Biotinylated Platinum(II) Ferrocenylterpyridine Complexes for Targeted Photoinduced Cytotoxicity

Article abstract of DOI:10.1021/acs.inorgchem.6b00680

Biotinylated platinum(II) ferrocenylterpyridine (Fc-tpy) complexes [Pt(Fc-tpy)(L¹)]Cl (1) and [Pt(Fc-tpy)(L²)]Cl (2), where HL¹ and HL² are biotin-containing ligands, were prepared, and their targeted photoinduc

Full text of DOI:10.1021/acs.inorgchem.6b00680

Article
pubs.acs.org/IC
Biotinylated Platinum(II) Ferrocenylterpyridine Complexes for
Targeted Photoinduced Cytotoxicity
Koushambi Mitra,† Abhijith Shettar,‡ Paturu Kondaiah,*,‡ and Akhil R. Chakravarty*,†
†Department of Inorganic and Physical Chemistry and ‡Department of Molecular Reproduction, Development and Genetics, Indian
Institute of Science, Bangalore 560012, India
S
* Supporting Information
ABSTRACT: Biotinylated platinum(II) ferrocenylterpyridine
(Fc-tpy) complexes [Pt(Fc-tpy)(L1)]Cl (1) and [Pt(Fc-
tpy)(L2)]Cl (2), where HL1 and HL2 are biotin-containing
ligands, were prepared, and their targeted photoinduced
cytotoxic effect in cancer cells over normal cells was studied.
A nonbiotinylated complex, [Pt(Fc-tpy)(L3)]Cl (3), was
prepared as a control to study the role of the biotin moiety
in cellular uptake properties of the complexes. Three
platinum(II) phenylterpyridine (Ph-tpy) complexes, viz.,
[Pt(Ph-tpy)(L1)]Cl (4), [Pt(Ph-tpy)(L2)]Cl (5), and [Pt-
(Ph-tpy)(L3)]Cl (6), were synthesized and explored to
understand the role of a metal-bound Fc-tpy ligand over Ph-
tpy as a photoinitiator. The Fc-tpy complexes displayed an
intense absorption band near 640 nm, which was absent in their Ph-tpy analogues. The Fc-tpy complexes (1 mM in 0.1 M
TBAP) showed an irreversible cyclic voltammetric anodic response of the Fc/Fc+ couple near 0.25 V. The Fc-tpy complexes
displayed photodegradation in red light of 647 nm involving the formation of a ferrocenium ion (Fc+) and reactive oxygen
species (ROS). Photoinduced release of the biotinylated ligands was observed from spectral measurements, and this possibly led
to the controlled generation of an active platinum(II) species, which binds to the calf-thymus DNA used for this study. The
biotinylated photoactive Fc-tpy complexes showed significant photoinduced cytotoxicity, giving a IC50 value of ∼7 μM in visible
light of 400−700 nm with selective uptake in BT474 cancer cells over HBL-100 normal cells. Furthermore, ferrocenyl complexes
resulted in light-induced ROS-mediated apoptosis, as indicated by DCFDA, annexin V/FITC staining, and sub-G1 DNA content
determined by fluorescent activated cell sorting analysis. The phenyl analogues 4 and 5 were photostable, served as DNA
intercalators, and demonstrated selective cytotoxicity in the cancer cells, giving IC50 values of ∼4 μM.
healthy cells.6−8 DDS reduces both the drug dosage and the
amount of drug circulating in body fluids. DDS can be suitably
1. INTRODUCTION
Site-directed delivery and controlled activation of an anticancer
designed with the knowledge that cancer cells overexpress
agent are the major challenges to potentiate its clinical
receptors to aid internalization of growth-promoting factors like
applications. The conventional treatment modalities such as
chemotherapy or radiotherapy suffer from high systemic
toxicity along with the associated risks of metastatic cancer
growth. Photodynamic therapy (PDT) has emerged as an
vitamins, glucose, peptides, antibodies, and hormones.9−12 For
example, drug conjugates with biotin, riboflavin, vitamin B12,
and folic acid are known to show cancer-cell-specific
uptake.13−15 We have chosen biotin, which is known as vitamin
alternate methodology in which a photosensitizer that is
H or vitamin B7 and is extensively used for tumor targeting, for
inactive and nontoxic in the dark selectively damages the
our study. In vitro and in vivo literature reports indicate that
photoexposed cancer cells, leaving the unexposed healthy cells
unaffected.1−5 In addition to selectivity, the efficacy of a PDT
agent can be increased by targeting cancer cells over normal
cells. Tumor cells with enhanced metabolic rates and leaky
vasculature have preferential accumulation of the photo-
uptake of biotin is facilitated by sodium-dependent multi-
vitamin transporters (SMVTs), which are overexpressed in
certain cancer cell lines.16−19 The strong affinity of biotin for
protein receptors like avidin and streptavidin (∼1014 M) is well
explored.20 Taxoids, gemcitabine, doxorubicin, annonaceous
sensitizers over normal cells. However, the presence of some
acetogenins, and p53 proteins, when conjugated to biotin, have
highly proliferating normal cells like red blood cells, gut
shown selective delivery to cancer cells without affecting their
cytotoxic effects.16,21,22 Kim and co-workers have reported
epithelia, bone marrow, and hair follicles enhances the
nonspecific uptake of the drug, thus requiring high drug
doses that lead to unwanted adverse effects. Thus, a targeted
drug-delivery system (DDS) assumes great importance for
augmented cellular uptake of the drug in cancer cells over
Received: March 18, 2016
© XXXX American Chemical Society
A
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several biotin−drug conjugates as tumor-targeted theranostic
and bioimaging probes.23−25
The present work stems from our interest to develop
platinum(II)-based PDT agents using a biotinylated acetylide
ligand for their targeted delivery to tumor cells rather than to
normal ones (Figure 1). We have chosen platinum as the metal
cytotoxicity of complex 1 in visible light (400−700 nm) in
human breast cancer cells (BT474) while remaining nontoxic in
the dark (IC50 value: 7 μM in light and >50 μM in the dark),
(ii) nominal toxicity in human normal breast cell lines HBL-
100 (IC50 in light: ∼45 μM), (iii) the Ph-tpy complexes
showing undesirable dark toxicity and no apparent PDT effect,
highlighting the importance of the Fc-tpy moiety in the
photoactivation process, (iv) the biotinylated Ph-tpy complex
showing significant cytotoxicity in the dark (IC50 value: ∼4
μM), indicating the importance of SMVT in enhancing the
cellular uptake, and (v) increased uptake of biotin-conjugated
complexes in cancer cells than in normal cells.
2. EXPERIMENTAL SECTION
Materials and Measurements. The chemicals and reagents were
procured from commercial sources. Dulbecco’s phosphate-buffered
saline (DPBS), calf-thymus (ct)-DNA, agarose (molecular biology
grade), catalase, superoxide dismutase (SOD), ethidium bromide
(EB), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide
(MTT), and 2′,7′-dichlorofluorescein diacetate (H2DCFH-DA) were
purchased from Sigma-Aldrich, USA. Potassium tetrachloroplatinate
and supercoiled (SC) pUC19 DNA (cesium chloride purified) were
obtained from Arora Matthey (India) and Bangalore Genie (India),
respectively. Tetrabutylammonium perchlorate (TBAP) was synthe-
sized using tetrabutylammonium bromide and perchloric acid. Ligands
HL1, 6-biotinylaminocaproic acid, and HL2 and the precursor
complexes [Pt(Fc-tpy)Cl]Cl and [Pt(Ph-tpy)Cl]Cl were synthesized
following reported methods.33,38,39 The ligands were synthesized using
amide-coupling reactions with anhydrous hydroxybenzotriazole
(HOBt) and 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide
(EDCI) as the coupling reagents. The crude product was column-
chromatographed to obtain pure ligands in moderate yields (∼40%).
Synthetic details and characterization data are given as Supporting
Information (Scheme S1 and Figures S1−S3).
