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Modifying charge and hydrophilicity of simple Ru(II) polypyridyl complexes radically alters biological activities: old complexes, surprising new tricks.
Compounds
capable of light-triggered cytotoxicity are appealing
potential therapeutics, because they can provide spatial and temporal
control over cell killing to reduce side effects in cancer therapy.
Two simple homoleptic Ru(II) polypyridyl complexes with almost-identical
photophysical properties but radically different physiochemical properties
were investigated as agents for photodynamic therapy (PDT). The two
complexes were identical, except for the incorporation of six sulfonic
acids into the ligands of one complex, resulting in a compound carrying
an overall −4 charge. The negatively charged compound exhibited
significant light-mediated cytotoxicity, and, importantly, the negative
charges resulted in radical alterations of the biological activity,
compared to the positively charged analogue, including complete abrogation
of toxicity in the dark. The charges also altered the subcellular
localization properties, mechanism of action, and even the mechanism
of cell death. The incorporation of negative charged ligands provides
a simple chemical approach to modify the biological properties of
light-activated Ru(II) cytotoxic agents.
## Introduction
Introduction Metal
complexes have been studied for decades as potential cytotoxic
agents, because of the unprecedented and continued success of cisplatin
as a chemotherapeutic. 1 , 2 Investigations into Ru(II) polypyridyl
complexes have been particularly extensive, because of their ease
of synthesis, appealing chemical, physical, and photophysical characteristics,
and their high affinities for nucleic acids. Most early studies focused
on characterizing the in vitro interactions of these
complexes with DNA, 3 − 6 and quantifying the potencies of the compounds both as traditional
cytotoxic agents and as light-activated agents for photodynamic therapy
(PDT) or phototherapy. In recent years, attention has shifted to understanding
the cellular localization 7 − 10 properties of Ru(II) complexes, along with their
mechanisms of cellular uptake 10 − 15 and cytotoxicity, providing a deeper understanding of how these
compounds elicit their biological activities. An attractive
feature of Ru(II) polypyridyl complexes that makes
them particularly useful for applications as biological probes and
effectors is the diversity of the chemical structures that are readily
available through modifications of the coordinated ligands. 16 However, most biological studies have focused
on complexes that carry an overall charge of +2 or greater. This significantly
limits the chemical structures and physical properties of the molecules
under investigation. A question that occurred to us was this: To what
degree could the biological properties of a chemically inert Ru(II)
complex be tuned by chemical modification of the ligands surrounding
the metal center? To address this question, we have investigated
the biological activities,
cellular uptake, localization, and mechanism of cell killing of two
simple Ru(II) complexes that are commonly used dyes for solar cell
research or biological staining, but have not been previously explored
as PDT agents. Ru(bathophenanthroline) 3 ( 1 ; see Chart 1 ) is a hydrophobic molecule
with a high DNA binding affinity, 17 , 18 while Ru(bathophenanthroline
disulfonate) 3 ( 2 ) is hydrophilic and possesses
a high affinity for proteins. 19 − 21 Both complexes are efficient
singlet oxygen ( 1 O 2 ) generators with the same
quantum yields for 1 O 2 production (Φ Δ ) and similar molar extinction coefficients (ε). 22 Both are luminescent, allowing for analysis
by fluorescence microscopy. However, although the photophysical properties
of the compounds are almost identical, the physical properties of
the two Ru(II) compounds are quite dissimilar (see Table 1 ). Compound 1 carries an overall charge of +2, while 2 has
an overall charge of −4. They also have very different hydrophilicities,
as indicated by their partition coefficient or log P values. Given the radically different physical properties of the
complexes, we anticipated differences in their biological effects
that could provide information for future rational design of light-activated
cytotoxic agents. Chart 1 Structures of Compounds 1 and 2 Table 1 Physical and Photophysical
Properties
of 1 and 2 property compound 1 compound 2 charge +2 –4 log P 1.8 ± 0.02 –2.2 ± 0.12 λ max (nm) 460 a 462 a ε (M –1 cm –1 ) 29,500 a 29, 300 a λ em (nm) 632 a 632 a Φ PL 0.101 b 0.176 c Φ Δ 0.42 a , d 0.43 a , d a From ref ( 22 ). b From ref ( 24 ). c From ref ( 25 ). d Determined in D 2 O. Considering the established dogma
of the field, it would be expected
that the negatively charged compound 2 would not enter
cells 23 and would suffer low efficacy,
while the positively charged, DNA-binding compound 1 would
prove the more effective PDT agent. Instead, our findings run counter
to this prediction. Here, we show large differences in potency, cellular
uptake, localization, and mechanism of cytotoxicity of these agents,
illustrating that radical modulation of biological properties is possible
with ligand modifications of simple homoleptic Ru(II) polypyridyl
complexes that are substitutionally inert. Most importantly, our results
also prove that a significantly greater range of charge states and
physical properties of Ru(II) complexes are compatible with potential
application as PDT agents.
## Results
Results DNA Damage and Cytotoxicity As both 1 and 2 are efficient catalysts
for the light-activated generation
of 1 O 2 (Φ Δ = 0.42, 0.43), 22 it was expected that the two compounds would
act as sensitizers for PDT. Accordingly, their DNA damaging properties
were assessed with pUC19 plasmid DNA and analyzed by gel electrophoresis
(Figure 1 ). Each compound was incubated with
plasmid and irradiated with 40 J/cm 2 of visible light (>400
nm) or kept in the dark. Compound 1 is known to bind
strongly with DNA, and precipitation of the DNA with the complex was
observed at concentrations above 31.3 μM both in the light and
in the dark (Figure 1 A). In contrast, 2 is a much more hydrophilic molecule, and the negatively
charged sulfonate functional groups were anticipated to cause electrostatic
repulsions between the complex and the negatively charged backbone
of the DNA. Consistent with low DNA affinity, 26 no DNA precipitation or smearing was observed with up to 500 μM
of 2 (Figure 1 B). When exposed
to light, both 1 and 2 induced single strand
DNA breaks, creating relaxed circular plasmid. This is likely due
to the photogeneration of 1 O 2 that mediates
the DNA damage. However, for 2 , the amount of relaxed
circle plasmid did not exhibit any concentration dependence above
125 μM, suggesting either a reduction in Φ Δ as the concentration of the complex is increased, or alternative
quenching mechanisms that impede DNA damage. 27 Figure 1 Agarose
gel electrophoresis of pUC19 with increasing concentrations
of (A) 1 and (B) 2 in the dark or irradiated
(λ > 400 nm). Lanes 1 and 12, DNA molecular weight standard;
lane 2, linear (reaction with EcoR1); lane 3, relaxed circle (reaction
with Cu(phen) 2 ); lanes 4 – 11, 0, 8.25,
16.5, 31.3, 62.5, 125, 250, and 500 μM compound. Cytotoxicity
dose response of (C) 1 and (D) 2 in the
dark (open squares) or irradiated (closed circles). HL60 cells were
incubated for 72 h with compound prior to quantification of viability. As both compounds are capable
of light-induced DNA damage, the
cytotoxicity of 1 and 2 were evaluated in
the A549 human non-small cell lung cancer, the HL60 human promyelocytic
leukemia, and the Jurkat human T lymphoblastoid cell lines in the
presence and absence of 7 J/cm 2 of >400 nm light. The
IC 50 values across the cell lines for 1 ranged
from
0.62 to 3.75 μM in the dark. Upon irradiation, potency was increased
to a range of 0.075 to 0.35 μM, resulting in an average phototoxicity
index (PI = IC 50 (dark)/IC 50 (light) of 10-
to 20-fold (see Figure 1 , Table 2 )). Table 2 Cytotoxicity IC 50 Values
(μM) in Various Cell Lines a IC 50 Value (μM) compound HL60, light HL60, dark A549, light A549, dark Jurkat, light Jurkat, dark PI b Jurkat 1 0.35 ± 0.18 3.75 ± 0.18 0.11 ± 0.02 0.62 ± 0.08 0.075 ± 0.004 1.63 ± 0.11 21.7 2 9.81 ± 1.09 >300 17.25 ± 9.82 >300 3.31 ± 0.36 >300 >90 cisplatin 3.1 ± 0.2 3.1 ± 0.2 3.4 ± 0.6 3.5 ± 0.2 0.5 ± 0.07 0.5 ± 0.07 1 a IC 50 values are averages
from three measurements. b The phototoxicity index (PI) is
the ratio of the dark and light IC 50 values. In marked contrast to the high toxicity
of 1 in the
absence of light, no toxicity was observed in the dark with 2 across all cell lines at concentrations up to 300 μM.
