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Cite this: RSC Adv., 2022, 12, 5878

Direct analysis of the actin-ﬁlament formationeﬀect in photodynamic therapy†
Atsushi Taninaka,ab Shunta Ugajin,a Hiromi Kurokawa,c Yu Nagoshi,aMayuka Kamiyanagi,a Osamu Takeuchi,a Hirofumi Matsuic and Hidemi Shigekawa*aPhotodynamic therapy (PDT) is a method in which a photosensitizer is administered in vivo and irradiatedwith light to generate reactive oxygen species (ROS), thereby causing the selective death of cancer cells.
Since PDT is a noninvasive cancer treatment method with few adverse eﬀects, it has attracted
considerable attention and is increasingly used. In PDT, there are two dominant processes based on theactin ﬁlament (A-ﬁlament) formation eﬀect: the destruction of cells by necrosis and vascular shutdown.
Despite the importance of its ﬁne control, the mechanism of the reaction process from the generationof reactive oxygen by photoinduction inducing the formation of A-ﬁlament and its polymerization toform stress ﬁbers (S-ﬁbers) has not yet been clariﬁed because, for example, it has been diﬃcult to
directly observe and quantify such processes in living cells by conventional methods. Here, we havecombined atomic force microscopy (AFM) with other techniques to reveal the mechanism of the Aﬁlament and S-ﬁber formation processes that underlie the cell death process due to PDT. First, it was
conﬁrmed that activation of the small G protein RhoA, which is a signal that induces an increase in Aﬁlament production, begins immediately after PDT treatment. The production of A-ﬁlament did notincrease with increasing light intensity when the amount of light was large. Namely, the activation of
RhoA reached an equilibrium state in about 1 min: however, the production of A-ﬁlament and its
polymerization continued. The observed process corresponds well with the change in the amount ofphosphorylated myosin-light chains, which induce A-ﬁlament polymerization. The increase in the elastic
Received 23rd December 2021
Accepted 11th February 2022

modulus of cells following the formation of S-ﬁber was conﬁrmed by AFM for the ﬁrst time. The
distribution of generated A-ﬁlament and S-ﬁber was consistent with the photosensitizer distribution. PDT
increases A-ﬁlament production, and when the ROS concentration is high, blebbing occurs and cells die,

DOI: 10.1039/d1ra09291j

but when it is low, cell death does not occur and S-ﬁber is formed. That is, it is expected that vascular

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shutdown can be controlled eﬃciently by adjusting the amount of photosensitizer and the light intensity.

Cancer is a disease that now aﬀects one in two people during
their lives and is still on the rise. Therefore, there is an urgent
need to establish a treatment method while also improving thesurvival rate and the quality of life aer treatment. Photodynamic therapy (PDT) is a method in which a photosensitizer isadministered in vivo and irradiated with light to generate
reactive oxygen species (ROS), thereby causing the selective
death of cancer cells.1–6 Since PDT is a noninvasive cancer
treatment method with few adverse eﬀects, it has been attracting attention and is being increasingly used to increase thequality of life of patients post-treatment. Previous studies haveshown that PDT has two main eﬀects. (1) Cancer cells and newaFaculty of Pure and Applied Sciences, University of Tsukuba, 305-8573 Ibaraki, Japan.
E-mail: hidemi@ims.tsukuba.ac.jpbc

TAKANO Co., LTD., Miyada-mura, Kamiina-gun, Nagano, 399-4301 Japan

Faculty of Medicine, University of Tsukuba, 305-8575 Ibaraki, Japan† Electronic supplementary
10.1039/d1ra09291j

information

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available.

See

DOI:

blood vessels produced by angiogenesis are destroyed bynecrosis and necroptosis induced by ROS.1,7–9 (2) The process ofaltering vascular cells that supply nutrients to cancer cells andthe cells around them causes the vascular shutdown andnecrosis of cancer cells.10–15 To use PDT eﬀectively, it is necessary to correctly understand the mechanisms of these eﬀects,
but unclear points in the mechanisms remain.
Various photosensitizers have been developed, most ofwhich are molecules with a porphyrin ring.1,3,16,17 In PDT, thisporphyrin ring is photoexcited to generate singlet oxygen, whichis converted to other ROS such as OH radicals and H2O2.1,3,16,17
In process (1) above, the main mechanism is the destruction of
the cytoplasm and cell membrane by ROS.3,11 On the other
hand, in process (2), cell destruction is considered to be caused
by cell–cell interaction and cell contraction due to an increase in
the numbers of intracellular actin laments (A-laments)
around the tumor (cancer tissue).10–15 Despite the importanceof its ne control, the mechanism of the reaction process fromthe generation of active oxygen by photoinduction inducing the© 2022 The Author(s). Published by the Royal Society of ChemistryPaperformation of A-lament and their polymerization to form stressbers (S-bers) (Fig. S1† for more details) has not yet beenclaried because, for example, it has been diﬃcult to directly
observe and quantify the A-lament and S-ber formation
processes in living cells by conventional methods.18,19
Recently, important mechanisms such as those of blebbing20
and the RhoA–Rho-associated protein kinase (ROCK)
pathway21,22 have been investigated and have attracted attention. It was claimed that the A-lament formation occurred
within 5 minutes aer PDT on the basis of the measurement ofphosphorylation of myosin and that the blood ow decreased in
the experiments using mice. However, the details of the reaction
process, such as its relationship with the activation of RhoA byactive oxygen and whether the cells are actually hardened owing
to the formation of A-lament, have not been conrmed.
Therefore, to clarify the mechanism for the further use of PDT,an urgent task is to introduce and develop a new method foranalysis.
Here, we have combined atomic force microscopy (AFM)with other techniques and analyzed the mechanism of the Alament and S-ber formation processes that underlies thecell death process due to PDT.

Experimental
In recent years, as a nondestructive method for analyzing thestate of cytoskeletons such as A-lament and microtubules
without staining, AFM has been used to measure the shape andmechanical properties of cells.23–31 Dynamic mode AFM can beused to measure the viscoelasticity of cells and observe Alament with a high elastic modulus.23,25–33 Another analysis
method is to determine the local elastic modulus by measuringforce curves.23,28,34–38 Although the method takes a longer time, it
is superior to viscoelasticity measurement by dynamic mode
AFM for the quantication of the elastic modulus because it
directly observes the mechanical properties of cells.
In addition, a uorescent probe (SPY555-actin, Cytoskeleton,
Inc.) that enables the direct imaging of A-lament in living cells
has recently been developed.39,40 It has low background noiseand can be used in multiple colors. Since the excitation wavelength of SPY555-actin (555 nm) has a small absorption coeﬃcient for taraporphyrin sodium,2,3,10,17 which is a secondgeneration photosensitizer, the eﬀect of uorescence observation on the analysis of A-lament dynamics induced by PDT is
reduced. It has been reported that SPY555-actin stabilizes Alament; thus the A-lament concentration naturally
increases when this probe is used,41 making quantitative analysis diﬃcult. However, depending on the cells and measurement conditions, the eﬀect of external stimuli such as PDT
becomes greater than that of stabilization. Under this condition, it becomes possible to use this probe with the method ofusing AFM, as carried out in this experiment.
When a photosensitizer taken into cells is irradiated withlight, singlet oxygen is generated and changes to ROS, which
activates RhoA. This change in the transformation process ofthe protein RhoA, a signal molecule of the mechanism, can beanalyzed by western blotting.

© 2022 The Author(s). Published by the Royal Society of Chemistry

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In this study, we have combined these techniques to directly
analyze the A-lament and S-ber formation processes. Namely,
to reveal the details of the mechanism of cell death due toexternal stress, a dominant eﬀect of PDT, we investigated therelationship between the activation of RhoA by reactive oxygen,
which triggers myosin phosphorylation, the increase in Alament concentration, and the increase in elastic modulus(Fig. S1† for more details).

Sample preparation and measurement
The cells used were rat gastric mucosa-derived cancer-likemutant RGK1 cells.41 Fluorescence observation, time-lapse
observation, and light irradiation were performed with an
IX83 system (Olympus Corp.) equipped with a stage top incubator (Tokai Hit Co., Ltd.). For AFM observation, an Asylum
Research MFP-3D-BIO atomic force microscope (Oxford
Instruments) was installed on an IX71 system (Olympus Corp.)
to measure the elastic modulus of the same cells before andaer light irradiation. Aer AFM observation with the IX71
system, light irradiation was performed with the IX83 system,
then the sample was moved to the IX71 system again, the same
cells were found by phase contrast microscopy observation, andAFM observation was performed (Methods for more details).