Elemental analysis, high-performance liquid chromatography
(HPLC), UV−visible, emission, IR, molar conductivity, and mass
(MS) and NMR spectral measurements were done using equipment
that was reported earlier.34 Cyclic voltammograms were recorded
using an EG&G PAR model 253 VersaStat potentiostat/galvanostat
equipped with electrochemical analysis software 270. A three-electrode
setup, consisting of a glassy carbon working electrode, a platinum wire
counter electrode, and a saturated calomel (SCE) pseudoreference
electrode, was used [Caution! mercury and its salts, being highly toxic,
were handled with proper safety precautions]. Potentials were finally
corrected versus Fc/Fc+ by keeping ferrocene as an internal standard.
Solid-state emission spectra were recorded using an Edinburgh
FLS920 spectrometer. A Becton Dickinson fluorescent activated cell
sorting (BD-FACS) Verse configured with three lasers (405 nm, 40
mW; 488 nm, 20 mW; and 640 nm, 40 mW) and an eight-parameter
Windows 7 operating system with BD-FACS suite software for
acquisition and analysis was used for FACS analysis. An inductively
coupled plasma mass spectroscopy (ICP-MS) method was used to
evaluate the platinum content by ICP-MS, Thermo X series II. A
Waters Alliance System equipped with EMPOWER 3 software (Waters
Corp., Milford, MA; 2695 separation module, 2996 photodiode-array
detector) was employed to perform HPLC. The 30 μL solutions (5
mM) were run in aqueous methanol (MeOH) while the MeOH
percentage was maintained as 95% for 17 min and 5% for 8 min. A
reversed-phase column (Lichrosphere 60, RP-select B, 5 mm, flow rate
of 1 mL min−1) was used, and all of the samples were detected at 275
nm.
Figure 1. Molecular structures of complexes 1−6.
considering the successful use of this metal-based drug, viz.,
cisplatin, carboplatin, and oxaliplatin, as a novel chemo-
therapeutic agent.26−28 Although few platinum(IV) complexes
are used in photoactivated chemotherapy, the potential of
platinum(II) complexes in PDT is yet to be realized.29−35
Platinum complexes show excellent photophysical properties
with enhanced triplet-state population upon excitation due to
spin−orbit coupling, and photoactive platinum complexes
could serve as attractive candidates for PDT. In addition,
ligand dissociation from ruthenium and platinum complexes in
their excited states often results in dual photochemotherapeutic
effects, which offer an oxygen-independent mechanistic action
suitable for the treatment of hypoxic solid tumors.36,37 We have
earlier shown that platinum(II) ferrocenylterpyridine (Fc-tpy)
complexes are significantly photocytotoxic in cancer cells with
low dark toxicity.33 The Fc-tpy moiety upon binding to
platinum(II) behaves as an excellent photoinitiator within the
PDT spectral window (600−800 nm) and generates reactive
oxygen species (ROS), which causes cellular damage. With an
objective to develop the virtually unexplored PDT chemistry of
platinum(II) complexes, we have designed and synthesized new
ternary PtII(Fc-tpy) complexes of biotin-conjugated acetylide
ligands HL1 and HL2, viz., [Pt(Fc-tpy)(L1)]Cl (1) and [Pt(Fc-
tpy)(L2)]Cl (2), where HL1 is 6-[5-(2-oxohexahydro-1H-
thieno[3,4-d]imidazol-4-yl)pentanamido]-N-(prop-2-yn-1-yl)-
hexanamide and HL2 is 5-(2-oxohexahydro-1H-thieno[3,4-
d]imidazol-4-yl)-N-(prop-2-yn-1-yl)pentanamide (Figure 1).
Complex 1 has an acetylide biotin unit bound to the PtII(Fc-
tpy) moiety, while it is separated with a 6-aminocaproic acid
linker in complex 2. A nonbiotinylated complex, [Pt(Fc-
tpy)(L3)]Cl (3), where HL3 is N-(prop-2-yn-1-yl)acetamide,
was prepared to investigate the role of the biotin unit in 1 and
2. Three other complexes, viz., [Pt(Ph-tpy)(L1)]Cl (4),
[Pt(Ph-tpy)(L2)]Cl (5), and [Pt(Ph-tpy)(L3)]Cl (6), were
prepared and studied to understand the role of the Fc-tpy and
Ph-tpy ligands as photosensitizers in 1−6. The significant
results from this study include (i) impressive photoinduced
Synthesis of Complexes 1−6. The complexes were prepared by
following a general synthetic procedure. To a N,N-dimethylformamide
(DMF) solution (2 mL) of the ligand (HL1, 94 mg, 0.24 mmol for 1
and 118 mg, 0.3 mmol for 4; HL2, 67 mg, 0.24 mmol for 2 and 83 mg,
0.3 mmol for 5; HL3, 30 mg, 0.3 mmol for 3 and 40 mg, 0.41 mmol for
6) was added freshly distilled triethylamine (5 mL), and the solution
was stirred for 1 h under a nitrogen atmosphere. To this solution was
B
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added the precursor complex [Pt(Fc-tpy)Cl]Cl (0.08 g, 0.12 mmol)
for 1−3 and [Pt(Ph-tpy)Cl]Cl (0.08 g, 0.14 mmol) for 4−6, taken in
dry DMF (8 mL) along with a catalytic amount of CuI (5 mg). The
resulting mixture was stirred for 48 h in the dark and then filtered. The
desired complex was precipitated by adding diethyl ether, dried, and
reprecipitated from an acetonitrile (MeCN) solution by adding diethyl
ether to obtain the product in an analytically pure form. The
characterization data are given below.
3129 s, 3069 s, 2485 m, 2415 m, 2173 m, 2076 m, 2025 m, 1673 s,
1653 s, 1605 s, 1541 m, 1477 m, 1415 m, 1244 m, 1029 w, 900 w, 768
s, 681 s, 590 w, 472 w, 438, m. ΛM, S m2 M−1 in DMF at 25 °C: 69.
[Pt(Ph-tpy)(L3)]Cl (6). Yield: 88 mg, 92%. Anal. Calcd for
C26H21ClN4OPt (6): C, 49.10; H, 3.33; N, 8.81. Found: C, 48.96;
H, 3.49; N, 8.72. ESI-MS in MeOH: m/z 600.1364 ([M − Cl]+,
1
100%). H NMR (400 MHz, DMSO-d6): δ 9.03 (m, 2H), 8.83 (m,
4H), 8.49 (m, 2H), 8.21 (b, 3H), 7.95 (m, 2H), 7.83 (m, 2H), 7.69
(m, 1H), 4.12 (d, 2H, J = 4 Hz), 1.90 (s, 3H). UV−visible in 10%
DMSO−DPBS (pH = 7.2) [λmax, nm (ε, M−1 cm−1)]: 397 (3670), 330
(10300), 285 (18400), 260 (23300). IR data in the solid phase
(cm−1): 3328 s, 3272 s, 3231 s, 3051 m, 3002 m, 2160 s, 2104 m, 2025
m, 1645 s, 1606 s, 1528 m, 1476 m, 1415 s, 1248 m, 1029 m, 906 w,
768 s, 681 m, 592 m, 494 m. ΛM, S m2 M−1 in DMF at 25 °C: 73.
Solubility. The complexes were sparingly soluble in MeOH,
MeCN, and ethyl acetate. They were soluble in DMF and dimethyl
sulfoxide (DMSO). They were insoluble in diethyl ether and
hydrocarbon solvents.
Theoretical Calculations. The transitions in the visible region
were obtained by performing linear-response time-dependent density
functional theory (TDDFT) at the B3LYP/LanL2DZ level of theory
using the G09 suite programs.33,40−43
DNA Binding and Cleavage Experiments. ct-DNA (5 mL, 500
μM) was treated with complexes 1−6 (50 μM) in 10% DMSO−DPBS
(pH = 7.2). The platinum contents in the isolated DNA in the dark
and under photoirradiated conditions were evaluated by ICP-MS
methods. The detailed procedures for DNA melting and viscosity
experiments are provided in the Supporting Information.
