← Back
Bombesin-Targeted Delivery of β-Carboline-Based Ir(III) and Ru(II) Photosensitizers for a Selective Photodynamic Therapy of Prostate Cancer.
Despite advances
in Ir(III)
and Ru(II) photosensitizers
(PSs),
their lack of selectivity for cancer cells has hindered their use
in photodynamic therapy (PDT). We disclose the synthesis and characterization
of two pairs of Ir(III) and Ru(II) polypyridyl complexes bearing two
β-carboline ligands (N^N’) functionalized with −COOMe
( L1 ) or −COOH ( L2 ), resulting in
PSs of formulas [Ir(C^N) 2 (N^N’)]Cl ( Ir-Me: C^N = ppy, N^N’ = L1 ; Ir-H: C^N
= ppy, N^N’ = L2 ) and [Ru(N^N) 2 (N^N’)](Cl) 2 ( Ru-Me: N^N = bpy, N^N’ = L1; Ru-H: N^N = bpy, N^N’ = L2 ). To enhance their selectivity
toward cancer cells, Ir-H and Ru-H were
coupled to a bombesin derivative ( BN3 ), resulting in
the metallopeptides Ir-BN and Ru-BN . Ir(III)
complexes showed higher anticancer activity than their Ru(II) counterparts,
particularly upon blue light irradiation, but lacked cancer cell selectivity.
In contrast, Ir-BN and Ru-BN exhibited selective
photocytoxicity against prostate cancer cells, with a lower effect
against nonmalignant fibroblasts. All compounds generated ROS and
induced severe mitochondrial toxicity upon photoactivation, leading
to apoptosis. Additionally, the ability of Ir-Me to oxidize
NADH was demonstrated, suggesting a mechanism for mitochondrial damage.
Our findings indicated that the conjugation of metal PSs with BN3 creates efficient PDT agents, achieving selectivity through
targeting bombesin receptors and local photoactivation.
## Introduction
Introduction In the last few decades, cancer has become
a leading cause of death
worldwide, with a significant impact on our society. Specifically,
prostate cancer is the second most common cancer and the fifth leading
cause of cancer death in men, with an estimated 1.4 million new cases
and 375000 deaths worldwide by 2020. The main treatment options for
localized prostate cancer are surgery and radiotherapy. For cases
of recurrent or metastatic disease, the most used therapies are androgen
deprivation and chemotherapy. 1 − 3 However, one of the main challenges
in the management of these patients is the development of drug resistance. 4 , 5 New strategies are being developed to overcome these limitations
resulting in an increased attention toward photodynamic therapy (PDT).
This minimal invasive and clinically approved therapy is based on
the use of a photosensitizer (PS) which is electronically excited
under light irradiation to form a short-lived excited state (PS*)
that can react with molecular oxygen to generate reactive oxygen species
(ROS). Ultimately, these ROS damage essential biomolecules and kill
cancer cells, leading to the disappearance of the target tumor. 6 Moreover, because of the spatiotemporal control
on the excitation of the PS, this therapy is highly precise compared
to chemotherapy. 7 Ideally, the PS must
display excellent photostability, low cytotoxicity in dark conditions,
absorption bands in the therapeutic window (600–850 nm), 8 selective accumulation in the tumor tissue, efficient
ROS generation, and fast clearance from the body. 9 , 10 Nonetheless,
clinically approved PSs, such as Photofrin, exhibit limitations and
are far from ideal. 11 As a result, there
is a need to develop new PSs that pave the way toward a wider clinical
use of PDT. During the last few years, there have been significant
advances
in developing novel Ir(III) and Ru(II) polypyridyl PSs. 12 , 13 These complexes display characteristics that make them suitable
for being used in PDT: (1) They display absorption bands in the visible
region; (2) the presence of a heavy metal center allows high spin–orbit
coupling constants leading to a fast and efficient population of the
triplet excited states, which results in longer lifetimes and higher
singlet oxygen quantum yields; 14 (3) they
usually display high photostability; 15 (4)
their photophysical and biological properties can be easily modulated
through the functionalization or modification of the ligands; 12 , 13 (5) they can generate different ROS, and some of them are active
in hypoxic conditions; 16 , 17 (6) in the excited state, they
can act as strong oxidants or reductants reacting with a wide variety
of substrates. 18 , 19 More specifically, β-carboline-based
Ir(III) and Ru(II) complexes are being studied as potential PDT agents
with promising results, although they display high or moderate cytotoxicity
toward nonmalignant cells in dark conditions. 20 − 22 Hence, it is
important to develop strategies to specifically deliver these complexes
to tumor cells. Targeted delivery systems not only minimize the impact
of the anticancer agents on normal tissues but also optimize their
accumulation and activity at the tumor site, lowering the PS doses
and the number of light exposure cycles, which in turn limits the
patient photosensitivity. One successful strategy is the conjugation
of complexes to carrier peptides, whose receptors are overexpressed
in cancer cells. 23 , 24 Peptides offer several advantages
over other tumor-targeting agents, such as proteins and monoclonal
antibodies, highlighting their small size, high ability to penetrate
tumors, and good biocompatibility. Furthermore, they can be synthesized
and modified in a relatively simple manner. The use of carrier peptides
for targeted drug delivery has been shown to be effective in prostate
cancer, where a high percentage of primary tumors and bone metastases
have been found to overexpress the gastrin-releasing peptide receptor
(GRPR). 25 − 28 GRPR is also overexpressed in other human cancers, including breast,
colorectal, lung, or pancreatic, while it is poorly expressed in healthy
tissues, making it an attractive target for selective cancer treatment. 29 , 30 , 31 One of the natural ligands
of GRPR in humans is the gastrin-releasing
peptide (GRP), a neuropeptide that regulates multiple physiological
functions. Additionally, GRP acts as a mitogen and proangiogenic factor
in different cancers. 30 Bombesin (BN) is
a 14-amino acid peptide (Pyr-Gln-Arg-Leu-Gly-Asn-Gln-Trp-Ala-Val-Gly-His-Leu-Met-NH 2 ) that shares a homologous seven-amino acid C-terminal region
with GRP. Due to its high affinity for GRPR, BN has been demonstrated
to be an effective carrier peptide for targeted delivery of drugs
and diagnostic agents to GRPR-overexpressing tumors. 28 , 32 , 33 This strategy was initially employed
in nuclear medicine to deliver 99m Tc complexes to cancer
cells. 34 − 36 More recently, different PSs have been conjugated
to BN analogues in an effort to direct their photodynamic activity
against cancer cells, while minimizing off-target effects on healthy
tissues. 37 − 42 In a previous work, we synthesized a series of BN derivatives with
the aim of identifying the most effective sequence for delivering
Pt(II) and Ru(II) complexes to prostate cancer cells. Among the peptides
developed, Gln-Arg-Leu-Gly-Asn-Gln-Trp-Ala-Val-Gly-His-Leu-Leu-NH 2 ( BN3 ) exhibited the most effective tumor-targeting
properties, and the resulting BN3 -derived metallopeptides
displayed high anticancer activity and reduced toxicity toward nonmalignant
fibroblasts. 38 Inspired by our previous
works, and considering the need to develop
new metal-based PSs, we selected BN3 as a tumor-homing
peptide with the objective of improving the selectivity and biocompatibility
of our new β-carboline-based Ir(III) and Ru(II) PSs. Thus, we
aimed to endow the resulting metallopeptides with two levels of selectivity
in their anticancer action, the first derived from the targeting properties
of BN3 and the second associated with the photoactivation
ability of the metal fragments. In this study, we disclose the
conjugation of different β-carboline-based
Ir(III) and Ru(II) PSs to BN3 and the evaluation of the
effect of this peptide on the selectivity and photocytotoxicity of
the conjugated complexes. Particular attention is paid to the influence
of the metal fragment on the anticancer activity of the metallopeptides.
## Results
and Discussion
Results
and Discussion Synthesis of Ligands and Complexes Two pairs of Ir
and Ru complexes with two different N^N’ ligands ( L1 and L2 ) and general formulas [Ir(C^N) 2 (N^N’)]Cl
( Ir-Me: C^N = ppy, N^N’ = L1 ; Ir-H: C^N = ppy, N^N’ = L2 ) and [Ru(N^N) 2 (N^N’)](Cl) 2 ( Ru-Me: N^N =
bpy, N^N’ = L1; Ru-H: N^N = bpy, N^N’ = L2 ) have been prepared aiming to evaluate the effect of the
metal fragment on their photophysical and biological properties ( Schemes 1 and 2 ). Moreover, complexes Ir-H and Ru-H have been prepared with a −COOH group to allow their conjugation
to the bombesin derivative BN3 . The ancillary ligands
( L1 - L2 ) were synthesized in several steps
according to Scheme 1 . First, we reacted tryptamine and 5-bromo-2-pyridinecarboxaldehyde
to obtain pyridyl-β-carboline A , following a method
previously described for similar compounds. 43 Then, the pyrrolic N–H group was substituted with a methyl
group to obtain derivative B , employing NaH as the base
and MeI as the methylating agent. 22 Subsequently,
a Heck vinylation protocol was performed on the bromopyridyl ring
of B , using methyl acrylate to obtain L1 . Furthermore, the ester group was hydrolyzed in alkaline media and
neutralized with HCl to obtain L2 , bearing a −COOH
group. Scheme 1 Synthetic Route for the Synthesis of L1 and L2 Scheme 2 Synthesis and Molecular Structure
of Complexes Ir-Me , Ir-H , Ru-Me , and Ru-H The Ir(III) complexes of general
formula rac -[Ir(ppy) 2 (N^N’)]Cl
(ppy = 2-phenylpyridinate
and N^N’
= L1 or L2 ) were prepared through a bridge
splitting reaction between the dimeric Ir(III) precursor rac -[Ir(μ-Cl)(ppy)] 2 and the corresponding β-carboline-based
ligand ( L1 or L2 , using a molar ratio 1:2,
dimer:ligand) to obtain Ir-Me and Ir-H ,
respectively ( Scheme 2 ). Ru(II) polypyridyl complexes of general formula rac -[Ru(bpy) 2 (N^N’)](Cl) 2 (bpy = 2,2′-bipyridine
and N^N’ = L1 or L2 ) were prepared
by heating the Ru(II) precursor, rac- [Ru(bpy) 2 Cl 2 ], with L1 or L2 (1:1
molar ratio), to obtain Ru-Me and Ru-H ,
respectively ( Scheme 2 ). The metal complexes were obtained as racemic mixtures (Λ,Δ)
of the respective chloride salts. Ligands and complexes were fully
characterized by nuclear magnetic resonance (NMR), mass spectrometry,
and elemental analysis (see Figures S1–S32 ). In particular, 1 H NMR spectra of all the complexes
were recorded in DMSO- d 6 at 25 °C.
The spectra displayed two sets of resonances for the two nonequivalent
ppy/bpy ligands owing to the lack of symmetry shown by these derivatives.
All of them showed a singlet around 3.93 ppm attributed to the N-Me
group, which is shifted to lower field compared to the corresponding
singlet of the free ligand (3.60 ppm). The high-resolution electrospray
ionization mass spectrometry (HR ESI(+) MS) spectra exhibited peaks
with mass/charge ratios and isotopic distributions fully compatible
with those calculated for either the monocationic Ir complexes or
the dicationic Ru complexes. The analytical data obtained from elemental
analysis are in good agreement with expected values. Synthesis of
the Metallopeptides Metallopeptides Ir-BN and Ru-BN incorporating the Ir(III) or
the Ru(II) complex at the N-terminus of the peptide BN3 were synthesized on solid phase following a standard 9-fluorenylmethoxycarbonyl
(Fmoc)/ tert -butyl ( t Bu) strategy
( Scheme 3 ). 38 An aminomethyl ChemMatrix resin was used as
solid support to which the Fmoc-Rink-amide linker was incorporated
leading to C-terminal amidated peptides. The attachment of this linker
to the ChemMatrix resin was accomplished using N,N -diisopropylcarbodiimide (DIC) as coupling reagent, ethyl (hydroxyimino)cyanoacetate
(Oxyma) as additive, and dimethylformamide (DMF) as solvent. The
elongation of the peptide sequence was performed through sequential
steps of Fmoc group removal with piperidine/DMF (3:7) and coupling
of the corresponding amino acids using DIC and Oxyma in DMF. A Kaiser
test was performed to confirm the completion of the coupling reactions. 44 Once the peptide sequence was completed, the
N-terminal Fmoc group was removed, and the corresponding metal complex Ir-H or Ru-H was incorporated in the presence
of DIC and Oxyma in DMSO. Next, the resulting metallopeptides Ir-BN and Ru-BN were cleaved from the support
by acidolytic treatment with trifluoroacetic acid (TFA)/H 2 O/triisopropylsilane (TIS), purified by reversed-phase column chromatography,
analyzed by HPLC, and characterized by mass spectrometry. Ir-BN and Ru-BN were obtained in >99% HPLC purity (see Figures S33–44 ). Scheme 3 Solid-Phase Synthesis
of Metallopeptides Ir-BN and Ru-BN X-ray Diffraction The crystal structure
of [ Ir-Me ]PF 6 was resolved by single-crystal
X-ray diffraction.
A suitable single crystal was obtained by slow evaporation of an acetonitrile
solution of [ Ir-Me ]Cl in the presence of NH 4 PF 6 . The complex crystallizes in the monoclinic C 2/ c space group. Due to helical chirality,
the unit cell contains two pairs of enantiomers (Δ,Λ)
( Figure S45 ). The ORTEP diagram for the
molecular structure of Λ-[Ir-Me] + is shown in Figure 1 . Selected bond distances and angles are compiled in Table 1 , and important crystallographic
parameters are gathered in Table S3 . Figure 1 ORTEP diagram
for the molecular structure of Λ-[Ir-Me] + obtained by single-crystal X-ray diffraction.
Thermal ellipsoids are shown at the 30% probability level. The respective
Δ enantiomer, H atoms, and the PF 6 – counterion have been omitted for the sake of clarity. Table 1 Selected Bond Distances (Å) and
Coordination Angles for rac -[Ir-Me] + rac -[Ir-Me] + Distances rac -[Ir-Me] + Angles (deg) Ir(1)–N(1) 2.153 (4) N(1)Ir(1)N(2) 76.79 (14) Ir(1)–N(2) 2.159 (3) C(32)Ir(1)N(4) 80.62 (19) Ir(1)–N(4) 2.045 (4) C(43)Ir(1)N(5) 80.3 (2) Ir(1)–C(32) 2.021 (5) Ir(1)–N(5) 2.046 (4) Ir(1)–C(43) 2.021 (4) The molecular structure of [Ir-Me] + exhibited the expected pseudo-octahedral
geometry
with the
predictable trans -N,N and cis -C,C
disposition for the two cyclometalated ligands (ppy). The Ir–N
bond distances for the N^N’ ligand (2.153(4), 2.159(3) Å)
were longer than those determined for the ppy ligands (2.046(4), 2.045(45)
Å), due to the strong trans influence attributed
to the metal-bonded phenyl rings. Both Ir–C bond lengths were
nearly identical (2.021(5) and 2.021(5)) and within the expected range
for this kind of complexes. The bite angles of the chelate rings were
also standard, that is, 76.79° for L1 and 80.62°,
80.30° for the ppy ligands. The N^N’ ligand displayed
a high torsion angle (−18.55°) in comparison with the
ppy ligands (2.07° and 3.04°) revealing a lower degree of
coplanarity, due to the high steric hindrance between the N-Me group
and the pyridyl ring. Furthermore, the crystal structure exhibited
different weak hydrogen-bonding interactions between fluorine atoms
from the counterion and different hydrogen donor groups of the iridium
complex. Also, a weak hydrogen-bonding interaction between an oxygen
atom (O1) from L1 and one hydrogen (H23) from a ppy ligand
of a neighboring molecule was observed ( Figure S46 and Table S4 ). UV–Vis Spectroscopy The UV–vis
absorption
spectra of Ir-H , Ir-Me , Ru-H , and Ru-Me ( Figure 2 ) were recorded at room temperature for H 2 O:DMSO (99:1) solutions (10 –5 M). All the complexes
displayed strong absorption bands with maxima between 250 and 350
nm that were attributed to spin-permitted ligand centered transitions
( 1 LC, π → π*). In the visible region,
Ir(III) complexes showed a main absorption band with maxima around
420 nm that extends up to 550 nm and corresponds to mixed spin-allowed
and spin-forbidden metal to ligand charge transfer ( 1 MLCT
and 3 MLCT) and ligand to ligand charge transfer (LLCT)
transitions. By contrast, the Ru(II) complexes showed two absorption
bands in the visible region, with maxima at 430 and 500 nm and a tail
extended up to 600 nm. These bands were also ascribed to 1 MLCT and 3 MLCT and LLCT transitions. Therefore, the Ru(II)
derivatives exhibited red-shifted bands compared to their Ir(III)
analogues. Moreover, complexes with the methyl ester group Ir-Me / Ru-Me showed absorption bands slightly red-shifted
relative to the complexes with −COOH groups, Ir-H / Ru-H . Figure 2 Overlaid absorbance spectra of Ir(III)/Ru(II) complexes
in H 2 O:DMSO (99:1, v:v) (10 –5 M) at room
temperature.