Results and discussion
A-Filament formation probed by uorescence measurement
First, we look at the results of direct observation using uorescence. Fig. 1(a)–(e) show the change observed aer 5 minutelight irradiation: (a) a phase contrast image of RGK1 stainedwith SPY555-actin (Cytoskeleton, Inc.) and its uorescence
images (b) before, (c) immediately aer, (d) 5 minutes aer, and(e) 10 minutes aer the end of light irradiation. A light witha diameter of 1.4 mm was irradiated at the center of the images
in Fig. 1 for 5 minutes by applying a bandpass lter of 635–
675 nm (center wavelength 655 nm, intensity 34 mW cm2) toa white light source attached to a uorescence microscope. If
PDT enhances the production of A-lament, the uorescenceintensity should increase only in the light-irradiated area. In
fact, Fig. 1(c)–(e) show a higher uorescence intensity in thelight-irradiated area than Fig. 1(b). See Fig. S2† for the results of
the quantitative evaluation of the uorescence intensity.
Although ROS is generated only during light irradiation, theuorescence intensity increased aer the end of the irradiation,
as shown in Fig. 1(d) and (e). That is, aer the mechanism toactivate the production of A-lament by the generation of ROS
during the light irradiation (process (a) in Fig. S1†), the processof A-lament formation (process (c) in Fig. S1†) continued forthe next 10 minutes. In the paper we referred to [ref. 13], it has
been conrmed that myosin phosphorylation begins about 5
minutes aer the light irradiation is stopped, which is in good
agreement with our results.
To further investigate the A-lament formation process, a 10
minute irradiation experiment was conducted to compare theresults with that obtained by 5 minute irradiation. Fig. 1(f)–(j)respectively show the images obtained by observations similar

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Results obtained by irradiating RGK1 with light with a center wavelength of 655 nm and an intensity of 34 mW cm2 for 5 minutes ((a)–(e))
and 10 minutes ((f)–(j)). (a) and (f) Phase contrast images before light irradiation, and ﬂuorescence images (b) and (g) before light irradiation, (c)and (h) immediately after light irradiation, (d) and (i) 5 minutes after light irradiation, and (e) and (j) 10 minutes after light irradiation.
Fig. 1to those in Fig. 1(a)–(e). First, the uorescence intensity
increased immediately aer the light irradiation, as in the caseof irradiation for 5 minutes, and the uorescence intensity
continued to increase for 5 and 10 minutes aer the end of lightirradiation. Namely, the production of A-lament continuedaer the generation of ROS. Moreover, the uorescence intensity obtained immediately aer 10 minutes of irradiation(Fig. 1(g) and (h), +30%) was higher than those obtained
immediately aer (Fig. 1(b) and (c), +20%) and 5 minutes aer
irradiation for 5 minutes (Fig. 1(b)–(d), +22%) (see Fig. S2(a) and(b)†). The amount of A-lament generated was higher in thecase of 10 minute irradiation than in the case of 5 minuteirradiation. Therefore, the amount of A-lament production isconsidered to depend on the amount of ROS generated during
light irradiation.
To further understand the mechanism, the change due to
5 min irradiation was investigated in more detail. We compared
the changes in cell structure observed by optical microscopy
with that in uorescence intensity, as shown in Fig. 2. The
middle row shows phase contrast images, the upper row showstheir enlarged views, and the lower row shows their uorescence images. Five minutes aer light irradiation, no signicantchange was observed in the cell structure. Subsequently, theuorescence intensity gradually increased. Around 20 to 25
minutes aer irradiation, as shown in Fig. 2(e) and (f), respectively, the uorescence intensity sharply increased at the location where the cells showed blebbing and swelling (seeFig. S2(c)†). Blebbing is thought to occur where the cell
membrane is damaged by ROS during light irradiation. Therefore, the inux of the SPY555-actin uorescent probe in themedium into the cells might have occurred there.
To see the eﬀect of A-lament stabilization by the SPY555actin uorescent probe,41 we examined the uorescence intensity when only the probe was inserted (Fig. S3†). When PDT was
not performed, almost no change was observed even aer 2 h(Fig. S2(d)†), and it was conrmed that the probe had little

