Pyropheophorbide-a Methyl Ester-mediated Photosensitization
Activates Transcription Factor NF-
B through the Interleukin-1
Receptor-dependent Signaling Pathway*
Jean-Yves
Matroule
§,
Giuzeppina
Bonizzi¶
,
Patrice
Morlière**,
Nicole
Paillous
,
René
Santus**,
Vincent
Bours¶§§, and
Jacques
Piette
¶¶
From the
Laboratory of Virology, ¶ Laboratory of
Medical Chemistry, Institute of Pathology B23, University of
Liège, B-4000 Liège, Belgium, the ** Muséum
National d'Histoire Naturelle, F-75231 Paris Cedex 05, and

IMRCP Laboratory, CNRS-URA 470, University
of Toulouse, F-31062 Toulouse, France
 |
ABSTRACT |
Pyropheophorbide-a methyl ester (PPME) is a
second generation of photosensitizers used in photodynamic therapy. We
demonstrated that PPME photosensitization activated NF-
B
transcription factor in colon cancer cells. Unexpectedly, this
activation occurred in two separate waves, i.e. a rapid and
transient one and a second slower but sustained phase. The former was
due to photosensitization by PPME localized in the cytoplasmic membrane
which triggered interleukin-1 receptor internalization and the
transduction pathways controlled by the interleukin-1 type I receptor.
Indeed, TRAF6 dominant negative mutant abolished NF-
B activation by
PPME photosensitization, and TRAF2 dominant negative mutant was without
any effect, and overexpression of I
B kinases increased gene
transcription controlled by NF-
B. Oxidative stress was not likely
involved in the activation. On the other hand, the slower and sustained
wave could be the product of the release of ceramide through activation
of the acidic sphingomyelinase. PPME localization within the lysosomal
membrane could explain why ceramide acted as second messenger in
NF-
B activation by PPME photosensitization. These data will allow a better understanding of the molecular basis of tumor eradication by
photodynamic therapy, in particular the importance of the host cell
response in the treatment.
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INTRODUCTION |
Photodynamic therapy
(PDT)1 is a new cancer
treatment modality that selectively destroys malignant, premalignant,
and benign lesions in patients (1-3) and is initiated by the selective
accumulation of a photosensitizing agent in malignant tissue. The
sensitizer is harmless unless and until activated by light of the
appropriate wavelength. This results in a photochemical reaction
leading to tumor destruction through a combination of direct
photodamage to cancer cells as well as tumor stroma, especially the
microvasculature of the tumor bed and the surrounding tissues (4). The
majority of photodynamic agents is provided exogenously via intravenous injection with subsequent uptake of the drug by tissues. The only drug
currently approved for therapy is a porphyrin oligomer (Photofrin) which is highly effective but exhibits several drawbacks such as (i) a
tendency to cause prolonged skin photosensitivity; (ii) an activation
wavelength lower than that optimal for effective penetration through
tissue; and (iii) a poorly defined chemical composition that makes a
detailed understanding of its mode of action and pharmacokinetics
difficult. To address these issues, new photosensitizers are being
developed, and a number of new agents are now in clinical trials.
Several groups have recently reported the antitumor efficacy of
pheophorbide- and pyropheophorbide-based photodynamic therapy (5-7).
These compounds are chemically well characterized, absorb light above
600 nm, and produce less long term normal tissue phototoxicity than
Photofrin. In the pyropheophorbide-a series, either as methyl esters or
as carboxylic acids, photosensitizing efficacy increases with the
length of the alkyl ether side chain, but the alkyl ether derivatives,
although having similar photophysical properties (singlet oxygen and
fluorescence yields), exhibit remarkable differences in
photosensitizing efficiency (8). These results suggest that besides
hydrophobicity, steric factors and conformation of the alkyl side
chains influence localization in the cells.
There are several lines of evidence suggesting that singlet oxygen
(1O2) is the major damaging species in PDT
(9-12). Other reactive oxygen species (ROS) may also be involved in
the biological effects caused by PDT (13), particularly in the case of
porphyrin derivatives used as photosensitizers (8, 14). Photochemically
targeted 1O2 is mainly responsible for
cytotoxicity caused by PDT. If target cells are not destroyed,
photo-oxidative stress may modulate the activation of nuclear
transcription factors that regulate the expression of stress response
genes. The cellular redox state is known to be involved in regulation
of gene expression, and ROS may act as chemical messengers modulating
gene expression via the activation of transduction pathways (15, 16).
Currently, the biological effects resulting from PDT-induced changes in
gene expression and signal transduction are largely unknown. As a
further step toward understanding modulation of gene expression by PDT, we report here the mechanism of NF-
B activation in colon cancer cells by pyropheophorbide-a methyl ester (PPME), a second generation photosensitizer showing great promise in PDT. Since cytokine release during PDT may have important biological effects for surrounding cells,
we decided to focus our attention on transcription factor NF-
B
because it is a redox-activated transcription factor involved in the
control of genes encoding several important cytokines such as
interleukin (IL)-1, IL-2, and IL-6 and tumor necrosis factor (TNF)-
as well as chemokines such as IL-8, regulated on activation normal T
cell expressed and secreted, and macrophage inflammatory protein-1 (see
Ref. 17 for review).
NF-
B complexes bind DNA as dimers constituted from a family of
proteins designated as the Rel/NF-
B family. In mammals, this family
contains proteins p50, p52, p65 (RelA), RelB, and c-Rel (Rel) (18, 19).
These five proteins harbor a related, but non-identical 300-amino acid
long Rel homology domain that is responsible for dimerization, nuclear
translocation, and specific DNA-binding. In addition, RelA, RelB, and
c-Rel, but not p50 or p52, contain one or two transactivating domains.
p50 and p52 derive from cytoplasmic precursors named p105 and p100,
respectively. NF-
B complexes are sequestered in the cytoplasm of
most resting cells by inhibitory proteins belonging to the I
B family
(20-23). The members of the I
B family are I
B
, I
B
,
I
B
, p100, and p105.
Following various stimuli, including the interaction of TNF-
and
IL-1
with their receptors, I
B
is first phosphorylated on
serines 32 and 36, then ubiquitinated at lysines 21 and 22, and rapidly
degraded by the proteasome, allowing NF-
B nuclear translocation and
gene activation (24, 25). In the case of these two types of cytokines,
the signal transduction pathways leading to the phosphorylation and
degradation of I
B proteins have recently been clarified in HeLa and
L293 cells (26-30). It is included in a 700-900-kDa complex called
signalosome whose important partners are proteins associated to the
TNF-
or IL-1 receptors, NF-
B-inducing kinase, IKK-
, -
, and
-
(26-31). Pro-inflammatory cytokines such as TNF-
or IL-1
or
the bacterial outer membrane component (lipopolysaccharide) are potent
activators of NF-
B, which mediate several of their biological
activities such as stimulation of the transcription in lymphocytes
through the intracellular generation of oxidative stress (32, 33).
However, the assumption that a similar mechanism is effective in other
cell lines has not yet been demonstrated.
In this paper, we report that PPME-mediated PDT of colon cancer cells
activates NF-
B, by triggering the signaling pathway controlled by
the IL-1 receptor likely without involvement of ROS. By increasing IL-1
receptor internalization, PPME photosensitization also leads to the
activation of the acidic sphingomyelinase (SMase) with intracellular
release of ceramide. Furthermore, our data suggest that, as a
consequence of PPME-mediated PDT, surviving colon cancer cells could
influence surrounding cells by releasing factors whose encoding genes
are controlled by NF-
B.