SC pUC19 DNA (0.2 μg, 30 μM, 2686 base pairs) was
preincubated for 1 h with complexes 1−6 (20 μM in 10% DMF−
Tris-HCl buffer (50 mM), pH = 7.2, 37 °C) and then subjected to
photoexposure using an argon−krypton mixed-gas ion laser of 647 nm
for 1 h. The percentage of nicked-circular (NC) DNA was evaluated
using gel electrophoresis with a UVITEC Gel Documentation
System.44 The DNA photocleavage measurements were also done in
the presence of different additives such as ROS scavengers/quenchers
(NaN3, 4 mM; TEMP, 4 mM; DMSO, 8 μL; KI, 4 mM; catalase, 4
units; SOD, 4 units) using procedures that were reported earlier.34
MTT and DCFDA Assays. MTT, upon reduction by NAD(P)H-
dependent cellular oxidoreductase enzymes present in the cytoplasm
of metabolically active cells, forms formazan as a purple insoluble
product. The amount of formazan estimated by a spectral method
correlates with the number of live cells and provides a quantitative
measurement of the cytotoxicity of the compound. The IC50 values
were obtained from nonlinear regression using GraphPad Prism 5.
About 8000 cells of BT474 (human breast carcinoma) and HBL-100
(normal breast epithelial human cells) were preincubated for 24 h in
the dark with varying concentrations (50, 40, 30, 25, 20, 10, and 5 μM)
of complexes 1−6. Photoirradiation was done in a DPBS medium in
visible light of 400−700 nm using a Luzchem photoreactor (model
LZC-1, Ontario, Canada, 10 J cm−2). Data were obtained as the
average of three independent sets of experiments, all performed in
triplicate for each concentration. DCFDA assay was done to quantify
ROS on the oxidative production of fluorescent DCF (λem = 525 nm).
BT474 cells (∼2 × 105 cells plated in duplicate) were incubated in the
presence or absence of complexes 1−6 (concentration = IC50 values in
light) for 24 h in the dark with or without photoirradiation in DPBS
(400−700 nm, 1 h). Cells were trypsinized and treated with 1 μM
DCFDA for ∼15 min, and the stained cells were determined by flow
cytometry.
[Pt(Fc-tpy)(L1)]Cl (1). Yield: 110 mg, ∼88%. Anal. Calcd for
C44H48ClFeN7O3PtS: C, 50.75; H, 4.65; N, 9.42. Found: C, 50.55; H,
4.89; N, 9.40. ESI-MS in MeOH: m/z 1005.2538 ([M − Cl]+; 100%).
1H NMR (400 MHz, DMSO-d6): δ 9.14 (d, 2H), 8.82 (d, 2H), 8.68
(s, 2H), 8.55 (m, 2H), 8.19 (b, 1H), 7.89 (m, 2H), 7.72 (b, 1H), 6.45
(s, 1H), 6.30 (s, 1H), 5.43 (d, 2H), 4.80 (s, 2H), 4.22 (s, 1H), 4.18 (s,
5H), 4.17 (b, 3H), 3.07 (m, 1H), 2.99 (m, 2H), 2.80 (s, 1H), 2.5 (s,
1H), 2.15 (m, 2H), 2.13 (m, 2H), 1.49−1.25 (b, 12H) [b, broad; s,
singlet; d, doublet; m, multiplet]. UV−visible in 10% DMSO−DPBS
(pH = 7.2) [λmax, nm (ε, M−1 cm−1)]: 640 (2700), 455 (3400), 345
(10000), 282 (17000). IR data in the solid phase (cm−1): 3240 m,
3064 m, 2915 m, 2859 m, 2181 w, 2068 w, 1983 w, 1688 s, 1639 s,
1602 s, 1539 s, 1431 m, 1241 m, 780 w, 725 w, 669 w, 498 m, 470 m
[s, strong; m, medium; w, weak]. ΛM, S m2 M−1 in DMF at 25 °C: 65.
[Pt(Fc-tpy)(L2)]Cl (2). Yield: 95 mg, ∼85%. Anal. Calcd for
C38H37ClFeN6O2PtS: C, 49.17; H, 4.02; N, 9.05. Found: C, 48.88;
H, 4.22; N, 9.11. ESI-MS in MeOH: m/z 892.1653 ([M − Cl]+,
1
100%). H NMR (400 MHz, DMSO-d6): δ 8.90 (m, 2H), 8.77 (m,
2H), 8.59 (m, 2H), 8.47 (m, 2H), 8.22 (b, 1H), 7.95−7.88 (b, 2H),
6.45 (s, 1H), 6.38 (s, 1H), 5.44 (d, 2H, J = 12 Hz), 4.81 (d, 2H, J = 8
Hz), 4.29 (m, 1H), 4.23 (s, 5H), 4.20 (b, 3H), 3.09 (m, 1H), 2.81 (m,
1H), 2.58 (m, 1H), 2.18 (b, 2H), 1.60−1.11 (b, 6H). UV−visible in
10% DMSO−DPBS (pH = 7.2) [λmax, nm (ε, M−1 cm−1)]: 642
(2900), 462 (3400), 350 (9000), 265 (25000). IR data in solid phase
(cm−1): 3261 s, 3229 s, 3063 s, 2920 s, 1687 s, 1643 s, 1604 s, 1543 s,
1432 m, 1246 m, 1163 w, 783 w, 576 w, 501 m, 478 m. ΛM, S m2 M−1
in DMF at 25 °C: 70.
[Pt(Fc-tpy)(L3)]Cl (3). Yield: 80 mg, ∼89%. Anal. Calcd for
C30H25ClFeN4OPt: C, 48.44; H, 3.39; N, 7.53. Found: C, 48.58; H,
3.45; N, 7.41. ESI-MS in MeOH: m/z 708.1064 ([M − Cl]+, 100%).
1H NMR (400 MHz, DMSO-d6): δ 9.13 (m, 2H), 8.98 (m, 2H), 8.82
(b, 2H), 8.67 (b, 2H), 8.57 (b, 1H), 7.90 (b, 2H), 5.45 (d, 2H, J = 8
Hz), 4.83 (d, 2H, J = 12 Hz), 4.22 (s, 5H), 4.21 (s, 2H), 2.08 (s, 3H).
UV−visible in 10% DMSO−DPBS (pH = 7.2) [λmax, nm (ε, M−1
cm−1)]: 660 (2900), 457 (3600), 342 (10000), 285 (19000). IR data
in solid phase (cm−1): 3234 s, 3064 s, 2923 s, 2853 s, 1688 s, 1640 s,
1603 s, 1544 s, 1472 s, 1429 s, 1244 m, 1042 w, 780 w, 664 w, 580 w,
474 m. ΛM, S m2 M−1 in DMF at 25 °C: 83.
[Pt(Ph-tpy)(L1)]Cl (4). Yield: 100 mg, 72%. Anal. Calcd for
C40H44ClN7O3PtS (4): C, 51.47; H, 4.75; N, 10.50. Found: C,
51.38; H, 4.92; N, 10.42. ESI-MS in MeOH: m/z 897.2973 ([M −
1
Cl]+, 100%). H NMR (400 MHz, DMSO-d6): δ 9.05 (m, 2H), 8.88
(m, 2H), 8.54 (d, 2H, J = 8 Hz), 8.21 (b, 3H), 7.95 (b, 2H), 7.70 (b,
6H), 6.41 (s, 1H), 6.35 (s, 1H), 4.30 (s, 1H), 4.13 (s, 1H), 4.12 (s,
2H), 3.08 (m, 1H), 2.99 (m, 2H), 2.82 (b, 1H), 2.58 (s, 1H), 2.15 (m,
2H), 2.04 (m, 2H), 1.61−1.52 (b, 12H). UV−visible in 10% DMSO−
DPBS (pH = 7.2) [λmax, nm (ε, M−1 cm−1)]: 398 (4400), 285
(19000), 257 (23000). IR data in the solid phase (cm−1): 3400 s, 3334
s, 3236 s, 3214 s, 3062 m, 3000 m, 2939 m, 2888 m, 2856 m, 1690 s,
1649 s, 1610 s, 1531 m, 1480 m, 1412 m, 1245 m, 1103 w, 1032 w,
899 w, 774 s, 593 m, 572 m, 483 m, 462, m. ΛM, S m2 M−1 in DMF at
25 °C: 67.
