A fast and selective near-infrared fluorescent sensor for multicolor imaging of biological nitroxyl (HNO)

Article abstract of DOI:10.1021/ja500315x

The first near-infrared fluorescent turn-on sensor for the detection of nitroxyl (HNO), the one-electron reduced form of nitric oxide (NO), is reported. The new copper-based probe, CuDHX1, contains a dihydroxanthene (DHX) fluorophore and a cyclam derivative as a Cu(II) binding site. Upon reaction with HNO, CuDHX1 displays a five-fold fluorescence turn-on in cuvettes and is selective for HNO over thiols and reactive nitrogen and oxygen species. CuDHX1 can detect exogenously applied HNO in live mammalian cells and in conjunction with the zinc-specific, green-fluorescent sensor ZP1 can perform multicolor/multianalyte microscopic imaging. These studies reveal that HNO treatment elicits an increase in the concentration of intracellular mobile zinc.

Full text of DOI:10.1021/ja500315x

Article
pubs.acs.org/JACS
A Fast and Selective Near-Infrared Fluorescent Sensor for Multicolor
Imaging of Biological Nitroxyl (HNO)
Alexandra T. Wrobel, Timothy C. Johnstone, Alexandria Deliz Liang, Stephen J. Lippard,*
and Pablo Rivera-Fuentes*
Department of Chemistry, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139, United States
S
* Supporting Information
ABSTRACT: The first near-infrared fluorescent turn-on
sensor for the detection of nitroxyl (HNO), the one-electron
reduced form of nitric oxide (NO), is reported. The new
copper-based probe, CuDHX1, contains a dihydroxanthene
(DHX) fluorophore and a cyclam derivative as a Cu(II)
binding site. Upon reaction with HNO, CuDHX1 displays a
five-fold fluorescence turn-on in cuvettes and is selective for
HNO over thiols and reactive nitrogen and oxygen species.
CuDHX1 can detect exogenously applied HNO in live
mammalian cells and in conjunction with the zinc-specific,
green-fluorescent sensor ZP1 can perform multicolor/multianalyte microscopic imaging. These studies reveal that HNO
treatment elicits an increase in the concentration of intracellular mobile zinc.
a
Scheme 1. Possible Biosynthetic Pathways to HNO
INTRODUCTION
■
Nitric oxide (NO) is an important biological signaling molecule
present in the cardiovascular,1 immune,2 and nervous systems.3
NO is released by macrophages during the immune response,2
controls smooth muscle relaxation,4 and regulates neuro-
transmission.3,5 Dysregulation of NO homeostasis is associated
with numerous diseases, including diabetes,6 stroke,7 Alz-
heimer’s disease,8 and cancer.9 HNO, the one-electron reduced
and protonated derivative of NO, displays distinctive chemistry
and biochemistry from that of NO.10 Exogenously applied
HNO confers vasoprotective effects,11 increases heart muscle
contractility,12 and inhibits platelet aggregation.13 HNO also
exacerbates ischemia-related injury14 and induces neuro-
toxicity.15 Under biological conditions, HNO is both an
electrophile that oxidizes thiols and a nucleophile that can
coordinate and reduce metal ions.16 Thiolates that bind zinc are
a potential biological target of HNO. Nitrosation of thiolates
induces the release of mobile zinc,17 which may trigger a variety
of signaling pathways.18 Understanding these and other
downstream effects of exogenously applied HNO is essential
if nitroxyl donors are to be used as therapeutic agents.19
Endogenous generation of HNO has not been directly
observed in live organisms. Some proposed biosynthetic
pathways are shown in Scheme 1. HNO can be produced by
nitric oxide synthase (NOS). Under normal physiological
conditions, NOS catalyzes the conversion of arginine to
citrulline and NO in the presence of the cofactor tetrahy-
drobiopterin (THB).20,21 In the absence of THB, isolated
neuronal NOS produces HNO instead of NO.21 Additionally,
HNO can be generated by oxidation of hydroxylamine
(NH2OH) with heme-containing proteins.14 NO and HNO
interconvert in the presence of superoxide dismutase (SOD),22
aL-NHA = N-hydroxy-L-arginine; nNOS = neuronal nitric oxide
synthase; SOD = superoxide dismutase; THB = tetrahydrobiopterin.
and cytochrome c can reduce NO to HNO.23 Recently, HNO
production was detected in the reaction between HSNO and
H2S24 and in the heme-iron-catalyzed reduction of nitrite with
H2S.25 These results suggest that HNO can be produced under
biological conditions and might be associated with particular
physiological or pathological states. Improved methods for
detecting HNO selectively, with well-defined spatiotemporal
resolution in live cells, tissues, and animals, are needed to fully
understand the biology of nitroxyl.