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MATERIALS AND METHODS |
Chemicals and Reagents--
Pyropheophorbide methyl ester (PPME)
was from Sigma and was used without any further purification. A stock
solution was made in ethanol (1 mM) and kept in the dark at
20 °C. PPME was diluted in the culture medium just before use and
added to exponentially growing cells. The fluorescent probes used for
cellular localization studies were purchased from Molecular
Probes-Europe (Leiden, The Netherlands). Radiolabeled nucleotides were
from ICN (United Kingdom), and 125I-labeled IL-1
and
TNF-
were from NEN Life Science Products (United Kingdom) or
Amersham Pharmacia Biotech (United Kingdom). Deuterium oxide was 99.8%
pure from Merck (Germany). All other chemicals were of reagent grade.
Anti-I
B
monoclonal antibody was obtained from C. Dargemont (Curie
Institute, Paris); anti-I
B
was obtained from Santa Cruz
Biotechnology; anti-p100 monoclonal and anti-p105 polyclonal antibodies
were obtained from U. Siebelnist (National Institutes of Health, Bethesda).
Cell Culture--
The human colon carcinoma cell line HCT-116
was grown in McCoy's 5A medium (Life Technologies, Inc., United
Kingdom) supplemented with 10% fetal calf serum (FCS, Life
Technologies, Inc.). Before photosensitization with PPME, HCT-116 cells
were grown for 1 week in 2% FCS. HCT-116 cells overexpressing I
B
mutated either at serines 32 and 36 (S32A, S36A) or at tyrosine 42 (Y42F) were obtained after transfection with the corresponding
expression plasmids bearing a gene conferring resistance to neomycin.
After a classical selection procedure (500 µg/ml neomycin), cells
were cloned and expanded in the presence of neomycin (250 µg/ml).
Plasmids--
pNF-
B-Luc reporter construct contains 5
B
sites from the human immunodeficiency virus type-1 long terminal repeat
cloned upstream of the luciferase gene (Stratagene). pTRAF6, p
TRAF6-(289-522) (34), p
TRAF2-(87-501) (35), and pIKK-
or -
constructs were gifts from D. Goeddel (Tularik). pI
B
S32A,S36A
was obtained from A. Israel (Pasteur Institute, France) and
pI
B
Y42F was obtained from J-F. Peyron (Nice, France). All
plasmids were purified using Qiagen column chromatography (Qiagen, The
Netherlands), and integrity was checked by agarose gel electrophoresis.
Exposure of HCT-116 Cells to PPME Photosensitization--
Before
photosensitization with PPME, HCT-116 cells were cultivated for 1 week
in McCoys 5A medium with 2% FCS and incubated with 2 µM
PPME during the last 20 h. Prior to irradiation, HCT-116 cells
were washed twice with PBS and then irradiated with red light (
>600 nm) at a fluence rate of 160 watts/m2 in Petri dishes
covered with PBS. After irradiation, HCT-116 cells were put back in
culture at 37 °C in McCoys 5A medium supplemented with 2% FCS. Cell
survival was determined after 24 h using trypan blue exclusion.
Photosensitization with PPME was carried out to attain 50% cell
survival after 24 h.
Cellular Localization and
Microspectrofluorimetry--
Co-localization experiments and
intracellular fluorescence spectroscopy were carried out with a
proprietary microspectrofluorometer constructed around a Leitz
"Diavert" inverted microscope equipped with a heated stage which
was maintained at 37 ± 1 °C (36, 37). Fluorescence excitation
was performed over the whole microscope field with 405 nm light from a
100-watt super-high pressure mercury lamp (~1 watt/cm2
without neutral filter). A bidimensional adjustable slit in the primary
image plane limited the area in the microscopic field from which the
fluorescence could be collected through a selected filter. In the
topographic mode, fluorescence light was reflected by a mirror to the
bidimensional cooled CCD target (1024 × 1024 pixels), which was
coupled to signal recording and processing software (Photometrics,
Tucson, AZ). Images of 400 × 400 superpixels (2 × 2 binning) corresponding to a 46 × 46-µm field were recorded, and
the exposure time was equal to 4 s. In the spectrotopographic mode, 150 grooves/nm grating replaced the mirror. The slit was reduced
to a narrow strip and used as the entrance to the grating, delineating
in the object plane a 2-µm-wide strip along the whole field from
which fluorescence was collected. This slit provided a spectral
resolution of about 5 nm. Fluorescence photons received by the
bidimensional detector produced a spectrotopographic image. Such images
(335 × 335 superpixels) recorded with a 2 × 2 binning can
be interpreted as either a succession, in the x direction, of linear monochromatic images of a 2-µm-wide strip or a succession of spectra along the y axis (46 µm), each corresponding to
an area of 2 × 0.15 µm2 of the strip. Exponentially
growing HCT 116 cells were grown as monolayers in custom-made
glass-bottom chambers designed for use with this microspectrofluorometer.
Electrophoretic Mobility Shift Assay (EMSA)--
Nuclear
extracts were isolated as described by a rapid micropreparation
technique derived from the large scale procedure of Dignam et
al. (38) based on the use of a lysis with detergent (Nonidet P-40)
followed by high salt extraction of nuclei (39). Binding reactions were
performed for 25 min at room temperature with 7.5 µg of total protein
in 20 µl of 20 mM HEPES-KOH, pH 7.9, 75 mM
NaCl, 1 mM EDTA, 5% glycerol, 0.5 mM
MgCl2, 2 µg of acetylated bovine serum albumin, 4 µg of
poly(dI-dC)·poly(dI-dC) (Amersham Pharmacia Biotech, U. K.), 1 mM dithiothreitol, and 0.2 ng of 32P-labeled
oligonucleotides (Eurogentech, Belgium). Oligonucleotides were labeled
by end-filling with the Klenow fragment of Escherichia coli
DNA polymerase I (Boehringer Mannheim, Germany) with
[32P]dATP, [32P]dCTP (NEN Life Science
Products or ICN, United Kingdom), and cold dTTP + dGTP. Labeled probes
were purified by spin chromatography on G-25 columns. DNA-protein
complexes were separated from unbound probe on native 6%
polyacrylamide gels at 150 V in 0.25 M Tris, 0.25 M sodium borate, and 0.5 mM EDTA, pH 8.0. Gels
were vacuum-dried and exposed to Fuji x-ray film at
80 °C for
16-24 h. The amounts of specific complexes were determined either by
counting the radioactivity with a PhosphorImager (Molecular Dynamics)
or by photodensitometry (LKB, Sweden) of the autoradiography.
Supershift experiments were carried out as described (40), using the
same EMSA protocol as described above except for the gel concentration
being 4%. The sequences of the probes (Eurogentech, Belgium) used in
this work are as shown in Sequence 1.