However, compound 2 was effective in killing cells when
irradiated, with IC 50 values ranging from 3.3 μM
to 17.3 μM, consistent with the concentrations required for in vitro DNA damage. This provides for a large therapeutic
window, as no cell death is observed for samples in the absence of
irradiation. Surprisingly, not only was compound 1 toxic to cells
upon irradiation, it also induced cell death far more rapidly in HL60
cells than traditional DNA damaging agents such as cisplatin. Complete
cell death was observed within 2 h of irradiation with 1 (see Figure S1 in the Supporting Information ). The compound also induced cell death in the dark, but more slowly,
with ∼30% viable cells remaining at 24 h. In marked contrast
to 1 , compound 2 induced cell death after
irradiation only after a long delay, with 70% viable cells remaining
at 24 h and 55% remaining at 48 h. The significant disparity
in the potency, phototoxicity index (PI),
and rates of cell killing for the two compounds despite their equivalent
abilities to sensitize 1 O 2 strongly suggested
that they were acting through different cellular mechanisms, possibly
by interacting with different biological targets. As the compounds
are substitutionally inert, they are unlikely to covalently modify
biomolecules. This complicates target identification by isolation
of protein or nucleic acid components of the cells, so the subcellular
localization was investigated instead. Cellular Uptake and Subcellular
Localization Both 1 and 2 are emissive,
allowing for direct visualization
in cells. Flow cytometry and fluorescence microscopy were used to
provide relative uptake values, the time dependence of compound uptake,
and information on the subcellular localization of the compounds.
To optimize signal intensity, 1 was assayed at 5 μM
while 20 μM was required for 2 . Flow cytometry
with A549 cells revealed greater uptake of 1 compared
with 2 at time points of 2 and 24 h, with an 11.6-fold
and 8.2-fold difference in signal, respectively (see Figure S2 in the Supporting Information ). Between the time
points of 2 and 24 h, the average emission of cells incubated with 1 increased by 2.8-fold while the amount of 2 increased by 4-fold. This data was supported by direct quantification
of the number of ruthenium atoms per cell using inductively coupled
plasma–optical emission spectroscopy (ICP-OES; see Tables S1 and S2 in the Supporting Information ). The relative amount of compound 1 in cells versus
the cell media increased ∼4-fold from 2 h to 24 h, from 5.4%
to 22%. 28 Much less of compound 2 entered the cells, with a maximal uptake of 0.7% at 24 h. While
low, this degree of uptake of the −4-charged 2 is comparable to cisplatin, a neutral compound, dosed at the same
concentration (20 μM, 0.8% at 24 h). The emission of 1 and 2 was also measured in A549 cells using
an ApoTome microscope (the adherent cell line was chosen to facilitate
the required wash steps to allow for the use of fluorescent reporters).
The relative rates of uptake of each of the complexes were analyzed
as a function of time, and both 1 and 2 were
visible inside cells as early as 2 h after compound addition (see Figure S3 in the Supporting Information ), consistent
with the flow cytometry and ICP-OES results. Images were taken at
2, 8, 18, and 24 h, and intracellular levels of both complexes appeared
to plateau at 8 h. The uptake data showed good agreement between the
three techniques and two cell lines, suggesting similar behavior in
adherent and suspension cell lines. Differences in the intracellular
localization of 1 and 2 were evaluated by
determining colocalization
of luminescence of the compounds and fluorescent markers of organelles
in A549 cells. Overlap in signals between the compounds, cellular
nucleus, mitochondria, and lysosome was measured over a 24 h period.
Compounds 1 and 2 have very different localization
profiles, as exemplified by the imaging 8 h after dosing (Figure 2 ). Compound 1 substantially localized
to lysosomes and the mitochondria in the absence of light, while 2 remained primarily in the cytosol. Exposure to light did
have an impact on compound localization, where 1 induced
nuclear localization of both the mitochondrial and lysosome markers
(see Figures 2 A and 2 B, top), indicating that photoinduced damage mediated by 1 reduced the integrity of the nuclear membrane. In contrast, 2 was primarily observed in lysosomes after irradiation, and
did not co-localize with mitochondria (Figures 2 A and 2 B, bottom). In addition, irradiation
in the presence of 2 did not result in the appearance
of organelle markers in the nucleus, suggesting the nuclear membrane
remained intact. Neither 1 nor 2 was found
to associate with the plasma or nuclear membranes. Figure 2 ApoTome microscopy showing
subcellular localization of 1 and 2 at 8
h. Co-localization of 1 and 2 in mitochondria
or lysosomes is indicated by the apparent
yellow emission. (A) Mitotracker Green FM was used to image mitochondria.
(B) Lysotracker Green DND-26 was used to image lysosomes. Red color
denotes intrinsic emission of 1 and 2 , whereas
blue color denotes Hoechst staining of the nucleus. The yellow color
occurs due to overlap of the red emission from the ruthenium complexes
and green emission of the organelle-specific dyes, indicating colocalization.
Compound 1 localizes in both the mitochondria and the
lysosomes while 2 was not predominantly found in either
organelle. Mitochondrial Function
and Time Dependence for Cell Death As compound 1 appeared to localize within mitochondria,
it was hypothesized that this may account for its high toxicity in
the absence of irradiation. To determine if either 1 or 2 caused a reduction in mitochondrial function, mitochondrial
membrane potential was measured using tetramethylrhodamine ethyl ester
(TMRE) (see Figure 3 ). TMRE is a cationic dye
that accumulates in active mitochondria as a result of the negative
membrane potential (Δψm). Inactive or depolarized mitochondria
exhibit a decreased membrane potential, and TMRE does not localize
in these organelles. Compound 1 induced rapid and complete
depolarization of mitochondria both in the dark and upon irradiation
(Figure 3 B, 3 C). However,
surprisingly, cell viability did not parallel mitochondrial potential.
While mitochondrial function was completely impaired at 2 h post treatment
with 1 in the dark, viability decreased slowly, with
44% ± 6% viable cells remaining after 24 h. In contrast, both
mitochondrial function and cell viability fell to 0% within 2 h of
irradiation. Thus, while 1 completely impedes mitochondrial
function within 2 h even in the absence of light, irradiation induced
additional damage that results in rapid cell death, along with the
loss of mitochondrial function. Figure 3 (A) Example images and quantification
of the emission of tetramethylrhodamine
ethyl ester (TMRE) in the presence and absence of 1 and 2 . Compound 1 does not show increased TMRE emission
over background emission of 1 , while compound 2 does not add to the TMRE emission. Mitochondrial potential and cell
viability of A549 cells as a function of time for (B) 1 , in the dark; (C) 1 , irradiated; (D) 2 , in the dark; and (E) 2 , irradiated. TMRE was used
to quantify membrane potential; values are relative to a no-compound
control value of 100. In contrast to compound 1 , 2 did
not
significantly reduce the mitochondrial potential over a 24 h period
either when irradiated or kept in the dark. Irradiation did reduce
cell viability to 70% ± 12% at 2 h and 59% ± 4% after 24
h, but this occurred without a significant decrease in the relative
mitochondrial potential (Figure 3 E), indicating
that 2 does not act through inhibition of mitochondrial
function. These results strongly suggest that mitochondrial failure
plays a role in the dark toxicity of 1 , and the lack
of mitochondrial localization and inhibition may explain the comparatively
low dark toxicity of 2 . Mechanism of Cell Death Most compounds used for PDT
that generate singlet oxygen induce apoptosis. Given the different
cellular localization properties and time profiles for cell death
induced by 1 and 2 , the mechanism of cell
death was investigated. Indicators of apoptotic cell death (activation
of PARP and caspase 3 through proteolysis) were determined in HL60
cells treated with either 1 or 2 (Figure 4 A). The known apoptotic-inducing compounds, cisplatin
and doxorubicin, were run in parallel. Compound 1 induced
the proteolytic activation of both PARP and caspase 3 within 2 h of
irradiation (Figure 4 A). In the absence of
light, PARP and caspase 3 were observed with 1 , but only
after 18 and 24 h, and to a lesser degree. In the absence of irradiation,
the amount of inactive procaspase 3 did not change. Exposure
of 2 to light also induced PARP cleavage as early as
2 h, but unlike 1 , increasing amounts of cleaved PARP
were observed over the course of 24 h (Figure 4 A). The increase in the level of activated caspase 3 also occurred
on a slower time scale than PARP cleavage, with the protein initially
observed at 18 and 24 h. This suggests that the irradiated samples
undergo apoptosis that is not primarily signaled through caspase 3.