Inset: zoom of the region between 500 and 700 nm. Emission and Photophysical Properties The emission
spectra of the metal complexes in deoxygenated solutions of H 2 O:DMSO (99:1, v:v, 10 –5 M) were recorded
at 25 °C, using λ ex of 405 nm for Ir-Me / Ir-H and λ ex of 450 nm for Ru-Me / Ru-H ( Figure 3 ). All the complexes displayed a single broad emission band
with maxima between 651 and 720 nm and large Stokes shifts (λ em – λ ex > 230 nm), which corroborated
the phosphorescent nature of the emission. The Ru(II) complexes exhibited
higher emission intensities, and their respective maxima are red-shifted
compared to those of the Ir(III) analogues. Moreover, Ir-Me / Ru-Me featured lower emission intensities compared
to Ir-H / Ru-H , respectively. Figure 3 Overlaid emission spectra
of Ir(III)/Ru(II) complexes
in deoxygenated H 2 O:DMSO (99:1, v:v, 10 –5 M) at 25 °C. The excited state lifetimes
(τ) and the photoluminescence
quantum yields were also determined for the new metal complexes ( Table 2 ). The Ir PSs displayed
two-component τ values of 2 ns [20%]/93 ns [80%] and 23 ns [51%]/145
ns [49%] for Ir-Me and Ir-H , respectively.
By contrast, the Ru(II) complexes exhibited one-component τ
values of 74 and 186 ns for Ru-Me and Ru-H , respectively. These values compare well with those of similar complexes
and are compatible with a triplet nature for the emissive states.
The photoluminescence quantum yields (Φ PL ) of Ir(III)/Ru(II)
were very low with values ranging from 1.1 to 4.13%. Table 2 Photophysical Properties of the Metal
Complexes in H 2 O:DMSO (99:1, v:v) at 25 °C under a
Nitrogen Atmosphere Compound λ ex (nm) λ em (nm) τ (ns) [contribution (%)] Φ PL (%) Ir-Me 405 676 2 [20] 93 [80] 3.50 Ir-H 405 651 23 [51] 145
[49] 4.13 Ru-Me 450 720 74 1.10 Ru-H 450 688 186 3.13 Determination of p K a Ir-H and Ru-H are susceptible to deprotonation
in aqueous media due to the presence of the acidic −COOH group.
In fact, the actual protonation state of these complexes at the different
physiological pH values defines their global charge and correspondingly
influences their solubility, their photophysical properties, and their
cellular uptake. 45 − 47 Thus, the p K a values
of Ir-H and Ru-H in their excited states
were experimentally determined by monitoring the variations of their
emission intensity at specific wavelengths versus different pH values
and fitting the data ( I em,λ /pH)
to a sigmoidal equation. In good agreement with similar complexes
reported in the literature, 45 , 46 there was a gradual
decrease in the emission intensity of Ir-H and Ru-H with decreasing pH ( Figures 4 and S50 ). The
p K a values for Ir-H and Ru-H in their excited states were 3.81 and 4.51, respectively.
These values can be considered as an acceptable approximation to the
values of the respective ground states. Therefore, we concluded that Ir-H and Ru-H adopt their deprotonated zwitterionic
forms in the whole range of physiological pHs. This means that Ir-H assumes a zwitterionic neutral form, while Ru-H adopts a zwitterionic monocationic structure. In both cases, there
is charge separation, and as a result, we predict that the cellular
uptake ability of these complexes could be reduced, in comparison
to that of their relatives Ir-Me and Ru-Me . Figure 4 p K a determination. (A) Overlaid emission
spectra of Ir-H in H 2 O:DMSO (99:1, v:v) (10 –5 M) at different pH values ranging from pH 7 to 2
(HCl titration). (B) Plot of the emission intensity of Ir-H at λ 651 nm in H 2 O:DMSO (99:1, v:v) (10 –5 M) as a function of pH (2–7) at 25 °C. (C) Scheme showing
the acid–base equilibrium for Ir-H . Photostability of Ir(III)/Ru(II) Complexes Photostability
is a desirable feature for PDT agents, since otherwise, their efficiency
as PSs can be decreased owing to so-called photobleaching. Thus, the
photostability of aerated solutions (1.5 × 10 –2 M, DMSO- d 6 :D 2 O, 3:2, v:v)
of all our complexes was evaluated by 1 H NMR. The solutions
were exposed to blue light irradiation (λ irr = 460
nm, 24 W), and the evolution of the respective samples was monitored
by 1 H NMR at different times (0, 6, and 24 h) at room temperature.
To our delight, no symptoms of photodegradation were observed, confirming
that all the complexes exhibited outstanding photostability under
the aforementioned conditions ( Figures 5 and S47–S49 ). Figure 5 Evolution of
the aromatic region of the 1 H NMR spectra
of Ru-H (1.5 × 10 –2 M) in DMSO- d 6 :D 2 O (3:2, v:v) under blue light
irradiation (LED, λ = 460 nm, 24 W) at different times. Photocatalytic Generation of Singlet Oxygen Singlet
oxygen is thought to be the main cytotoxic species in type II PDT
processes. Therefore, we determined the ability of Ir-Me and Ru-Me to generate 1 O 2 under
photocatalytic conditions, as illustrative examples of our PSs. As
shown in Figures 6 and S51 , we monitored by means of UV–vis spectroscopy
the oxidation of 9,10-anthracenediyl bis(methylene)dimalonic acid
(ABDA, 8x 10 –5 M) in the presence of atmospheric
oxygen, using Ir-Me (10 –5 M) as the
PS and H 2 O:DMSO (1:1) as the solvent system under blue
light exposure (λ ir = 460 nm). ABDA is a very specific
probe for 1 O 2 , since it reacts selectively with
the in situ produced 1 O 2 to generate the respective
endoperoxide, which is characterized by a loss of π-extended
aromaticity and the disappearance of several absorption bands in the
UV–vis region (i.e., 379 nm). Moreover, the singlet oxygen
quantum yield values ( Φ Δ ) of Ir-Me and Ru-Me were determined using rose bengal
( Φ Δ = 0.75) as the reference.
As expected, both Ir-Me and Ru-Me exhibited
photocatalytic activity in the generation of 1 O 2 , but their respective Φ Δ were
low, 4 and 9%, in agreement with its moderate excited state lifetimes
(93 and 74 ns). A control experiment in the absence of PS was also
performed resulting in a negligible conversion. Figure 6 Photobleaching of ABDA
(8 × 10 –5 M) in the
presence of Ir-Me (10 –5 M) using H 2 O:DMSO (1:1) under blue light irradiation (460 nm, 24 W) during
100 min at room temperature. Photocatalytic Oxidation of NADH NADH is an enzymatic
cofactor playing a main role as an electron donor in the mitochondrial
electron transport chain. Hence, it is considered a potential molecular
target for those anticancer drugs that cause oxidative stress in mitochondria.
Indeed, several research groups have recently established a relationship
between the photocatalytic oxidation of NADH to NAD + , the
depolarization of the mitochondrial membrane and apoptotic cell death. 48 , 49 Aiming to prove this hypothesis for Ir-Me , as
a model PS, we monitored by UV–vis spectroscopy the photocatalytic
oxidation of an aerated solution of NADH (0.1 mM) in the presence
of Ir-Me (5 μM) using H 2 O:DMSO (99:1)
under blue light irradiation (λ ir = 460 nm) for 15
min. As a result, we observed the decrease of the band due to NADH
at 338 nm ( Figure 7 ). This evolution is compatible with the photocatalyzed formation
of NAD + . Two control experiments in the absence of either Ir-Me ( Figure S52 ) or light ( Figure S53 ) corroborated the photocatalytic character
of this transformation. Figure 7 Evolution of the UV–vis spectra during
the photocatalytic
oxidation of NADH (100 μM) in the presence of Ir-Me (5 μM) in aerated H 2 O:DMSO (99:1) under blue light
irradiation (460 nm, 24 W) at room temperature (15 min). Effect on Cell Viability The effect of Ir-H , Ir-Me , Ru-H , and Ru-Me on
cell viability was evaluated in the A549 lung cancer and PC-3 prostate
cancer cell lines by MTT assays. PC-3 cells were specifically chosen
due to their high expression levels of GRPR on their surface, allowing
for further evaluation of the activity of the BN3 -derived
metallopeptides Ir-BN and Ru-BN . 38 , 51 In contrast, A549 cells express low levels of GRPR. 52 MRC-5 fibroblasts were used as a nonmalignant cell model. 53 Table 3 reports the half maximal inhibitory concentration (IC 50 ) values obtained after treating the cells either under dark
conditions or upon exposure to blue light at a dose of 24.1 J cm –2 . The photocytotoxicity index (PI), defined as the
IC 50,dark /IC 50,light , is provided for each compound. Table 3 Photocytotoxic Effects of Complexes Ir-H , Ir-Me , Ru-H , and Ru-Me and
of Metallopeptides Ir-BN and Ru-BN on Different
Cell Lines in the Dark and after Blue Light Irradiation IC 50 a (μM) A549 PC-3 MRC-5 Dark Light PI b Dark Light PI b Dark Light PI b Ir-H 35.0 ± 5.4 3.32 ± 1.1 10.5 32.2 ± 1.9 9.23 ± 0.63 3.5 39.1 ± 2.4 6.46 ± 0.801 6.1 Ir-Me 0.471 ± 0.031 0.0334 ± 0.0068 14.2 1.17 ± 0.29 0.0723 ± 0.031 16.2 1.12 ± 0.31 0.0313 ± 0.019 35.8 Ru-H 67.8 ± 6.7 56.7 ± 1.6 1.2 70.4 ± 2.7 34.8 ± 4.3 2.0 53.9 ± 5.4 43.5 ± 4.5 1.2 Ru-Me 74.7 ± 2.03 33.8 ± 5.2 2.2 76.8 ± 2.1 48.6 ± 0.5 1.6 62.5 ± 2.6 43.7 ± 7.2 1.4 Ir-BN 68.3 ± 15.8 12.6 ± 1.3 5.4 35.3 ± 3.8 8.51 ± 0.8 4.1 80.9 ± 1.3 28.7 ± 5.9 2.8 Ru-BN 49.5 ± 11.9 23.6 ± 11.2 2.1 41.3 ± 1.7 4.11 ± 1.2 10.1 >100 >100 n.d. Cisplatin 5.99 ± 1.2 n.d. n.d. 5.55 ± 0.97 n.d. n.d. 5.34 ± 0.38 n.d. n.d. a Cells were incubated with the
compounds for 4 h at 37°C and then kept in the dark or exposed
to blue light irradiation for 1 h (460 nm, 24.1 J cm –2 ). Cell viability was assessed after 48 h of treatment by MTT assays.
Data represent the mean ± SD of at least three independent experiments,
each performed in triplicate. b Photocytotoxic index (PI) = IC 50, dark /IC 50, light . n.d.: not determined. The Ir(III) complexes exhibited a greater effect on
cell viability
than their Ru(II) counterparts in all cell lines. Among them, Ir-Me displayed the highest anticancer activity, with IC 50 values of 0.471 μM in A549 cells and 1.17 μM
in PC-3 cells in dark conditions, which were lower than those of cisplatin
under the same experimental conditions. Conversely, Ir-H displayed IC 50,dark values above 30 μM, revealing
that the esterification of the β-carboline ligand with a methyl
group significantly enhanced the biological activity of the resulting Ir-Me complex. Upon blue light irradiation, the cytotoxicity
of Ir-Me increased by 14.2-fold in A549 cells and 16.2-fold
in PC-3 cells, resulting in IC 50,light values of 33.4 nM
and 72.3 nM, respectively. The anticancer activity of Ir-H also increased after light exposure by 10.5-fold in A549 cells and
3.49-fold in PC-3 cells, with IC 50,light values decreasing
to the low micromolar range. It should be noted that Ir-H and Ir-Me displayed similar IC 50 values
in MRC-5 fibroblasts as in cancer cells, both in the dark and upon
irradiation, evidencing their nondiscriminatory effects between nonmalignant
and cancer cells. Regarding the Ru(II) complexes, Ru-H and Ru-Me showed very moderate anticancer activity
in dark conditions, which
was not significantly enhanced upon exposure to blue light despite
having higher light absorption at 460 nm than the Ir (III) complexes
( Figure 2 ). Moreover,
they exhibited similar toxicity levels against MRC-5 fibroblasts,
as observed with the Ir(III) complexes. The cytotoxic characterization
of Ir-H , Ir-Me , Ru - H , and Ru-Me was further
complemented by hemolysis assays, which determined whether they could
induce the rupture of red blood cell membranes, leading to the release
of hemoglobin. 54 Both Ir(III) and Ru(II)
complexes exhibited hemolysis levels of less than 1% both in the absence
and in the presence of blue light irradiation. This finding indicates
that the complexes are nontoxic to red blood cells and have good blood
compatibility for future clinical applications. Finally, the
photocytotoxic activity of the BN3 -derived
metallopeptides, Ir-BN and Ru-BN , was evaluated.
In PC-3 cells, Ir-BN exhibited similar activity relative
to the precursor complex, Ir-H , both in the dark and
upon activation with blue light. However, in MRC-5 fibroblasts, the
IC 50,dark and IC 50,light values for Ir-BN were 2.1- and 4.4-fold higher, respectively, than those of Ir-H ( Table 3 ). It should be noted that Ir-BN exhibited 2.3-fold
and 1.9-fold higher cytotoxicity in PC-3 cells compared to MRC-5 fibroblasts
and A549 cancer cells with low GRPR expression, respectively. After
exposure to blue light, the selectivity for PC-3 cells increased to
3.4-fold with respect to MRC-5 fibroblasts. Regarding Ru-BN , the IC 50,dark and IC 50,light values in PC-3
cells were 1.7 and 8.5 times lower, respectively, than those of the
precursor Ru-H complex. In terms of selectivity, the
activity of Ru-BN in PC-3 cells was 1.2- and 5.7-fold
higher than in A549 cells under dark and irradiated conditions, respectively.