Received: January 11, 2014
Published: February 24, 2014
© 2014 American Chemical Society
4697
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
Chart 1. Chemical Structures and Photophysical Properties of Some Previously Reported and Newly Developed (CuDHX1)
Probes for HNO26−30
A number of fluorescent sensors that detect HNO in live
cells are available (Chart 1). One approach to HNO sensing
relies on the use of fluorophores functionalized with amines
that bind Cu(II). The fluorescence of these sensors is quenched
upon binding of the paramagnetic cupric ion. Upon exposure to
HNO, Cu(II) reduces to Cu(I), which renders the complex
diamagnetic and restores the fluorescence of the probe. Sensors
based on this principle include BODIPY-based complex
CuBOT1 (Chart 1),26,27 coumarin-based derivative CuCOT1
(Chart 1),28 and the CuBRNO series of benzoresorufin-based
sensors (Chart 1) that detect both HNO and NO.29 Recently,
the metal-free probe P-Rhod (Chart 1) was reported.30 This
sensor exploits the reactivity of HNO with triphenylphosphine
to produce an azaylide, which undergoes an intramolecular
nucleophilic attack that releases rhodol, a bright green
fluorophore. Despite their value, these probes have short-
comings associated with their lack of selectivity or short
emission wavelengths. It would be valuable to have a sensor
that not only detects HNO with high selectivity but would also
emit in the near-infrared (NIR) region, desired for in vivo
imaging because of deeper tissue penetration.31 Moreover,
sensors with narrow emission profiles at long wavelengths can
be used together with visible-light probes in multicolor
microscopy experiments.32,33 In the present article we describe
the design, synthesis, and implementation of CuDHX1 (Chart
1), a novel sensor that satisfies all of these requirements.
The structure of CuDHX1 comprises a cyclam binding site
for Cu(II) and a dihydroxanthene (DHX) NIR fluorophore.
We chose cyclam because Cu(II) complexes of this ligand react
very slowly with the interfering species H2S and glutathione
(GSH) under physiological conditions.34 The DHX fluoro-
phore was selected because it is a bright NIR emitter with
excellent biocompatibility.35 Here we report the results of our
studies using CuDHX1 to image HNO in live cells as well as
multicolor imaging experiments to investigate the ability of
Angeli’s salt, an HNO donor, to affect the levels of mobile zinc
in live cells.
EXPERIMENTAL SECTION
■
General Methods. All reactions were performed under a nitrogen
atmosphere unless otherwise specified. ZP1 was prepared as previously
described.36 Reagents were purchased from commercial sources and
used as received. Solvents were purified and degassed by standard
procedures. Nitric oxide was passed through an Ascarite column and a
6 ft coil containing silica gel at −78 °C to remove impurities and then
collected and stored under nitrogen in a gas storage bulb. NMR
spectra were acquired on a Varian Inova-500 or a Varian Mercury-300
1
instrument. H NMR chemical shifts are reported in ppm relative to
SiMe4 (δ = 0) and were referenced internally with respect to residual
protons in the solvent (δ = 3.31 for CD3OD or δ = 2.50 for DMSO-
d6). Coupling constants are reported in Hz. 13C NMR chemical shifts
are reported in ppm relative to SiMe4 (δ = 0) and were referenced
internally with respect to solvent signal (δ = 39.51 for DMSO-d6). 19
F
NMR chemical shifts are reported in ppm relative to CFCl3 (δ = 0)
and were referenced internally with respect to 2,2,2-trifluoroethanol (δ
= −77.03). Low-resolution mass spectra (LRMS) were acquired on an
Agilent 1100 Series LC/MSD Trap spectrometer (LCMS), using
electrospray ionization (ESI). High-resolution mass spectrometry
(HR-ESI-MS) was conducted by staff at the MIT Department of
Chemistry Instrumentation Facility on a Bruker Daltonics APEXIV 4.7
T FT-ICR-MS instrument. Silica gel 60 321 (0.015−0.040 mm) was
used for flash column chromatography. Semipreparative HPLC
separations were carried out on an Agilent 1200 HPLC instrument
with a multiwavelength detector and automated fraction collector
using a C18 reverse stationary phase (Zorbax-SB C18, 5 μm, 9.5 × 250
mm) and a mobile phase composed of two solvents (A: 0.1% (v/v)
trifluoroacetic acid (TFA) in H2O; B: 0.1% (v/v) TFA in CH3CN).