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I
B
, I
B
, p105, and p100 Detection--
I
B
,
I
B
, p105, and p100 inhibitory subunits were detected by Western
blot analysis using specific antibodies. Cytoplasmic extracts were
prepared at various times after the photosensitization by cell lysis
with a detergent (Nonidet P-40), pelleting the nuclei, and collecting
the supernatant fraction (41). Cytoplasmic proteins were added to a
loading buffer (10 mM Tris-HCl, pH 6.8, 1% SDS, 25%
glycerol, 0.1 mM
-mercaptoethanol, 0.03% bromphenol
blue), boiled, and electrophoresed on a 12% polyacrylamide-SDS gel and electro-transferred to Immobilon-P membranes (Millipore, United Kingdom). Filters were incubated with anti-I
B
(1:500 dilution), anti-I
B
(1:500 dilution), anti-p105 (1:1700 dilution), or
anti-p100 (1:2000 dilution) antibodies for 60 min at room temperature,
then with peroxidase-conjugated goat anti-rabbit IgG or anti-mouse IgG
(1:500 dilution) for 60 min at room temperature, and finally analyzed
using Amersham's enhanced chemiluminescence system (ECL) (Amersham
Pharmacia Biotech, United Kingdom) with Fuji x-ray film.
Cytokine Receptor Internalization--
Cytokine receptor
internalization was carried out as described (42). In short, HCT-116
cells were first washed in PBS, trypsinized, and counted. The cell
pellet was resuspended in 100 µl of binding buffer (McCoy's 5A-10%
FCS plus 0.2% bovine serum albumin) and 1 µl of
125I-labeled cytokine (20,000 Bq/assay) and incubated for
2 h at 4 °C with shaking before being placed at 37 °C to
initiate internalization. Cells were then centrifuged at 14,000 × g for 30 s, and the pellet was resuspended in 100 µl
of acidic buffer (50 mM glycine, pH 3.0) and placed on ice
for 90 s. Binding buffer (600 µl) was then added to the cells,
centrifuged at 14,000 × g for 30 s, and the supernatant recovered and counted. The pellet was resuspended in 70 µl of binding buffer and loaded on 300 µl of 20% (w/v) sucrose, 1% (w/v) bovine serum albumin. After centrifugation for 5 min at
14,000 × g, the pellet was recovered and counted.
Internalization was evaluated by determining the ratio between the
radioactivity measured in the cell pellet versus the total
radioactivity (outside + inside cells).
Transient Transfection Assays--
HCT-116 cells were grown in
6-well plates for 2 days in McCoy's 5A medium supplemented with 10%
FCS and transfected with 0.1 µg of
B-Luc reporter plasmid and
various amounts of expression plasmids. The total concentration of
plasmid was kept at 1 µg with pRC-CMV. Plasmids were mixed in
Opti-MEM (Life Technologies, Inc., United Kingdom), added to Fugene
liposomes (2 µl) (Boehringer Mannheim, Germany) for 15 min at room
temperature, and loaded on cells in 2 ml of McCoy's 5A containing 10%
FCS for 24 h. Then HCT-116 were treated either with TNF-
or
IL-1
for 24 h or with PPME (2 µM) for 6 h
and irradiated with red light (120 s). After irradiation, cells were
cultivated for 15 h and then washed twice in PBS, lysed for 15 min, and centrifuged at 15,000 × g for 4 min.
Luciferase activities corrected for protein amounts (Bio-Rad protein
assay) were measured in supernatants.
Ceramide Generation--
For total ceramide quantification,
HCT-116 cells were incubated before treatment for 24 h in McCoy's
5A medium with 2% FCS and 2 µCi/ml of
9,10-[3H]palmitic acid (ICN, United Kingdom). After lipid
extraction (43) and mild alkaline hydrolysis, the lower phase of the
Folch extract containing the labeled sphingolipids was evaporated.
Lipids were dissolved in chloroform/methanol (2:1, v/v), spotted, and separated on an analytical TLC. Ceramide was visualized by
I2 staining, and the corresponding spots were scraped and
quantified by liquid scintillation counting.
Quantification of Acidic and Neutral Sphingomyelinase
Activities--
Sphingomyelinase (SMase) assays were performed as
described previously (44, 45). HCT-116 cells were treated with or
without 150 units/ml TNF-
or with 2 µM PPME and light
and then incubated for various times. To measure acidic SMase activity,
50 µg of proteins were incubated for 2 h at 37 °C in a buffer
containing 250 mM sodium acetate, pH 5.0, 1 mM
EDTA, and 0.2 mCi/ml
[choline-methyl-14C]sphingomyelin (ICN, United
Kingdom; 20,000 Bq per assay). To measure neutral SMase activity, 100 µg of proteins were incubated for 2 h at 37 °C in 20 mM HEPES, pH 7.4, 1 mM MgCl2, and
0.2 mCi/ml [choline-methyl-14C]sphingomyelin
(Amersham Pharmacia Biotech, United Kingdom; 20,000 Bq per assay).
Released radioactive phosphocholine was extracted with
chloroform/methanol (2:1, v/v) and quantified by liquid scintillation counting.
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RESULTS |
PPME Localizes in Membranes and in Lysosomes--
Since
it has been previously demonstrated that many hydrophobic derivatives
of porphyrin localize at the cell membrane (46), we decided to
investigate whether PPME exhibited a similar cellular distribution.
Incubation of HCT-116 cells in the presence of 2 µM PPME
for 17 h allowed us to detect a strong PPME fluorescence at the
cytoplasmic membrane and in several internal compartments (Fig.
1B). To characterize the
nature of the intracellular compartments where PPME accumulated, cells
were co-incubated with PPME and various fluorescent probes. Using
rhodamine 123 as a fluorescent stain of mitochondria (47), it was
evident that no PPME accumulated inside the mitochondria, even after
long incubation times, e.g. overnight incubation (data not
shown). Cells were incubated with 1 µM PPME for 18 h
and then with 2 µM neutral red for the last 30 min prior
to irradiation. Under this condition, in the absence of a prolonged
wash, neutral red was expected to stain mainly the endoplasmic
reticulum-Golgi complex (48). Fig.
2A demonstrates that, whereas
the red fluorescence of PPME could be easily observed, practically no
fluorescence was recorded using a green filter that allowed detection
of neutral red emission (48). This was further illustrated in Fig.
2B with spectrotopographic images. At the onset of
irradiation, red fluorescence of PPME at 677 nm was observed without
any strong contribution in the 500-550 nm range. Twenty seconds after
the beginning of the irradiation, virtually all the PPME fluorescence
was bleached, but a significant increase in the green fluorescence with
a maximum toward 570 nm was observed. Such a maximum was in agreement
with neutral red localization in the endoplasmic reticulum-Golgi
complex (48). It was therefore suggested that neutral red as well as
some PPME co-localized in this environment, leading to the quenching of neutral fluorescence probably by singlet-singlet energy transfer from
neutral red to PPME. Irreversible PPME photobleaching inhibited the
energy transfer process and restored neutral red fluorescence. Cells
were also incubated with 0.25 µM PPME and 1 µM lucifer yellow for 18 h. Under these conditions
and as expected, the lysosomotropic dye lucifer yellow accumulated into
lysosomes as shown in Fig. 2C. This figure also demonstrated
that, in addition to weak uniform fluorescence, spots that perfectly
match those obtained with lucifer yellow fluorescence were observed.
Moreover, sequences of images obtained with a green filter to isolate
the lucifer yellow fluorescence from that of PPME showed a rapid
decrease of that fluorescence, thereby suggesting lysosome
destabilization since no loss occurred when cells were loaded with
lucifer yellow alone (data not shown). In order to confirm whether PPME
could localize in the endoplasmic reticulum and/or in the Golgi
apparatus, we incubated HCT-116 cells with 0.5 µM PPME
during 15 h and then with BCP (4 µM) during 15 min.