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eﬀect on stabilizing A-lament in this experiment. That is, theresults obtained in this experiment show the generation of Alament by PDT.
Activation of RhoA probed by western blotting
Next, we take a closer look at these processes. First, to conrm themechanism that increases the production of A-lament, thechange in the transforming protein RhoA, a signal molecule ofthe mechanism, was measured by western blotting. Fig. 3 shows
the change in the amount of RhoA (normalized by the amount ofb-tubulin) obtained by 1 min irradiation, 1 min irradiation and
4 min incubation, 5 min irradiation, and 15 min irradiation at
the light intensity determined for the system (0.0531 mW cm2)(original data are also shown in Fig. S4†). The control is the result
obtained without any treatment. When RhoA is activated by ROS
to be RhoA*, RhoA* binds to ROCK and also activates mammalian diaphanous-related formin (mDia), resulting in the promotion A-lament formation13,20–22,42,43 (see Fig. S1†). Namely,a decrease in RhoA concentration indicates A-lament formation.
The RhoA concentration decreased signicantly aer 1 min
irradiation and then appeared to decrease only slightly. That is,although the mechanism of A-lament formation continues aer
1 min, as observed in the uorescence measurement, the RhoA*
production process is considered to be in an equilibrium state
within the rst 1 min.
Relationship between amount of generated reactive oxygen
and A-lament production
Next, the relationship between the amount of ROS generated andA-lament production was investigated. Fig. 4 shows the resultsof uorescence observation aer irradiating a sample with lightof diﬀerent intensities (87, 187, and 497 mW cm2) for 1 min
while keeping the temperature at 37  C. At 497 mW cm2,
probably because the scattered irradiation light was strong, theuorescence intensity outside the light spot of 1.4 mm diameter© 2022 The Author(s). Published by the Royal Society of ChemistryPaper

RSC AdvancesFig. 2 Phase contrast images (middle row), their magniﬁcations (top row), and ﬂuorescence images (bottom row) of RGK1 when light witha central wavelength of 655 nm and an intensity of 34 mW cm2 was irradiated for 5 minutes. (a) Immediately, (b) 5 min, (c) 10 min, (d) 15 min, (e)20 min, (f) 25 min, and (g) 30 min after irradiation.

Fig. 3 (a) Typical image of western blotting (see Fig. S4† for all data).
(b) RhoA concentration measured by western blotting when thesample was irradiated with light with a wavelength of 660 nm and an
intensity of 0.0531 mW cm2 (n ¼ 5: n is the number of experiments).
The values are normalized by the amount of b-tubulin antibody.

increased to the same level as that where the light was exposed.
Irradiation increased uorescence, but the uorescence intensity
did not depend on the intensity of light (87 mW cm2, 15%, 187
mW cm2, 13%, 497 mW cm2, 15%). At 187 and 497 mW cm2,
there was little change in uorescence intensity immediatelyaer, 5 minutes aer, and 10 minutes aer irradiation (1.2%)(see Fig. S2(e)–(g)†).
The bottom row of Fig. 4 shows the results when light with anintensity of 34 mW cm2 was irradiated for 3 minutes. The© 2022 The Author(s). Published by the Royal Society of Chemistryuorescence intensity for 3 minutes of irradiation is about thesame as that when irradiating the sample with light of 87 mWcm2 for 1 min; namely, the actual amount of light is about thesame. In the case of 34 mW cm2 intensity, although the light wasirradiated for 3 minutes and then the sample was allowed to standat 37  C for 7 minutes (Fig. 4(t)), the uorescence intensity was
only slightly diﬀerent from that immediately aer the irradiation(see Fig. S2(h)†).
To see the details of the change in the uorescence describedabove, we magnied phase contrast images obtained with lightintensities of 497, 87, and 34 mW cm2, as shown in Fig. 5. Whenthe sample was irradiated at 497 mW cm2 for 1 min (Fig. 5(a)),
the cells showed necrosis and swelled and ruptured in about 15
minutes. On the other hand, when the sample was irradiated at
87 mW cm2 for 1 min (Fig. 5(b)), blebbing started about 15
minutes aer the light irradiation, which continued for morethan 6 h, and the cells swelled and burst (see Movie S1†). Thisphenomenon closely resembles the “death dance” observed forhuman embryonic stem cells (hESCs) owing to the excessive
activation of myosin.20 In the case of the irradiation at 34 mWcm2, no signicant changes were observed, which was alsoconrmed by the analysis of uorescence, as shown in Fig. S5(see Fig. S2(i)†). The eﬀect of light irradiation is considered to be
determined by the light intensity and irradiation time. Under theirradiation condition of 34 mW for 1 minute, A-lament gradually increased (Fig. S2(i)†), but the cells were not destroyed in therange measured up to about 24 h. Therefore, when PDT is performed with this amount of light, immediate death due tonecrosis does not occur, and it seems that this amount of light issuitable for producing a shutdown eﬀect.
On the basis of these results, it is considered that when lightis weakened and irradiated in PDT, the mechanism of myosin
activation by RhoA/ROCK and A-lament formation does not
destroy cells.11,13,14,22,43 Although the increase in A-lament