[Pt(Ph-tpy)(L2)]Cl (5). Yield: 97 mg, 79%. Anal. Calcd for
C34H33ClN6O2PtS (5): C, 49.79; H, 4.06; N, 10.25. Found: C,
49.58; H, 4.13; N, 10.11. ESI-MS in MeOH: m/z 784.1996 ([M −
Cl]+, 100%). 1H NMR (400 MHz, DMSO-d6): 9.12 (m, 2H) 9.03 (m,
2H), 8.87 (b, 2H), 8.55 (m, 2H), 8.19 (b, 3H), 7.89 (m, 2H), 7.69 (m,
3H), 6.41 (s, 1H), 6.35 (s, 1H), 4.65 (s, 1H), 4.18 (s, 1H), 4.11 (b,
2H), 3.11 (m, 1H), 2.80 (d, 1H, J = 4 Hz), 2.58 (b, 1H), 2.14 (m, 2H),
1.61−1.12 (b, 6H). UV−visible in 10% DMSO−DPBS (pH = 7.2)
[λmax, nm (ε, M−1 cm−1)]: 406 (5600), 330 (17000), 285 (27000), 258
(31000). IR data in the solid phase (cm−1): 3472 s, 3352 s, 3247 s,
Annexin V-FITC/PI Assay. BT474 cells (∼2 × 105) were
incubated (concentration = IC50 values in light) for 24 h followed
by light treatment in DPBS (400−700 nm, 10 J cm−2). Cells were
further kept for 12 h post-treatment and then trypsinized. Annexin V-
FITC (2 μL) and propidium iodide (PI; 1 μL) were then added to
clear suspensions of cells in 500 μL of a 1X binding buffer and kept in
the dark for 10 min. The percent cell population in each quadrant was
estimated by FACS analysis. Experiments were performed in duplicate
along with untreated controls for reference.
C
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Table 1. Selected Physicochemical Data of Complexes 1−6
1
2
3
4
5
6
a
λabs /nm (ε/
640 (2700), 455
(3400)
642 (2900), 462
(3400)
660 (2900), 457
(3600)
398 (4400), 285
(19000)
406 (5600), 330
(17000)
397 (3700), 330
(10300)
M−1cm−1
)
b
λemi /nm
635
635
640
639
640
Φ × 105
1.5
3.2
50
43
10
τ/μs
0.015
0.018
<0.01
0.2
0.25
0.09
c
IR: νN−H /cm−1
3240, 3064
1688, 1639
0.25
3229, 3063
1687, 1643
0.26
3234, 3064
1688, 1640
0.25
3214, 3062
1690, 1649
3247, 3129
1673, 1653
3272, 3231
1645, 1606
c
IR: νCO /cm−1
CV
d
Ef /V [ΔEp/mV]
−1.81 [95], −1.32
−1.82 [85], −1.30
−1.79 [80], −1.28
−1.76 [65], −1.20
−1.76 [80], −1.24
−1.76 [90], −1.24
[75]
[100]
[105]
[65]
[85]
[90]
e
ΛM /S m2 mol−1
65
70
b
83
67
69
73
a
In 10% DMSO−DPBS (pH = 7.2). In DCM at 298 K (excitation wavelength = 470 nm). The quantum yield was calculated using
c
d
[Ru(bpy)3](PF6)2 in MeCN as the standard (Φ = 0.062). IR spectra recorded in the solid phase. Peaks corresponding to cyclic voltamogramms of
1 mM sample in DMF−0.1 M TBAP. The potentials are recorded versus SCE (E0 = 0.244 V), and values are reported relative to Fc/Fc+ using
ferrocene (Ef = 0.437 V) as the internal standard. Ef = 0.5(Epa + Epc). ΔEp = (Epa − Epc), where Epa and Epc are the anodic and cathodic peak
e
potentials, respectively. Scan rate = 50 mV s−1. Molar conductivity in DMF.
Cell Cycle Analysis. Complexes 1−6 (concentration = IC50 values
in light) were added to the BT474 cells (∼2 × 105), incubated in the
dark for 24 h, and photoirradiated in visible light (400−700 nm, 1 h).
FACS analysis was done with samples (in duplicate) along with dark
and untreated controls. The percentage of cells in each cell-cycle phase
was obtained by data analysis using WinMDI, version 2.8.
Cellular Uptake from Platinum Estimation. To quantify the
total platinum uptake, about 106 BT474 cells were seeded in 60 mm
tissue culture dishes and treated with complexes 1−6 (50 μM) for 24
h. One identical plate was used to determine the protein content by
Bradford assay. Similar experiments were performed with or without
prior incubation of cells in the presence of biotin (100 μM) for 2 h.
The samples were run using ICP-MS for the platinum content along
with the standards (0, 100, 250, and 500 ppb fitted in a linear plot with
a correlation of 0.91). The platinum content obtained in ppb units was
then expressed as ng of Pt/μg of protein. All of the experiments were
performed in triplicate and with untreated controls.
coordinated Fc-tpy ligand. The biotinylated complexes 1, 2, 4,
and 5 showed additional signals in the regions 6.3−8.2 ppm
(for N−H protons) and 1.2−4.2 ppm (for aliphatic protons).
In contrast, the control complexes 3 and 6 displayed only a
singlet at 1.9 ppm corresponding to the CH3 group protons.
The 13C and 195Pt NMR spectra could not be recorded because
of the low solubility of the complexes in deuterated DMSO.
The molar conductance values of the complexes recorded in
DMF at 25 °C were in the range of 65−83 S m2 M−1,
suggesting their 1:1 electrolytic nature. Mass spectral analysis of
dilute MeOH solutions of the samples displayed a single peak
(respective m/z values for 1−6 are 1005.25, 892.16, 708.10,
897.29, 784.19, and 600.13) which is in accordance with that
expected for the ionized [M − Cl]+ species. The presence of
platinum and the unipositive charge were evident from the
isotopic distribution pattern (Figures S8−S13 in the Support-
ing Information).
3. RESULTS AND DISCUSSION
The UV−visible spectra of complexes 1−6 (50 μM) were
recorded in 10% DMSO−DPBS (pH = 7.2). Complexes 1−3
with the ferrocene unit showed strong absorption bands in the
visible region having maxima at 650 and 460 nm with respective
ε values of ∼2900 and ∼3500 M−1 cm−1 (Figures 2 and S14 in
Synthesis and General Properties. It was earlier
observed that the metal-bound chloride in [(R-tpy)PtCl]Cl
gets substituted by acetylide ligands, where R-tpy is a
terpyridine (tpy) ligand with a pendant R group.33 This
synthetic strategy is useful to attach a cancer-targeting moiety
to the metal-based photoinitiator. Biotinylated derivatives
containing a free carboxylic group were coupled with
propargylamine using anhydrous HOBt and EDCI to obtain
the biotinylated acetylide ligands (Scheme S1 in the Supporting
Information). The complexes were synthesized in ∼85% yields
by reacting the precursor [(R-tpy)PtCl]Cl (R = Fc for 1−3 and
Ph for 4−6) and ligands (HL1 for 1 and 4, HL2 for 2 and 5, and
HL3 for 3 and 6) in DMF using a catalytic amount of CuI and
excess triethylamine under a nitrogen atmosphere (Scheme S2
in the Supporting Information). The schematic drawings of 1−
6 and the ligands used are shown in Figure 1. The purity of the
complexes was checked by HPLC, ICP-MS, and CHN analysis
(Figure S4 in the Supporting Information). Selected character-
Figure 2. Visible absorption spectra of 1, 4, and Fc-tpy in 10%
DMSO−DPBS (pH = 7.2).