BCP emits a red fluorescence only in an aggregated form which is
promoted in the Golgi apparatus and in the Golgi vesicles, whereas in
the monomeric form it binds to internal membranes or to the cytoplasmic
membrane where it emits a green fluorescence (49). As shown in Fig.
2D, BCP emitted only a green fluorescence due to a
localization in the cytoplasmic membrane and in the endoplasmic reticulum. This probe turned out to perfectly co-localize with PPME
(Fig. 2D). These data were confirmed by using a carbocyanine derivative (DiOC6) which is a green fluorescent marker of the endoplasmic reticulum (50). As shown in Fig. 2F, incubation of HCT-116 cells with PPME as above and addition of DiOC6 (2 mg/ml) during 15 min allowed us to observe a clear co-localization of the two
fluorescent molecules in the endoplasmic reticulum. These data
unambiguously demonstrate that PPME accumulated at the cytoplasmic membrane but also in lysosomes and in the endoplasmic reticulum.

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Fig. 1.
A, chemical structure of PPME.
B, localization of PPME in HCT-116 cells. Cells were mixed
with 2 µM PPME in the dark and mounted on slides before
being observed by fluorescence microscopy ( exc = 515-560 nm) and under phase contrast microscopy. Arrows
indicated PPME localization at the cytoplasmic membranes or in several
internal compartments.
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Fig. 2.
A, cells were incubated with 1 µM PPME for 18 h and 2 µM neutral red
for the last 30 min before measurements. After incubation, cells were
washed twice with culture medium and left in culture medium without
serum and without phenol red for measurements. Left panel,
phase contrast image; middle panel, fluorescence image
recorded through a 645-nm cut-off filter. Maximum intensity was about
4500 counts. Right panel, fluorescence recorded through a
broad interference filter (540 ± 45 nm). Maximum intensity was
about 150 counts. Fluorescence excitation wavelength was 405 nm.
B, fluorescence spectra obtained from spectrotopographic
images. Cells were incubated with 1 µM PPME for 18 h
and 2 µM neutral red for the last 60 min. After
incubation, cells were washed twice with culture medium and left in
culture medium without serum and without phenol red for measurements.
Spectra were averaged from three adjacent pixels. Spectra were obtained
from spectrotopographic images recorded at the onset of the irradiation
(full line) and 20 s later (dashed line).
The fluorescence excitation and irradiation wavelength was 405 nm.
C, cells were incubated for 18 h with 0.25 µM PPME and 1 µM lucifer yellow. After
incubation, cells were washed twice with culture medium and left in
culture medium without serum and without phenol red for measurements.
Left panel, phase contrast image; middle panel,
fluorescence image recorded through a 645-nm cut-off filter. Maximum
intensity was about 600 counts. Right panel, fluorescence
recorded through a broad interference filter (540 ± 45 nm).
Maximum intensity was about 1000 counts. Fluorescence excitation
wavelength was 405 nm. D, cells were incubated for 18 h
with 0.25 µM PPME and 4 µM BodipyCeram.
After incubation, cells were washed twice with culture medium and left
in culture medium without serum and without phenol red for
measurements. Left panel, phase contrast image; middle
panel, fluorescence image recorded through a 645-nm cut-off
filter. Maximum intensity was about 954 counts. Right panel,
fluorescence recorded through a broad filter (536 nm). Maximum
intensity was about 240 counts. Fluorescence excitation wavelength was
404 nm. E, cells were incubated for 18 h with 0.25 µM PPME and carbocyanine (DiOC, 2 mg/ml). After
incubation, cells were washed twice with culture medium and left in
culture medium without serum and without phenol red for measurements.
Left panel, phase contrast image; middle panel,:
fluorescence image recorded through a 645-nm cut-off filter. Maximum
intensity was about 1252 counts. Right panel, fluorescence
recorded through a broad filter (536 nm). Maximum intensity was about
6567 counts. Fluorescence excitation wavelength was 404 nm.
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NF-
B Activation Occurs in Two Separate Waves--
Determining
whether photodynamic photosensitizers such as PPME which localized in
cytoplasmic and in internal membranes could activate NF-
B was of
interest because it could lead to important information on a possible
immunomodulation by tumor cells treated by such a photoactive drug. To
this end, HCT-116 cells were incubated for 17 h in the dark with 2 µM PPME and then irradiated with red light. Nuclear
extracts were prepared at various times after photosensitization and
analyzed by EMSA. As shown in Fig.
3A, an important retarded band
appeared transiently after irradiation with its maximal intensity observed between 10 and 30 min. The intensity of the band decreased, almost disappearing after 1 h. A second wave of NF-
B activation could then be observed 2 h after irradiation (Fig. 3A).
Contrary to the first wave of activation, the second appeared slowly
and was sustained up to 24 h. This is the first demonstration that an inducing agent could activate NF-
B in two separate waves as follows: a rapid and transient phase followed by a slower and sustained
one. Competition experiments carried out with a wild-type or a mutated
unlabeled NF-
B probe demonstrated that the upper retarded band was
specific, whereas the lower one was not (Fig. 3B). To
determine whether the second phase of activation was due to a
post-transcriptional mechanism, HCT-116 cells were preincubated with
cycloheximide for 60 min before being photosensitized by PPME. As shown
in Fig. 3A, the intensity of the NF-
B band was not
decreased by cycloheximide, demonstrating that the two waves of NF-
B
activation were due to a post-transcriptional activation mechanism.
Using antibodies directed against the various members of the
Rel/NF-
B family (p50, RelA, c-Rel, RelB, and p52), we also observed
that the retarded complex involved the classical p50/RelA heterodimer
(Fig. 3C).

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Fig. 3.
The effect of PPME photoreaction on
B-DNA binding activities in HCT-116 cells.
A, induction of a nuclear B enhancer DNA-binding protein
after treatment of HCT-116 cells with 2 µM PPME for
17 h and 200 s of irradiation with red light. HCT-116 cells were
also preincubated for 60 min in the presence of cycloheximide
(CHX) at 25 µg/ml. Nuclear extracts were prepared at
various times after irradiation, and equal amounts of protein were
mixed with a 32P-labeled B probe. Samples were loaded on
6% native polyacrylamide gels and electrophoresed at 150 V. The
autoradiogram of the gel is shown, and the arrow indicates
the position of the specific complex; the nonspecific DNA binding was
noted, n.s. (the position of the free probe was not shown).
B, competition analysis. Nuclear extracts from HCT-116 cells
photosensitized with 2 µM PPME were prepared 240 min
after the light treatment and analyzed by EMSA as in A.
Nuclear extracts were either mixed directly with the
32P-labeled probe or with the labeled probe in the presence
of 5 or 20 molar excess of wild-type or mutated B probe.
C, immunoreactivity of the PPME-inducible protein- B
enhancer complex. Nuclear extracts from HCT-116 cells photosensitized
as above were either mixed directly with the 32P-labeled
B probe or incubated with antisera specific for p50, RelA, c-Rel,
RelB, and p52 before being mixed with the 32P-labeled B
probe. Samples were then loaded on a 6% polyacrylamide native gel and
electrophoresed. D, transient transfection assay of HCT-116
cells with a B-Luc reporter construct. HCT-116 cells were
transfected with 0.1 µg of B-Luc before being treated with 150 units/ml IL-1 or 150 units/ml TNF- for 24 h or with 2 µM PPME with or without light and then left in culture
for 15 h. Luciferase activity was then measured in cell extracts
and expressed as fold increase over untreated cells.