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Fig. 4 Eﬀect of PDT on RGK1 obtained by irradiation with light with a center wavelength of 655 nm and intensities of 87 mW cm2 ((a)–(e)), 187
mW cm2 ((f)–(j)), and 497 mW cm2 ((k)–(o)) for 1 minute. (a), (f) and (k) Phase contrast images before irradiation, (b), (g) and (l) ﬂuorescence
images before irradiation, (c), (h) and (m) ﬂuorescence images immediately after irradiation, (d), (i) and (n) ﬂuorescence images after 4 minutes,
and (e), (j) and (o) ﬂuorescence images after 9 minutes. (p)–(t) Similar images obtained by irradiation with an intensity of 34 mW cm2 for 3
minutes.

production depends on the amount of ROS generated, it isconsidered that the ROS involved in the increase in A-lament
production has a threshold amount that can be tolerated bycells. That is, even if a large amount of ROS is generated ina short time, ROS exceeding the threshold amount is considered to destroy cells regardless of the increase in A-lament
production. It is necessary to reduce the amount of light suﬃciently to induce the vascular shutdown eﬀect of PDT withoutcausing cell destruction. In fact, in previous studies investigating the activation of myosin, experiments were conducted at
low light intensities.13,14,44

wavelength 545 nm, intensity 235 mW cm2) was carried out forner control. In addition, the uorescence observation of Alament was performed in the same manner as in Fig. 1 and 2,
and similar results were obtained, as shown in Fig. S7 and S8,†
respectively. Considering the diﬀerence in absorption coeﬃcient
shown in Fig. S6,† the light intensity in the case of 235 mW cm2
irradiation at 545 nm is 235 mW cm2  0.132 z 31 mW cm2,
namely, abut 10% lower than 34 mW cm2, the intensity used in
the case of a 655 nm wavelength, and the expected results wereobtained (see Fig. S2(a)–(c) and (j)–(l)†). There is a tendency thatthe amount of generated A-lament increased with increasing
the amount of light irradiation.

Wavelength dependence of PDT eﬀect
The wavelength and intensity of the excitation light are adjusted
by combining the light source and lters. Our system can select655 nm or 545 nm, which are used in ordinary uorescencemicroscopes. However, for example, at 655 nm, it is limited to theexcitation intensities shown in Fig. 4. Therefore, we tried towiden the measurement range by using 545 nm. Fig. S6† shows
the ultraviolet-visible absorption spectrum of the aqueous taraporphyrin sodium solution. As previously described, since theexcitation wavelength of SPY555-actin (555 nm) has a small
absorption coeﬃcient for taraporphyrin sodium, the eﬀect ofuorescence observation on the analysis of A-lament dynamics
induced by PDT is reduced. Therefore, instead of using light at
655 nm, which has a large absorption coeﬃcient for taraporphyrin sodium, irradiation at 535–555 nm (center

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AFM measurement
Next, the local elastic modulus of the cells was measured by
AFM. As shown in Fig. S1,† S-bers are formed by the A-lamentbound to desmosomes and aﬀect the elastic modulus of cells.
Therefore, information on the A-lament formation process canbe analyzed by measuring the elastic modulus of cells. That is,
the state of A-lament in a cell can be known from the change inelastic modulus, and thereby, information on the state of thecell including damage can be obtained.23,28,34–38 Biochemical
experiments are usually conducted at 37  C to maintain cell
activities. However, since the aim of this experiment was toaccurately measure the elastic modulus, experiments were
performed at room temperature (25  C) to reduce the change
in the measurement conditions caused by cell activities.

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Time-lapse phase contrast images of RGK1 irradiated with light with a center wavelength of 655 nm and intensities of (a) 497 mW cm2, (b)87 mW cm2, and (c) 34 mW cm2 for 1 minute.
Fig. 5

Fig. 6 shows AFM images of the sample and the force curves
measured at each point (A–F) in the images. Hysteresis wasobserved on the cells, showing that they are alive. These
measurements were performed over a region of 100 mm  100
mm on a grid of 64 points  64 points to obtain an elastic
modulus map (see Methods for details).
Fig. 7(a)–(d) show phase contrast and topographic (topo)
images of typical cells in the sample before and aer irradiation
with light of 535–555 nm wavelength (center wavelength

545 nm, intensity 550 mW cm2) for 5 minutes. From Fig. 7(a)and (b), it can be seen that no blebbing occurred in the cells andthat the cells were not necrotic despite the light irradiation. In
fact, as can be seen from Fig. 6, force curves with hysteresis wereobtained near the nucleus and the cell membrane with theapproach and retraction of the cantilever, whereas a force curve
without hysteresis was obtained on the dish. Namely, the cells
were shown to have viscoelastic properties (stress relaxation)
before and aer PDT, conrming that they were not necrotized.