Conversely, cells treated with 2 and protected from light
did not display PARP or caspase 3 cleavage, which is consistent with
viability measurements indicating no cytotoxicity in the absence of
light. The level of procaspase 3 also did not change over 24 h, further
confirming a lack of cytotoxicity under these conditions. Since 2 showed strong PARP induction without significant
caspase 3 activation, as compared to 1 and cisplatin
or doxorubicin, a mechanism of cell death through necrosis was explored.
The level of an alternate 55 kDa PARP fragment was determined by immunoblot
as a marker for necrosis ( see Figure S6 in the
Supporting Information ), with 10% (v/v) ethanol used as a positive
control. 29 Exposure of HL60 cells to 1 produced this fragment at significant levels both when protected
from light and when irradiated, which is consistent with necrosis.
Cisplatin and doxorubicin also produced this cleavage product, indicating
that some cells had progressed into necrosis ( Figure S6 in the Supporting Information ). In contrast, cells
exposed to 2 , both in the presence and absence of irradiation,
produced a lower level of the 55 kDa PARP fragment, similar to the
untreated cells, suggesting necrosis is not a significant cell death
pathway for this compound. Figure 4 (A) Western blot analysis of cleaved PARP, caspase
3, and pro-caspase
3. GAPDH is used as a loading control. (B) Agarose gel electrophoresis
of genomic DNA harvested from HL60 cells after treatment with various
agents. Lanes 1 and 9, ladder; lane 2, no compound control; lane 3,
10% EtOH, 24 h (necrosis control); lane 4, cisplatin, 24 h (apoptosis
control); lane 5, 1 , irradiated, 8 h; lane 6, 1 , in the dark, 8 h; lane 7, 2 , irradiated, 24 h; and
lane 8, 2 , in the dark, 24 h. To support the assessment of the disparate mechanisms of
cell death
induced by 1 and 2 , the degradation pattern
of genomic DNA was investigated. DNA laddering is observed as a result
of DNA fragmentation stemming from the executionary phase of apoptosis.
In contrast, necrotic cell death lacks this characteristic laddering
effect, allowing differentiation between these two mechanisms. HL60
cells were exposed to the Ru(II) complexes, cisplatin, and 10% ethanol,
followed by genomic DNA isolation and resolution by gel electrophoresis.
As expected, the apoptosis inducer, cisplatin, initiated DNA fragmentation,
resulting in a laddering pattern on the gel (Figure 4 B). This laddering was absent in the cells treated with ethanol
and compounds 1 and 2 in the dark. However,
both compounds 1 and 2 displayed similar
laddering patterns as cisplatin when irradiated, suggesting apoptosis
is a contributing cell death pathway for both compounds when irradiated.
In contrast, given the cytotoxicity of 1 in the dark
and the presence of the 55 kDa PARP fragment, it appears that necrotic
cell death is a significant pathway for 1 . For compound 2 , the absence of DNA laddering is a result of the lack of
cytotoxicity of the negatively charged compound. Flow cytometry
was also employed to further corroborate the assessment
of the mechanism of cell death using Annexin V/propidium iodide (PI)
or Annexin V/Hoechst staining to differentiate apoptotic versus necrotic
cell death. Fluorescent Annexin V conjugates recognize the translocation
of phosphatidylserine to the outer leaflet of the cell membrane during
apoptosis. PI and Hoechst were used as nuclear stains to distinguish
between live and dead cells; while PI is most commonly used, Hoechst
was also applied, because of spectral interference of 1 and 2 with PI. The presence of significant populations
of PI positive cells in the absence of Annexin V staining for 1 both in light and in the dark demonstrate that necrosis
is a significant cell death pathway for this compound (see Figures S7 and S8 in the Supporting Information ; 5% and 14%, respectively, at 2 h). In contrast, compound 2 produced large populations of Annexin V positive cells (55%
at 24 h), showing an apoptotic pathway (see Figures
S9 and S10 in the Supporting Information ). Similarly, Hoechst
staining was consistent with necrosis as a contributing pathway for 1 in the dark and in light, while 2 induced apoptosis.
## DNA Damage and Cytotoxicity
DNA Damage and Cytotoxicity As both 1 and 2 are efficient catalysts
for the light-activated generation
of 1 O 2 (Φ Δ = 0.42, 0.43), 22 it was expected that the two compounds would
act as sensitizers for PDT. Accordingly, their DNA damaging properties
were assessed with pUC19 plasmid DNA and analyzed by gel electrophoresis
(Figure 1 ). Each compound was incubated with
plasmid and irradiated with 40 J/cm 2 of visible light (>400
nm) or kept in the dark. Compound 1 is known to bind
strongly with DNA, and precipitation of the DNA with the complex was
observed at concentrations above 31.3 μM both in the light and
in the dark (Figure 1 A). In contrast, 2 is a much more hydrophilic molecule, and the negatively
charged sulfonate functional groups were anticipated to cause electrostatic
repulsions between the complex and the negatively charged backbone
of the DNA. Consistent with low DNA affinity, 26 no DNA precipitation or smearing was observed with up to 500 μM
of 2 (Figure 1 B). When exposed
to light, both 1 and 2 induced single strand
DNA breaks, creating relaxed circular plasmid. This is likely due
to the photogeneration of 1 O 2 that mediates
the DNA damage. However, for 2 , the amount of relaxed
circle plasmid did not exhibit any concentration dependence above
125 μM, suggesting either a reduction in Φ Δ as the concentration of the complex is increased, or alternative
quenching mechanisms that impede DNA damage. 27 Figure 1 Agarose
gel electrophoresis of pUC19 with increasing concentrations
of (A) 1 and (B) 2 in the dark or irradiated
(λ > 400 nm). Lanes 1 and 12, DNA molecular weight standard;
lane 2, linear (reaction with EcoR1); lane 3, relaxed circle (reaction
with Cu(phen) 2 ); lanes 4 – 11, 0, 8.25,
16.5, 31.3, 62.5, 125, 250, and 500 μM compound. Cytotoxicity
dose response of (C) 1 and (D) 2 in the
dark (open squares) or irradiated (closed circles). HL60 cells were
incubated for 72 h with compound prior to quantification of viability. As both compounds are capable
of light-induced DNA damage, the
cytotoxicity of 1 and 2 were evaluated in
the A549 human non-small cell lung cancer, the HL60 human promyelocytic
leukemia, and the Jurkat human T lymphoblastoid cell lines in the
presence and absence of 7 J/cm 2 of >400 nm light. The
IC 50 values across the cell lines for 1 ranged
from
0.62 to 3.75 μM in the dark. Upon irradiation, potency was increased
to a range of 0.075 to 0.35 μM, resulting in an average phototoxicity
index (PI = IC 50 (dark)/IC 50 (light) of 10-
to 20-fold (see Figure 1 , Table 2 )). Table 2 Cytotoxicity IC 50 Values
(μM) in Various Cell Lines a IC 50 Value (μM) compound HL60, light HL60, dark A549, light A549, dark Jurkat, light Jurkat, dark PI b Jurkat 1 0.35 ± 0.18 3.75 ± 0.18 0.11 ± 0.02 0.62 ± 0.08 0.075 ± 0.004 1.63 ± 0.11 21.7 2 9.81 ± 1.09 >300 17.25 ± 9.82 >300 3.31 ± 0.36 >300 >90 cisplatin 3.1 ± 0.2 3.1 ± 0.2 3.4 ± 0.6 3.5 ± 0.2 0.5 ± 0.07 0.5 ± 0.07 1 a IC 50 values are averages
from three measurements. b The phototoxicity index (PI) is
the ratio of the dark and light IC 50 values. In marked contrast to the high toxicity
of 1 in the
absence of light, no toxicity was observed in the dark with 2 across all cell lines at concentrations up to 300 μM.