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In order to ascertain whether the NF-
B complex induced by PPME and
light was transcriptionally active, we transiently transfected HCT-116
cells with a
B-Luc reporter plasmid construct. As shown in Fig.
3D, transfected HCT-116 cells incubated with PPME in the dark did not give rise to detectable
B-driven transcriptional activity, whereas photosensitization of the transfected HCT-116 cells
allowed us to detect increased transcriptional activity (4-fold),
demonstrating that the p50/RelA heterodimer found in cell nuclei by
EMSA was transcriptionally active. On the other hand, classical NF-
B
inducers such as TNF-
or IL-1
led to a 5- and 10-fold increase in
the
B-driven transcriptional activity, respectively (Fig.
3D).
Western blot analysis of cytoplasmic extracts from HCT-116 cells
photosensitized by PPME also showed a rapid and transient decrease in
the amount of the I
B
molecule after 30 min (Fig. 4A). Analysis of the band
intensity revealed that about 50% of the cytoplasmic I
B
pool was
degraded after 30 min. Similarly to what was observed by EMSA (see Fig.
3A), the I
B
degradation is transient and the
cytoplasmic I
B
pool was replenished after 1 h before
starting again to decrease (Fig. 4A). Concomitantly to what
was observed by EMSA, this second wave of I
B
degradation was slow
and sustained. After 24 h almost all the I
B
pool was degraded. These data unambiguously demonstrated that NF-
B activation by PPME and red light involved I
B
degradation in two different waves, likely involving different mechanisms. The fate of three other
inhibitory Rel proteins was also followed by Western blots. As shown in
Fig. 4, B-D, neither I
B
, p105, nor p100
inhibitors were degraded following PPME photosensitization, showing
that in HCT-116 cells most of the NF-
B activation mechanism was
controlled by I
B
degradation.

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Fig. 4.
Analysis of the NF- B
inhibitory proteins after treatment of HCT-116 cells with PPME and
light. Cytoplasmic extracts were prepared at various times after
the photoreaction and analyzed by Western blot with different
antibodies. A, I B ; B, I B ;
C, p105; and D, p100. The arrows
indicate the position of the various inhibitory proteins.
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Singlet oxygen has frequently been involved as secondary messenger in
cells treated by photodynamic therapy (9-12). In order to evaluate the
role of this reactive oxygen species in NF-
B activation, HCT-116
cells were irradiated in PBS where H2O was replaced by
D2O, since singlet oxygen lifetime is increased by this
isotopic substitution (51). EMSA analysis revealed that isotopic
substitution significantly increased the intensity of the p50-RelA
complex found in the nucleus of HCT-116 cells photosensitized by PPME
(Fig. 5A). Interestingly, this
increase in band intensity was mainly detectable during the first rapid
and transient phase (Fig. 5A). However, these data by
themselves could not directly implicate singlet oxygen as a mediator of
NF-
B activation because isotopic replacement could modify other
physicochemical parameters such as excited state lifetime (51). In
order to clarify this point, we decided to investigate the effects of
several antioxidant molecules (e.g. singlet oxygen
quenchers) on NF-
B activation by PPME photosensitization. Many
studies have shown that antioxidants blocked NF-
B activation by ROS
such as hydrogen peroxide (14) but also when pro-inflammatory cytokines
were used as stimuli (15). The chosen antioxidants were as follows: (i)
classical hydrophilic molecules such as
N-acetyl-L-cysteine (40 and 50 mM added 60 min before irradiation), pyrrolidine 9-dithiocarbamate (100-500 µM added 60 min before irradiation), and Trolox
(a water-soluble derivative of vitamin E, 500 µM added 90 min before irradiation); and (ii) lipophilic antioxidants such as
vitamin E (100 and 400 µM added either 60 min or 24 h before irradiation) and vitamin E acetate (400 µM added
24 h before irradiation). As shown in Fig. 5B, vitamin
E acetate and all other of these antioxidant molecules did not exhibit
inhibitory effects on NF-
B activation in HCT-116 cells by PPME
photosensitization indicating that the intracellular release of ROS was
probably not involved in NF-
B activation. To determine further
whether ROS were involved or not in NF-
B activation by PPME
photosensitization, lipoperoxides were measured by TBARs assay. Under
experimental conditions leading to maximal NF-
B activation, there
was no lipoperoxide detectable in treated HCT-116 cells, confirming
that NF-
B activation by PPME photosensitization did not involve ROS
generation and membrane peroxidation.

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Fig. 5.
A, EMSA analysis of NF- B activity in
nuclear extracts of HCT-116 cells treated with 2 µM PPME
and light in the presence or absence of deuterium oxide. EMSA were
carried out as in Fig. 3, and the arrows indicate the
specific NF- B complex and a nonspecific band (n.s.).
B, EMSA analysis of NF- B activity in nuclear extracts of
HCT-116 cells treated with 2 µM PPME and light in the
presence or absence of -tocopherol (vitamin E) acetate (400 µM added 24 h before irradiation). EMSA were carried
out as in Fig. 3, and the arrows indicate the specific
NF- B complex and a nonspecific band (n.s.).
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PPME Photosensitization Promotes IL-1 Receptor
Internalization--
Detection in HCT-116 cells photosensitized by
PPME of a rapid and transient activation of NF-
B could mimic what
was observed in many cell types after treatment with cytokines. To
determine whether PPME photosensitization mimicked cell activation by
cytokines, we investigated the effects of PPME photosensitization on
cytokine receptor in terms of receptor internalization. We first looked at IL-1 receptor internalization. As shown in Fig.
6A, addition of radiolabeled
IL-1
to HCT-116 allowed us to observe its internalization within 30 min of incubation at 37 °C (lanes 1-3). This
internalization could be competed out by preincubation of cells with
unlabeled IL-1
(Fig. 6A, lane 4). When HCT-116 cells were
incubated with PPME for 20 h at 37 °C (lane 5) or
for 15 min at 4 °C (lane 6) and then irradiated and
pulsed with radiolabeled IL-1
, there was a significant increase in
IL-1 receptor internalization (Fig. 6A). When cells were
treated in a similar manner but preincubated with unlabeled IL-1
before PPME was added to HCT-116 cells, no internalization could be
detected (Fig. 6A, lane 7), demonstrating that PPME
photosensitization promoted IL-1 type I receptor internalization.

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Fig. 6.
A, PPME promotes IL-1 receptor
internalization in HCT-116 cells. HCT-116 cells were incubated for
2 h at 4 °C with radiolabeled IL-1 and then placed for 0 min
(lane 1), 15 min (lane 2), or 30 min (lane
3) at 37 °C to allow internalization of the receptor.