Specific purification protocols are described below for each compound.
IUPAC names of all compounds are provided and were determined
using CS ChemBioDrawUltra 12.0.
Synthesis of (E)-2-(2-(6-hydroxymethyl)-2,3-dihydro-1H-xanthen-
4-yl)vinyl)-3,3-dimethyl-1-propyl-3H-indol-1-ium iodide (1). A sol-
ution of IR780 (300 mg, 0.45 mmol) in dry DMF (5 mL) was added
to 5-(hydroxymethyl)benzene-1,3-diol (63 mg, 0.45 mmol). Triethyl-
amine (0.6 mL, 4.49 mmol) was added, and the mixture was stirred at
4698
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
110 °C for 30 min. The solvent was evaporated under reduced
pressure and the solid was purified by column chromatography (SiO2;
CH2Cl2/CH3OH 9:1) to give the product as a dark blue solid (195
176.87. 19F NMR (282 MHz, CD3OD): −75.18. HR-ESI-MS. Calcd
for [C46H60N5O2]+: 714.4742, found: 714.4758.
Spectroscopic Methods. All aqueous solutions were prepared
using deionized water with resistivity 18.2 mΩ cm−1, obtained using a
Milli-Q water purification system. All solvents were supplied by
Aldrich and used as received. Piperazine-N,N′-bis(2-ethanesulfonic
acid) (PIPES) and 99.999% KCl were purchased from Calbiochem.
Stock solutions of compound 3 and ligand DHX1 in DMSO were
prepared in the 1−8 mM range and stored at −20 °C in 1 mL aliquots
and thawed immediately before each experiment. The copper
complexes were prepared by dissolving the ligand in 1 mL of
CH3OH, adding 1 equiv of CuCl2 to each solution, and stirring
overnight. The solvent was evaporated, and the resulting sensor was
redissolved in 1 mL DMSO and stored at −20 °C. All spectroscopic
measurements were conducted in aqueous buffer containing 50 mM
PIPES (pH 7.0) and 100 mM KCl, with the exception of those
performed in CH3OH and at varying pH values. UV−vis spectra were
acquired using a Cary 50 spectrometer using quartz cuvettes from
Starna (1 cm path length). Fluorescence spectra were acquired on a
Photon Technology International fluorimeter. All measurements were
conducted on solutions maintained at 25 °C by circulating water baths.
Extinction coefficients of the ligands and sensors were determined by
using 1−3 μM solutions in aqueous buffer. Fluorescence quantum
yields were determined using the same solutions, λex = 650 nm.
Fluorescence emission spectra were integrated from 660 to 900 nm.
Quantum yields were referenced to IR780, which has a reported
quantum yield of 0.076 in CH3OH, λex = 725 nm.37
Anaerobic Sample Preparation. Degassed aliquots of the stock
solutions of each sensor were brought into an anaerobic chamber
under a nitrogen atmosphere (O2 < 1 ppm) dedicated to work with
aqueous solutions (hereafter called the “wet box”). Solutions
containing 5 μM Cu-3 and 2 μM CuDHX1 were prepared using 2
mL of either degassed aqueous buffer or CH3OH in gastight cuvettes.
Angeli’s salt (Na2N2O3, Cayman Chemical) was used as the source of
HNO, because it decomposes rapidly (t1/2 = 3 min) at pH = 7 to
produce HNO and NaNO2.38 Solutions of Angeli’s salt (4 mM) were
prepared in the wet box in degassed 10 mM NaOH (2 mL) and
brought out of the wet box in a gastight syringe. NO gas was removed
from the wet box in a gastight syringe and injected into the headspace
of each gastight cuvette before measuring fluorescence.