Fig. 6 AFM images and force curves obtained at diﬀerent points in the images. (A) and (D) Are near the cell nucleus, (B) and (E) are on the cellbody, and (C) and (F) are at the bottom of the dish.

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Fig. 7 Results obtained by irradiating RGK1 with light with a center wavelength of 545 nm and an intensity of 550 mW cm2 for 5 minutes ((a)–(f))
and 10 minutes ((g)–(l)): phase contrast images (a) and (g) before light irradiation and (b) and (h) after light irradiation, topographic images (c) and(i) before light irradiation and (d) and (j) after light irradiation, and local elasticity maps (e) and (k) before light irradiation and (f) and (l) after lightirradiation.

The elastic modulus was estimated by tting the force
curves, measured at each point of the cell during the approachof the AFM tip, with the Hertz contact mechanical model,45 andthe elastic modulus maps obtained before and aer PDT are

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shown in Fig. 7(e) and (f), respectively. Fig. 7(g)–(l) show theresults obtained aer 10 minutes irradiation. From the phase
contrast (Fig. 7(g) and (h)) and topo (Fig. 7(i) and (j)) images, it
can be seen that the cells are not necrotic, as in the case of 5

© 2022 The Author(s). Published by the Royal Society of ChemistryPaperminutes irradiation (Fig. 7(a)–(d)). Fig. 7(l) shows a higher
elastic modulus than Fig. 7(k). In addition, the amount of
increase in elastic modulus in Fig. 7(l) is larger than that inFig. 7(f). From this, it is considered that the A-lament
production increases with the irradiation, and the productionof F-actin depends on the amount of ROS generated.
From the comparison before and aer PDT, it was found that
although the elastic modulus of the cells increased locally, it did
not increase uniformly in all cells. This indicates that the cells
are not in the same state and A-lament is in various development states. The variation may be owing to the diﬀerences inthe division cycle and the degree of diﬀerentiation of individual
cells. To estimate the relative increase in elastic modulus asa result of PDT in consideration of the development of Alament in each cell, the elastic modulus of each cell wasmeasured before and aer PDT, and the average elastic
modulus was estimated.
Fig. 8(a) shows the average elastic modulus near the cell
nucleus and in the entire cell excluding the nucleus (cell body)
before and aer PDT. Before and aer 5 minutes irradiation, theaverage elastic modulus near the cell nucleus was estimated tobe 2.63 and 2.91 kPa, respectively. The average elastic modulus of
the cell body was estimated to be 13.9 and 15.7 kPa, respectively.

RSC Advances
The mean elastic modulus increased slightly, by approximately
11% (1.11 times) near the cell nucleus and by approximately 13%(1.13 times) for the cell body. On the other hand, before and aer
10 minute irradiation, the average elastic modulus near the cell
nucleus was estimated to be 2.58 and 3.38 kPa, and the average
elastic modulus of the cell body was estimated to be 11.3 and21.4 kPa, respectively.
The relative increase in average elastic modulus is larger for
10 minutes irradiation than for 5 minutes irradiation; therelative increase in average elastic modulus near the cell
nucleus is approximately 31% (1.31 times) and that of the cellbody is approximately 89% (1.89 times). These are in good
agreement with the results observed by uorescence measurement (Fig. S2(j) and (k), (a) and (b)†).
Fig. 8(b) shows the average elastic modulus when the cells towhich taraporphyrin sodium was not added were irradiated.
There is little diﬀerence between the cases of 5 and 10 min
irradiation. The relative increase in the average elastic modulusof the cell body was estimated to be 16% (1.16 times) and 26%(1.26 times) for the cases of 5 and 10 min irradiation, respectively, both results showing a slight increase. One of the causes
is considered to be the eﬀect of stress due to AFM measurement.