Lane 4, HCT-116 cells were preincubated for 15 min with
unlabeled IL-1 before addition of radiolabeled IL-1 and incubated
a further 15 min at 37 °C. Lane 5, HCT-116 cells were
pretreated for 20 h at 37 °C with 2 µM PPME and
illuminated before addition of radiolabeled IL-1 . Lane 6, HCT-116 cells were pretreated for 15 min at 4 °C with 2 µM PPME and illuminated before addition of radiolabeled
IL-1 . Lane 7 is identical to lane 6, except
that HCT-116 cells were incubated first with cold IL-1 for 15 min at
4 °C. B, PPME does not influence TNF- receptor
internalization. HCT-116 cells were incubated for 2 h at 4 °C
with radiolabeled TNF- before being placed at 37 °C for 0 min
(lane 1) or 30 min (lane 2). Lane 3,
HCT-116 cells were preincubated with unlabeled TNF- before being
treated as in lane 2. Lane 4, HCT-116 cells were
preincubated for 20 h at 37 °C with 2 µM PPME and
illuminated before being incubated in the presence of radiolabeled
TNF- for 2 h at 4 °C. Lane 5 is identical to
lane 4, except that preincubation with PPME was carried out
at 4 °C for 15 min. Internalization was determined as the ratio
between the amount of the radiolabeled cytokine taken up by cells and
that remaining outside cells.
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Since other cytokine receptors could be involved in the reaction, we
measured the effect of PPME photosensitization on TNF receptor. As
shown in Fig. 6B, addition of radiolabeled TNF-
to
HCT-116 cells made it possible to follow TNF receptor internalization within 30 min at 37 °C (lane 2). Again, preincubation
with unlabeled TNF-
significantly decreased internalization (Fig.
6B, lane 3). Conversely to what was observed with IL-1
receptor, PPME photosensitization did not promote TNF receptor
internalization (Fig. 6B, lanes 4 and 5),
demonstrating that its effect was specific to the IL-1 receptor.
IL-1 Signaling Mediates NF-
B Activation by PPME
Photosensitization--
The demonstration that PPME photosensitization
could favor IL-1 receptor internalization yielding a rapid and
transient activation of NF-
B prompted us to determine whether PPME
photosensitization activates NF-
B through the IL-1 signaling
pathway. To investigate this, HCT-116 cells were transfected with a
B-Luc reporter plasmid construct together with various plasmids
expressing mutant or wild-type TRAF6 protein, which is associated with
the IL-1 receptor (34). HCT-116 cells were then either photosensitized
with PPME or treated with IL-1
or TNF-
. As shown in Fig.
7A, the expression of TRAF6 in
HCT-116 cells prior to stimulation did not significantly modify
B-driven transcriptional activity in cells photosensitized with PPME
or treated with IL-1
. In cells treated with TNF-
, TRAF6
overexpression slightly inhibited
B-driven transcriptional activity
(Fig. 7A). However, when HCT-116 cells were co-transfected with a dominant negative version of TRAF6 (
TRAF6), there was a clear
down-regulation of NF-
B transactivation in cells treated with
IL-1
or photosensitized by PPME, i.e. luciferase
activities were lower than 50% of the control with 1 µg of
TRAF6
(Fig. 7A). In the case of HCT-116 treated with TNF-
,
co-transfection with
TRAF6 led also to a dose-dependent
decrease in transcriptional activity, but luciferase activity remained
close to the control value at the lowest
TRAF6 concentrations (0.1 and 0.25 µg). These data showed that PPME photosensitization and
IL-1
stimulation induced similar effects on transcription controlled
by a
B gene promoter, whereas TNF-
induced
B-dependent transcription through a distinct
pathway.

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Fig. 7.
HCT-116 cells transient transfection assays
with 0.1 µg of B-Luc
reporter plasmid in the presence of plasmids expressing various
signaling proteins. Luciferase activities were measured on cell
extracts and expressed as a percentage of the luciferase activity
obtained in cells induced either by TNF- or IL-1 or by PPME
photosensitization. A, co-transfection was carried out
either with 0.1 µg of TRAF6 expression plasmid or with increasing
amounts of a plasmid expressing a dominant negative version of TRAF6
(from 0.1 to 1 µg). Cells were treated either with 150 units/ml
IL-1 for 24 h or with 150 units/ml TNF- for 24 h or
treated with 2 µM PPME and irradiated for 2 min before
being replaced in culture for 15 h. B, co-transfection
was carried out with either 0.1 µg of TRAF2 expression plasmid or
with increasing amounts of a plasmid expressing a dominant negative
version of TRAF2 (from 0.1 to 1 µg). Cells were treated either with
IL-1 or TNF- or with PPME and light as in A.
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The role of TRAF2, an adaptor molecule associated with the TNF
receptor, was also investigated in the same assay (35). As shown in
Fig. 7B, expression of a TRAF2 dominant negative protein (
TRAF2) decreased
B-driven transcriptional activity in HCT-116 cells treated with TNF-
in a dose-dependent manner,
whereas there was no effect on cells photosensitized by PPME or treated
by IL-1
. These data were in agreement with the experiments where the
receptor internalization was measured and confirmed that PPME
photosensitization could not recruit members of the TNF-signaling pathway.
Recently, I
B kinases (IKK-
and -
) have been identified as the
main cellular kinases carrying out I
B
phosphorylation on serine
residues 32 and 36 after cell induction with pro-inflammatory cytokines
(IL-1
and TNF-
) (26-29). In order to verify whether IKKs could
be activated by PPME photosensitization, HCT-116 cells were
co-transfected by the
B-luciferase reporter plasmid and a plasmid
expressing either IKK-
or IKK-
or IKK-
and -
. As shown in
Fig. 8A, overexpression of
IKK-
or -
in unstimulated HCT-116 cells increased
B-driven
transcriptional activity. Co-overexpression of IKK-
and -
yielded
a synergistic activation giving rise to 11-fold induction of the
B-driven transcription (Fig. 8A). Similarly, overexpression of IKK-
or -
or IKK-
plus -
increased in a dose-dependent fashion
B-driven transcriptional activity
in HCT-116 cells treated with either TNF-
(Fig. 8B) or
IL-1
(data not shown). Similarly, when HCT-116 cells were
transfected to overexpress IKKs before being photosensitized with PPME,
there was a significant increase of the
B-driven transcription; this
effect was particularly significant when both IKK-
and -
were
co-expressed (Fig. 8C). To confirm the involvement of IKKs
in the PPME-mediated NF-
B activation, one HCT-116 cell line
overexpressing I
B
mutated at serines 32 and 36 (S32A,S36A) was
constructed (MUT4). In addition, an HCT-116 cell line overexpressing
I
B
mutated at tyrosine 42 (Y42F) was also generated (Tyr-42).
These two cell lines were then photosensitized as described above, and
nuclear extracts were prepared after various times to be analyzed by
EMSA. As shown in Fig. 8D, a classical NF-
B complex can
be visualized in wild-type HCT-116 cells and in HCT-116 Tyr-42 cells.
On the other hand, in the cell line (MUT4) overexpressing I
B
S32A,S36A, there was no NF-
B complex induced by photosensitization
demonstrating again that NF-
B activation by PPME required IKKs and
I
B
phosphorylation on serines 32 and 36. These data led to
identifying IKKs as the main I
B
kinases stimulated by PPME
photosensitization.

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Fig. 8.
HCT-116 cells transient transfection assays
with 0.1 µg of B-Luc
reporter plasmid in the presence of plasmids expressing
I B kinases
(IKK- or - ).
A, co-transfection was carried out with increasing amounts
of IKK- or IKK- or IKK- plus - expression plasmids (0.1 and
0.3 µg). Luciferase activities were measured on cell extracts and
expressed as fold increase versus untransfected cells.