Cyclic Voltammetry. Cyclic voltammograms were measured in a
three-electrode cell with a 2.0 mm diameter glassy carbon working
electrode, a platinum auxiliary electrode, and Ag/Ag+ pseudoreference
electrode in acetonitrile. The solvent contained 0.1 M n-Bu4NPF6 as
the supporting electrolyte. The measurements were performed at
room temperature with a VersaSTAT3 potentiostat from Princeton
Applied Research operated with V3 studio software. Measurements
were carried out at a scan rate of 200 mV s−1 on quiescent solutions
that had been sparged with N2 for 5 min. All data were referenced to
the Fc/Fc+ couple as an internal standard.
Electron Paramagnetic Resonance (EPR) Spectroscopy. Low-
temperature X-band EPR spectra (77 K, 9 GHz) were collected with a
Bruker EMS spectrometer equipped with an ER 4199HS cavity and a
Gunn diode microwave source. EPR samples were prepared
anaerobically. Solid DHX1 and 3 were brought into a glovebox. 0.8
equiv of Cu(MeCN)4BF4 were added to 400 μM DHX1 or 3 in 350
μL CH3OH, stirred overnight, and brought out of the glovebox in
sealed EPR tubes. Angeli’s salt (100 equiv) was prepared in 10 mM
NaOH anaerobically and brought out of the wet box in a gastight
syringe. When CuCl2 was used to prepare CuDHX1 for the HNO
reactivity EPR studies, different results were observed (Figure S21).
For the NO reactivity test, degassed aliquots from the stock solution of
CuDHX1 were brought into a wet box. Solutions containing 400 μM
CuDHX1 were prepared in 350 μL of degassed CH3OH and brought
out of the wet box in sealed EPR tubes. NO gas was taken out of the
wet box in a gastight syringe and injected into the headspace of the
EPR tube. Spectra were simulated in Matlab using the solid-state/
frozen-solution functionality (‘pepper’) implemented in EasySpin.39
Analyte Selectivity Studies. Selectivity of the fluorescence turn-
on toward biologically relevant analytes was determined by comparing
1
3
mg, yield 76%). H NMR (300 MHz, CD3OD): 1.04 (t, J = 7.4 Hz,
3H), 1.80 (s, 6H), 1.89−1.96 (m, 4H), 2.69−2.82 (m, 4H), 4.26 (t, 3J
= 7.2 Hz), 4.79 (s, 2H), 6.40 (d, 3J = 14.7 Hz, 1H), 6.74 (d, 4J = 2 Hz,
1H), 6.91 (d, 4J = 2 Hz, 1H), 7.36−7.41 (m, 1H), 7.45−7.49 (m, 2H),
3
7.60−7.63 (m, 1H), 7.70 (s, 1H), 8.69 (d, J = 14.5 Hz, 1H). LRMS
(ESI). Calcd for [C29H32NO3]+: 442.2, found 442.2.
Synthesis of (E)-2-(2-(8-(chloromethyl)-6-hydroxy-2,3-dihydro-
1H-xanthen-4-yl)vinyl)-3,3-dimethyl-1-propyl-3H-indol-ium iodide
(2). Thionyl chloride (75 μL, 1.03 mmol) and dry pyridine (83 μL,
1.03 mmol) were dissolved in dry CH2Cl2 (1 mL) and cooled to 0 °C.
Compound 1 (195 mg, 0.34 mmol) was dissolved in dry CH2Cl2 (1
mL) and dry DMF (0.1 mL) and added slowly to the mixture of
pyridine and thionyl chloride. After 30 min, H2O (0.1 mL) was added,
and the mixture was dried with Na2SO4. The solvent was evaporated
under reduced pressure. The dark blue solid was dissolved in CH2Cl2/
CH3OH (9:1) and filtered through a plug of SiO2 with CH2Cl2/
CH3OH (9:1, 200 mL). The solvent was evaporated, and the crude
product was used immediately. LRMS (ESI). Calcd for
[C29H31ClNO2]+: 460.2, found 460.1.