However, compound 2 was effective in killing cells when
irradiated, with IC 50 values ranging from 3.3 μM
to 17.3 μM, consistent with the concentrations required for in vitro DNA damage. This provides for a large therapeutic
window, as no cell death is observed for samples in the absence of
irradiation. Surprisingly, not only was compound 1 toxic to cells
upon irradiation, it also induced cell death far more rapidly in HL60
cells than traditional DNA damaging agents such as cisplatin. Complete
cell death was observed within 2 h of irradiation with 1 (see Figure S1 in the Supporting Information ). The compound also induced cell death in the dark, but more slowly,
with ∼30% viable cells remaining at 24 h. In marked contrast
to 1 , compound 2 induced cell death after
irradiation only after a long delay, with 70% viable cells remaining
at 24 h and 55% remaining at 48 h. The significant disparity
in the potency, phototoxicity index (PI),
and rates of cell killing for the two compounds despite their equivalent
abilities to sensitize 1 O 2 strongly suggested
that they were acting through different cellular mechanisms, possibly
by interacting with different biological targets. As the compounds
are substitutionally inert, they are unlikely to covalently modify
biomolecules. This complicates target identification by isolation
of protein or nucleic acid components of the cells, so the subcellular
localization was investigated instead.
## Cellular Uptake and Subcellular
Localization
Cellular Uptake and Subcellular
Localization Both 1 and 2 are emissive,
allowing for direct visualization
in cells. Flow cytometry and fluorescence microscopy were used to
provide relative uptake values, the time dependence of compound uptake,
and information on the subcellular localization of the compounds.
To optimize signal intensity, 1 was assayed at 5 μM
while 20 μM was required for 2 . Flow cytometry
with A549 cells revealed greater uptake of 1 compared
with 2 at time points of 2 and 24 h, with an 11.6-fold
and 8.2-fold difference in signal, respectively (see Figure S2 in the Supporting Information ). Between the time
points of 2 and 24 h, the average emission of cells incubated with 1 increased by 2.8-fold while the amount of 2 increased by 4-fold. This data was supported by direct quantification
of the number of ruthenium atoms per cell using inductively coupled
plasma–optical emission spectroscopy (ICP-OES; see Tables S1 and S2 in the Supporting Information ). The relative amount of compound 1 in cells versus
the cell media increased ∼4-fold from 2 h to 24 h, from 5.4%
to 22%. 28 Much less of compound 2 entered the cells, with a maximal uptake of 0.7% at 24 h. While
low, this degree of uptake of the −4-charged 2 is comparable to cisplatin, a neutral compound, dosed at the same
concentration (20 μM, 0.8% at 24 h). The emission of 1 and 2 was also measured in A549 cells using
an ApoTome microscope (the adherent cell line was chosen to facilitate
the required wash steps to allow for the use of fluorescent reporters).
The relative rates of uptake of each of the complexes were analyzed
as a function of time, and both 1 and 2 were
visible inside cells as early as 2 h after compound addition (see Figure S3 in the Supporting Information ), consistent
with the flow cytometry and ICP-OES results. Images were taken at
2, 8, 18, and 24 h, and intracellular levels of both complexes appeared
to plateau at 8 h. The uptake data showed good agreement between the
three techniques and two cell lines, suggesting similar behavior in
adherent and suspension cell lines. Differences in the intracellular
localization of 1 and 2 were evaluated by
determining colocalization
of luminescence of the compounds and fluorescent markers of organelles
in A549 cells. Overlap in signals between the compounds, cellular
nucleus, mitochondria, and lysosome was measured over a 24 h period.
Compounds 1 and 2 have very different localization
profiles, as exemplified by the imaging 8 h after dosing (Figure 2 ). Compound 1 substantially localized
to lysosomes and the mitochondria in the absence of light, while 2 remained primarily in the cytosol. Exposure to light did
have an impact on compound localization, where 1 induced
nuclear localization of both the mitochondrial and lysosome markers
(see Figures 2 A and 2 B, top), indicating that photoinduced damage mediated by 1 reduced the integrity of the nuclear membrane. In contrast, 2 was primarily observed in lysosomes after irradiation, and
did not co-localize with mitochondria (Figures 2 A and 2 B, bottom). In addition, irradiation
in the presence of 2 did not result in the appearance
of organelle markers in the nucleus, suggesting the nuclear membrane
remained intact. Neither 1 nor 2 was found
to associate with the plasma or nuclear membranes. Figure 2 ApoTome microscopy showing
subcellular localization of 1 and 2 at 8
h. Co-localization of 1 and 2 in mitochondria
or lysosomes is indicated by the apparent
yellow emission. (A) Mitotracker Green FM was used to image mitochondria.
(B) Lysotracker Green DND-26 was used to image lysosomes. Red color
denotes intrinsic emission of 1 and 2 , whereas
blue color denotes Hoechst staining of the nucleus. The yellow color
occurs due to overlap of the red emission from the ruthenium complexes
and green emission of the organelle-specific dyes, indicating colocalization.
Compound 1 localizes in both the mitochondria and the
lysosomes while 2 was not predominantly found in either
organelle.
## Mitochondrial Function
and Time Dependence for Cell Death
Mitochondrial Function
and Time Dependence for Cell Death As compound 1 appeared to localize within mitochondria,
it was hypothesized that this may account for its high toxicity in
the absence of irradiation. To determine if either 1 or 2 caused a reduction in mitochondrial function, mitochondrial
membrane potential was measured using tetramethylrhodamine ethyl ester
(TMRE) (see Figure 3 ). TMRE is a cationic dye
that accumulates in active mitochondria as a result of the negative
membrane potential (Δψm). Inactive or depolarized mitochondria
exhibit a decreased membrane potential, and TMRE does not localize
in these organelles. Compound 1 induced rapid and complete
depolarization of mitochondria both in the dark and upon irradiation
(Figure 3 B, 3 C). However,
surprisingly, cell viability did not parallel mitochondrial potential.
While mitochondrial function was completely impaired at 2 h post treatment
with 1 in the dark, viability decreased slowly, with
44% ± 6% viable cells remaining after 24 h. In contrast, both
mitochondrial function and cell viability fell to 0% within 2 h of
irradiation. Thus, while 1 completely impedes mitochondrial
function within 2 h even in the absence of light, irradiation induced
additional damage that results in rapid cell death, along with the
loss of mitochondrial function. Figure 3 (A) Example images and quantification
of the emission of tetramethylrhodamine
ethyl ester (TMRE) in the presence and absence of 1 and 2 . Compound 1 does not show increased TMRE emission
over background emission of 1 , while compound 2 does not add to the TMRE emission. Mitochondrial potential and cell
viability of A549 cells as a function of time for (B) 1 , in the dark; (C) 1 , irradiated; (D) 2 , in the dark; and (E) 2 , irradiated. TMRE was used
to quantify membrane potential; values are relative to a no-compound
control value of 100. In contrast to compound 1 , 2 did
not
significantly reduce the mitochondrial potential over a 24 h period
either when irradiated or kept in the dark. Irradiation did reduce
cell viability to 70% ± 12% at 2 h and 59% ± 4% after 24
h, but this occurred without a significant decrease in the relative
mitochondrial potential (Figure 3 E), indicating
that 2 does not act through inhibition of mitochondrial
function. These results strongly suggest that mitochondrial failure
plays a role in the dark toxicity of 1 , and the lack
of mitochondrial localization and inhibition may explain the comparatively
low dark toxicity of 2 .
