Olivier, N.; Honigmann, A.; Yang, G.; Plass, T.; Mueller, V.; Reymond, L.; Corrêa, I.R., Jr.; Luo,Z.-G.; et al. A Near-Infrared Fluorophore for Live-Cell Super-Resolution Microscopy of Cellular Proteins. Nat. Chem. 2013, 5,
132–139. [CrossRef]
Sneddon, I.N. The Relation between Load and Penetration in the Axisymmetric Boussinesq Problem for a Punch of Arbitrary
Profile. Int. J. Eng. Sci. 1965, 3, 47–57. [CrossRef]
Urban, C.F.; Lourido, S.; Zychlinsky, A. How Do Microbes Evade Neutrophil Killing? Cell. Microbiol. 2006, 8, 1687–1696.
[CrossRef]