1
ization data are given in Table 1. The H NMR spectra of the
complexes were recorded and assigned to the respective
protons (Figures S5−S7 in the Supporting Information). All
of the complexes showed peaks in the region 7.7−9.1 ppm,
which were assignable to the aromatic protons of the terpyridyl
moiety. For complexes 1−3, two doublets at 5.4 and 4.8 ppm
and a singlet at 4.2 ppm were observed, which is characteristic
of the protons of the cyclopentadienyl rings of the metal-
the Supporting Information). These bands are characteristic of
the metal-bound Fc-tpy ligand.33,45 The coordination to
platinum metal is important for observing the band in the
red-light region (∼650 nm) because it lowers the energy gap
(HOMO−LUMO) by extensive conjugation. In contrast, the
uncoordinated ligand Fc-tpy displays only a shoulder at 480 nm
(ε, 2070 M−1 cm−1) under similar conditions, emphasizing the
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need for metal coordination to observe the low-energy spectral
band. An intense band within the spectral window of 600−800
nm is important in PDT considering greater tissue penetration
of red light.46 The Ph-tpy complexes 4−6 displayed absorption
bands at ∼395 nm (ε, ∼ 4500 M−1 cm−1). In the solid state at
298 K, the phenyl complexes 4−6 displayed broad emission
bands in the red region (700−735 nm; Table 1 and Figure S15
in the Supporting Information). The ferrocene moiety acts as
an efficient quencher of the MLCT and LC states, thereby
resulting in no emission of complexes 1−3 at room
temperature or at 77 K. This is in accordance with reductive
quenching by the ferrocene unit reported for the Fc-tpy
complexes.47−49 The phenyl complexes 4−6 showed weak
unstructured luminescence at ∼640 and 710 nm (Φ ∼ 0.0005
and τ ∼ 0.2 μs) in dichloromethane solutions. The ferrocenyl
complexes 1 and 2 showed very weak luminescence at around
635 nm (Φ ∼ 0.3 × 10−4 and τ ∼ 0.015 μs). The complexes
showed no significant luminescence in MeCN, DMSO, or
DMF or in aqueous DMSO. The IR spectral signals were
observed in the ranges 3000−3250 cm−1 for amide N−H
stretch, 2850−2900 cm−1 for CC and C−H stretches, and
1540−1690 cm−1 for CO, CC, and C−N stretches (Figure
S16 in the Supporting Information).50 Out-of-plane N−H
frequencies were observed at 700 cm−1.
Electrochemical Studies. The redox properties of the
ferrocene (Fc)-appended complexes 1−3 were studied using
1.0 mM solutions in DMF−0.1 M TBAP. The potentials versus
SCE were corrected with reference to Fc/Fc+ (Ef = 0.437 V).
The Fc/Fc+ couples appeared as an irreversible to quasi-
reversible response near 0.25 V (Table 1 and Figure S17 in the
Supporting Information).51 The anodic response was found to
undergo a positive shift of ∼90 mV compared to the free Fc-tpy
ligand. This is attributed to the increasing inductive effect of the
electron-withdrawing tpy on metal chelation. This also indicates
stabilization of the FeII redox state of Fc-tpy on binding to PtII,
which is difficult to chemically oxidize compared to free Fc-tpy
and ferrocene. The irreversibility of the peaks may be due to
electrode poisoning, adsorption, or instability of the Fc+ moiety
in the complex. A cyclic voltammetric (CV) study using
ferrocene and a free Fc-tpy ligand with same electrode
configuration gave expected reversible peaks, thus eliminating
the possibility of electrode poisoning or surface adsorption
processes (Figure S18a,b in the Supporting Information). To
probe the stability of the oxidized species, we carried out
chronoamperometric studies by setting the voltage at +0.4 V
(vs Fc/Fc+). The UV−visible spectra recorded after certain
time intervals indicated the loss of Fc-centered bands at 480
and 590 nm, suggesting degradation of the oxidized species
(S18c in the Supporting Information). All of the complexes
exhibited cathodic responses near −1.3 and −1.8 V, which are
attributed to two successive one-electron-reduction processes
involving the tpy moiety. The reversible reduction of the
coordinated tpy moiety is relatively unperturbed by the
attached ferrocenyl or phenyl moiety. The free ligands Ph-tpy
or Fc-tpy did not show such reduction (Figure S18d in the
Supporting Information). The biotinylated ligands were also
redox-inactive within the voltage window. The phenyl
analogues 4−6 showed no oxidation peaks due to the lack of
a Fc moiety. The results suggest that the ferrocenyl complexes
could undergo oxidation upon photoexposure, with their
phenyl analogues being inactive.
properties of the Fc-tpy complexes 1−3 with theoretical studies
using the B3LYP/LanL2DZ level of theory with the Gaussian
G09 suite of programs.40−43 The coordinates for the basic
structures of 1−3 were taken from the crystal structure of
[Pt(Fc-tpy)Cl]Cl and then modified as required by adding the
acetylide fragments. These coordinates were utilized to obtain
the respective energy-minimized structures by DFT methods
(Figure S19 and Table S1 in the Supporting Information). In
the optimized structure of complex 1, the ferrocenyl ring and
the aliphatic chain containing the biotin moiety adopted close
configurations because of the flexibility of the long alkyl chains.
Interestingly, a chemically significant N−H····π interaction was
observed in the optimized structure of 1. The distance of
hydrogen of the amide group of the biotin unit from the
unsubstituted Cp ring of the Fc unit was 2.53 Å, thereby
signifying the presence of weak interactions, which stabilized
the molecule in such a configuration. To further understand the
localizations and contributions of the molecular orbitals
responsible for such transitions, we performed TDDFT on
these energy-minimized ground-state geometries. Selected
transitions within the 400−900 nm range along with their
oscillator strengths (f) are listed in Table S2 in the Supporting
Information. The highest probable transition was at 592.04 nm
for 1 (f = 0.0270), 592.25 nm for 2 (f = 0.0276), and 590.99
nm for 3 (f = 0.0302). This transition was assignable to the
ligand moiety (Fc-tpy) to platinum(II) metal charge transfer
(LMCT; ∼40%), viz., from the d orbitals of Fe of Fc
(HOMO−3 for 1, HOMO−1 for 2, and HOMO for 3) to the
π orbitals of Pt-tpy (LUMO), and intraligand charge transfer
(ILCT; ∼45%), viz., from the d orbitals of Fe in Fc (HOMO−4
for 1, HOMO−3 for 2, and HOMO−1 for 3) to the π orbitals
of Cp rings (LUMO+6; Figures 3 and S20 and S21 in the
Supporting Information). The absence of such transitions in
free Fc-tpy suggests the important contribution of the low-lying
orbitals from the metal-coordinated ligands for the realization
of strong absorptions in the low-energy visible spectral region.
Computations also revealed strong absorptions at 515 and 465
nm for all of the complexes with respective f values of ca. 0.02
3
3
Figure 3. Frontier molecular orbitals of complex 1 involved in
transitions in the red light region as obtained from TDDFT
calculations using the B3LYP/LanL2DZ level of theory. Color code:
purple, platinum; red, oxygen; blue, nitrogen; black, carbon; yellow,
sulfur; gray, hydrogen. The orbitals are drawn using a contour value of
0.03.
TDDFT Calculations. We attempted to rationalize the
strong absorptions in the visible region and the photophysical
E
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and 0.01. Interestingly, these transitions were characterized by
charge transfer (L′MCT and IL′CT; ∼90%) involving the
triple-bonded ligands (HOMO−9 and HOMO−7 for 1,
HOMO−5 and HOMO−7 for 2, and HOMO−2 and
HOMO−4 for 3) and Fc-tpy-Pt (LUMO and LUMO+1;
Figure 3 and S18 and S19 in the Supporting Information). The
comparatively low energy gap of transitions (ΔE = −1.5 eV)
compared to free ferrocene (ΔE = −5.5 eV) or Fc-tpy (−4.2
eV) makes them better photoinitiators capable of responding to
longer-wavelength irradiation. Additionally, it was observed that
the orbitals of the ferrocenyl (Fc) moiety and that of Pt−−L′
comprise mainly antibonding interactions and, hence, are
dissociative in nature. This suggests probable loss of the
ferrocenyl unit, which also triggers Pt−C(acetylide) bond
dissociation in the excited states of these photodegradation-
prone molecules.52,53
excited state can be stabilized in the lower oxidation state of 2+
in the presence of π-acidic ligands like phen. We can thus infer
that metal chelators like phen can leach out iron from the
complex in its photoexcited state and form stable binary
species. The effect of the addition of excess glutathione (GSH)
was explored because GSH is highly abundant in cancer cells
(∼15 mM). When the complexes were photoirradiated in the
presence of GSH (15 mM in 10% DMSO−DPBS), the
absorption bands at longer wavelengths showed instant decay
and the solution turned yellow, giving two new bands at 575
and 580 nm (Figure S27 in the Supporting Information). The
mass spectral pattern of these solutions did not show the
formation of any species of discrete composition.