## Mechanism of Cell Death
Mechanism of Cell Death Most compounds used for PDT
that generate singlet oxygen induce apoptosis. Given the different
cellular localization properties and time profiles for cell death
induced by 1 and 2 , the mechanism of cell
death was investigated. Indicators of apoptotic cell death (activation
of PARP and caspase 3 through proteolysis) were determined in HL60
cells treated with either 1 or 2 (Figure 4 A). The known apoptotic-inducing compounds, cisplatin
and doxorubicin, were run in parallel. Compound 1 induced
the proteolytic activation of both PARP and caspase 3 within 2 h of
irradiation (Figure 4 A). In the absence of
light, PARP and caspase 3 were observed with 1 , but only
after 18 and 24 h, and to a lesser degree. In the absence of irradiation,
the amount of inactive procaspase 3 did not change. Exposure
of 2 to light also induced PARP cleavage as early as
2 h, but unlike 1 , increasing amounts of cleaved PARP
were observed over the course of 24 h (Figure 4 A). The increase in the level of activated caspase 3 also occurred
on a slower time scale than PARP cleavage, with the protein initially
observed at 18 and 24 h. This suggests that the irradiated samples
undergo apoptosis that is not primarily signaled through caspase 3.
Conversely, cells treated with 2 and protected from light
did not display PARP or caspase 3 cleavage, which is consistent with
viability measurements indicating no cytotoxicity in the absence of
light. The level of procaspase 3 also did not change over 24 h, further
confirming a lack of cytotoxicity under these conditions. Since 2 showed strong PARP induction without significant
caspase 3 activation, as compared to 1 and cisplatin
or doxorubicin, a mechanism of cell death through necrosis was explored.
The level of an alternate 55 kDa PARP fragment was determined by immunoblot
as a marker for necrosis ( see Figure S6 in the
Supporting Information ), with 10% (v/v) ethanol used as a positive
control. 29 Exposure of HL60 cells to 1 produced this fragment at significant levels both when protected
from light and when irradiated, which is consistent with necrosis.
Cisplatin and doxorubicin also produced this cleavage product, indicating
that some cells had progressed into necrosis ( Figure S6 in the Supporting Information ). In contrast, cells
exposed to 2 , both in the presence and absence of irradiation,
produced a lower level of the 55 kDa PARP fragment, similar to the
untreated cells, suggesting necrosis is not a significant cell death
pathway for this compound. Figure 4 (A) Western blot analysis of cleaved PARP, caspase
3, and pro-caspase
3. GAPDH is used as a loading control. (B) Agarose gel electrophoresis
of genomic DNA harvested from HL60 cells after treatment with various
agents. Lanes 1 and 9, ladder; lane 2, no compound control; lane 3,
10% EtOH, 24 h (necrosis control); lane 4, cisplatin, 24 h (apoptosis
control); lane 5, 1 , irradiated, 8 h; lane 6, 1 , in the dark, 8 h; lane 7, 2 , irradiated, 24 h; and
lane 8, 2 , in the dark, 24 h. To support the assessment of the disparate mechanisms of
cell death
induced by 1 and 2 , the degradation pattern
of genomic DNA was investigated. DNA laddering is observed as a result
of DNA fragmentation stemming from the executionary phase of apoptosis.
In contrast, necrotic cell death lacks this characteristic laddering
effect, allowing differentiation between these two mechanisms. HL60
cells were exposed to the Ru(II) complexes, cisplatin, and 10% ethanol,
followed by genomic DNA isolation and resolution by gel electrophoresis.
As expected, the apoptosis inducer, cisplatin, initiated DNA fragmentation,
resulting in a laddering pattern on the gel (Figure 4 B). This laddering was absent in the cells treated with ethanol
and compounds 1 and 2 in the dark. However,
both compounds 1 and 2 displayed similar
laddering patterns as cisplatin when irradiated, suggesting apoptosis
is a contributing cell death pathway for both compounds when irradiated.
In contrast, given the cytotoxicity of 1 in the dark
and the presence of the 55 kDa PARP fragment, it appears that necrotic
cell death is a significant pathway for 1 . For compound 2 , the absence of DNA laddering is a result of the lack of
cytotoxicity of the negatively charged compound. Flow cytometry
was also employed to further corroborate the assessment
of the mechanism of cell death using Annexin V/propidium iodide (PI)
or Annexin V/Hoechst staining to differentiate apoptotic versus necrotic
cell death. Fluorescent Annexin V conjugates recognize the translocation
of phosphatidylserine to the outer leaflet of the cell membrane during
apoptosis. PI and Hoechst were used as nuclear stains to distinguish
between live and dead cells; while PI is most commonly used, Hoechst
was also applied, because of spectral interference of 1 and 2 with PI. The presence of significant populations
of PI positive cells in the absence of Annexin V staining for 1 both in light and in the dark demonstrate that necrosis
is a significant cell death pathway for this compound (see Figures S7 and S8 in the Supporting Information ; 5% and 14%, respectively, at 2 h). In contrast, compound 2 produced large populations of Annexin V positive cells (55%
at 24 h), showing an apoptotic pathway (see Figures
S9 and S10 in the Supporting Information ). Similarly, Hoechst
staining was consistent with necrosis as a contributing pathway for 1 in the dark and in light, while 2 induced apoptosis.
## Discussion
Discussion The goal of phototherapy is to achieve cell death
in cancerous
or abnormal tissues that are irradiated, while protecting healthy
tissues that are not exposed to light. As a result, the compounds
developed for this application should possess large “therapeutic
windows” where the toxicity in the dark is minimized. This
has proven to be challenging for many inorganic agents developed for
PDT applications. The rational design of new and more efficacious
compounds would be facilitated by (1) a better understanding of the
mechanisms of action that induce dark toxicity for promising PDT agents,
and (2) the identification of chemical features that eliminate dark
toxicity. The investigations of the biological activities of these
two simple Ru(II) complexes provide guidance for both approaches toward
improved PDT agents. While compound 1 demonstrated
notable potency when
irradiated (0.075–0.35 μM, depending on cell line), the
toxicity of the compound in the dark (0.62 to 3.75 μM) reduces
its potential as a PDT agent. Imaging studies showed both mitochondrial
and lysosomal localization, and assessment of mitochondrial potential
indicated that 1 immediately inhibits mitochondrial function.
However, the disconnect between the time dependence of the inhibition
of mitochondrial function and the reduction in cell viability in the
dark shows that disruption of mitochondrial membrane potential does
not lead to rapid cell death. It is possible that the cells treated
with 1 and kept in the dark survive for several hours,
despite the complete abrogation of mitochondrial function, due to
the Warburg effect, where cancer cells exhibit a reduced reliance
on oxidative phosphorylation and increased dependence on glycolysis
for energy production. After 72 h, cell death is complete for 1 in the dark.
The obliteration of mitochondrial function would explain the observation
of necrotic cell death in this case. 30 In
contrast, when 1 is irradiated, it appears that a combination
of necrotic and apoptotic pathways are activated. Cell death is so
rapid (with most cells undergoing death at 2 h, as indicated by Trypan
Blue staining) that necrosis is likely to be a primary pathway for
a majority of cells. The breakdown of membrane integrity is apparent
not only for the plasma membrane, but also for the membranes of organelles,
as cells treated with 1 and irradiated showed nonspecific
nuclear localization of both Mitotracker and Lysotracker. The high
dark toxicity, mitochondrial localization, and induction of necrotic
death pathways may reduce the potential of compounds structurally
similar to 1 for PDT. In marked contrast, compound 2 was found to possess
several features that encourage further exploration of derivatives
or similar compounds. While uptake was low, it was comparable to that
of cisplatin, despite the overall charge of −4. Most importantly, 2 exhibited IC 50 values on the order of 3.3–17.3
μM when exposed to light, with no observed toxicity in the dark.
A slight increase in intracellular accumulation was observed upon
irradiation, possibly due to induction of plasma membrane damage that
facilitated compound uptake. Once inside the cell, the compound remained
in the cytosol, with no observable localization to the mitochondria
or inhibition of mitochondrial function. Furthermore, the light-induced
cell death mediated by 2 occurred by apoptosis, in contrast
to the mitochondrial targeting of 1 , which resulted in
necrosis. One possibility to explore is that PDT compounds that avoid
mitochondrial localization will exhibit lower toxicity in the absence
of light than those that associate with the mitochondria. It is anticipated
that structural modifications that result in a modest increase in
cellular uptake could sufficiently drive down potency to make improved
derivatives of 2 that maintain large therapeutic windows.