B, co-transfection was carried out with increasing amounts
of IKK- or IKK- or IKK- plus - expression plasmids (0.1 and
0.3 µg) before being treated with TNF- (150 units/ml) as in Fig.
7. Luciferase activities were measured on cell extracts and expressed
as percentage of the luciferase activity in HCT-116 cells treated with
TNF- alone. C, co-transfection was carried out with
increasing amounts of IKK- or IKK- or IKK- plus -
expression plasmids (0.1 and 0.3 µg) before being photosensitized
with PPME as in Fig. 7. Luciferase activities were measured on cell
extracts and expressed as percentage of the luciferase activity in
HCT-116 cells photosensitized with PPME alone. D, EMSA
analysis of NF- B activity in nuclear extracts of HCT-116 cells
(WT), of HCT-116 cells overexpressing I B mutated at
serines 32 and 36 (MUT4), and of HCT-116 cells
overexpressing I B mutated at tyrosine 42 (Y42). These
cells were grown in similar conditions before being treated with 2 µM PPME and light. EMSA were carried out as in Fig. 3,
and the arrows indicate the specific NF- B complex and a
nonspecific band (n.s.).
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NF-
B Activation by PPME Involves Ceramide
Generation--
Ceramide is a second messenger involved in signaling
pathways following TNF-
or IL-1
stimulation (44, 52). Ceramide production after cellular stimulation with pro-inflammatory cytokines results from sphingomyelin hydrolysis catalyzed by either the acidic or
the neutral sphingomyelinase (SMase). It was reported that these two
enzymes are linked to distinct pathways following interaction of
TNF-
with TNF receptor 1 (53) or IL-1
to its type 1 receptor
(52). Since PPME photosensitization utilized IL-1 signaling proteins to
activate NF-
B, ceramide production was measured at various times
after photosensitization. As shown in Fig.
9A, total ceramide
significantly increased as early as 10 and 30 min after
photosensitization, reaching its maximal value after 2 h before
gradually declining at longer times. A similar behavior was recorded in
HCT-116 cells treated with TNF-
(Fig. 9A). These data
demonstrated that ceramide generation occurred soon after
photosensitization but remained elevated for at least 6 hours,
indicating that ceramide could be considered as being a messenger of
the second wave of NF-
B induction by PPME photosensitization.

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Fig. 9.
A, ceramide production in HCT-116 cells
treated either with 150 units/ml TNF- or with PPME and light.
HCT-116 cells were labeled with 2 µCi/ml
9,10-[3H]palmitic acid. After 24 h labeling, the
medium was removed; the cells were stimulated for the indicated times,
and total cellular ceramide was determined. Values are indicated as
fold stimulation of the control (unstimulated cells). B,
activation of the acidic sphingomyelinase activity in HCT-116 cells
treated either with 150 units/ml TNF- or PPME and light. Values are
indicated as fold induction of the control (unstimulated cells).
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Since ceramide release could be attributed to either acidic or neutral
SMase activation, these two enzymatic activities were determined in
HCT-116 cells treated with TNF-
or photosensitized by PPME. From the
data presented in Fig. 9B, it is evident that both TNF-
and PPME photosensitization led to acidic SMase activation. This
activation was transient and appeared somewhat earlier than the release
of ceramide. On the other hand, neutral SMase activity was measured on
extracts from HCT-116 cells treated with 150 units/ml TNF-
or
photosensitized with PPME. Although the basal activity of neutral SMase
was rather low in HCT-116 cells, it was increased 1.3-fold after 15 min
of treatment with TNF-
(data not shown). However, no neutral SMase
activation could be detected after PPME photosensitization (data not
shown), demonstrating that ceramide generation following
photosensitization was totally due to acidic SMase activation.
To demonstrate further the involvement of acidic SMase and ceramide in
NF-
B activation by PPME photosensitization, two experiments were
carried out. First, chloroquine (100 µM), an acidic SMase inhibitor, was added 60 min prior to HCT-116 cell photosensitization with PPME, and NF-
B activation was evaluated by EMSA both 30 min and
24 h after photosensitization. As shown in Fig.
10A, chloroquine addition
significantly decreased NF-
B activation during the first (30 min)
and second activation wave (24 h), showing that acidic SMase could be
considered as a mediator of the response to PPME photosensitization.
Second, C2-ceramide was also added to HCT-116 cells to
mimic their release by the activation of acidic SMase. As shown in Fig.
10B, NF-
B activation could be clearly visualized by
addition of C2-ceramide on HCT-116 cells. NF-
B
activation was clearly observed both at a short and long time after
stress, emphasizing the role of ceramide as second messenger in PPME
photosensitization.

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Fig. 10.
A, effect of chloroquine (100 µM) on NF- B activation in HCT-116 cells
photosensitized by PPME. Chloroquine was added to the cells 60 min
prior to PPME photosensitization (carried out as in Fig. 2), and
nuclear extracts were prepared after 30 min or 24 h. NF- B
induction was measured by EMSA and expressed in fold induction compared
with the control (non-photosensitized cells). The intensities of the
specific band were measured by phosphorimaging. B,
C2-ceramide induces NF- B in HCT-116 cells. Cells were
treated with 30 µM C2-ceramide, and nuclear
extracts were analyzed by EMSA after various times (from 0 to 24 h). The specificity of the complex was determined by competition with a
50-fold excess of unlabeled wild-type or mutated probe (data not
shown).
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 |
DISCUSSION |
In this paper, we have shown that pyropheophorbide-a methyl ester,
a second generation photosensitizer, is a strong activator of the
NF-
B transcription factor. The underlying mechanism is rather
uncommon; PPME localizes in membranes and promotes IL-1 receptor
internalization upon photosensitization, triggering the transduction
machinery linked to the IL-1 receptor. This NF-
B activation is
transient and does not require oxidative stress. After 2 h, a
second wave of NF-
B action gradually appeared lasting up to 24 h. During these two phases, ceramides were generated, suggesting that
these lipids act as second messengers to activate NF-
B at longer
times. We postulate that PPME located in the lysosomal membrane is
responsible for ceramide generation, since acidic SMase, a lysosomal
enzyme, is activated by the photosensitizing action of PPME.
Investigating how pyropheophorbide-a can modulate gene expression is of
great importance, because tumor eradication by PDT will likely depend
not only on an efficient tumor cell killing but also on the adaptation
of tumor cells surviving treatment. Since many important genes involved
in the control of the immune system and in the inflammatory reaction
are controlled by NF-
B, we have decided to pay attention to the
mechanisms by which PPME photosensitization activates NF-
B.
Photosensitization has already been shown to induce NF-
B in T
lymphocytes (41, 54) and in other cell types (55). Although the
mechanisms have not yet been clarified, oxidative stress mediated by
photosensitization is likely implicated since antioxidants inhibit
NF-
B activation or cellular oxidation products can be detected.