Importantly, Ru-BN exhibited no cytotoxicity in MRC-5
fibroblasts even at concentrations as high as 100 μM, in both
dark and irradiated conditions. The photocytotoxic effects of Ru-H , Ru-Me , and Ru-BN were also
evaluated following photoactivation
with red light (λ ir = 655 nm), prompted by the red-shifted
band observed in the absorption spectra of the Ru (II) complexes ( Figure 2 ). Notably, red light
offers superior tissue penetration compared to blue light, potentially
enabling treatment of deeper tumors. 55 As
shown in Table 4 , red
light had a low impact on the activity of both Ru(II) complexes, with
PIs below 1.3 in both PC-3 and MRC-5 cells. In contrast, Ru-BN displayed markedly higher photocytotoxic activity against PC-3 cells,
with a PI of 7.3, while it displayed no cytotoxicity toward MRC-5
fibroblasts. Table 4 Photocytotoxic Effects of Ru-H , Ru-Me , and Ru-BN after Red Light Irradiation a PC-3 MRC-5 IC 50,red light (μM) PI IC 50,red light (μM) PI Ru-H 54.3 ± 4.7 1.3 62.3 ± 7.1 0.9 Ru-Me 63.1 ± 5.8 1.2 66.6 ± 4.8 0.9 Ru-BN 5.66 ± 1.2 7.3 >100 -- a Cells were incubated with the
compounds for 4 h at 37 °C and then kept in the dark or exposed
to red light irradiation for 1 h (655 nm, 24.1 J cm –2 ). Cell viability was assessed after 48 h of treatment by MTT assays.
Data represent the mean ± SD of at least three independent experiments,
each performed in triplicate. PI: photocytotoxic index = IC 50, dark /IC 50,red light . To examine the role of BN3 as a carrier
peptide for
specifically targeting PC-3 cells, the influence of blocking the bombesin
receptors on the anticancer efficacy of the metallopeptides was assessed.
PC-3 cells were treated with Ir-BN and Ru-BN at concentrations near their IC 50,light (10 and 5 μM,
respectively) in the presence of increasing bombesin concentrations
(0, 10, 50, and 100 μM) to competitively inhibit GRPR binding.
Following irradiation with blue light, the viability of the cells
was evaluated. The photocytotoxic activity of both Ir-BN and Ru-BN was significantly attenuated in a bombesin
concentration-dependent manner, with approximately a 25% reduction
in the presence of bombesin at 10 μM and a more pronounced inhibition
of approximately 35% at 50 and 100 μM ( Figure S54 ). These results strongly suggest that the cytotoxic effect
of Ir-BN and Ru-BN is mediated, at least
in part, by their interaction with GRPR. Overall, these findings
support the efficacy of BN3 conjugation in enhancing
the selectivity of PSs toward cancer cells
overexpressing GRPR. This is also evidenced by photoselectivity indexes
(IC 50,dark in nonmalignant cells/IC 50,light in
GRPR overexpressing cancer cells) of 9.5 for Ir-BN after
activation with blue light and exceeding 24.3 and 17.6 for Ru-BN after activation with blue and red light, respectively. Inhibition
of Colony Formation The effect of the metallopeptides Ir-BN and Ru-BN as well as of the complexes Ir-H and Ru-H on the viability of PC-3 cells
was next investigated through clonogenic assays, which determine the
fraction of cells that survive treatment and retain the ability to
generate new colonies. 56 This is a crucial
feature of metastatic cancer cells, which need to proliferate in distant
tissues to create secondary tumors. For this purpose, PC-3 cells were
treated with the compounds at an equimolar concentration of 5 μM
with or without photoactivation with blue light. Ten days later, the
number of cells that survived and were able to grow and form colonies
was assessed in comparison to control cells. Cells treated with cisplatin
were used as a positive control. None of the compounds inhibited the
clonogenic activity of the cells in the absence of light irradiation
( Figure 8 ). However,
upon blue light irradiation, the number of colonies decreased by 27.3%,
34.7%, and 50.4% in cells exposed to Ir-H , Ir-BN , and Ru-BN , respectively. No effect of Ru-H on the clonogenic capability of the cells was observed at 5 μM,
which is consistent with its high IC 50,light value ( Table 3 ). Figure 8 Clonogenic assay. PC-3
cells were treated with the indicated compounds
at 5 μM in the dark and with blue light irradiation. Control
cells were treated with the medium alone. Cisplatin was used as a
positive control. Cells were incubated for 10 days to allow colony
formation. (A) Images of the colonies. (B) Bar charts representing
the percentage of colonies versus control cells after each treatment
(mean ± SD of 3 experiments). ** p < 0.01;
*** p < 0.001 versus control cells. Intracellular ROS Generation The photocytotoxic effects
of the complexes and metallopeptides were further elucidated by evaluating
their ability to photogenerate ROS at the cellular level. PC-3 cells
were treated with the compounds at the respective IC 50,light and irradiated with blue light. Subsequently, ROS generation was
measured using the 2′,7′-dichlorodihydrofluorescein
diacetate (H 2 DCFDA) probe, which diffuses into cells and
is oxidized by ROS to produce a green fluorescent signal. As illustrated
in Figure 9 A, in all
cases, ROS levels were significantly increased by more than 2-fold
in comparison to the control cells. Ir-Me exhibited the
most potent ROS-generating activity, with a 4.5-fold increase. It
is worth noting that, despite Ir-Me exhibiting relatively
low efficiency as a photocatalyst for the generation of 1 O 2 , its effective cellular uptake could counterbalance
this limitation (see Figure 10 ), enabling effective ROS generation inside the cells. Nevertheless,
to ascertain the potential contributions of other ROS to the photocytotoxic
activity, the capacity of the compounds to undergo type I PDT processes,
resulting in the production of superoxide anions (O 2 •– ), was examined using a specific fluorescent
probe that emits an orange signal upon interaction with this radical. Ir-H , Ir-Me , and Ir-BN induced the
most significant increase in O 2 •– levels, with fold changes of 2.4, 3.5, and 3.4, respectively, compared
to control cells. In contrast, the Ru complexes exhibited lower increments
( Figure 9 B). These
findings highlight the diverse mechanisms of ROS generation employed
by these compounds, in particular the Ir complexes, which may contribute
to their overall photocytotoxic efficacy. Figure 9 Intracellular ROS generation.
PC-3 cells were treated with the
indicated complexes and metallopeptides at their respective IC 50,light for 4 h, followed by blue light irradiation for 1
h (460 nm, 24.1 J cm –2 ). The elevation of general
ROS (A) and superoxide anion (B) levels was determined with specific
probes by flow cytometry. Bars represent the mean fold increase (±standard
deviation) relative to control untreated cells from three independent
experiments. * p < 0.05; ** p < 0.01, *** p < 0.001 compared to control
cells. Figure 10 Cellular internalization. PC-3 cells
were incubated for
4 h with
the indicated compounds at 5 μM, and the Ir (A) or Ru (B) content
per 1 × 10 6 cells was quantified by ICP-MS. Bars represent
the mean value of three replicates ± SD. * p < 0.05; *** p < 0.001 versus Ir-H or Ru-H . Internalization Studies The intracellular accumulation
of the PSs is a crucial requirement to achieve a photocytotoxic effect.
This is because the ROS generated by the PSs have a very limited radius
of action and only operate in the cellular compartment or organelle
in which they are generated. 57 In particular,
the absence of hemolytic activity observed in both Ir(III) and Ru(II)
complexes suggests a mechanism of action that does not involve cell
membrane disruption. This further emphasizes the importance of cellular
uptake for their cytotoxic activity. Additionally, when using tumor-targeting
ligands, such as BN3 , the intracellular uptake of the
metallopeptides depends on the level of expression of the target receptor
on the cell surface and the internalization mechanism, which may differ
from that of the free complex. Therefore, the internalization of complexes Ir-H , Ir-Me , Ru - H ,
and Ru-Me and of metallopeptides Ir-BN and Ru-BN in PC-3 cells was determined by quantifying the intracellular
metal content by inductively coupled plasma mass spectrometry (ICP-MS)
( Figure 10 ). After
4 h treatment with the compounds at 5 μM, the amount of Ir in
cells exposed to Ir-H was 3.1 ± 0.7 ng per million
cells, whereas in the case of its conjugated form Ir-BN , it increased 5.7-fold to 17.9 ± 5.6 ng per million cells.
This result showed that conjugation to BN3 enhances the
cellular uptake of the Ir(III) complex. Notably, Ir-Me exhibited the highest internalization, resulting in 266.4 ±
13.9 ng Ir per million cells, consistent with its superior anticancer
activity. We speculate that the higher ability of Ir-Me to accumulate within the cells is consistent with its lipophilic
character (vide infra). By contrast, Ir-H exhibited a
poor intracellular uptake likely due to the presence of the negatively
charged −COO – group in its deprotonated zwitterionic
form. It should be noted that Ir-Me has a significantly
higher internalization capacity than the Ir-BN conjugate.
This discrepancy can be attributed to the lipophilic nature of Ir-Me , which enables its passive diffusion across the cell
membrane. Consequently, Ir-Me can be more rapidly and
directly internalized by cells. Conversely, Ir-BN entry
involves receptor-mediated endocytosis, a slower, energy-dependent
process that often results in lower intracellular concentrations and
can be influenced by variations in the endocytic pathway activity. In the case of cells treated with the Ru(II) complexes, the intracellular
metal contents were markedly lower, with 0.329 ng per million cells
and 0.326 ng per million for Ru-H and Ru-Me , respectively. We believe that the scarce accumulation of the Ru
derivatives is due to their lower lipophilicity (vide infra), since
in the physiological pH range, Ru-H shows a monocationic
zwitterionic form and Ru-Me a dicationic nature. These
charged states may hinder passive diffusion across the cell membrane.
However, the amount of intracellular Ru significantly increased 14-fold
to 4.46 ng/million cells for Ru-BN . Overall, these
results demonstrate that the conjugation of the
complexes to BN3 not only enhances their selectivity
for tumor cells but also increases the intracellular accumulation
of the conjugated complexes Ir-H and Ru-H , leading to increased cytotoxic activity, particularly for the Ru(II)
derivative. Lipophilicity and Self-Aggregation Studies In order
to obtain a better understanding of the cellular uptake abilities
of these complexes, we carried out experiments to determine their
lipophilicity and self-aggregation properties. First, the n -octanol/PBS partition coefficient values, log P oct/PBS , of the metal PSs were experimentally determined to quantify their
lipophilicity ( Figure S56 ). Indeed, the
shake flask method was employed, and the metal content was measured
in both phases by UV/vis spectroscopy. Thus, we established that both
Ru derivatives exhibited low lipophilicity in agreement with either
the dicationic nature of Ru-Me (log P oct/PBS = −0.43) or the monocationic zwitterionic character of Ru-H (log P oct/PBS = −1.10) at physiological
pH. These results are consistent with the low cellular uptake shown
by these derivatives. By contrast, both Ir derivatives show
higher lipophilicity relative
to their Ru congeners, which is rationally explained as a result of
the monocationic nature of Ir-Me and the neutral zwitterionic
character of Ir-H at physicological pH. Indeed, no significant
differences can be observed in the lipophilicity between Ir-Me (log P oct/PBS = 2.03) and Ir-H (log P oct/PBS = 1.80). Therefore, we speculate that the pronounced
divergent cellular uptake behavior experimentally observed for Ir-H and Ir-Me could be explained as a result
of the electronic repulsion between the negatively charged −COO – group in Ir-H and the negative charge
of the phosphate groups present in the cellular membrane. In other
words, we assume that the presence of the −COO – group in the zwitterionic form of Ir-H at physiological
pH values would hinder its uptake through the cell membrane by passive
diffusion. 58 The possible aggregation
properties (self-assembly behavior) were
investigated for the lipophilic derivative Ir-Me through
the respective analysis of the Lambert–Beer law using UV–vis
spectroscopy in H 2 O:DMSO (99:1). More specifically, the
absorbance of Ir-Me at λ = 417 nm was plotted versus
concentration in the range between 1 and 40 μM. An excellent
linear fitting was obtained confirming that there is no significant
deviation from the Lambert–Beer law ( Figures S57 and S58 ). Therefore, we conclude that Ir-Me does not form aggregates under these conditions. Intracellular
Distribution The intracellular distribution
of the compounds was then investigated by confocal microscopy. Preliminary
flow cytometry experiments revealed that significant intracellular
fluorescence was detectable only in cells exposed to Ir-Me , consistent with the higher intracellular accumulation of this compound
( Figure S55 ). Accordingly, confocal
microscopy images showed a strong red fluorescence signal from Ir-Me inside the cells, as shown in Figure 11 . The complex displayed a high degree of
colocalization with the mitochondrial dye MitoView Green, as evidenced
by the yellow signal in the merged image (Pearson correlation coefficient
(PCC): 0.832). In contrast, a lower degree of overlap was observed
with the LysoTracker Green DND-26 lysosomal dye (PCC: 0.599), indicating
that the complex predominantly accumulates within the mitochondria
and exhibits a lower distribution within the endolysosomal system.
This is compatible with the monocationic and lipophilic nature of Ir-Me and has been previously reported for other mitochondria-targeting
monocationic bis-cyclometalated Ir(III) complexes. 59 − 63 Figure 11 Confocal images of A549 cells after 1 h of incubation
with Ir-Me at 1 μM (red). MitoView Green or LysoTracker
Green
DND-26 (green) were used for colocalization studies with mitochondria
or lysosomes, respectively. Colocalization is shown in yellow in the
merged image. Scale bars represent 20 μm. Mitochondrial Targeted Activity Given the high degree
of accumulation of Ir-Me in mitochondria, the potential
of complexes Ir-H , Ir-Me , Ru - H , and Ru-Me and of metallopeptides Ir-Bn and Ru-BN to induce mitochondrial dysfunction
was next investigated. Mitochondria are the primary source of ATP
and anabolites for cellular metabolism and are involved in the regulation
of key cellular functions, including redox status and cell signaling.
Consequently, they play a crucial role in both cell growth and cell
death regulation. 64 PC-3 cells were incubated
with the compounds at the corresponding IC 50,light for
4 h and subsequently exposed to blue light irradiation for 1 h. Subsequently,
mitochondria were labeled with MitoTracker Red CMXRos, a lipophilic
cationic red fluorescent dye that accumulates within healthy mitochondria
due to their negative mitochondrial membrane potential (MMP). 65 Cell nuclei were counterstained in blue using
Hoechst 33342 to localize the cells. Confocal microscopy images revealed
a strong red fluorescence emission from mitochondria in untreated
control cells ( Figure 12 A). Conversely, exposure of cells to the photoactivated complexes
and metallopeptides led to a notable attenuation of the mitochondrial
red fluorescence, which was particularly evident in the case of the
complex Ir-H and of the metallopeptides Ir-BN and Ru-BN . These results strongly suggest that these
complexes exert their photodynamic activity by inducing mitochondrial
membrane depolarization, a hallmark of mitochondrial dysfunction. 65 Figure 12 Effect on mitochondria function. PC-3 cells were incubated
with
the indicated compounds at the corresponding IC 50,light for 4 h at 37 °C and then exposed to blue light for 1 h. Cells
incubated with the medium alone served as the negative control. (A)
Confocal microscopy images of the cells. Nuclei were stained blue
with Hoechst (λ ex : 350 nm; λ em :
461 nm), and mitochondria were labeled with MitoTracker Red CMXRos
(λ ex : 579 nm; λ em : 599 nm). Insets
show a 3× magnification of the cells. Scale bars represent 50
μm in 1× images and 20 μm in 3× images. (B)
Flow cytometry analysis of the effect of the complexes on mitochondrial
membrane potential. The percentage of cells exhibiting JC-10 green
(λ em : 529 nm) and orange fluorescence (λ em : 590 nm) is represented (mean ± SD of three independent
experiments). A decrease in the percentage of cells emitting orange
fluorescence indicates a loss of MMP. p < 0.05
and ** p < 0.01 compared to control cells. Flow cytometry analysis using the mitochondrial
specific dye JC-10
confirmed this effect. The fluorescence of JC-10 undergoes a reversible
change from green to orange as MMP increases, due to the formation
of JC-10 aggregates within polarized mitochondria. This property allows
for the simultaneous detection of healthy and depolarized mitochondria.