Fig. 8 Changes in the average elastic modulus of RGK1 measured by AFM when the irradiation time was 5 minutes (red) and 10 minutes (blue): (a)with taraporphyrin sodium (TS) and (b) without. The number of cells measured, n, was 28, 33, 31, and 31 for (a) 5 min irradiation, (a) 10 min
irradiation, (b) 5 min irradiation, and (b) 10 min irradiation, respectively. First, as explained in Methods, the elastic modulus value was quantiﬁed foreach cell by averaging the measured values at each point in the nucleus and peripheral parts imaged in Fig. 7. First, as explained in Methods, theelastic modulus value was quantiﬁed for each cell by averaging the measured values at each point in the nucleus and peripheral parts imaged inFig. 7. The value of each bar graph is the average of the same processing for n cells measured under each condition. (c) Phase contrast image of
RGK1. (d) Fluorescence image of taraporphyrin sodium in RGK1 (shown by color of magenta). Excitation wavelength: 635–675 nm, ﬂuorescentwavelength: 696–736 nm for (c) and (d).

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Paper

Fig. 9 Schematic model explaining the PDT eﬀect based on A-ﬁlament production. TS: taraporphyrin sodium, ROS: reactive oxygen species,
MLC: myosin light chain, ROCK: Rho-associated protein kinase.

The elastic modulus of cells is mainly determined by thetension of S-bers formed by A-laments bound to desmosome(see Fig. S1†). Therefore, the details of the change from F-actin
to F-stress can be evaluated using uorescence and AFM. In
a previous study,13 it was observed that myosin phosphorylation
progressed within 5 minutes aer light irradiation, and then the
total amount decayed over 1 hour. The result that the elastic
modulus hardly changed in 5 minutes and changed greatly in
10 minutes corresponds well to the process in which A-lamentbinds to desmosome and becomes S-ber by myosin phosphorylation. The larger relative increase in the average elastic
modulus of the cell body is considered to be due to more Alament formation in the area. As shown in Fig. 8(c) and (d),taraporphyrin sodium is widely distributed in the cells except
near the nucleus.

5886 | RSC Adv., 2022, 12, 5878–5889

The change in elastic modulus is more clearly observed thanthat in uorescence measurement, indicating that the sensitivity of AFM measurement is suﬃciently high for a quantitative
analysis. Because AFM has nanoscale spatial resolution, further
applications in this eld such as evaluation of the characteristics of each cell are expected.
Model for explaining PDT based on F-actin formation
From the above results, the eﬀects of PDT using taraporphyrinsodium on cells are considered as follows. As shown in Fig. 9,
ROS is generated from taraporphyrin sodium taken up by cells
and activates RhoA.1,13,14,22 The activated RhoA binds toROCK,21,46 and the mechanism of myosin phosphorylation andA-lament production immediately begins to operate.13,14,21,22
The formation of F-actin even aer the light irradiation© 2022 The Author(s). Published by the Royal Society of ChemistryPapercorresponds well to the change in the amount of phosphorylated myosin. In addition, as revealed by the PL image and theAFM elastic modulus map, the distribution of the photosensitizer that produces the ROS triggering the A-lament formation
process is important.
On the other hand, ROS also has a mechanism of inhibiting
RhoA activation21 (nitric oxide activates cGK. See Fig. S1† forcomparison). As a result, the activation of RhoA equilibrates ata certain concentration depending on the amount of irradiation
light. Namely, there is an upper limit to the extent to which cells
can relieve oxidative stress, that is, the amount of active oxygenthat activates RhoA is considered to depend on the equilibrium
state determined by the light intensity. Active oxygen thatexceeds the threshold directly destroys cells in a short time.
Even if the cells do not die instantly, if the amount of active
oxygen is large, blebbing starts, and cells swell aer about 60
minutes and then rupture. Therefore, to necrotize cancer cells
by impaired blood ow, it is necessary to reduce the amount oflight to suppress blebbing and generate the A-lament requiredfor impaired blood ow. Recently, PDT has also been applied toretina treatment,47–49 and future developments are expected byadjusting the process revealed in this study.

Conclusion
The F-actin formation process, which is the dominant eﬀect of
PDT, was analyzed in detail using a uorescent probe that can beused to observe A-lament in living cells, by western blotting
analysis of RhoA activation, and by local elastic modulus
measurement by AFM. First, it was conrmed that activation of
the small G protein RhoA, which is a signal that induces anincrease in A-lament production, begins immediately aer PDT
treatment. The production of the A-lament did not increase withthe light intensity when the amount of light was large. Namely,
the activation of RhoA reached an equilibrium state in about
1 min; however, the production of A-lament and its polymerization continued. The observed process corresponds well to thechange in the amount of the phosphorylated myosin-lightchains, which induce A-lament polymerization. The increasein the elastic modulus of cells following the formation of S-ber
was conrmed using AFM for the rst time. The distribution ofgenerated A-lament and S-ber was consistent with the photosensitizer distribution. PDT increases A-lament production, andwhen the ROS concentration is high, blebbing occurs and cells
die, but when it is low, cell death does not occur and S-ber isformed. That is, it is expected that vascular shutdown can becontrolled eﬃciently by adjusting the amount of photosensitizer
and the light intensity. We have succeeded in demonstrating that
we can extract previously hidden information by combining
physical methods with biochemical methods.