1
It is apparent from the H NMR analysis in DMSO-d6 that
the complexes released the unsubstituted cyclopentadienyl ring
(Cp, C5H5) of the ferrocenyl moiety upon photoirradiation,
with new peaks appearing as a multiplet at 6.5−6.4 ppm and a
singlet at 2.7 ppm (Figures 4 and S28 and S29 in the
Mechanism of Photodegradation. The stability of the
complexes was studied in different solvents in the presence of
additives under photoirradiation and in dark conditions. The
solvents, additives, and extensive conjugation in the complexes
were found to have a profound impact on the photodegradation
processes of ferrocenyl complexes. Complexes 1−6 (50 μM)
were found to be stable in 10% DMSO−DPBS (pH = 7.2) or in
DMSO for 48 h in the dark. However, the Fc-tpy complexes 1−
3 degraded upon photoexposure to broad-band visible light of
400−700 nm (Luzchem photoreactor, 10 J cm−2) or to red
laser light of 647 nm (50 mW). The mechanistic pathway and
photoproducts were investigated by UV−visible absorption, 1H
NMR, and ESI-MS spectral methods. They exhibited solvent-
dependent changes in the absorption spectra when monitored
as a function of the photoirradiation time. The complexes upon
photoirradiation in 10% DMSO−DPBS (50 μM, pH = 7.2)
showed a gradual decrease of the intensity of the absorption at
∼650 and 460 nm after 1 h of photoexposure time (Figure S22
in the Supporting Information). In pure DMSO or DMF
solvent, the irradiated complexes showed a red shift of the band
from 580 to 640 nm with a large increment in the ε value (from
6770 to 21460 M−1 cm−1) upon just 30 s of light exposure
(Figure S23 in the Supporting Information). The band
gradually flattened upon continuous photolysis, and the green
solution turned initially to sky blue, then to deep blue, and
finally black (Figure S23d in the Supporting Information). This
rapid spectral change was attributed to solvent charge-transfer
transitions in the oxidized Fc moiety, which are favored in
electron-acceptor solvents like DMSO or DMF.54 A similar
rapid loss of the visible spectral band was visible upon the
addition of ceric ammonium nitrate (CAN) as an oxidizing
agent, turning the solution to yellow (the color of excess CAN),
indicating the involvement of a photooxidative process (Figure
S23 in the Supporting Information). The addition of 1,10-
phenanthroline (phen) to the 30 min photoirradiated solution
of the complex gave a red solution showing two new bands at
520 and 430 nm, which are assignable to a tris-phen complex of
iron(II) (Figure S24 in the Supporting Information).55 Mass
analysis of these samples showed the presence of a peak at m/z
298.18, confirming the formation of a FeII-phen species (Figure
S25 in the Supporting Information). Similar absorption bands
at 520 and 430 nm appeared when a solution of ferric chloride
was treated with phen, turning the solution red (Figure S26 in
the Supporting Information). Mass spectral analysis of these
diluted samples also exhibited a prominent peak at m/z 298.18,
thereby indicating the formation of a tris-phen complex of
iron(II). This experiment confirmed that the iron(III) from the
1
Figure 4. Time-dependent photoinduced (400−700 nm) H NMR
spectral changes of complex 1 (5 mM in DMSO-d6) as indicated in the
figure (L = light of 400−700 nm, 10 J cm−2, D = dark, and m =
photoexposure time in min).
Supporting Information). To further support the leaching out
of the Cp ring,56,57 we performed a control photolysis
experiment with 1,1′-bis(diphenylphosphino)ferrocene in
UVA light of 395 nm. In this Fc derivative, the substituted
Cp ring has high molecular weight and, hence, the photorelease
can be detected by mass spectra. After 5 h of photoexposure,
the faint yellow color of the DMSO solution intensified. NMR
and mass spectral evidence pointed out the release of
cyclopenta-1,3-dien-1-yldiphenylphosphine oxide (Figures S30
and S31 in the Supporting Information). The phosphorus in
the released species was found to be oxidized, which confirms
the generation of ROS (the m/z value for the protonated
species is 267.09). It is a favorable argument to conclude that
complexes 1−3 followed an analogous mechanism upon
photoexposure at longer wavelengths and shorter irradiation
times, which is due to the presence of extensive conjugation
and metal coordination. Another interesting observation from
the NMR studies was the release of the triple-bonded ligands
upon prolonged photoexposure of the complex (Figures 4 and
S28 and S29 in the Supporting Information). The peak
appearing at 2.2 ppm upon light exposure of complex 3 is
assignable to the acetylenic proton of CCH, indicating
dissociation of the Pt−acetylide bond. These samples were then
diluted with HPLC-grade MeOH and subjected to mass
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analysis. The respective m/z 395.25 and 282.13 [assigned to
free ligands, (M + H)+] proved cleavage of the Pt−C(acetylide)
bond in light (Figures S32a,b in the Supporting Information).
The photoinduced detachment of the biotin ligand is of interest
because this provides an effective way of delivering the
complexes with the help of biotin receptors and then releasing
the active phototoxin with the help of light. A peak of m/z
485.7 corresponding to [Pt(tpy)Cl]+ was observed in the mass
spectrum of the photodegraded complex 3 (Figure S32c in the
Supporting Information). All of the experiments when repeated
in darkness showed unaltered spectra, implying the importance
of light activation for achieving degradation of the complex. In
summary, these results altogether suggest a photoredox process
to be operative in the light-promoted degradation of complexes
1−3 having a redox-active extensively conjugated Fc-tpy moiety
(Scheme S3 in the Supporting Information). In contrast, the
phenyl analogues showed significant photostability.
μM, 2686 base pairs) was studied using red light of 647 nm
(Figure 5). This wavelength was chosen considering the 640
Platinum Estimation. Photorelease of the biotinylated
ligands led to the formation of an active platinum(II) species
that can bind to ct-DNA. To explore such a possibility, we
treated two identical sets of ct-DNA (5 mL, 500 μM) with
complexes 1−3 (50 μM). One such set was photoirradiated in
visible light (400−700 nm) for 1 h, while the other was kept in
darkness. The DNA was then precipitated with an excess of
cold ethanol and isolated to measure the amount of platinum
covalently bound to DNA. Complexes 1−3 showed prominent
enhancement (∼500 pg of Pt/μg of DNA) in DNA-bound
platinum amounts only after irradiation of the samples (Table
S3 in the Supporting Information). The phenyl analogues,
however, did not show any significant increase of platinated
DNA in darkness or upon exposure to light.
DNA Binding and Cleavage Studies. The DNA melting
experiments were done to explore the effect of the complexes
(10 μM) on the melting temperatures (Tm) of ct-DNA (100
μM; Figure S33a in the Supporting Information). Intercalation
of the planar aromatic molecules between the DNA base pairs
stabilizes the duplex DNA, which is reflected in an increase of
the melting temperature (ΔTm). The ΔTm values of ∼0.5 °C
for 1−3 indicate the lack of DNA intercalation of the Fc-tpy
complexes. In contrast, the phenyl analogues 4−6 gave a ΔTm
value of ∼5 °C, suggesting an intercalative mode of their
binding to ct-DNA. EB as a DNA intercalator gave a ΔTm value
of 11 °C under similar conditions. The viscosity of ct-DNA is
known to be significantly altered in the presence of DNA
intercalators or groove binders. Viscometric data suggested
intercalative properties of the Ph-tpy complexes 4−6. It was
also observed from the viscosity plot of (η/η0)1/3 versus
[complex]/[DNA], where η and η0 are the viscosities of DNA
in the presence and absence of the complex, that the Fc-tpy
complexes caused only minor changes in the viscosity,
indicating their inability to bind to ct-DNA in an intercalative
manner (Figure S33b in the Supporting Information).