In control cells, orange fluorescence emission corresponding to healthy
mitochondria was detected in 63.0 ± 10.6% of cells ( Figure 12 B). However, treatment
with the photoactivated compounds significantly reduced the population
of cells emitting orange fluorescence to 16.6 ± 9.2%, 21.2 ±
4.9%, and 16.0 ± 9.5% for Ir-H , Ir-Me , and Ir-BN , respectively, and to 28.2 ± 7.8%,
19.7 ± 7.6%, and 23.0 ± 10.4% for Ru-H , Ru-Me , and Ru-BN , respectively ( Figure 12 B), confirming mitochondrial
membrane depolarization. Consistent with the microscopy experiments,
the most notable effect on MMP was observed with complex Ir-H . Ir-BN and Ru-BN metallopeptides also
induced significant mitochondrial membrane depolarization, indicating
that conjugation with BN3 does not hinder this effect.
Furthermore, the complexes’ ability to photocatalytically oxidize
NADH, a primary electron donor for the mitochondrial electron transport
chain, may contribute to the mitochondrial damage. Cell Death
Mechanism Mitochondria play an important
role in the regulation of apoptosis. Alterations in the MMP can cause
the release of apoptotic factors, such as cytochrome c, which activates
the caspase cascade, leading to programmed cell death. 66 , 67 In order to elucidate whether the toxic effect of the compounds
at the mitochondrial level triggers cell death by apoptosis, a dual
annexin V/propidium iodide labeling experiment was performed. The
initial stages of apoptosis are characterized by alterations in the
symmetry of phospholipids in the cytoplasmic membrane, which can be
detected using annexin V (An). The late stages also include disruption
of the cell membrane, permitting the penetration of propidium iodide
(PrI), which emits red fluorescence when bound to DNA. In contrast,
the cell membrane of necrotic cells becomes readily permeable to PrI
but does not exhibit phospholipid translocations. This allows the
discrimination among viable cells (An–/PrI−), necrotic
cells (An–/PrI+), and early-(An+/PrI−) and late-stage
(An+/PrI+) apoptotic cells. PC-3 cells were treated with the photoactivated
complexes and metallopeptides at five times the corresponding IC 50,light , and 24 h later, samples were analyzed using flow
cytometry. In all cases, the treatment resulted in a significant increase
in the number of apoptotic cells, particularly in the late stage of
apoptosis, compared to control cells ( Figure 13 ). A higher percentage of apoptotic cells
was detected upon treatment with Ir(III) complexes than Ru(II) complexes,
reaching a 11.9% of cells in early apoptosis and 41.7% of cells in
late apoptosis in the case of Ir-Me . In contrast, the
population of necrotic cells was not increased by any treatment. Overall,
these results indicated that the photodynamic activity of both the
complexes and metallopeptides promotes a regulated cell death by apoptosis. Figure 13 Cell
death mechanism. PC-3 cells were treated with the photoactivated
compounds at five times the corresponding IC 50,light . After
24 h, the percentage of viable, necrotic, early apoptotic, and late
apoptotic cells was analyzed by flow cytometry using propidium iodide
and annexin V staining. Untreated cells were used as controls. Bars
represent the mean ± standard deviation from three independent
experiments. * p < 0.05 and ** p < 0.01 versus control cells.
## Synthesis of Ligands and Complexes
Synthesis of Ligands and Complexes Two pairs of Ir
and Ru complexes with two different N^N’ ligands ( L1 and L2 ) and general formulas [Ir(C^N) 2 (N^N’)]Cl
( Ir-Me: C^N = ppy, N^N’ = L1 ; Ir-H: C^N = ppy, N^N’ = L2 ) and [Ru(N^N) 2 (N^N’)](Cl) 2 ( Ru-Me: N^N =
bpy, N^N’ = L1; Ru-H: N^N = bpy, N^N’ = L2 ) have been prepared aiming to evaluate the effect of the
metal fragment on their photophysical and biological properties ( Schemes 1 and 2 ). Moreover, complexes Ir-H and Ru-H have been prepared with a −COOH group to allow their conjugation
to the bombesin derivative BN3 . The ancillary ligands
( L1 - L2 ) were synthesized in several steps
according to Scheme 1 . First, we reacted tryptamine and 5-bromo-2-pyridinecarboxaldehyde
to obtain pyridyl-β-carboline A , following a method
previously described for similar compounds. 43 Then, the pyrrolic N–H group was substituted with a methyl
group to obtain derivative B , employing NaH as the base
and MeI as the methylating agent. 22 Subsequently,
a Heck vinylation protocol was performed on the bromopyridyl ring
of B , using methyl acrylate to obtain L1 . Furthermore, the ester group was hydrolyzed in alkaline media and
neutralized with HCl to obtain L2 , bearing a −COOH
group. Scheme 1 Synthetic Route for the Synthesis of L1 and L2 Scheme 2 Synthesis and Molecular Structure
of Complexes Ir-Me , Ir-H , Ru-Me , and Ru-H The Ir(III) complexes of general
formula rac -[Ir(ppy) 2 (N^N’)]Cl
(ppy = 2-phenylpyridinate
and N^N’
= L1 or L2 ) were prepared through a bridge
splitting reaction between the dimeric Ir(III) precursor rac -[Ir(μ-Cl)(ppy)] 2 and the corresponding β-carboline-based
ligand ( L1 or L2 , using a molar ratio 1:2,
dimer:ligand) to obtain Ir-Me and Ir-H ,
respectively ( Scheme 2 ). Ru(II) polypyridyl complexes of general formula rac -[Ru(bpy) 2 (N^N’)](Cl) 2 (bpy = 2,2′-bipyridine
and N^N’ = L1 or L2 ) were prepared
by heating the Ru(II) precursor, rac- [Ru(bpy) 2 Cl 2 ], with L1 or L2 (1:1
molar ratio), to obtain Ru-Me and Ru-H ,
respectively ( Scheme 2 ). The metal complexes were obtained as racemic mixtures (Λ,Δ)
of the respective chloride salts. Ligands and complexes were fully
characterized by nuclear magnetic resonance (NMR), mass spectrometry,
and elemental analysis (see Figures S1–S32 ). In particular, 1 H NMR spectra of all the complexes
were recorded in DMSO- d 6 at 25 °C.
The spectra displayed two sets of resonances for the two nonequivalent
ppy/bpy ligands owing to the lack of symmetry shown by these derivatives.
All of them showed a singlet around 3.93 ppm attributed to the N-Me
group, which is shifted to lower field compared to the corresponding
singlet of the free ligand (3.60 ppm). The high-resolution electrospray
ionization mass spectrometry (HR ESI(+) MS) spectra exhibited peaks
with mass/charge ratios and isotopic distributions fully compatible
with those calculated for either the monocationic Ir complexes or
the dicationic Ru complexes. The analytical data obtained from elemental
analysis are in good agreement with expected values.
## Synthesis of
the Metallopeptides
Synthesis of
the Metallopeptides Metallopeptides Ir-BN and Ru-BN incorporating the Ir(III) or
the Ru(II) complex at the N-terminus of the peptide BN3 were synthesized on solid phase following a standard 9-fluorenylmethoxycarbonyl
(Fmoc)/ tert -butyl ( t Bu) strategy
( Scheme 3 ). 38 An aminomethyl ChemMatrix resin was used as
solid support to which the Fmoc-Rink-amide linker was incorporated
leading to C-terminal amidated peptides. The attachment of this linker
to the ChemMatrix resin was accomplished using N,N -diisopropylcarbodiimide (DIC) as coupling reagent, ethyl (hydroxyimino)cyanoacetate
(Oxyma) as additive, and dimethylformamide (DMF) as solvent. The
elongation of the peptide sequence was performed through sequential
steps of Fmoc group removal with piperidine/DMF (3:7) and coupling
of the corresponding amino acids using DIC and Oxyma in DMF. A Kaiser
test was performed to confirm the completion of the coupling reactions. 44 Once the peptide sequence was completed, the
N-terminal Fmoc group was removed, and the corresponding metal complex Ir-H or Ru-H was incorporated in the presence
of DIC and Oxyma in DMSO. Next, the resulting metallopeptides Ir-BN and Ru-BN were cleaved from the support
by acidolytic treatment with trifluoroacetic acid (TFA)/H 2 O/triisopropylsilane (TIS), purified by reversed-phase column chromatography,
analyzed by HPLC, and characterized by mass spectrometry. Ir-BN and Ru-BN were obtained in >99% HPLC purity (see Figures S33–44 ). Scheme 3 Solid-Phase Synthesis
of Metallopeptides Ir-BN and Ru-BN
## X-ray Diffraction
X-ray Diffraction The crystal structure
of [ Ir-Me ]PF 6 was resolved by single-crystal
X-ray diffraction.
A suitable single crystal was obtained by slow evaporation of an acetonitrile
solution of [ Ir-Me ]Cl in the presence of NH 4 PF 6 . The complex crystallizes in the monoclinic C 2/ c space group. Due to helical chirality,
the unit cell contains two pairs of enantiomers (Δ,Λ)
( Figure S45 ). The ORTEP diagram for the
molecular structure of Λ-[Ir-Me] + is shown in Figure 1 . Selected bond distances and angles are compiled in Table 1 , and important crystallographic
parameters are gathered in Table S3 . Figure 1 ORTEP diagram
for the molecular structure of Λ-[Ir-Me] + obtained by single-crystal X-ray diffraction.
Thermal ellipsoids are shown at the 30% probability level. The respective
Δ enantiomer, H atoms, and the PF 6 – counterion have been omitted for the sake of clarity. Table 1 Selected Bond Distances (Å) and
Coordination Angles for rac -[Ir-Me] + rac -[Ir-Me] + Distances rac -[Ir-Me] + Angles (deg) Ir(1)–N(1) 2.153 (4) N(1)Ir(1)N(2) 76.79 (14) Ir(1)–N(2) 2.159 (3) C(32)Ir(1)N(4) 80.62 (19) Ir(1)–N(4) 2.045 (4) C(43)Ir(1)N(5) 80.3 (2) Ir(1)–C(32) 2.021 (5) Ir(1)–N(5) 2.046 (4) Ir(1)–C(43) 2.021 (4) The molecular structure of [Ir-Me] + exhibited the expected pseudo-octahedral
geometry
with the
predictable trans -N,N and cis -C,C
disposition for the two cyclometalated ligands (ppy). The Ir–N
bond distances for the N^N’ ligand (2.153(4), 2.159(3) Å)
were longer than those determined for the ppy ligands (2.046(4), 2.045(45)
Å), due to the strong trans influence attributed
to the metal-bonded phenyl rings. Both Ir–C bond lengths were
nearly identical (2.021(5) and 2.021(5)) and within the expected range
for this kind of complexes. The bite angles of the chelate rings were
also standard, that is, 76.79° for L1 and 80.62°,
80.30° for the ppy ligands. The N^N’ ligand displayed
a high torsion angle (−18.55°) in comparison with the
ppy ligands (2.07° and 3.04°) revealing a lower degree of
coplanarity, due to the high steric hindrance between the N-Me group
and the pyridyl ring. Furthermore, the crystal structure exhibited
different weak hydrogen-bonding interactions between fluorine atoms
from the counterion and different hydrogen donor groups of the iridium
complex. Also, a weak hydrogen-bonding interaction between an oxygen
atom (O1) from L1 and one hydrogen (H23) from a ppy ligand
of a neighboring molecule was observed ( Figure S46 and Table S4 ).
## UV–Vis Spectroscopy
UV–Vis Spectroscopy The UV–vis
absorption
spectra of Ir-H , Ir-Me , Ru-H , and Ru-Me ( Figure 2 ) were recorded at room temperature for H 2 O:DMSO (99:1) solutions (10 –5 M). All the complexes
displayed strong absorption bands with maxima between 250 and 350
nm that were attributed to spin-permitted ligand centered transitions
( 1 LC, π → π*). In the visible region,
Ir(III) complexes showed a main absorption band with maxima around
420 nm that extends up to 550 nm and corresponds to mixed spin-allowed
and spin-forbidden metal to ligand charge transfer ( 1 MLCT
and 3 MLCT) and ligand to ligand charge transfer (LLCT)
transitions. By contrast, the Ru(II) complexes showed two absorption
bands in the visible region, with maxima at 430 and 500 nm and a tail
extended up to 600 nm. These bands were also ascribed to 1 MLCT and 3 MLCT and LLCT transitions. Therefore, the Ru(II)
derivatives exhibited red-shifted bands compared to their Ir(III)
analogues. Moreover, complexes with the methyl ester group Ir-Me / Ru-Me showed absorption bands slightly red-shifted
relative to the complexes with −COOH groups, Ir-H / Ru-H . Figure 2 Overlaid absorbance spectra of Ir(III)/Ru(II) complexes
in H 2 O:DMSO (99:1, v:v) (10 –5 M) at room
temperature.
Inset: zoom of the region between 500 and 700 nm.
## Emission and Photophysical Properties
Emission and Photophysical Properties The emission
spectra of the metal complexes in deoxygenated solutions of H 2 O:DMSO (99:1, v:v, 10 –5 M) were recorded
at 25 °C, using λ ex of 405 nm for Ir-Me / Ir-H and λ ex of 450 nm for Ru-Me / Ru-H ( Figure 3 ). All the complexes displayed a single broad emission band
with maxima between 651 and 720 nm and large Stokes shifts (λ em – λ ex > 230 nm), which corroborated
the phosphorescent nature of the emission. The Ru(II) complexes exhibited
higher emission intensities, and their respective maxima are red-shifted
compared to those of the Ir(III) analogues. Moreover, Ir-Me / Ru-Me featured lower emission intensities compared
to Ir-H / Ru-H , respectively. Figure 3 Overlaid emission spectra
of Ir(III)/Ru(II) complexes
in deoxygenated H 2 O:DMSO (99:1, v:v, 10 –5 M) at 25 °C. The excited state lifetimes
(τ) and the photoluminescence
quantum yields were also determined for the new metal complexes ( Table 2 ). The Ir PSs displayed
two-component τ values of 2 ns [20%]/93 ns [80%] and 23 ns [51%]/145
ns [49%] for Ir-Me and Ir-H , respectively.
By contrast, the Ru(II) complexes exhibited one-component τ
values of 74 and 186 ns for Ru-Me and Ru-H , respectively. These values compare well with those of similar complexes
and are compatible with a triplet nature for the emissive states.
The photoluminescence quantum yields (Φ PL ) of Ir(III)/Ru(II)
were very low with values ranging from 1.1 to 4.13%. Table 2 Photophysical Properties of the Metal
Complexes in H 2 O:DMSO (99:1, v:v) at 25 °C under a
Nitrogen Atmosphere Compound λ ex (nm) λ em (nm) τ (ns) [contribution (%)] Φ PL (%) Ir-Me 405 676 2 [20] 93 [80] 3.50 Ir-H 405 651 23 [51] 145
[49] 4.13 Ru-Me 450 720 74 1.10 Ru-H 450 688 186 3.13
## Determination of p
Determination of p K a Ir-H and Ru-H are susceptible to deprotonation
in aqueous media due to the presence of the acidic −COOH group.