## Conclusions
Conclusions Given the high binding affinities of most Ru(II) complexes for
DNA ( K b > 10 6 ), it was previously
believed that the compounds would preferentially localize in the nucleus,
and indeed several do. 7 , 8 , 31 − 33 However, recent fluorescence and electron microscopy
studies have shown localization of several Ru(II) compounds in the
mitochondria, suggesting it is a common target. 34 , 35 Other reports indicate membrane accumulation and disruption, 10 , 36 along with apoptosis pathways that are mediated by the mitochondria. 37 , 38 Gasser and co-workers have shown in a recent report that this mitochondrial
localization was, in fact, required for cytotoxicity for a lead compound
in a structure–activity relationship study of a family of Ru(II)
polypyridyl complexes. 39 While mitochondrial
accumulation results in cytotoxicity, this
mechanism is not likely to be compatible with a PDT-type approach
where such redox-active compounds are required to be essentially nontoxic
in the absence of photons. Alternatively, targeting moieties such
as nuclear localization signals can be conjugated to the coordinated
ligands to affect the affinity of the complex for biological molecules
and regulate cellular uptake and subcellulation localization. 14 , 39 , 40 This approach requires significant
chemical modifications, and the targeting often fails to increase
cytotoxicity. While microscopy is a powerful tool to assess
compound localization,
imaging experiments can cause relocalization of compounds that induce
production of 1 O 2 or perform other photochemical
reactions when exposed to light. 41 Previous
studies on porphyrins used for PDT applications also have demonstrated
this phenomenon, 42 including uptake and
relocalization of an anionic tetrasulfonated porphyrin. 43 For this reason, it is important to perform
imaging using a minimum of light exposure, and to probe for compound
relocalization by comparing to conditions where the treated cells
have been exposed to significant light doses. This current study
shows that simple ligand modifications produce
complexes with divergent physical properties and, correspondingly,
different biological activities. Compound 1 , despite
its high DNA affinity, localizes to the mitochondria and induces rapid
membrane depolarization and necrotic cell death. It is possible that
this will be a common problem for compounds containing the bathophenanthroline
ligand. Compound 2 , despite its overall charge of −4,
is taken up into cancer cells to a sufficient degree to mediate light-induced
cell death through an apoptotic pathway. The absence of mitochondrial
localization may be the factor that eliminates the dark toxicity of
this Ru(II) complex. The incorporation of the sulfonic acids into
the ligands is likely responsible for the alteration in subcellular
localization, suggesting a possible general approach to reducing dark
toxicity for other Ru(II) complexes developed for applications in
phototherapy.
## Experimental Section
Experimental Section Materials Ru(bathophenanthroline) 3 ( 1 ) and Ru(bathophenanthroline disulfonate) 3 ( 2 ) were synthesized using previously established
procedures. 20 , 44 All cell lines were purchased
from ATCC. Cell culture media, heat-inactivated
fetal bovine serum (FBS), 4–20% tris-glycine precast gels,
Dulbecco’s phosphate buffered saline (DPBS), penicillin-streptomycin
solution (pen-strep), and 0.4% Trypan Blue solution were from Invitrogen.
35 mm wide, 4-compartment CELLview cell culture dishes were obtained
from USA Scientific. Serum supreme was from Lonza. Hoechst 33342,
Lysotracker Green DND-26 and Mitotracker Green FM were purchased from
Invitrogen. Propidium iodide (PI) and FITC-Annexin V were obtained
from BD Science. Trimethylrhodamine ethyl ester (TMRE) was purchased
from Sigma–Aldrich. Antibodies for PARP-1, procaspase 3, and
GAPDH were from Santa Cruz Biotechnology, Inc., while cleaved PARP
and cleaved caspase 3 was from Cell Signaling Technology. RIPA buffer
was purchased from Santa Cruz Biotechnology, Inc. Clarity Western
ECL Substrate was from Bio-Rad. An apoptotic DNA-ladder kit was purchased
from Roche Applied Science. DNA Gel Electrophoresis Compounds
were mixed with 40
μg/mL pUC19 plasmid DNA in 10 mM potassium phosphate buffer,
pH 7.4. To determine the effect of light, samples were irradiated
with light (>400 nm) from a 200 W light source for total light
doses
of 40 J/cm 2 . Samples were then incubated for 12 h at room
temperature in the dark. Single- and double-strand DNA break controls
were prepared, and the DNA samples were resolved on agarose gels,
as described previously. 45 In brief,
samples were resolved on a 1% agarose gels prepared in tris-acetate
buffer with 0.3 μg of plasmid/lane. The gels were stained with
0.5 μg/mL ethidium bromide in tris-acetate buffer at room temperature
for 40 min, destained with tris-acetate buffer, and imaged on a ChemiDoc
MP System (Bio-Rad). Cell Cytotoxicity Determination Human alveolar adenocarcinoma
cell line A549, Human promyelocytic leukemia cell line HL60, and Human
T lymphocyte cell line Jurkat cells were maintained in media supplemented
with 10% FBS and 50 U/mL pen-strep at 37 °C with 5% CO 2 , with DMEM used for A549 cells and IMDM and RPMI 1640 used for HL60
and Jurkat cells, respectively. Cells were assayed in Opti-MEM supplemented
with 1% serum supreme and 50 U/mL pen-strep and seeded into 96 well
plates at a density of 1.5 × 10 3 cells/well for A549,
2 × 10 4 cells/well for HL60, and 1 × 10 4 cells/well for Jurkat followed by a 6 h incubation at 37 °C,
5% CO 2 . Cells were then dosed with serial dilutions of
compound and incubated for 18 h. They were then irradiated with 7
J/cm 2 light (>400 nm) in 30 s pulses or kept in the
dark.
Cell viability was determined 72 h later by measuring the conversion
of resazurin to resorufin, 45 using a SpectraFluor
Plus Plate Reader (Tecan). Intracellular Measurement of Ru Complexes
by Flow Cytometry A549 cells were seeded in Opti-MEM with
1% serum supreme at a concentration
of 2 × 10 5 cells/ml in 25 cm 2 cell culture
flasks and incubated overnight. A concentration of 5 μM of 1 and 20 μM of 2 were added to the cells
and incubated for 2 or 24 h protected from light; the concentration
of 20 μM was selected for 2 to correspond with
the IC 50 of the complex when irradiated with light. A concentration
of 5 μM was used for 1 for compatibility with fluorescent
imaging. After 2 and 24 h the media was removed, cells were washed
twice with DPBS, trypsinized, and collected by centrifugation at 125
× g for 4 min. The cells were resuspended in
Opti-MEM, filtered through 40-μm cell strainers, and analyzed
on a FACSCalibur (Becton–Dickenson) with an excitation wavelength
of 488 nm and emission measured at 650 nm. A minimum of 30,000 events
were measured for each sample. ApoTome Structured Illumination
Imaging of Ru Complex Uptake The ApoTome microscope was used
to resolve fine features of cellular
structure. This instrument averages the fluorescence of three separate
images to greatly reduce out-of-plane fluorescence. A549 cells were
seeded in 35 mm, four-compartment CELLview culture dishes at a density
of 5 × 10 4 cells per compartment in a 500 μL
volume and incubated for 24 h in Opti-MEM containing 1% FBS, followed
by the addition of 5 μM 1 or 20 μM 2 and time points measured at 2, 8, 18, or 24 h. Media was
removed at each time point, cells rinsed with DPBS, and incubated
in Opti-MEM with 16 μM Hoechst 33342, 16 μM Hoechst and
0.15 μM Lysotracker Green DND-26, or 16 μM Hoechst and
0.2 μM Mitotracker Green FM. Cells were incubated for 30 min
then washed three times with DPBS and imaged at 50× magnification
using an ApoTome structured illumination fluorescent microscope (Carl
Zeiss AG). Mitochondrial Membrane Potential Measurement A549 cells
were seeded at 2 × 10 4 cells/well in 24-well plates