1O2 is known to be the main ROS produced by
photosensitization (56) and is proposed by several authors (57-59) as
a second messenger in gene activation in human skin fibroblasts
irradiated by UV-A. Because its lifetime can be significantly increased
by deuterium substitution, the observation that NF-
B translocation
is greater in a medium where H2O is substituted by
D2O suggests that 1O2 could be
involved in the activation mechanism. However, the role of
1O2 as mediator in NF-
B activation by PPME
photosensitization could be considered unlikely because (i) none of the
tested antioxidants (hydrophilic and hydrophobic) capable of quenching
1O2 inhibit NF-
B activation and (ii) no
lipoperoxide can be detected in photosensitized HCT-116 cells. The
increased NF-
B activation by PPME photosensitization after
D2O substitution could then be explained by an increased
lifetime of the PPME excited state which has already been observed for
other photosensitizers (51). This would involve a radical mechanism
implicating either an electron or a charge transfer between a PPME
excited state and membrane proteins such as the IL-1 type 1 receptor or
the acidic SMase. Although a so-called type I reaction would be
implicated to explain NF-
B action by PPME photosensitization, we
cannot totally rule out that part of the photochemical mechanism could
be due to 1O2. Indeed, the lack of inhibition
by a lipophilic antioxidant such as vitamin E could also be explained
by a subcellular concentration too low to efficiently compete with the
reaction between the PPME excited state and membrane receptor.
Definitive proof of 1O2 involvement in NF-
B
activation could only come from directly measuring its emission at 1268 nm. However, 1O2 detection by infrared emission
on cells is currently not feasible.
One of the main contributions of this work is to unambiguously show
that PPME photosensitization can specifically mobilize the IL-1
transduction pathway leading to NF-
B activation. This is
demonstrated by (i) significant internalization of the IL-1 receptor
after photosensitization with PPME; (ii) the inhibition of NF-
B
activation by expression of TRAF6 dominant negative, a protein linked
to the IL-1 receptor; (iii) the absence of down-regulation by
expression of TRAF2 dominant negative mutant protein; and (iv) the
increased NF-
B activity when IKK-
, IKK-
, and IKK-
plus -
are overexpressed in photosensitized cells. The type of IL-1 receptor
whose internalization is increased by PPME photosensitization is likely
to be the type I receptor because the IL-1 type II receptor is not
capable of transducing signals and mainly acts as a decoy receptor
(60). Interestingly, such an internalization cannot be recorded with
the TNF receptor. One explanation could be that PPME cannot promote TNF
receptor trimerization, which is required to both transduce signals and
promote internalization (61). Whereas the molecular mechanism by which
PPME promotes IL-1 receptor internalization is unknown, it could be
interesting to determine whether this effect is restricted to receptors
that do not require homo- or heterodimerization for their functioning.
A modification of cell-surface receptor by PPME photosensitization has
recently been published (62); curiously, these authors reported that a
mixture between pheorbide-a and PPME can reduce binding of cytokines to
their receptors (TNF-
, IL-8, complement factor 5a, and epidermal growth factor). As these data were obtained (i) by measuring cytokine binding to their receptor on neutrophils and (ii) with a mixture of
photosensitizers, they reinforce the idea that these compounds can
interact with cell-surface receptors and either inactivate them or
trigger transduction pathways. The localization of photosensitizers into cellular membranes is therefore expected to be an important determinant controlling the susceptibility of surface proteins. For
example, di-hematoporphyrin ether was reported to affect the binding of
antibodies to high affinity Fc receptors but not to other surface
molecules (63), and methyl pheophorbide-b inhibited the binding of
several cytokines to their receptor but not the binding of TNF-
(64). Collectively, these data support the hypothesis that a particular
membrane environment, as well as the nature of the receptors, may
determine whether the receptor function is activated or inhibited and,
therefore, the extent of cell response to PDT.
This new concept is important because it will greatly influence our
understanding of the cellular response to PDT not only in terms of
tumor cell survival but also of the modification of tumor environment
by the release of mediators. As PDT not only reduces tumor burden but
also induces inflammation, it is proposed that recruitment of activated
macrophages to the inflamed tumor is an important factor in complete
tumor eradication (65). In this respect, study of the mechanism of
NF-
B activation by PDT is important because many genes encoding
cytokines and chemokines are controlled by this factor. Work is now in
progress in our laboratory to determine the nature of the genes that
are up- and down-regulated by PPME photosensitization. An answer to
these questions will lead to a better understanding of the role of host cell response in the antitumor effect of PDT and how the immune response can potentiate antitumor immunity. Recently, it has been demonstrated in a BALB/c mouse model that PDT delivered to normal and
tumor tissue in vivo causes marked changes in the expression of IL-6 and IL-10 but not of TNF-
, suggesting that the general inflammatory response to PDT may be mediated by IL-6 (66). Because cytokine genes are controlled by NF-
B and by other transcription factors such as AP-1, c-EBP, CREB, etc., the differential regulation of
cytokine genes by PDT could be due to the better inducibility of
several transcription factors as opposed to others which could be
weakly or not at all regulated by PDT. For example, Photofrin has been
shown to lead to a strong and prolonged activation of c-Jun and c-Fos
(67), demonstrating that genes having both NF-
B-and AP-1-responsive
elements in their promoter are prone to be up-regulated by PDT.
This paper has shown that NF-
B can be considered as a main
transcription factor activated by PDT. Mechanism of PPME-mediated NF-
B activation is rather unique because it resembles the response elicited by IL-1 but differs from the response initiated by ROS generation. Membrane localization of PPME is responsible for this peculiar response; the effect on IL-1 receptor and the rapid NF-
B activation is due to PPME in the cytoplasmic membrane, but the slow and
sustained NF-
B activation is likely explained by lysosomal PPME
localization giving rise to the production of ceramide by activation of
the acidic SMase. As many hydrophobic photosensitizers can target the
lysosomal membrane (36), it could be postulated that ceramide
production could constitute a general response to PDT. Recently,
ceramide has been shown to be involved in the PDT-mediated apoptosis of
mouse lymphoma cells (68), demonstrating the importance of lipid second
messengers both in gene activation and in apoptosis. Since NF-
B has
recently been shown to be involved in the control of apoptosis induced
by cytokines (68-71) and DNA-damaging drugs (72, 73), the data
reported in this paper will add to the understanding of how PPME
promotes apoptosis in tumor cells.
 |
ACKNOWLEDGEMENT |
We thank A-C. Helin for the MUT4 cell line.
 |
FOOTNOTES |
*
This work was supported in part by grants from the Belgian
National Fund for Scientific Research (NFSR, Brussels, Belgium), from
the Concerted Action Program (University of Liège), and from
Télévie (NFSR, Brussels, Belgium).The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
§
Research fellow from the Fonds de Recherche Industrielle et
Agricole (Brussels, Belgium).
Supported by the European Community (Biotech Program).
§§
Research associate from the Belgian National Fund for Scientific Research.
¶¶
Research director from the Belgian National Fund for
Scientific Research. To whom correspondence should be addressed:
Laboratory of Virology, Institute of Pathology B23, University of
Liège, B-4000 Liège, Belgium. Tel.: 32-4-366.24.42; Fax:
32-4-366.24.33; E-mail: jpiette{at}ulg.ac.be.
The abbreviations used are:
PDT, photodynamic
therapy; PPME, pyropheophorbide-a methyl ester; 1O2, singlet oxygen; ROS, reactive oxygen
species; IL, interleukin; TNF-
, tumor necrosis factor
; IKK, I
B kinase; SMase, sphingomyelinase, FCS, fetal calf serum; DiOC6, 3,3'-dihexylpxacarbocyanine iodide; EMSA, electrophoretic mobility
shift assay.
 |
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