In fact, the actual protonation state of these complexes at the different
physiological pH values defines their global charge and correspondingly
influences their solubility, their photophysical properties, and their
cellular uptake. 45 − 47 Thus, the p K a values
of Ir-H and Ru-H in their excited states
were experimentally determined by monitoring the variations of their
emission intensity at specific wavelengths versus different pH values
and fitting the data ( I em,λ /pH)
to a sigmoidal equation. In good agreement with similar complexes
reported in the literature, 45 , 46 there was a gradual
decrease in the emission intensity of Ir-H and Ru-H with decreasing pH ( Figures 4 and S50 ). The
p K a values for Ir-H and Ru-H in their excited states were 3.81 and 4.51, respectively.
These values can be considered as an acceptable approximation to the
values of the respective ground states. Therefore, we concluded that Ir-H and Ru-H adopt their deprotonated zwitterionic
forms in the whole range of physiological pHs. This means that Ir-H assumes a zwitterionic neutral form, while Ru-H adopts a zwitterionic monocationic structure. In both cases, there
is charge separation, and as a result, we predict that the cellular
uptake ability of these complexes could be reduced, in comparison
to that of their relatives Ir-Me and Ru-Me . Figure 4 p K a determination. (A) Overlaid emission
spectra of Ir-H in H 2 O:DMSO (99:1, v:v) (10 –5 M) at different pH values ranging from pH 7 to 2
(HCl titration). (B) Plot of the emission intensity of Ir-H at λ 651 nm in H 2 O:DMSO (99:1, v:v) (10 –5 M) as a function of pH (2–7) at 25 °C. (C) Scheme showing
the acid–base equilibrium for Ir-H .
## Photostability of Ir(III)/Ru(II) Complexes
Photostability of Ir(III)/Ru(II) Complexes Photostability
is a desirable feature for PDT agents, since otherwise, their efficiency
as PSs can be decreased owing to so-called photobleaching. Thus, the
photostability of aerated solutions (1.5 × 10 –2 M, DMSO- d 6 :D 2 O, 3:2, v:v)
of all our complexes was evaluated by 1 H NMR. The solutions
were exposed to blue light irradiation (λ irr = 460
nm, 24 W), and the evolution of the respective samples was monitored
by 1 H NMR at different times (0, 6, and 24 h) at room temperature.
To our delight, no symptoms of photodegradation were observed, confirming
that all the complexes exhibited outstanding photostability under
the aforementioned conditions ( Figures 5 and S47–S49 ). Figure 5 Evolution of
the aromatic region of the 1 H NMR spectra
of Ru-H (1.5 × 10 –2 M) in DMSO- d 6 :D 2 O (3:2, v:v) under blue light
irradiation (LED, λ = 460 nm, 24 W) at different times.
## Photocatalytic Generation of Singlet Oxygen
Photocatalytic Generation of Singlet Oxygen Singlet
oxygen is thought to be the main cytotoxic species in type II PDT
processes. Therefore, we determined the ability of Ir-Me and Ru-Me to generate 1 O 2 under
photocatalytic conditions, as illustrative examples of our PSs. As
shown in Figures 6 and S51 , we monitored by means of UV–vis spectroscopy
the oxidation of 9,10-anthracenediyl bis(methylene)dimalonic acid
(ABDA, 8x 10 –5 M) in the presence of atmospheric
oxygen, using Ir-Me (10 –5 M) as the
PS and H 2 O:DMSO (1:1) as the solvent system under blue
light exposure (λ ir = 460 nm). ABDA is a very specific
probe for 1 O 2 , since it reacts selectively with
the in situ produced 1 O 2 to generate the respective
endoperoxide, which is characterized by a loss of π-extended
aromaticity and the disappearance of several absorption bands in the
UV–vis region (i.e., 379 nm). Moreover, the singlet oxygen
quantum yield values ( Φ Δ ) of Ir-Me and Ru-Me were determined using rose bengal
( Φ Δ = 0.75) as the reference.
As expected, both Ir-Me and Ru-Me exhibited
photocatalytic activity in the generation of 1 O 2 , but their respective Φ Δ were
low, 4 and 9%, in agreement with its moderate excited state lifetimes
(93 and 74 ns). A control experiment in the absence of PS was also
performed resulting in a negligible conversion. Figure 6 Photobleaching of ABDA
(8 × 10 –5 M) in the
presence of Ir-Me (10 –5 M) using H 2 O:DMSO (1:1) under blue light irradiation (460 nm, 24 W) during
100 min at room temperature.
## Photocatalytic Oxidation of NADH
Photocatalytic Oxidation of NADH NADH is an enzymatic
cofactor playing a main role as an electron donor in the mitochondrial
electron transport chain. Hence, it is considered a potential molecular
target for those anticancer drugs that cause oxidative stress in mitochondria.
Indeed, several research groups have recently established a relationship
between the photocatalytic oxidation of NADH to NAD + , the
depolarization of the mitochondrial membrane and apoptotic cell death. 48 , 49 Aiming to prove this hypothesis for Ir-Me , as
a model PS, we monitored by UV–vis spectroscopy the photocatalytic
oxidation of an aerated solution of NADH (0.1 mM) in the presence
of Ir-Me (5 μM) using H 2 O:DMSO (99:1)
under blue light irradiation (λ ir = 460 nm) for 15
min. As a result, we observed the decrease of the band due to NADH
at 338 nm ( Figure 7 ). This evolution is compatible with the photocatalyzed formation
of NAD + . Two control experiments in the absence of either Ir-Me ( Figure S52 ) or light ( Figure S53 ) corroborated the photocatalytic character
of this transformation. Figure 7 Evolution of the UV–vis spectra during
the photocatalytic
oxidation of NADH (100 μM) in the presence of Ir-Me (5 μM) in aerated H 2 O:DMSO (99:1) under blue light
irradiation (460 nm, 24 W) at room temperature (15 min).
## Effect on Cell Viability
Effect on Cell Viability The effect of Ir-H , Ir-Me , Ru-H , and Ru-Me on
cell viability was evaluated in the A549 lung cancer and PC-3 prostate
cancer cell lines by MTT assays. PC-3 cells were specifically chosen
due to their high expression levels of GRPR on their surface, allowing
for further evaluation of the activity of the BN3 -derived
metallopeptides Ir-BN and Ru-BN . 38 , 51 In contrast, A549 cells express low levels of GRPR. 52 MRC-5 fibroblasts were used as a nonmalignant cell model. 53 Table 3 reports the half maximal inhibitory concentration (IC 50 ) values obtained after treating the cells either under dark
conditions or upon exposure to blue light at a dose of 24.1 J cm –2 . The photocytotoxicity index (PI), defined as the
IC 50,dark /IC 50,light , is provided for each compound. Table 3 Photocytotoxic Effects of Complexes Ir-H , Ir-Me , Ru-H , and Ru-Me and
of Metallopeptides Ir-BN and Ru-BN on Different
Cell Lines in the Dark and after Blue Light Irradiation IC 50 a (μM) A549 PC-3 MRC-5 Dark Light PI b Dark Light PI b Dark Light PI b Ir-H 35.0 ± 5.4 3.32 ± 1.1 10.5 32.2 ± 1.9 9.23 ± 0.63 3.5 39.1 ± 2.4 6.46 ± 0.801 6.1 Ir-Me 0.471 ± 0.031 0.0334 ± 0.0068 14.2 1.17 ± 0.29 0.0723 ± 0.031 16.2 1.12 ± 0.31 0.0313 ± 0.019 35.8 Ru-H 67.8 ± 6.7 56.7 ± 1.6 1.2 70.4 ± 2.7 34.8 ± 4.3 2.0 53.9 ± 5.4 43.5 ± 4.5 1.2 Ru-Me 74.7 ± 2.03 33.8 ± 5.2 2.2 76.8 ± 2.1 48.6 ± 0.5 1.6 62.5 ± 2.6 43.7 ± 7.2 1.4 Ir-BN 68.3 ± 15.8 12.6 ± 1.3 5.4 35.3 ± 3.8 8.51 ± 0.8 4.1 80.9 ± 1.3 28.7 ± 5.9 2.8 Ru-BN 49.5 ± 11.9 23.6 ± 11.2 2.1 41.3 ± 1.7 4.11 ± 1.2 10.1 >100 >100 n.d. Cisplatin 5.99 ± 1.2 n.d. n.d. 5.55 ± 0.97 n.d. n.d. 5.34 ± 0.38 n.d. n.d. a Cells were incubated with the
compounds for 4 h at 37°C and then kept in the dark or exposed
to blue light irradiation for 1 h (460 nm, 24.1 J cm –2 ). Cell viability was assessed after 48 h of treatment by MTT assays.
Data represent the mean ± SD of at least three independent experiments,
each performed in triplicate. b Photocytotoxic index (PI) = IC 50, dark /IC 50, light . n.d.: not determined. The Ir(III) complexes exhibited a greater effect on
cell viability
than their Ru(II) counterparts in all cell lines. Among them, Ir-Me displayed the highest anticancer activity, with IC 50 values of 0.471 μM in A549 cells and 1.17 μM
in PC-3 cells in dark conditions, which were lower than those of cisplatin
under the same experimental conditions. Conversely, Ir-H displayed IC 50,dark values above 30 μM, revealing
that the esterification of the β-carboline ligand with a methyl
group significantly enhanced the biological activity of the resulting Ir-Me complex. Upon blue light irradiation, the cytotoxicity
of Ir-Me increased by 14.2-fold in A549 cells and 16.2-fold
in PC-3 cells, resulting in IC 50,light values of 33.4 nM
and 72.3 nM, respectively. The anticancer activity of Ir-H also increased after light exposure by 10.5-fold in A549 cells and
3.49-fold in PC-3 cells, with IC 50,light values decreasing
to the low micromolar range. It should be noted that Ir-H and Ir-Me displayed similar IC 50 values
in MRC-5 fibroblasts as in cancer cells, both in the dark and upon
irradiation, evidencing their nondiscriminatory effects between nonmalignant
and cancer cells. Regarding the Ru(II) complexes, Ru-H and Ru-Me showed very moderate anticancer activity
in dark conditions, which
was not significantly enhanced upon exposure to blue light despite
having higher light absorption at 460 nm than the Ir (III) complexes
( Figure 2 ). Moreover,
they exhibited similar toxicity levels against MRC-5 fibroblasts,
as observed with the Ir(III) complexes. The cytotoxic characterization
of Ir-H , Ir-Me , Ru - H , and Ru-Me was further
complemented by hemolysis assays, which determined whether they could
induce the rupture of red blood cell membranes, leading to the release
of hemoglobin. 54 Both Ir(III) and Ru(II)
complexes exhibited hemolysis levels of less than 1% both in the absence
and in the presence of blue light irradiation. This finding indicates
that the complexes are nontoxic to red blood cells and have good blood
compatibility for future clinical applications. Finally, the
photocytotoxic activity of the BN3 -derived
metallopeptides, Ir-BN and Ru-BN , was evaluated.
In PC-3 cells, Ir-BN exhibited similar activity relative
to the precursor complex, Ir-H , both in the dark and
upon activation with blue light. However, in MRC-5 fibroblasts, the
IC 50,dark and IC 50,light values for Ir-BN were 2.1- and 4.4-fold higher, respectively, than those of Ir-H ( Table 3 ). It should be noted that Ir-BN exhibited 2.3-fold
and 1.9-fold higher cytotoxicity in PC-3 cells compared to MRC-5 fibroblasts
and A549 cancer cells with low GRPR expression, respectively. After
exposure to blue light, the selectivity for PC-3 cells increased to
3.4-fold with respect to MRC-5 fibroblasts. Regarding Ru-BN , the IC 50,dark and IC 50,light values in PC-3
cells were 1.7 and 8.5 times lower, respectively, than those of the
precursor Ru-H complex. In terms of selectivity, the
activity of Ru-BN in PC-3 cells was 1.2- and 5.7-fold
higher than in A549 cells under dark and irradiated conditions, respectively.
Importantly, Ru-BN exhibited no cytotoxicity in MRC-5
fibroblasts even at concentrations as high as 100 μM, in both
dark and irradiated conditions. The photocytotoxic effects of Ru-H , Ru-Me , and Ru-BN were also
evaluated following photoactivation
with red light (λ ir = 655 nm), prompted by the red-shifted
band observed in the absorption spectra of the Ru (II) complexes ( Figure 2 ). Notably, red light
offers superior tissue penetration compared to blue light, potentially
enabling treatment of deeper tumors. 55 As
shown in Table 4 , red
light had a low impact on the activity of both Ru(II) complexes, with
PIs below 1.3 in both PC-3 and MRC-5 cells. In contrast, Ru-BN displayed markedly higher photocytotoxic activity against PC-3 cells,
with a PI of 7.3, while it displayed no cytotoxicity toward MRC-5
fibroblasts. Table 4 Photocytotoxic Effects of Ru-H , Ru-Me , and Ru-BN after Red Light Irradiation a PC-3 MRC-5 IC 50,red light (μM) PI IC 50,red light (μM) PI Ru-H 54.3 ± 4.7 1.3 62.3 ± 7.1 0.9 Ru-Me 63.1 ± 5.8 1.2 66.6 ± 4.8 0.9 Ru-BN 5.66 ± 1.2 7.3 >100 -- a Cells were incubated with the
compounds for 4 h at 37 °C and then kept in the dark or exposed
to red light irradiation for 1 h (655 nm, 24.1 J cm –2 ). Cell viability was assessed after 48 h of treatment by MTT assays.
Data represent the mean ± SD of at least three independent experiments,
each performed in triplicate. PI: photocytotoxic index = IC 50, dark /IC 50,red light . To examine the role of BN3 as a carrier
peptide for
specifically targeting PC-3 cells, the influence of blocking the bombesin
receptors on the anticancer efficacy of the metallopeptides was assessed.
PC-3 cells were treated with Ir-BN and Ru-BN at concentrations near their IC 50,light (10 and 5 μM,
respectively) in the presence of increasing bombesin concentrations
(0, 10, 50, and 100 μM) to competitively inhibit GRPR binding.
Following irradiation with blue light, the viability of the cells
was evaluated. The photocytotoxic activity of both Ir-BN and Ru-BN was significantly attenuated in a bombesin
concentration-dependent manner, with approximately a 25% reduction
in the presence of bombesin at 10 μM and a more pronounced inhibition
of approximately 35% at 50 and 100 μM ( Figure S54 ). These results strongly suggest that the cytotoxic effect
of Ir-BN and Ru-BN is mediated, at least
in part, by their interaction with GRPR. Overall, these findings
support the efficacy of BN3 conjugation in enhancing
the selectivity of PSs toward cancer cells
overexpressing GRPR. This is also evidenced by photoselectivity indexes
(IC 50,dark in nonmalignant cells/IC 50,light in
GRPR overexpressing cancer cells) of 9.5 for Ir-BN after
activation with blue light and exceeding 24.3 and 17.6 for Ru-BN after activation with blue and red light, respectively.