and incubated for 18 h, followed by the addition of 5 μM 1 or 20 μM 2 . They were incubated for an
additional 8 h, and then irradiated with light as described for cell
cytotoxicity measurements or kept in the dark. The cells were incubated
for an additional 2, 8, 18, or 24 h, washed with DPBS, followed by
the addition of 0.5 μM TMRE in Opti-MEM, and incubated for 30
min. The cells were then washed twice with DPBS and imaged. The change
in TMRE signal was determined by the difference in fluorescence between
compound-treated cells in the presence and absence of TMRE. Fluorescence
was measured at 10× magnification using an ApoTome microscope
in normal fluorescence mode. Images were processed and the average
cell fluorescence was calculated using ImageJ software. Samples dosed
with 1 and 2 were compared to untreated
A549 cells ( n = 3) to give the relative mitochondrial
potential. Cell Viability As a Function of Time HL60 cells were
seeded in Opti-MEM at a density of 1 × 10 6 cells/mL
and incubated for 2 h. The cells were then dosed 1 or 2 , incubated for 8 h, irritated as above or protected from
light, and incubated for 2, 8, 18, 24, or 48 h. As HL60 cells grow
in suspension, Trypan Blue staining was employed in place of resazurin
to simplify and accelerate cell viability analysis. At each time point,
a 10 μL cell suspension was mixed with an equal amount of Trypan
Blue solution and cell viability determined by manual counting with
a hemocytometer. Apoptosis Marker Immunoblotting HL60 cells were cultured,
dosed with 1 and 2 , then irradiated as above
or protected from light. Cells were harvested 0, 2, 8, 18, or 24 h
after treatment, pelleted by centrifugation at 124 × g for 5 min, washed twice with DPBS, and lysed in RIPA buffer
supplemented with 5 mM sodium pyrophosphate for 15 min on ice. The
insoluble fraction was removed by centrifugation at 20,800 × g for 10 min at 4 °C. The supernatant was collected
and the protein concentration determined by BCA assay. Ten micrograms
(10 μg) of protein was loaded onto 4–20% tris-glycine
gels, followed by transfer to nitrocellulose membranes. After blocking
with 5% nonfat milk in DPBS with 0.1% Tween20 (PBST) for 1 h at room
temperature, the membrane was immunoblotted with PARP-1 at a 1:500
dilution, procaspase 3, cleaved caspase 3, and cleaved PARP at 1:1,000
dilutions, or GAPDH at a 1:2,000 dilution in 5% nonfat milk overnight
at 4 °C. Immunoblots were washed with PBST for 10 min four times
and incubated for 1 h with secondary antibodies at a 1:10,000 dilution
for GAPDH and at a 1:5,000 dilution for all other antibodies. Detection
was performed using Clarity Western ECL substrate and imaged with
the ChemiDoc MP System. DNA Laddering Gel Electrophoresis HL60 cells were cultured,
dosed with 1 and 2 , then photoactivated
or protected from light as detailed above. Cells were harvested 2
h after photoactivation for 1 or after 24 h for 2 , pelleted by centrifugation at 124 × g for 5 min, washed twice with DPBS, and prepared with an apoptotic
DNA-ladder kit as per manufacturer instructions. Gel electrophoresis
was carried out using a 1% agarose gel containing 0.5 μg/mL
ethidium bromide with 1 μg DNA loaded per lane and run for 90
min at 75 V. Gel imaging was performed with the ChemiDoc MP System. Quantification of Metal Complex Uptake by ICP-OES HL60
cells were seeded in Opti-MEM at a density of 1 × 10 6 cells/mL in 25 cm 2 cell culture flasks, cultured overnight,
then dosed with 5 μM 1 , 20 μM 2 , or 20 μM cisplatin. Cells treated with Ru complexes were
incubated for 8 h, protected from light before irradiating or kept
in the dark. Cells were collected after 2 and 24 h by centrifugation
at 124 × g for 5 min. The cell media was transferred
to separate 15-mL centrifuge tubes, and cells were washed twice with
DPBS. Two milliliters (2 mL) of concentrated HNO 3 was added
to media samples while cell pellets were resuspended in 5 mL of 20%
(v/v) HNO 3 . All samples were heated at 110 °C for
3 h. After digestion, the volume of all cell samples was adjusted
to 5 mL and media samples were adjusted to 10 mL with deionized (DI)
water. The metal content was analyzed using a VISTA-PRO CCD simultaneous
inductively coupled plasma optical emission spectrometer (Varian,
Inc.) with detection at 240.272, 245.657, and 267.876 nm for ruthenium
and 214.424, 217.468, and 265.945 nm for platinum, with a replicate
reading time of 60 s. Yttrium (1 ppm) in 1% nitric acid was employed
as an internal standard. The percentage intracellular metal ratio
in 10 7 cells was calculated by normalizing the metal amount
to 10 7 cells, and then divided by the total metal amount
in both media and cells. Cell Death by Flow Cytometry HL60
cells were cultured,
dosed with 1 and 2 , then irradiated or protected
from light. Cells were harvested 2 and 24 h after treatment, pelleted
by centrifugation at 124 × g for 5 min, washed
twice with DPBS, stained for 15 min with FITC-Annexin V and PI or
FITC-Annexin V and Hoechst 33342, because of the interference between
the emission of 1 and PI. Cells were analyzed with a
FACSCalibur (Becton–Dickenson). A minimum of 20,000 events
were measured for each sample.
## Materials
Materials Ru(bathophenanthroline) 3 ( 1 ) and Ru(bathophenanthroline disulfonate) 3 ( 2 ) were synthesized using previously established
procedures. 20 , 44 All cell lines were purchased
from ATCC. Cell culture media, heat-inactivated
fetal bovine serum (FBS), 4–20% tris-glycine precast gels,
Dulbecco’s phosphate buffered saline (DPBS), penicillin-streptomycin
solution (pen-strep), and 0.4% Trypan Blue solution were from Invitrogen.
35 mm wide, 4-compartment CELLview cell culture dishes were obtained
from USA Scientific. Serum supreme was from Lonza. Hoechst 33342,
Lysotracker Green DND-26 and Mitotracker Green FM were purchased from
Invitrogen. Propidium iodide (PI) and FITC-Annexin V were obtained
from BD Science. Trimethylrhodamine ethyl ester (TMRE) was purchased
from Sigma–Aldrich. Antibodies for PARP-1, procaspase 3, and
GAPDH were from Santa Cruz Biotechnology, Inc., while cleaved PARP
and cleaved caspase 3 was from Cell Signaling Technology. RIPA buffer
was purchased from Santa Cruz Biotechnology, Inc. Clarity Western
ECL Substrate was from Bio-Rad. An apoptotic DNA-ladder kit was purchased
from Roche Applied Science.
## DNA Gel Electrophoresis
DNA Gel Electrophoresis Compounds
were mixed with 40
μg/mL pUC19 plasmid DNA in 10 mM potassium phosphate buffer,
pH 7.4. To determine the effect of light, samples were irradiated
with light (>400 nm) from a 200 W light source for total light
doses
of 40 J/cm 2 . Samples were then incubated for 12 h at room
temperature in the dark. Single- and double-strand DNA break controls
were prepared, and the DNA samples were resolved on agarose gels,
as described previously. 45 In brief,
samples were resolved on a 1% agarose gels prepared in tris-acetate
buffer with 0.3 μg of plasmid/lane. The gels were stained with
0.5 μg/mL ethidium bromide in tris-acetate buffer at room temperature
for 40 min, destained with tris-acetate buffer, and imaged on a ChemiDoc
MP System (Bio-Rad).
## Cell Cytotoxicity Determination
Cell Cytotoxicity Determination Human alveolar adenocarcinoma
cell line A549, Human promyelocytic leukemia cell line HL60, and Human
T lymphocyte cell line Jurkat cells were maintained in media supplemented
with 10% FBS and 50 U/mL pen-strep at 37 °C with 5% CO 2 , with DMEM used for A549 cells and IMDM and RPMI 1640 used for HL60
and Jurkat cells, respectively. Cells were assayed in Opti-MEM supplemented
with 1% serum supreme and 50 U/mL pen-strep and seeded into 96 well
plates at a density of 1.5 × 10 3 cells/well for A549,
2 × 10 4 cells/well for HL60, and 1 × 10 4 cells/well for Jurkat followed by a 6 h incubation at 37 °C,
5% CO 2 . Cells were then dosed with serial dilutions of
compound and incubated for 18 h. They were then irradiated with 7
J/cm 2 light (>400 nm) in 30 s pulses or kept in the
dark.