Intercalation within the base pairs also results in an
enhancement of the emission intensity due to trapping of the
molecule in the hydrophobic DNA core. Only the phenyl
complexes 4−6 showed notable enhancement of the
luminescence intensity in DNA-buffer solutions, which further
demonstrated the DNA intercalating ability of PtII(Ph-tpy)
(Figure S34 in the Supporting Information).58
Figure 5. Photoinduced cleavage of pUC19 DNA in the presence of
Fc-tpy complexes 1−3 (20 μM in 10% DMF buffer) in light (red bars,
Ar−Kr continuous-wave laser source, 647 nm, 50 mW) and dark
(black bars) as shown in the figure. Different additives added: NaN3 or
TEMP (4 mM), KI (4 mM), or DMSO (8 μL), catalase, and SOD (4
units).
nm absorption maxima of the photoactive Fc-tpy complexes 1−
3. The complexes (20 μM in 10% DMF buffer, pH = 7.2)
showed photoinduced SC DNA cleavage, resulting in the
formation of ∼50% NC form. The presence of singlet oxygen
quenchers like TEMP and NaN3 (4 mM) failed to inhibit DNA
cleavage, thus excluding the formation of any singlet oxygen
(1O2) as the ROS. The formation of NC DNA was, however,
reduced to ∼25% in the presence of hydroxyl radical
scavengers, viz., KI (4 mM) and DMSO (8 μL). The DNA
photocleavage activity was also decreased (∼20% NC) in the
presence of SOD (4 units) and catalase (4 units), suggesting
the formation of hydroxyl radicals (•OH) via superoxide and
hydrogen peroxide as the ROS.
Cell Viability Studies. Photodegradation of the Fc-tpy
complexes, leading to the generation of radical species
prompted us to investigate the antiproliferative properties of
the complexes in cancer cells in light (400−700 nm, 10 J cm−2)
and in the dark. With the complexes having pendant biotin
units, we selected the biotin-receptor positive cells, viz., human
breast carcinoma cancer cells (BT474), to assess the cellular
activity. Human normal breast epithelial cells (HBL-100) were
taken as the reference to validate the selective uptake of these
complexes. Colorimetric MTT assay, based on an estimation of
the resulting purple formazan by spectral methods, was used to
determine the IC50 values of the complexes (Figures 6 and S35
in the Supporting Information). The results were compared
with those known for related photoactive platinum(II) and
tpy−metal complexes (Table 2).33a,34,35,60,61 Complex 1
showed photoinduced cytotoxicity in BT474 cells, giving an
IC50 value of ∼7 μM in visible light (400−700 nm), while
remaining essentially nontoxic in the dark (IC50: >50 μM).
Complexes 2 and 3 gave IC50 values of ∼16 μM upon light
exposure. This implies that the presence of a spacer moiety in
complex 1 resulted in better cellular uptake, which can be
correlated with its better cellular toxicity. Upon comparison, it
was observed that complex 1 exhibited comparable photo-
cytotoxicity with less dark toxicity than the previously reported
complexes. However, the other complexes did not have any
The possibility of any ROS generation upon photo-
degradation of the complexes was explored from the DNA
photocleavage experiments using complexes 1−3 and different
additives.59 The photocleavage of SC pUC19 DNA (0.2 μg, 30
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Table 3. Cellular Uptake of Complexes in BT474 Cells
Expressed as ng of Pt/μg of Protein
a
b
complex
without biotin
with biotin
0.0
untreated
0.0
1
2
3
4
5
6
11.15 0.82
3.83 0.20
2.77 0.52
10.65 1.06
8.65 1.17
7.56 0.85
2.48 0.13
1.80 0.15
2.14 0.15
5.42 0.57
4.67 0.33
6.91 0.64
a
BT474 cells treated with the complexes (50 μM) for 12 h in the dark.
Cells preincubated with biotin (100 μM) for 2 h and then treated
b
with the complexes (50 μM) for 12 h in the dark. The platinum
content estimated by the ICP-MS method.
Figure 6. Cell viability plots in BT474 cells preexposed to biotinylated
complexes 1, 2, 4, and 5 for 24 h in the dark followed by either
photoirradiation (10 J cm−2, 400−700 nm) or darkness.
showed lower cellular uptake (∼2.8 and 7.5 ng of Pt/μg of
protein, respectively). It was also observed that prior treatment
of the cells with biotin (100 μM) for 2 h suppressed cellular
uptake of the biotinylated complexes 1, 2, 4, and 5. The uptake
of 1 (11 ng of Pt/μg of protein) was reduced by presaturation
of biotin receptors to 2.5 ng of Pt/μg of protein. Similar results
were also obtained for complex 4, which showed reduced
uptake to half upon prior treatment of the cells with biotin. In
contrast, the nonbiotinylated complexes 3 and 6 showed similar
cellular uptake values in the absence or presence of biotin. The
complexes did not react with the externally added biotin, as
evidenced from the unaltered mass and NMR spectra, and this
ruled out any possibility of interaction of the complex with
biotin, which may lead to the lowering of cellular uptake. It is
thus concluded that the biotin-receptor-mediated uptake of the
complexes was responsible for their potent toxicity selectively
in the cancer cells.
ROS and Cellular Apoptosis. Light-promoted generation
of ROS is an essential feature of PDT agents. Cellular ROS
production by complexes 1−3 was examined by 2,7-
dichlorofluorescein diacetate (DCFH-DA) assay (Figure 7).
This cell-permeant nonfluorescent dye gets oxidized by cellular
ROS to produce 2,7-dichlorofluorescein (DCF), which emits at
525 nm (λexc = 488 nm). The emission intensity of DCF was
quantified by using FACS techniques, and it gave an estimate to
the amount of ROS generated within the cells. Complexes 1−3
showed DCF formation (mean DCF intensity ∼3000 au) in
BT474 cells only upon photoexposure to light of 400−700 nm
but not in the dark (mean DCF intensity ∼900 au) (Figure
tumor-targeting moieties attached to them, which rendered
them unspecific in action. In contrast, the phenyl analogues
displayed high dark toxicity, as is known for several tpy
complexes of platinum(II) possibly because of their better
DNA intercalating properties, as predicted from the DNA
melting and viscosity data.33b,62−65 Complex 4 gave an IC50
value of 4 μM in light, which is comparable to that of cisplatin.
The nonbiotinylated control complex 6 showed ∼5-fold lower
toxicity compared to 4 with an IC50 value of 24 μM.
Interestingly, the biotin complexes 1, 2, 4, and 5 showed low
cytotoxicity in the normal cell line HBL-100 (IC50: ∼40 μM),
while the control complexes 3 and 6, lacking the biotin moiety,
showed toxicity similar to that observed in the cancer cells
(IC50: ∼25 μM). The MTT assay data highlight the importance
of the biotin moiety toward achieving selective therapeutic
activity in the cancer cells.
Biotinylated compounds are well-known for their increased
cellular uptake via streptavidin−avidin receptors that are
overexpressed in cancer cells.16−18 Thus, we hypothesized
that the biotin complexes resulted in better cytotoxicity because
of enhanced cellular uptake compared to their control
counterparts. To justify our argument, we further carried out
cellular uptake studies by a quantified estimation of the total
platinum uptake within the cells. The results given in Table 3
show the highest cellular platinum amount (∼10−11 ng of Pt/
μg of protein) of complexes 1 and 4. Complexes 3 and 6
a
b
Table 2. Photocytotoxicity of 1−6 As Determined from MTT Assay along with Selected Metal Complexes for Comparison
BT474
HBL-100
c
d
c
d
complex
IC50 (light) (μM)
IC50 (dark) (μM)
IC50 (light) (μM)
IC50 (dark) (μM)
1
7.7 2.5
16.7 3.4
16.9 3.8
4.3 2.7
13.7 3.4
20.7 5.5
>50
>50
45.2 5.2
42.6 4.6
24.5 3.5
20.2 2.2
28.5 5.0
22.4 4.3
>50
>50
2
>50
>50
3
>50
>50
4
15.5 3.1
17.5 5.2
24.1 4.2
>50
27.3 5.4
30 4.3
29.5 4.2
>50
5
6
HL1
cisplatin
4.4 2.3
15.6 5.2
a
The IC50 values (μM) obtained from MTT assay in BT474 and HBL-100 cells pretreated with complexes in the dark for 24 h, either followed by
light exposure (400−700 nm, 10 J cm−2) or kept in the dark. The IC50 values (μM) in HaCaT cells of [Pt(Fc-tpy)Cl]Cl,33a [Pt(pap)(an-cat)],34
b
[Pt(NH3)2(cur)]NO3,35 and [VO(Fc-tpy)(py-acac)]ClO460 (in MCF-7 cells) and platinum(IV) diazido61 are 12.2 ( 0.5), 4.4 ( 0.2), 13 ( 4), 2.1
c
( 0.3), and 19.5 ( 6.1) in visible light and 74.8 ( 1.1), 43.5 ( 0.8), >200, 38.5 ( 1.1), and 70.2 ( 6.1) in the dark, respectively. In light of 400−
700 nm (exposure time of 1 h). In the dark.
d
H
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Article
Figure 7. DCFH-DA assay showing generation of ROS in BT474 cells
treated with complexes 1 and 4 for 24 h in the dark and exposed to
light (red cylinders) or in the dark (black cylinders). The y axis
denotes the mean DCF intensity (in arbitrary units) obtained with the
cells alone/cells + DCFDA/cells + complexes + DCFH-DA (light,
400−700 nm, 10 J cm−2, or in the dark).