Methods
Sample preparation
RGK1 cultured in a ask was placed in a dish with a diameter of
60 mm and allowed to stand in an incubator for one day. The dishwas administered with 50 mL of 0.69 g L1 taraporphyrin sodium© 2022 The Author(s). Published by the Royal Society of Chemistry

RSC Advances
PBS solution and allowed to stand in the incubator again for 22 to
24 h. The medium in the dish was replaced, then used as a sample.
Fluorescence observation
A uorescent probe (SPY555-actin, Cytoskeleton, Inc.) was used
for the direct imaging of A-lament in living cells. It has low
background noise and multiple colors are available. In the case of
GFP, genetic manipulation is required to use it for living cells,
and when it is used for antibody staining or living cells, it isinjected by making a hole. SPY555-actin can probe the state of Alament production while cells are still alive.39,40 SPY555-actin
was dissolved in 50 mL of anhydrous dimethyl sulfoxide(DMSO), and 2 mL of the solution was added to the sample. Thenthe sample was allowed to stand in an incubator with a temperature of 37  C for 60 minutes. Aer that, the dish was transferred
to a stage top incubator (manufactured by Tokai Hit Co., Ltd.),and uorescence observation was performed using a uorescence microscope in an environment of 37  C, saturated water
vapor, and 5% CO2 (535–555 nm bandpass lter). Aer observing
the uorescence, the bandpass lter was replaced with a 635–
675 nm bandpass lter, and the sample was irradiated with lightto examine the eﬀect of PDT. A mercury white light source (U-HG
LGPS manufactured by Olympus) was used. Then uorescenceobservation was performed again. In the case of AFM measurement of the elastic modulus (Fig. 6–8), a 535–555 nm bandpass
lter was used to nely adjust of light intensity for PDT, as
explained in the text. The exposure times were 2 and 11.7 s,
respectively, for 20 and 4 (5 area) magnications.
Western blotting analysis
A light pigtail with a wavelength of 660 nm was output froma semiconductor laser (Thorlabs: LP660-SF50) and was expandedwith an optical ber to a diameter of 60 mm, and light with anintensity of 0.0531 mW cm2, determined using the system, wasirradiated from the bottom of the sample at 37  C. Aer lightirradiation, proteins were extracted from the sample using RIPA
buﬀer, and western blotting was performed. The primary antibody RhoA (67B9) rabbit mAb (Cell Signaling Technology) wasused for the detection of RhoA, and b-tubulin antibody (ProteinTech) was used for quantitative standardization (normalization).
Modulus measurement
An Asylum Research MFP-3D-BIO atomic force microscope
manufactured by Oxford Instruments was used with an
Olympus Biolever BL-AC40TS-C2 cantilever (silicon nitride,
spring constant 0.1 N m1, half-apex angle 17.5 ). The force
curve was measured in a region of 100 mm  100 mm with a gridof 64 points  64 points, and the local elastic modulus was
obtained from the force curve at each point using Hertz's
equation and used as an elastic modulus map. When using
Hertz's equation, the elastic modulus of the cantilever was
290 GPa and Poisson's ratio of the sample was 0.5. About vecells existed in the region of 100 mm  100 mm, and the average
elastic modulus was calculated for each cell by dividing the cell
into two regions, namely, the vicinity of the nucleus and theremaining part of the cell (cell body). The values near the

RSC Adv., 2022, 12, 5878–5889 | 5887

RSC Advances
nucleus were averaged within a circular region having half thecell height as the diameter. The values of the other parts wereaveraged over the entire area except for the nucleus and the dishpart in each elastic modulus map. Here, the region with anelastic modulus of 1 MPa or more was dened as the dishportion.

Author contributions
A. T. developed the system and took data with S. U., H. K., Y. N.,
and M. K. H. M. and O. T. provided technical advice. H. S. organized and supervised the project and edited the paper with A. T.

Conﬂicts of interest
There are no conicts to declare.

Acknowledgements
We acknowledge the nancial supports of a Grant-in-Aid for
Scientic Research (JP17H06088, JP19H02591, JP19K16854)
from Japan Society for the Promotion of Science and Japan Core
Research Evolutional Science and Technology (CREST)(JPMJCR1875).

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