## Inhibition
of Colony Formation
Inhibition
of Colony Formation The effect of the metallopeptides Ir-BN and Ru-BN as well as of the complexes Ir-H and Ru-H on the viability of PC-3 cells
was next investigated through clonogenic assays, which determine the
fraction of cells that survive treatment and retain the ability to
generate new colonies. 56 This is a crucial
feature of metastatic cancer cells, which need to proliferate in distant
tissues to create secondary tumors. For this purpose, PC-3 cells were
treated with the compounds at an equimolar concentration of 5 μM
with or without photoactivation with blue light. Ten days later, the
number of cells that survived and were able to grow and form colonies
was assessed in comparison to control cells. Cells treated with cisplatin
were used as a positive control. None of the compounds inhibited the
clonogenic activity of the cells in the absence of light irradiation
( Figure 8 ). However,
upon blue light irradiation, the number of colonies decreased by 27.3%,
34.7%, and 50.4% in cells exposed to Ir-H , Ir-BN , and Ru-BN , respectively. No effect of Ru-H on the clonogenic capability of the cells was observed at 5 μM,
which is consistent with its high IC 50,light value ( Table 3 ). Figure 8 Clonogenic assay. PC-3
cells were treated with the indicated compounds
at 5 μM in the dark and with blue light irradiation. Control
cells were treated with the medium alone. Cisplatin was used as a
positive control. Cells were incubated for 10 days to allow colony
formation. (A) Images of the colonies. (B) Bar charts representing
the percentage of colonies versus control cells after each treatment
(mean ± SD of 3 experiments). ** p < 0.01;
*** p < 0.001 versus control cells.
## Intracellular ROS Generation
Intracellular ROS Generation The photocytotoxic effects
of the complexes and metallopeptides were further elucidated by evaluating
their ability to photogenerate ROS at the cellular level. PC-3 cells
were treated with the compounds at the respective IC 50,light and irradiated with blue light. Subsequently, ROS generation was
measured using the 2′,7′-dichlorodihydrofluorescein
diacetate (H 2 DCFDA) probe, which diffuses into cells and
is oxidized by ROS to produce a green fluorescent signal. As illustrated
in Figure 9 A, in all
cases, ROS levels were significantly increased by more than 2-fold
in comparison to the control cells. Ir-Me exhibited the
most potent ROS-generating activity, with a 4.5-fold increase. It
is worth noting that, despite Ir-Me exhibiting relatively
low efficiency as a photocatalyst for the generation of 1 O 2 , its effective cellular uptake could counterbalance
this limitation (see Figure 10 ), enabling effective ROS generation inside the cells. Nevertheless,
to ascertain the potential contributions of other ROS to the photocytotoxic
activity, the capacity of the compounds to undergo type I PDT processes,
resulting in the production of superoxide anions (O 2 •– ), was examined using a specific fluorescent
probe that emits an orange signal upon interaction with this radical. Ir-H , Ir-Me , and Ir-BN induced the
most significant increase in O 2 •– levels, with fold changes of 2.4, 3.5, and 3.4, respectively, compared
to control cells. In contrast, the Ru complexes exhibited lower increments
( Figure 9 B). These
findings highlight the diverse mechanisms of ROS generation employed
by these compounds, in particular the Ir complexes, which may contribute
to their overall photocytotoxic efficacy. Figure 9 Intracellular ROS generation.
PC-3 cells were treated with the
indicated complexes and metallopeptides at their respective IC 50,light for 4 h, followed by blue light irradiation for 1
h (460 nm, 24.1 J cm –2 ). The elevation of general
ROS (A) and superoxide anion (B) levels was determined with specific
probes by flow cytometry. Bars represent the mean fold increase (±standard
deviation) relative to control untreated cells from three independent
experiments. * p < 0.05; ** p < 0.01, *** p < 0.001 compared to control
cells. Figure 10 Cellular internalization. PC-3 cells
were incubated for
4 h with
the indicated compounds at 5 μM, and the Ir (A) or Ru (B) content
per 1 × 10 6 cells was quantified by ICP-MS. Bars represent
the mean value of three replicates ± SD. * p < 0.05; *** p < 0.001 versus Ir-H or Ru-H .
## Internalization Studies
Internalization Studies The intracellular accumulation
of the PSs is a crucial requirement to achieve a photocytotoxic effect.
This is because the ROS generated by the PSs have a very limited radius
of action and only operate in the cellular compartment or organelle
in which they are generated. 57 In particular,
the absence of hemolytic activity observed in both Ir(III) and Ru(II)
complexes suggests a mechanism of action that does not involve cell
membrane disruption. This further emphasizes the importance of cellular
uptake for their cytotoxic activity. Additionally, when using tumor-targeting
ligands, such as BN3 , the intracellular uptake of the
metallopeptides depends on the level of expression of the target receptor
on the cell surface and the internalization mechanism, which may differ
from that of the free complex. Therefore, the internalization of complexes Ir-H , Ir-Me , Ru - H ,
and Ru-Me and of metallopeptides Ir-BN and Ru-BN in PC-3 cells was determined by quantifying the intracellular
metal content by inductively coupled plasma mass spectrometry (ICP-MS)
( Figure 10 ). After
4 h treatment with the compounds at 5 μM, the amount of Ir in
cells exposed to Ir-H was 3.1 ± 0.7 ng per million
cells, whereas in the case of its conjugated form Ir-BN , it increased 5.7-fold to 17.9 ± 5.6 ng per million cells.
This result showed that conjugation to BN3 enhances the
cellular uptake of the Ir(III) complex. Notably, Ir-Me exhibited the highest internalization, resulting in 266.4 ±
13.9 ng Ir per million cells, consistent with its superior anticancer
activity. We speculate that the higher ability of Ir-Me to accumulate within the cells is consistent with its lipophilic
character (vide infra). By contrast, Ir-H exhibited a
poor intracellular uptake likely due to the presence of the negatively
charged −COO – group in its deprotonated zwitterionic
form. It should be noted that Ir-Me has a significantly
higher internalization capacity than the Ir-BN conjugate.
This discrepancy can be attributed to the lipophilic nature of Ir-Me , which enables its passive diffusion across the cell
membrane. Consequently, Ir-Me can be more rapidly and
directly internalized by cells. Conversely, Ir-BN entry
involves receptor-mediated endocytosis, a slower, energy-dependent
process that often results in lower intracellular concentrations and
can be influenced by variations in the endocytic pathway activity. In the case of cells treated with the Ru(II) complexes, the intracellular
metal contents were markedly lower, with 0.329 ng per million cells
and 0.326 ng per million for Ru-H and Ru-Me , respectively. We believe that the scarce accumulation of the Ru
derivatives is due to their lower lipophilicity (vide infra), since
in the physiological pH range, Ru-H shows a monocationic
zwitterionic form and Ru-Me a dicationic nature. These
charged states may hinder passive diffusion across the cell membrane.
However, the amount of intracellular Ru significantly increased 14-fold
to 4.46 ng/million cells for Ru-BN . Overall, these
results demonstrate that the conjugation of the
complexes to BN3 not only enhances their selectivity
for tumor cells but also increases the intracellular accumulation
of the conjugated complexes Ir-H and Ru-H , leading to increased cytotoxic activity, particularly for the Ru(II)
derivative.
## Lipophilicity and Self-Aggregation Studies
Lipophilicity and Self-Aggregation Studies In order
to obtain a better understanding of the cellular uptake abilities
of these complexes, we carried out experiments to determine their
lipophilicity and self-aggregation properties. First, the n -octanol/PBS partition coefficient values, log P oct/PBS , of the metal PSs were experimentally determined to quantify their
lipophilicity ( Figure S56 ). Indeed, the
shake flask method was employed, and the metal content was measured
in both phases by UV/vis spectroscopy. Thus, we established that both
Ru derivatives exhibited low lipophilicity in agreement with either
the dicationic nature of Ru-Me (log P oct/PBS = −0.43) or the monocationic zwitterionic character of Ru-H (log P oct/PBS = −1.10) at physiological
pH. These results are consistent with the low cellular uptake shown
by these derivatives. By contrast, both Ir derivatives show
higher lipophilicity relative
to their Ru congeners, which is rationally explained as a result of
the monocationic nature of Ir-Me and the neutral zwitterionic
character of Ir-H at physicological pH. Indeed, no significant
differences can be observed in the lipophilicity between Ir-Me (log P oct/PBS = 2.03) and Ir-H (log P oct/PBS = 1.80). Therefore, we speculate that the pronounced
divergent cellular uptake behavior experimentally observed for Ir-H and Ir-Me could be explained as a result
of the electronic repulsion between the negatively charged −COO – group in Ir-H and the negative charge
of the phosphate groups present in the cellular membrane. In other
words, we assume that the presence of the −COO – group in the zwitterionic form of Ir-H at physiological
pH values would hinder its uptake through the cell membrane by passive
diffusion. 58 The possible aggregation
properties (self-assembly behavior) were
investigated for the lipophilic derivative Ir-Me through
the respective analysis of the Lambert–Beer law using UV–vis
spectroscopy in H 2 O:DMSO (99:1). More specifically, the
absorbance of Ir-Me at λ = 417 nm was plotted versus
concentration in the range between 1 and 40 μM. An excellent
linear fitting was obtained confirming that there is no significant
deviation from the Lambert–Beer law ( Figures S57 and S58 ). Therefore, we conclude that Ir-Me does not form aggregates under these conditions.
## Intracellular
Distribution
Intracellular
Distribution The intracellular distribution
of the compounds was then investigated by confocal microscopy. Preliminary
flow cytometry experiments revealed that significant intracellular
fluorescence was detectable only in cells exposed to Ir-Me , consistent with the higher intracellular accumulation of this compound
( Figure S55 ). Accordingly, confocal
microscopy images showed a strong red fluorescence signal from Ir-Me inside the cells, as shown in Figure 11 . The complex displayed a high degree of
colocalization with the mitochondrial dye MitoView Green, as evidenced
by the yellow signal in the merged image (Pearson correlation coefficient
(PCC): 0.832). In contrast, a lower degree of overlap was observed
with the LysoTracker Green DND-26 lysosomal dye (PCC: 0.599), indicating
that the complex predominantly accumulates within the mitochondria
and exhibits a lower distribution within the endolysosomal system.
This is compatible with the monocationic and lipophilic nature of Ir-Me and has been previously reported for other mitochondria-targeting
monocationic bis-cyclometalated Ir(III) complexes. 59 − 63 Figure 11 Confocal images of A549 cells after 1 h of incubation
with Ir-Me at 1 μM (red). MitoView Green or LysoTracker
Green
DND-26 (green) were used for colocalization studies with mitochondria
or lysosomes, respectively. Colocalization is shown in yellow in the
merged image. Scale bars represent 20 μm.
## Mitochondrial Targeted Activity
Mitochondrial Targeted Activity Given the high degree
of accumulation of Ir-Me in mitochondria, the potential
of complexes Ir-H , Ir-Me , Ru - H , and Ru-Me and of metallopeptides Ir-Bn and Ru-BN to induce mitochondrial dysfunction
was next investigated. Mitochondria are the primary source of ATP
and anabolites for cellular metabolism and are involved in the regulation
of key cellular functions, including redox status and cell signaling.
Consequently, they play a crucial role in both cell growth and cell
death regulation. 64 PC-3 cells were incubated
with the compounds at the corresponding IC 50,light for
4 h and subsequently exposed to blue light irradiation for 1 h. Subsequently,
mitochondria were labeled with MitoTracker Red CMXRos, a lipophilic
cationic red fluorescent dye that accumulates within healthy mitochondria
due to their negative mitochondrial membrane potential (MMP). 65 Cell nuclei were counterstained in blue using
Hoechst 33342 to localize the cells. Confocal microscopy images revealed
a strong red fluorescence emission from mitochondria in untreated
control cells ( Figure 12 A). Conversely, exposure of cells to the photoactivated complexes
and metallopeptides led to a notable attenuation of the mitochondrial
red fluorescence, which was particularly evident in the case of the
complex Ir-H and of the metallopeptides Ir-BN and Ru-BN . These results strongly suggest that these
complexes exert their photodynamic activity by inducing mitochondrial
membrane depolarization, a hallmark of mitochondrial dysfunction. 65 Figure 12 Effect on mitochondria function. PC-3 cells were incubated
with
the indicated compounds at the corresponding IC 50,light for 4 h at 37 °C and then exposed to blue light for 1 h. Cells
incubated with the medium alone served as the negative control. (A)
Confocal microscopy images of the cells. Nuclei were stained blue
with Hoechst (λ ex : 350 nm; λ em :
461 nm), and mitochondria were labeled with MitoTracker Red CMXRos
(λ ex : 579 nm; λ em : 599 nm). Insets
show a 3× magnification of the cells. Scale bars represent 50
μm in 1× images and 20 μm in 3× images. (B)
Flow cytometry analysis of the effect of the complexes on mitochondrial
membrane potential. The percentage of cells exhibiting JC-10 green
(λ em : 529 nm) and orange fluorescence (λ em : 590 nm) is represented (mean ± SD of three independent
experiments). A decrease in the percentage of cells emitting orange
fluorescence indicates a loss of MMP. p < 0.05
and ** p < 0.01 compared to control cells. Flow cytometry analysis using the mitochondrial
specific dye JC-10
confirmed this effect. The fluorescence of JC-10 undergoes a reversible
change from green to orange as MMP increases, due to the formation
of JC-10 aggregates within polarized mitochondria. This property allows
for the simultaneous detection of healthy and depolarized mitochondria.
In control cells, orange fluorescence emission corresponding to healthy
mitochondria was detected in 63.0 ± 10.6% of cells ( Figure 12 B). However, treatment
with the photoactivated compounds significantly reduced the population
of cells emitting orange fluorescence to 16.6 ± 9.2%, 21.2 ±
4.9%, and 16.0 ± 9.5% for Ir-H , Ir-Me , and Ir-BN , respectively, and to 28.2 ± 7.8%,
19.7 ± 7.6%, and 23.0 ± 10.4% for Ru-H , Ru-Me , and Ru-BN , respectively ( Figure 12 B), confirming mitochondrial
membrane depolarization. Consistent with the microscopy experiments,
the most notable effect on MMP was observed with complex Ir-H . Ir-BN and Ru-BN metallopeptides also
induced significant mitochondrial membrane depolarization, indicating
that conjugation with BN3 does not hinder this effect.
Furthermore, the complexes’ ability to photocatalytically oxidize
NADH, a primary electron donor for the mitochondrial electron transport
chain, may contribute to the mitochondrial damage.
## Cell Death
Mechanism
Cell Death
Mechanism Mitochondria play an important
role in the regulation of apoptosis. Alterations in the MMP can cause
the release of apoptotic factors, such as cytochrome c, which activates
the caspase cascade, leading to programmed cell death. 66 , 67 In order to elucidate whether the toxic effect of the compounds
at the mitochondrial level triggers cell death by apoptosis, a dual
annexin V/propidium iodide labeling experiment was performed. The
initial stages of apoptosis are characterized by alterations in the
symmetry of phospholipids in the cytoplasmic membrane, which can be
detected using annexin V (An). The late stages also include disruption
of the cell membrane, permitting the penetration of propidium iodide
(PrI), which emits red fluorescence when bound to DNA. In contrast,
the cell membrane of necrotic cells becomes readily permeable to PrI
but does not exhibit phospholipid translocations. This allows the
discrimination among viable cells (An–/PrI−), necrotic
cells (An–/PrI+), and early-(An+/PrI−) and late-stage
(An+/PrI+) apoptotic cells. PC-3 cells were treated with the photoactivated
complexes and metallopeptides at five times the corresponding IC 50,light , and 24 h later, samples were analyzed using flow
cytometry. In all cases, the treatment resulted in a significant increase
in the number of apoptotic cells, particularly in the late stage of
apoptosis, compared to control cells ( Figure 13 ). A higher percentage of apoptotic cells
was detected upon treatment with Ir(III) complexes than Ru(II) complexes,
reaching a 11.9% of cells in early apoptosis and 41.7% of cells in
late apoptosis in the case of Ir-Me . In contrast, the
population of necrotic cells was not increased by any treatment. Overall,
these results indicated that the photodynamic activity of both the
complexes and metallopeptides promotes a regulated cell death by apoptosis. Figure 13 Cell
death mechanism. PC-3 cells were treated with the photoactivated
compounds at five times the corresponding IC 50,light . After
24 h, the percentage of viable, necrotic, early apoptotic, and late
apoptotic cells was analyzed by flow cytometry using propidium iodide
and annexin V staining. Untreated cells were used as controls. Bars
represent the mean ± standard deviation from three independent
experiments. * p < 0.05 and ** p < 0.01 versus control cells.