S36a in the Supporting Information). In contrast, complexes
4−6 showed ROS generation (mean DCF intensity ∼2500 au)
in both dark and light conditions because of their dark toxicity
(Figure S36b in the Supporting Information).
The selective uptake of annexin V-FITC and PI divides the
total cell population into four quadrants. PI stains the necrotic
cells (upper left), annexin V-FITC stains the early apoptotic
cells (lower right), late apoptotic cells uptake both dyes (upper
right), and live cells are impermeable to both dyes (lower left).
BT474 was pretreated with the complexes for 24 h, exposed to
light for 1 h, and then subjected to annexin V-FITC/PI assay
after a 12 h recuperation period (Figures 8 and S37 in the
Supporting Information). Complexes 1−3 triggered the
respective 65, 50, and 46% of early apoptosis upon light
(400−700 nm) exposure. The observed early apoptotic cell
population in the dark was 26% for 1, 12% for 2, and 22% for 3,
indicating the importance of light exposure to observe
apoptotic cell death. Complexes 4−6 were almost equally
active in the dark and light in the absence of any PDT effect,
resulting in 80, 75, and 70% early apoptosis, respectively. The
cell cycle profiling assay in BT474 cells, based on the amount of
DNA stained by PI, quantified the sub-G1 DNA content. The
Fc-tpy complexes 1−3 showed substantial sub-G1 arrest
(∼30%) only upon photoirradiation with visible light (400−
700 nm), while resulting in only ∼15% cell death in darkness.
The Ph-tpy complexes 4−6 showed almost equal sub-G1
population (∼25−65%) in both dark and light conditions
(Figure S38 in the Supporting Information).
Figure 8. Annexin V-FITC/PI assay showing percent cell population
of BT474 cells pretreated with the complexes (concentration = IC50
values in light) for 24 h in the dark: cells alone (a); cells + PI +
annexin (b); complex 1 in the dark (c) and upon photoexposure (d);
complex 4 in the dark (e) and upon photoexposure (f). Quadrants:
lower left, live cells; lower right, early apoptosis; upper right, late
apoptosis; upper left, dead cells.
biotinylated ligand upon photoactivation leads to the delivery
of the phototoxin selectively into the cancer cells. Complex 1
showed potent light-mediated anticancer effects selectively in
the cancer cells, giving a IC50 value of ∼7 μM. The high toxicity
is attributed to the synergistic cytotoxic effects of the
photoreleased active platinum species and ROS. The phenyl
counterpart 4 paralleled the behavior of cisplatin under similar
conditions, giving a IC50 value of 4 μM. The reduced activity of
the nonbiotinylated counterparts and the reduced cellular
uptake of the complexes in the presence of externally added
biotin exemplify the important role biotin plays in achieving
selective uptake of the complexes in the cancer cells. While the
Fc-tpy complexes 1 and 2 define dual-action photochemother-
apeutic agents, their phenyl analogues 4 and 5 act as cancer-
cell-selective chemotherapeutic agents.
ASSOCIATED CONTENT
* Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.inorg-
chem.6b00680.
4. CONCLUSIONS
■
S
Biotinylated platinum(II) complexes of the Fc-tpy ligand show
the PDT effect in visible light and specific accumulation into
the cancer cells over the normal cells. The Ph-tpy complexes, in
contrast, do not show any apparent PDT effect. In addition, the
Ph-tpy complexes show significant cellular toxicity in the dark.
The Fc-tpy ligand alone is inactive in visible light, while the
same upon binding to platinum(II) displays photoinduced
cytotoxicity. The Fc-tpy complexes 1−3 were found to release
biotinylated ligands in red light (647 nm, 50 mW), and this has
possibly led to photoinitiated covalent DNA binding by an
active platinum species. This is significant because premature
activation of platinum(II) drugs like cisplatin and its analogues
leads to side effects, and a spatially controlled release of the
active species is desired. Furthermore, the release of the
Details of the experimental methods and measurements,
reaction schemes (Schemes S1 and S2), mechanistic
aspect (Scheme S3), NMR (Figures S1, S2, and S5−S7),
ESI-MS (Figures S3 and S8−S13), HPLC chromato-
grams (Figure S4), UV−visible data (Figure S14),
emission data (Figure S15), IR spectra (Figure S16),
cyclic voltammograms (Figures S17 and S18), optimized
structure and frontier molecular orbitals (Figures S19−
S21), photostability (Figures S22−S32), DNA melting
and viscosity (Figure S33), luminescence (Figure S34),
I
DOI: 10.1021/acs.inorgchem.6b00680
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Inorganic Chemistry
MTT assay (Figure S35), FACS analysis (Figures S36−
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38), Cartesian coordinates and electronic transitions in
the complexes (Tables S1 and S2), and platinum content
in ct-DNA (Table S3) (PDF)
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Med. Chem. 2014, S, 1−8.
AUTHOR INFORMATION
Corresponding Authors
(17) Vadlapudi, A. D.; Vadlapatla, R. K.; Pal, D.; Mitra, A. K. AAPS J.
2012, 14, 832−842.
■
(18) Vadlapudi, A. D.; Vadlapatla, R. K.; Mitra, A. K. Curr. Drug
Targets 2012, 13, 994−1003.
*E-mail: paturu@mrdg.iisc.ernet.in. Tel.: +91-80-22932688.
Fax: +91-80-3600999.
(19) Saha, S.; Majumdar, R.; Hussain, A.; Dighe, R. R.; Chakravarty,
A. R. Philos. Trans. R. Soc., A 2013, 371, 20120190.
(20) Sun, Q.; Qian, J.; Tian, H.; Duan, L.; Zhang, W. Chem. Commun.
2014, 50, 8518−8521.
*E-mail: arc@ipc.iisc.ernet.in. Tel.: +91-80-22932533. Fax:
+91-80-23600683.
Author Contributions
(21) Vineberg, J. G.; Wang, T.; Zuniga, E. S.; Ojima, I. J. Med. Chem.
2015, 58, 2406−2416.
The manuscript was written through contributions of all
authors. All authors have given approval to the final version of
the manuscript.
(22) Vineberg, J. G.; Zuniga, E. S.; Kamath, A.; Chen, Y.-J.; Seitz, J.
D.; Ojima, I. J. Med. Chem. 2014, 57, 5777−5791.
(23) Maiti, S.; Park, N.; Han, J. H.; Jeon, H. M.; Lee, J. H.; Bhuniya,
S.; Kang, C.; Kim, J. S. J. Am. Chem. Soc. 2013, 135, 4567−4572.
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J. S. Chem. Commun. 2014, 50, 7690−7693. (b) Park, S.; Kim, E.; Kim,
W. Y.; Kang, C.; Kim, J. S. Chem. Commun. 2015, 51, 9343−9345.
(25) Bhuniya, S.; Maiti, S.; Kim, E.-J.; Lee, H.; Sessler, J. L.; Hong, K.
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Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank the Department of Science and Technology (DST),
Government of India (GOI), for financial support (Grants SR/
S5/MBD-02/2007 and EMR/2015/000742). A.R.C. thanks the
DST for a J. C. Bose national fellowship and the Alexander von
Humboldt Foundation, Germany, for donation of an electro-
chemical system. K.M is a recipient of a BMS fellowship. A.S. is
a recipient of a Dr. D. S. Kothari postdoctoral fellowship of
UGC. P.K. received financial support from the Department of
Biotechnology, GOI. We are thankful to Ms. Urvashi for FACS
analysis. We thank Dr. R. Chakrabarti, Centre for Earth
Sciences, of our institute for the ICP-MS facility. We thank Ms.
Sharada for her help in recording solid-state emission.
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