Cell viability was determined 72 h later by measuring the conversion
of resazurin to resorufin, 45 using a SpectraFluor
Plus Plate Reader (Tecan).
## Intracellular Measurement of Ru Complexes
by Flow Cytometry
Intracellular Measurement of Ru Complexes
by Flow Cytometry A549 cells were seeded in Opti-MEM with
1% serum supreme at a concentration
of 2 × 10 5 cells/ml in 25 cm 2 cell culture
flasks and incubated overnight. A concentration of 5 μM of 1 and 20 μM of 2 were added to the cells
and incubated for 2 or 24 h protected from light; the concentration
of 20 μM was selected for 2 to correspond with
the IC 50 of the complex when irradiated with light. A concentration
of 5 μM was used for 1 for compatibility with fluorescent
imaging. After 2 and 24 h the media was removed, cells were washed
twice with DPBS, trypsinized, and collected by centrifugation at 125
× g for 4 min. The cells were resuspended in
Opti-MEM, filtered through 40-μm cell strainers, and analyzed
on a FACSCalibur (Becton–Dickenson) with an excitation wavelength
of 488 nm and emission measured at 650 nm. A minimum of 30,000 events
were measured for each sample.
## ApoTome Structured Illumination
Imaging of Ru Complex Uptake
ApoTome Structured Illumination
Imaging of Ru Complex Uptake The ApoTome microscope was used
to resolve fine features of cellular
structure. This instrument averages the fluorescence of three separate
images to greatly reduce out-of-plane fluorescence. A549 cells were
seeded in 35 mm, four-compartment CELLview culture dishes at a density
of 5 × 10 4 cells per compartment in a 500 μL
volume and incubated for 24 h in Opti-MEM containing 1% FBS, followed
by the addition of 5 μM 1 or 20 μM 2 and time points measured at 2, 8, 18, or 24 h. Media was
removed at each time point, cells rinsed with DPBS, and incubated
in Opti-MEM with 16 μM Hoechst 33342, 16 μM Hoechst and
0.15 μM Lysotracker Green DND-26, or 16 μM Hoechst and
0.2 μM Mitotracker Green FM. Cells were incubated for 30 min
then washed three times with DPBS and imaged at 50× magnification
using an ApoTome structured illumination fluorescent microscope (Carl
Zeiss AG).
## Mitochondrial Membrane Potential Measurement
Mitochondrial Membrane Potential Measurement A549 cells
were seeded at 2 × 10 4 cells/well in 24-well plates
and incubated for 18 h, followed by the addition of 5 μM 1 or 20 μM 2 . They were incubated for an
additional 8 h, and then irradiated with light as described for cell
cytotoxicity measurements or kept in the dark. The cells were incubated
for an additional 2, 8, 18, or 24 h, washed with DPBS, followed by
the addition of 0.5 μM TMRE in Opti-MEM, and incubated for 30
min. The cells were then washed twice with DPBS and imaged. The change
in TMRE signal was determined by the difference in fluorescence between
compound-treated cells in the presence and absence of TMRE. Fluorescence
was measured at 10× magnification using an ApoTome microscope
in normal fluorescence mode. Images were processed and the average
cell fluorescence was calculated using ImageJ software. Samples dosed
with 1 and 2 were compared to untreated
A549 cells ( n = 3) to give the relative mitochondrial
potential.
## Cell Viability As a Function of Time
Cell Viability As a Function of Time HL60 cells were
seeded in Opti-MEM at a density of 1 × 10 6 cells/mL
and incubated for 2 h. The cells were then dosed 1 or 2 , incubated for 8 h, irritated as above or protected from
light, and incubated for 2, 8, 18, 24, or 48 h. As HL60 cells grow
in suspension, Trypan Blue staining was employed in place of resazurin
to simplify and accelerate cell viability analysis. At each time point,
a 10 μL cell suspension was mixed with an equal amount of Trypan
Blue solution and cell viability determined by manual counting with
a hemocytometer.
## Apoptosis Marker Immunoblotting
Apoptosis Marker Immunoblotting HL60 cells were cultured,
dosed with 1 and 2 , then irradiated as above
or protected from light. Cells were harvested 0, 2, 8, 18, or 24 h
after treatment, pelleted by centrifugation at 124 × g for 5 min, washed twice with DPBS, and lysed in RIPA buffer
supplemented with 5 mM sodium pyrophosphate for 15 min on ice. The
insoluble fraction was removed by centrifugation at 20,800 × g for 10 min at 4 °C. The supernatant was collected
and the protein concentration determined by BCA assay. Ten micrograms
(10 μg) of protein was loaded onto 4–20% tris-glycine
gels, followed by transfer to nitrocellulose membranes. After blocking
with 5% nonfat milk in DPBS with 0.1% Tween20 (PBST) for 1 h at room
temperature, the membrane was immunoblotted with PARP-1 at a 1:500
dilution, procaspase 3, cleaved caspase 3, and cleaved PARP at 1:1,000
dilutions, or GAPDH at a 1:2,000 dilution in 5% nonfat milk overnight
at 4 °C. Immunoblots were washed with PBST for 10 min four times
and incubated for 1 h with secondary antibodies at a 1:10,000 dilution
for GAPDH and at a 1:5,000 dilution for all other antibodies. Detection
was performed using Clarity Western ECL substrate and imaged with
the ChemiDoc MP System.
## DNA Laddering Gel Electrophoresis
DNA Laddering Gel Electrophoresis HL60 cells were cultured,
dosed with 1 and 2 , then photoactivated
or protected from light as detailed above. Cells were harvested 2
h after photoactivation for 1 or after 24 h for 2 , pelleted by centrifugation at 124 × g for 5 min, washed twice with DPBS, and prepared with an apoptotic
DNA-ladder kit as per manufacturer instructions. Gel electrophoresis
was carried out using a 1% agarose gel containing 0.5 μg/mL
ethidium bromide with 1 μg DNA loaded per lane and run for 90
min at 75 V. Gel imaging was performed with the ChemiDoc MP System.
## Quantification of Metal Complex Uptake by ICP-OES
Quantification of Metal Complex Uptake by ICP-OES HL60
cells were seeded in Opti-MEM at a density of 1 × 10 6 cells/mL in 25 cm 2 cell culture flasks, cultured overnight,
then dosed with 5 μM 1 , 20 μM 2 , or 20 μM cisplatin. Cells treated with Ru complexes were
incubated for 8 h, protected from light before irradiating or kept
in the dark. Cells were collected after 2 and 24 h by centrifugation
at 124 × g for 5 min. The cell media was transferred
to separate 15-mL centrifuge tubes, and cells were washed twice with
DPBS. Two milliliters (2 mL) of concentrated HNO 3 was added
to media samples while cell pellets were resuspended in 5 mL of 20%
(v/v) HNO 3 . All samples were heated at 110 °C for
3 h. After digestion, the volume of all cell samples was adjusted
to 5 mL and media samples were adjusted to 10 mL with deionized (DI)
water. The metal content was analyzed using a VISTA-PRO CCD simultaneous
inductively coupled plasma optical emission spectrometer (Varian,
Inc.) with detection at 240.272, 245.657, and 267.876 nm for ruthenium
and 214.424, 217.468, and 265.945 nm for platinum, with a replicate
reading time of 60 s. Yttrium (1 ppm) in 1% nitric acid was employed
as an internal standard. The percentage intracellular metal ratio
in 10 7 cells was calculated by normalizing the metal amount
to 10 7 cells, and then divided by the total metal amount
in both media and cells.
## Cell Death by Flow Cytometry
Cell Death by Flow Cytometry HL60
cells were cultured,
dosed with 1 and 2 , then irradiated or protected
from light. Cells were harvested 2 and 24 h after treatment, pelleted
by centrifugation at 124 × g for 5 min, washed
twice with DPBS, stained for 15 min with FITC-Annexin V and PI or
FITC-Annexin V and Hoechst 33342, because of the interference between
the emission of 1 and PI. Cells were analyzed with a
FACSCalibur (Becton–Dickenson). A minimum of 20,000 events
were measured for each sample.