## Conclusions
Conclusions In conclusion, we have proved that the
conjugation of Ru(II)- or
Ir(III)-based PSs to the bombesin derivative BN3 can
be employed successfully to develop efficient PDT agents with two
possible levels of selectivity in their anticancer action: (1) the
first level would be provided by the targeting ability of BN3 toward cancer cells that overexpress BN receptors; (2) the second
level is based on the local activation of the metal fragments of our
PSs upon photoirradiation of the tumors. Our Ru(II) and Ir(III)
complexes were designed to combine remarkable
photocatalytic properties in the generation of ROS and suitable functional
groups on the N^N’ ligand for either enable peptide conjugation
(−COOH) or facilitate the cellular internalization (−COOMe).
Thus, following straightforward synthetic approaches, we have demonstrated
that it is possible to obtain the complexes Ru-H , Ru-Me , Ir - H , and Ir-Me and their respective metallopeptides Ru-BN and Ir-BN . Moreover, we have shown that the free complexes absorb
visible light efficiently and that they are phosphorescent, although
with low emission quantum yields. We also established that all the
metal complexes are photostable and that Ru-H and Ir-H adopt their deprotonated zwitterionic forms in the whole
range of physiological pHs. Regarding their biological properties,
our findings indicate that
Ir(III) PSs exhibit greater cytotoxicity than their Ru(II) congeners
in the dark against A549 and PC-3 cancer cells but also against nonmalignant
fibroblasts. In particular, Ir-Me was found to exhibit
the highest anticancer potency. Upon blue light irradiation, the cytotoxicity
of both Ir(III) PSs was significantly enhanced, with Ir-Me reaching PI values above 14 in cancer cells. Consistently, internalization
studies demonstrated that Ir-Me accumulated more efficiently
than Ir-H in the cells. Overall, these results revealed
that esterification with a methyl group avoids the formation of the
carboxylate group (−COO – ), increasing the
cellular internalization of the monocationic Ir-Me complex,
and therefore its cytotoxicity, relative to its zwitterionic congener, Ir-H . The Ru(II) PSs, Ru-H and Ru-Me , exhibited low cytotoxicity both in the dark and upon photoactivation,
which is consistent with their low lipophilicity and low cellular
internalization, with minor differences between them. We also
disclosed that binding of Ir-H to BN3 notably
enhances its selectivity toward cancer cells overexpressing
GRPR, as reflected by the lower IC 50 values of the resulting
metallopeptide Ir-BN in PC-3 cells compared to A549 cells,
which exhibit low GRPR expression, and importantly, to noncancerous
MRC-5 fibroblasts. Furthermore, conjugation of Ru-H to BN3 has been demonstrated to markedly enhance the accumulation
and photocytotoxic efficacy of the resulting metallopeptide Ru-BN i n PC-3 cancer cells, while exerting no toxicity on
noncancerous fibroblasts. The attenuation of the photocytotoxic activity
of Ir-BN and Ru-RN in the presence of bombesin
supports that the activity of the metallopeptides is at least partially
mediated by binding to GRPR. Hence, these results prove that conjugation
of the metal PSs to BN3 as tumor-targeting peptide is
a promising strategy in terms of selectivity and photocytotoxic activity. Clonogenic assays confirmed the light-dependent cytotoxic activity
of Ir-H , Ir-BN , Ru-H , and Ru-BN . Following blue light irradiation, the colony-forming
ability of treated cells was significantly reduced, indicating effective
photocytotoxicity. Hemolysis assays demonstrated that the complexes
are nontoxic to red blood cells, suggesting good blood biocompatibility.
These experiments also indicated that they do not cause damage to
cell membranes. Regarding the mechanism of action, experimental evidence
indicates that the complexes accumulate within mitochondria, where
they can generate 1 O 2 and O 2 •– and oxidize NADH in a photocatalytic manner.
This subsequently causes depolarization of the mitochondrial membrane
and the initiation of apoptosis. Overall, this study validates
the potential of exploiting specific
alterations in human cancer cells, such as overexpression of the bombesin
receptor, to design novel PDT agents with superior selectivity and
potency. Our findings indicate that BN3 could effectively
deliver metal-based PSs to tumor tissues, enhancing their accumulation
and minimizing off-target effects. This targeted approach holds significant
promise for the development of more effective and selective PDT strategies.
## Experimental Procedures
Experimental Procedures General Information All synthetic manipulations for
Ir(III) and Ru(II) complexes were carried out under an atmosphere
of dry, oxygen-free nitrogen using standard Schlenk techniques. The
solvents were dried and distilled under a nitrogen atmosphere before
use. Elemental analyses were performed with a Thermo Fisher Scientific
EA Flash 2000 Elemental Microanalyzer. UV–vis absorption was
measured in a Jasco V-750 UV–visible spectrophotometer. Fluorescence
steady-state and lifetime measurements were performed in an FLS980
(Edinburgh Instruments) fluorimeter with Xenon Arc Lamp 450 W and
TCSPC laser, respectively. Photoluminescence quantum yields was determined
by using FLS980 (Edinburgh Instruments) with Xenon Arc Lamp 450 W
and Red PMT Sphere as a detector. HR-ESI(+) mass spectra were recorded
with an Agilent LC-MS system (1260 Infinity LC/6545 Q-TOF MS spectrometer)
using dichloromethane (DCM) as a sample solvent and H 2 O
(0.1% formic acid)/MeOH (0.1% formic acid) and 30:70 as the mobile
phase. The experimental m / z values
are expressed in Da and were compared with the m / z values for monoisotopic fragments. NMR spectra were recorded
at 298 K on Bruker Avance III (300.130 MHz for 1 H; 75.468
MHz for 13 C). 1 H NMR spectra were acquired with
32 scans into 32 k data points over a spectral width of 16 ppm. 1 H and 13 C{ 1 H} chemical shifts were internally
referenced to TMS via the residual 1 H and 13 C signals of DMSO- d 6 (δ = 2.50
ppm and δ = 39.52 ppm), according to the values reported by
Fulmer et al. 68 Chemical shift values (δ)
are reported in ppm and coupling constants (J) in hertz. The splitting
of proton resonances in the reported 1 H NMR data is defined
as s = singlet, d = doublet, t = triplet, q = quartet, m = multiplet,
and bs = broad singlet. 2D NMR spectra were recorded using standard
pulse sequences. All NMR data processing was carried out using MestReNova
version 10.0.2. Cell Culture The PC-3 human prostate
cancer, A549 basal
lung adenocarcinoma, and MRC-5 lung fibroblast cell lines were obtained
from the American Type Culture Collection (ATCC). Cells were cultured
in Dulbecco’s modified Eagle’s medium (DMEM) (Corning)
supplemented with 10% fetal bovine serum (FBS) (Gibco-BRL) and 1% l -glutamine-penicillin-streptomycin (Cultek). Cells were maintained
in a Heracell 150 incubator (Thermofisher Scientific) at 37 °C
in a 5% CO 2 atmosphere. Mycoplasma contamination was regularly
monitored with the Mycoplasma Gel Detection Kit (Biotools). Photocytotoxic
Activity Stock solutions of the complexes
and metallopeptides were initially prepared at a concentration of
5 mM in DMSO and then diluted in sterile distilled water to achieve
a final concentration of 1 mM, with a resulting DMSO concentration
of 20% v/v. Cells were seeded into 96-well plates at a density of
2500 cells/well for A549, 3500 cells/well for PC-3, and 5500 cells/well
for MRC-5. Following 24 h of incubation for attachment, cells were
treated in triplicate with freshly prepared working solutions of the
compounds in the culture medium at concentrations ranging from 0.001
to 100 μM. The maximum DMSO concentration in the working solutions
was 2% (v/v). The working concentrations for each compound were adjusted
based on their cytotoxicity. Following a 4 h incubation period to
allow internalization of the compounds into the cells, the plates
were kept in dark conditions or exposed to blue (460 nm) or red (655
nm) light for 1 h using an LED system (LuxLight), providing a total
dose of 24.1 J cm –2 . All plates were incubated in
the dark for an additional 43 h. Cells were then washed with phosphate-buffered
saline (PBS), and 100 μL of fresh culture medium containing
10% of 3-[4,5-dimethylthiazol-2-yl]-2,5 diphenyl tetrazolium bromide
(MTT) solution (0.5 mg/mL) (Sigma- Aldrich) was added to each well.
After a 2 h incubation period, the formazan crystals were dissolved
in DMSO, and the absorbance was measured at 570 nm using a Multiscan
Plate Reader (Synergy 4, Biotek, Winooski, USA). The concentration
causing a 50% reduction in cell viability (IC 50 ) was determined
for each compound with the Gen5 software (BioTek). At least three
independent experiments were conducted for each compound. For
bombesin receptor blocking experiments, PC-3 cells seeded in 96-well
plates were treated with bombesin (Thermo Scientific) at increasing
concentrations (0, 10, 50, and 100 μM) for 30 min, followed
by the addition of Ir-BN or Ru-BN at final
concentrations of 10 or 5 μM, respectively. For each bombesin
concentration, control cells without metallopeptide treatment were
included to account for a possible effect of bombesin on cell proliferation.
After 4 h of incubation, the treatments were removed, and the cells
were exposed to blue light for 1 h. 43 h later, MTT assays were carried
out. The effects of the treatments were calculated by comparing the
absorbance of treated cells with that of cells exposed to the same
concentration of bombesin without metallopeptides. Each treatment
was performed in duplicate, and two independent experiments were performed. Clonogenic Assay PC-3 cells were seeded at a density
of 100000 cells per well in 12-well plates and allowed to adhere for
24 h. The cells were treated with the compounds at their respective
IC 50,light for 4 h and subsequently irradiated with blue
light or kept in the dark for one hour. The treatments were then removed,
and the cells were washed with PBS, harvested by trypsinization, and
counted using a Novocyte flow cytometer (Agilent Technologies). Subsequently,
3000 cells were seeded in 5 cm culture dishes and incubated for 10
days to allow colonies to form. A colony was defined to consist of
at least 50 cells. Cells treated with cisplatin (5 μM) were
used as a positive control, and cells treated with the culture medium
alone were used as a negative control. Dishes were washed with PBS,
and cells were fixed and stained with 1% methylene blue in 70% ethanol.
Images of the dishes were obtained with the Alpha Innotech Imaging
System (Alpha Innotech), and colony counting was performed using Fiji
ImageJ software. Each compound was evaluated in triplicate. Hemolysis
Assay Hemolysis assays were performed using
commercially available porcine blood preserved in sodium polyphosphate
as an anticoagulant (Norfrisa, Spain). Blood was diluted with PBS
to a concentration of 5%, and red blood cells (RBCs) were obtained
by centrifugation. Subsequently, 150 μL of the RBC suspension
was incubated with 150 μL of each compound at its respective
IC 50,light for 4 h, followed by 1 h of incubation under
blue light irradiation or in the dark with agitation on an orbital
shaker. Samples treated with PBS were used as negative controls, and
a solution of PBS with 0.2% Tween was used as a positive control to
induce 100% RBC lysis. All treatments were performed in duplicates.
The samples were then centrifuged, and 80 μL of the supernatant
was diluted with an equal volume of water and added to a 96-well plate.
Hemoglobin release in the supernatant was measured at 540 nm using
a Synergy 4 plate reader (Biotek). The optical density (OD) values
obtained from samples treated with the compounds (ODtest) were normalized
relative to the positive (ODpos) and negative (ODneg) control samples
to obtain the hemolysis ratio (HR) using the following equation: ROS Generation PC-3 cells were seeded at a density
of 100000 cells per well in 12-well plates and incubated overnight.
The compounds were added to the cells at their respective IC 50,light and incubated for 4 h to allow internalization. The treatments were
removed, and after washing with PBS, the 2′,7′-dichlorodihydrofluorescein
diacetate (H 2 DCFDA) probe (Sigma-Aldrich) at 10 μM
was added to each well. Cells were then irradiated with blue light
(24.1 J cm –2 ) or kept in the dark for 1 h. Cells
were subsequently harvested by trypsinization, and the median fluorescence
emission of 10000 cells was measured using a Novocyte flow cytometer
equipped with NovoExpress software. The fluorescence fold increase
relative to untreated control cells was calculated. Each experiment
was performed in triplicate for each complex and cell line. A similar protocol was followed to evaluate superoxide production
using the ROS-ID Superoxide Detection Kit (Enzo Life Sciences) according
to the manufacturer’s instructions. Cellular Internalization PC-3 cells were seeded in
6-well plates at a density of 2 million cells/well. After 24 h, cells
were incubated with the compounds at a concentration of 5 μM
for 4 h. Untreated cells were used as a negative control. Cells were
washed with PBS and harvested by trypsinization. The number of cells
in each sample was determined using a Novocyte flow cytometer (Agilent
Technologies). The samples were centrifuged to obtain the cell pellet,
and the complex content was subsequently assessed using inductively
coupled plasma mass spectrometry (ICP-MS). To this end, the cell pellets
were dissolved in 400 μL of 69% v/v concentrated nitric acid
and heated at 60 °C overnight. The digested samples were allowed
to cool and diluted until 5 mL with Milli-Q water. The Ir or Ru content
was analyzed in Agilent 7500c ICP-MS located at the Technical Research
Services of the University of Girona. Standards were freshly prepared
in Milli-Q water containing the same proportion of HNO 3 (8%) before each experiment. The concentrations used for the calibration
curve were approximately 0, 3, 9, 17, 35, 66, and 100 μg.kg –1 . Rhodium was added to all samples and standards as
the internal standard at a concentration around 10 μg.kg –1 . The isotopes detected were 193 Ir, 101 Ru, and 103 Rh, respectively. Measures were carried
out in triplicate. 193 Ir/ 103 Rh and 101 Ru / 103 Rh signal ratios were corrected with the real exact
Rh concentration in each sample and standard. The amount of metal
in each sample was normalized to the cell number. Three independent
samples were analyzed for each complex. Confocal Microscopy The subcellular distribution of
the complexes was assessed by confocal fluorescence microscopy. A549
cells were selected for these experiments as they have an extended
cytoplasm, which facilitates the visualization of different organelles.
Cells were plated on glass-bottom 8-well chamber slides (Ibidi) at
a density of 50000 cells per well. After 24 h, the cells were treated
with Ir-Me at 1 μM in DMEM without phenol red.
Untreated cells were used as the negative control. To assess the colocalization
with specific organelles, MitoView Green (Biotium) (excitation/emission:
490/523 nm) and LysoTracker Green DND-26 (Thermo Fisher Scientific)
(excitation/emission: 504/511 nm) dyes were used at a concentration
of 100 nM to label mitochondria and lysosomes, respectively. After
1 h of incubation at 37 °C, cells were washed with cold PBS and
immediately imaged using a Nikon A1R confocal microscope. Images were
analyzed using NIS-Elements AR (Nikon, Japan) and ImageJ software.
Colocalization was assessed by the Pearson correlation coefficient,
using the JACoP plugin. 69 Mitochondrial
Damage Confocal microscopy: PC-3 cells
were plated on glass-bottom 8-well chamber slides at a density of
75000 cells per well. 24 h later, the cells were treated with the
compounds at their respective IC 50,light for 4 h, followed
by exposure to blue light for 1 h or incubation in the dark. Untreated
cells were used as the negative control. The cells were then rinsed
three times with PBS and incubated with the MitoTracker Red CMXRos
dye (Molecular Probes) (excitation/emission: 579/599 nm) at a concentration
of 200 nM in phenol red-free DMEM for 30 min at 37 °C. Cell nuclei
were stained blue using Hoechst 33342 dye (Invitrogen) (excitation/emission:
350/461 nm) diluted 1:4000. Images were captured using a Nikon A1R
confocal microscope and analyzed using ImageJ software. Flow
cytometry assessment of the mitochondrial membrane depolarization:
PC-3 cells were seeded on 12-well plates at a density of 100000 cells
per well. 24 h later, cells were treated with the compounds as described
above. Cells were subsequently harvested using trypsinization and
incubated with JC-10 dye (Deltaclon) according to the manufacturer’s
protocol. A Novocyte flow cytometer was employed to analyze the fluorescence
emission of JC-10 in 10000 cells. Fluorescence was detected at 590
nm (FL2) to identify the percentage of cells with healthy mitochondria
and at 529 nm (FL1) to determine the percentage of cells with depolarized
mitochondria. Each compound was evaluated in three independent experiments,
and the mean and standard deviation of the results were represented. Apoptosis Assays PC-3 cells were seeded into 12-well
plates at a density of 100000 cells/well. After 24 h, cells were treated
with compounds at a concentration five times the corresponding IC 50 under blue light irradiation. Cisplatin at 25 μM was
used as the positive control. 24 h later, cells were harvested by
trypsinization and stained with the Vybrant Apoptosis Assay Kit (Molecular
Probes) following the manufacturer’s protocol. The samples
were immediately analyzed using a Novocyte flow cytometer. Annexin-FITC
staining was detected at a wavelength of 520 nm, and propidium iodide
was detected at 617 nm. The fluorescence emission of 10000 cells per
sample was measured, and the percentages of live, early apoptotic,
late apoptotic, and necrotic cell populations were determined.
## General Information
General Information All synthetic manipulations for
Ir(III) and Ru(II) complexes were carried out under an atmosphere
of dry, oxygen-free nitrogen using standard Schlenk techniques. The
solvents were dried and distilled under a nitrogen atmosphere before
use. Elemental analyses were performed with a Thermo Fisher Scientific
EA Flash 2000 Elemental Microanalyzer. UV–vis absorption was
measured in a Jasco V-750 UV–visible spectrophotometer. Fluorescence
steady-state and lifetime measurements were performed in an FLS980
(Edinburgh Instruments) fluorimeter with Xenon Arc Lamp 450 W and
TCSPC laser, respectively. Photoluminescence quantum yields was determined
by using FLS980 (Edinburgh Instruments) with Xenon Arc Lamp 450 W
and Red PMT Sphere as a detector. HR-ESI(+) mass spectra were recorded
with an Agilent LC-MS system (1260 Infinity LC/6545 Q-TOF MS spectrometer)
using dichloromethane (DCM) as a sample solvent and H 2 O
(0.1% formic acid)/MeOH (0.1% formic acid) and 30:70 as the mobile
phase. The experimental m / z values
are expressed in Da and were compared with the m / z values for monoisotopic fragments. NMR spectra were recorded
at 298 K on Bruker Avance III (300.130 MHz for 1 H; 75.468
MHz for 13 C). 1 H NMR spectra were acquired with
32 scans into 32 k data points over a spectral width of 16 ppm. 1 H and 13 C{ 1 H} chemical shifts were internally
referenced to TMS via the residual 1 H and 13 C signals of DMSO- d 6 (δ = 2.50
ppm and δ = 39.52 ppm), according to the values reported by
Fulmer et al. 68 Chemical shift values (δ)
are reported in ppm and coupling constants (J) in hertz. The splitting
of proton resonances in the reported 1 H NMR data is defined
as s = singlet, d = doublet, t = triplet, q = quartet, m = multiplet,
and bs = broad singlet. 2D NMR spectra were recorded using standard
pulse sequences. All NMR data processing was carried out using MestReNova
version 10.0.2.
## Cell Culture
Cell Culture The PC-3 human prostate
cancer, A549 basal
lung adenocarcinoma, and MRC-5 lung fibroblast cell lines were obtained
from the American Type Culture Collection (ATCC). Cells were cultured
in Dulbecco’s modified Eagle’s medium (DMEM) (Corning)
supplemented with 10% fetal bovine serum (FBS) (Gibco-BRL) and 1% l -glutamine-penicillin-streptomycin (Cultek). Cells were maintained
in a Heracell 150 incubator (Thermofisher Scientific) at 37 °C
in a 5% CO 2 atmosphere. Mycoplasma contamination was regularly
monitored with the Mycoplasma Gel Detection Kit (Biotools).
## Photocytotoxic
Activity
Photocytotoxic
Activity Stock solutions of the complexes
and metallopeptides were initially prepared at a concentration of
5 mM in DMSO and then diluted in sterile distilled water to achieve
a final concentration of 1 mM, with a resulting DMSO concentration
of 20% v/v. Cells were seeded into 96-well plates at a density of
2500 cells/well for A549, 3500 cells/well for PC-3, and 5500 cells/well
for MRC-5. Following 24 h of incubation for attachment, cells were
treated in triplicate with freshly prepared working solutions of the
compounds in the culture medium at concentrations ranging from 0.001
to 100 μM. The maximum DMSO concentration in the working solutions
was 2% (v/v). The working concentrations for each compound were adjusted
based on their cytotoxicity. Following a 4 h incubation period to
allow internalization of the compounds into the cells, the plates
were kept in dark conditions or exposed to blue (460 nm) or red (655
nm) light for 1 h using an LED system (LuxLight), providing a total
dose of 24.1 J cm –2 . All plates were incubated in
the dark for an additional 43 h. Cells were then washed with phosphate-buffered
saline (PBS), and 100 μL of fresh culture medium containing
10% of 3-[4,5-dimethylthiazol-2-yl]-2,5 diphenyl tetrazolium bromide
(MTT) solution (0.5 mg/mL) (Sigma- Aldrich) was added to each well.
After a 2 h incubation period, the formazan crystals were dissolved
in DMSO, and the absorbance was measured at 570 nm using a Multiscan
Plate Reader (Synergy 4, Biotek, Winooski, USA). The concentration
causing a 50% reduction in cell viability (IC 50 ) was determined
for each compound with the Gen5 software (BioTek). At least three
independent experiments were conducted for each compound. For
bombesin receptor blocking experiments, PC-3 cells seeded in 96-well
plates were treated with bombesin (Thermo Scientific) at increasing
concentrations (0, 10, 50, and 100 μM) for 30 min, followed
by the addition of Ir-BN or Ru-BN at final
concentrations of 10 or 5 μM, respectively. For each bombesin
concentration, control cells without metallopeptide treatment were
included to account for a possible effect of bombesin on cell proliferation.
After 4 h of incubation, the treatments were removed, and the cells
were exposed to blue light for 1 h. 43 h later, MTT assays were carried
out. The effects of the treatments were calculated by comparing the
absorbance of treated cells with that of cells exposed to the same
concentration of bombesin without metallopeptides. Each treatment
was performed in duplicate, and two independent experiments were performed.
## Clonogenic Assay
Clonogenic Assay PC-3 cells were seeded at a density
of 100000 cells per well in 12-well plates and allowed to adhere for
24 h. The cells were treated with the compounds at their respective
IC 50,light for 4 h and subsequently irradiated with blue
light or kept in the dark for one hour. The treatments were then removed,
and the cells were washed with PBS, harvested by trypsinization, and
counted using a Novocyte flow cytometer (Agilent Technologies). Subsequently,
3000 cells were seeded in 5 cm culture dishes and incubated for 10
days to allow colonies to form. A colony was defined to consist of
at least 50 cells. Cells treated with cisplatin (5 μM) were
used as a positive control, and cells treated with the culture medium
alone were used as a negative control. Dishes were washed with PBS,
and cells were fixed and stained with 1% methylene blue in 70% ethanol.
Images of the dishes were obtained with the Alpha Innotech Imaging
System (Alpha Innotech), and colony counting was performed using Fiji
ImageJ software. Each compound was evaluated in triplicate.
## Hemolysis
Assay
Hemolysis
Assay Hemolysis assays were performed using
commercially available porcine blood preserved in sodium polyphosphate
as an anticoagulant (Norfrisa, Spain). Blood was diluted with PBS
to a concentration of 5%, and red blood cells (RBCs) were obtained
by centrifugation. Subsequently, 150 μL of the RBC suspension
was incubated with 150 μL of each compound at its respective
IC 50,light for 4 h, followed by 1 h of incubation under
blue light irradiation or in the dark with agitation on an orbital
shaker. Samples treated with PBS were used as negative controls, and
a solution of PBS with 0.2% Tween was used as a positive control to
induce 100% RBC lysis. All treatments were performed in duplicates.
The samples were then centrifuged, and 80 μL of the supernatant
was diluted with an equal volume of water and added to a 96-well plate.
Hemoglobin release in the supernatant was measured at 540 nm using
a Synergy 4 plate reader (Biotek). The optical density (OD) values
obtained from samples treated with the compounds (ODtest) were normalized
relative to the positive (ODpos) and negative (ODneg) control samples
to obtain the hemolysis ratio (HR) using the following equation:
## ROS Generation
ROS Generation PC-3 cells were seeded at a density
of 100000 cells per well in 12-well plates and incubated overnight.
The compounds were added to the cells at their respective IC 50,light and incubated for 4 h to allow internalization. The treatments were
removed, and after washing with PBS, the 2′,7′-dichlorodihydrofluorescein
diacetate (H 2 DCFDA) probe (Sigma-Aldrich) at 10 μM
was added to each well. Cells were then irradiated with blue light
(24.1 J cm –2 ) or kept in the dark for 1 h. Cells
were subsequently harvested by trypsinization, and the median fluorescence
emission of 10000 cells was measured using a Novocyte flow cytometer
equipped with NovoExpress software. The fluorescence fold increase
relative to untreated control cells was calculated. Each experiment
was performed in triplicate for each complex and cell line. A similar protocol was followed to evaluate superoxide production
using the ROS-ID Superoxide Detection Kit (Enzo Life Sciences) according
to the manufacturer’s instructions.
## Cellular Internalization
Cellular Internalization PC-3 cells were seeded in
6-well plates at a density of 2 million cells/well. After 24 h, cells
were incubated with the compounds at a concentration of 5 μM
for 4 h. Untreated cells were used as a negative control. Cells were
washed with PBS and harvested by trypsinization. The number of cells
in each sample was determined using a Novocyte flow cytometer (Agilent
Technologies). The samples were centrifuged to obtain the cell pellet,
and the complex content was subsequently assessed using inductively
coupled plasma mass spectrometry (ICP-MS). To this end, the cell pellets
were dissolved in 400 μL of 69% v/v concentrated nitric acid
and heated at 60 °C overnight. The digested samples were allowed
to cool and diluted until 5 mL with Milli-Q water. The Ir or Ru content
was analyzed in Agilent 7500c ICP-MS located at the Technical Research
Services of the University of Girona. Standards were freshly prepared
in Milli-Q water containing the same proportion of HNO 3 (8%) before each experiment. The concentrations used for the calibration
curve were approximately 0, 3, 9, 17, 35, 66, and 100 μg.kg –1 . Rhodium was added to all samples and standards as
the internal standard at a concentration around 10 μg.kg –1 . The isotopes detected were 193 Ir, 101 Ru, and 103 Rh, respectively. Measures were carried
out in triplicate. 193 Ir/ 103 Rh and 101 Ru / 103 Rh signal ratios were corrected with the real exact
Rh concentration in each sample and standard. The amount of metal
in each sample was normalized to the cell number. Three independent
samples were analyzed for each complex.
## Confocal Microscopy
Confocal Microscopy The subcellular distribution of
the complexes was assessed by confocal fluorescence microscopy. A549
cells were selected for these experiments as they have an extended
cytoplasm, which facilitates the visualization of different organelles.
Cells were plated on glass-bottom 8-well chamber slides (Ibidi) at
a density of 50000 cells per well. After 24 h, the cells were treated
with Ir-Me at 1 μM in DMEM without phenol red.
Untreated cells were used as the negative control. To assess the colocalization
with specific organelles, MitoView Green (Biotium) (excitation/emission:
490/523 nm) and LysoTracker Green DND-26 (Thermo Fisher Scientific)
(excitation/emission: 504/511 nm) dyes were used at a concentration
of 100 nM to label mitochondria and lysosomes, respectively. After
1 h of incubation at 37 °C, cells were washed with cold PBS and
immediately imaged using a Nikon A1R confocal microscope. Images were
analyzed using NIS-Elements AR (Nikon, Japan) and ImageJ software.
Colocalization was assessed by the Pearson correlation coefficient,
using the JACoP plugin. 69
## Mitochondrial
Damage
Mitochondrial
Damage Confocal microscopy: PC-3 cells
were plated on glass-bottom 8-well chamber slides at a density of
75000 cells per well. 24 h later, the cells were treated with the
compounds at their respective IC 50,light for 4 h, followed
by exposure to blue light for 1 h or incubation in the dark. Untreated
cells were used as the negative control. The cells were then rinsed
three times with PBS and incubated with the MitoTracker Red CMXRos
dye (Molecular Probes) (excitation/emission: 579/599 nm) at a concentration
of 200 nM in phenol red-free DMEM for 30 min at 37 °C. Cell nuclei
were stained blue using Hoechst 33342 dye (Invitrogen) (excitation/emission:
350/461 nm) diluted 1:4000. Images were captured using a Nikon A1R
confocal microscope and analyzed using ImageJ software. Flow
cytometry assessment of the mitochondrial membrane depolarization:
PC-3 cells were seeded on 12-well plates at a density of 100000 cells
per well. 24 h later, cells were treated with the compounds as described
above. Cells were subsequently harvested using trypsinization and
incubated with JC-10 dye (Deltaclon) according to the manufacturer’s
protocol. A Novocyte flow cytometer was employed to analyze the fluorescence
emission of JC-10 in 10000 cells. Fluorescence was detected at 590
nm (FL2) to identify the percentage of cells with healthy mitochondria
and at 529 nm (FL1) to determine the percentage of cells with depolarized
mitochondria. Each compound was evaluated in three independent experiments,
and the mean and standard deviation of the results were represented.
## Apoptosis Assays
Apoptosis Assays PC-3 cells were seeded into 12-well
plates at a density of 100000 cells/well. After 24 h, cells were treated
with compounds at a concentration five times the corresponding IC 50 under blue light irradiation. Cisplatin at 25 μM was
used as the positive control. 24 h later, cells were harvested by
trypsinization and stained with the Vybrant Apoptosis Assay Kit (Molecular
Probes) following the manufacturer’s protocol. The samples
were immediately analyzed using a Novocyte flow cytometer. Annexin-FITC
staining was detected at a wavelength of 520 nm, and propidium iodide
was detected at 617 nm. The fluorescence emission of 10000 cells per
sample was measured, and the percentages of live, early apoptotic,
late apoptotic, and necrotic cell populations were determined.