Synthesis of (E)-2-(2-(8-((1,4,8,11-tetraazacyclotetradecan-1-yl)-
methyl)-6-hydroxy-2,3-dihydro-1H-xanthen-4-yl)vinyl)-3,3-dimeth-
yl-1-propyl-3H-indol-1-ium trifluoroacetate (3). A solution of crude
compound 2 in dry CH3CN (3 mL) was added to 1,4,8,11-
tetraazacyclotetradecane (cyclam, 136 mg, 0.68 mmol). Diisopropyl-
ethylamine (0.12 mL, 0.68 mmol) was added, and the mixture was
heated to reflux. After 30 min, the solvent was evaporated, and the
residue was purified by RP-HPLC according to the following protocol:
constant flow rate 3 mL min−1; isocratic flow 2% B, 0−5 min; gradient,
35−95% B, 10−25 min. The product was collected between 16.5−16.8
min. All equivalent fractions recovered from independent runs were
combined and lyophilized to dryness to yield the TFA salt of
1
compound 3 (46 mg, 18% over two steps). Mp: 110−120 °C. H
NMR (500 MHz, DMSO-d6): 0.96 (t, 3J = 7.4 Hz, 3H), 1.72 (s, 7H),
1.81 (m, 7H), 2.67 (m, 10H), 3.10 (m, 10H), 3.80 (s, 2H), 4.35 (t, 3J
3
3
= 7.1 Hz, 2H), 6.53 (d, J = 15 Hz, 1H), 6.92 (m, 1H), 7.42, (t, J =
7.7 Hz, 1H), 7.51 (t, 3J = 8.3 Hz, 1H), 7.66 (d, 3J = 8.0 Hz, 1H), 7.72
(d, 3J = 7.7 Hz, 2H), 8.55 (d, J = 15 Hz, 1H). 13C NMR (125 MHz,
3
DMSO-d6): 11.08, 20.10, 20.92, 23.60, 27.59, 28.55, 40.44, 45.95,
49.56, 50.24, 101.59, 103.73, 113.12, 113.38, 113.71, 115.75, 115.96,
118.11, 120.48, 122.76, 125.85, 126.85, 127.96, 128.97, 136.75, 141.65,
141.88, 144.50, 154.73, 158.13, 158.42, 158.65, 158.90, 160.62, 161.12,
177.01. 19F NMR (282 MHz, CD3OD): −75.31. HR-ESI-MS. Calcd
for [C39H54N5O2]+: 624.4273, found: 624.4259.
Synthesis of (E)-2-(2-(8-((8-benzyl-1,4,8,11-tetraazacyclotetrade-
can-1-yl)methyl)-6-hydroxy-2,3-dihydro-1H-xanthen-4-yl)vinyl)-3,3-
dimethyl-1-propyl-3H-indol-1-ium trifluoroacetate (DHX1). A sol-
ution of compound 3 (46 mg, 0.06 mmol) in CH3CN (3 mL) and
(bromomethyl)benzene (4 μL, 0.03 mmol) were combined.
Diisopropylethylamine (22 μL, 0.12 mmol) was added, and the
reaction mixture stirred at room temperature for 2 h. The solvent was
evaporated, and the product was purified by RP-HPLC according to
the following protocol: constant flow rate 3 mL min−1; isocratic flow
2% B, 0−5 min; gradient, 35−95% B, 10−25 min. The product was
collected between 18.1−18.5 min. All equivalent fractions recovered
from independent runs were combined and lyophilized to dryness to
yield the TFA salt of compound DHX1 (14.2 mg, 28%). Mp: 115−120
°C. 1H NMR (500 MHz, DMSO-d6): 0.97 (t, 3J = 7 Hz, 3H), 1.72 (s,
6H), 1.80 (m, 6H), 2.06 (m, 2H), 2.66 (m, 3H), 2.72 (m, 5H), 3.14
3
3
(m, 8H), 3.83 (s, 3H), 4.35 (t, J = 7 Hz, 2H), 6.52 (d, J = 15 Hz,
3
1H), 6.91 (m, 1H), 7.00 (m, 1H), 7.40 (m, 6H), 7.50 (t, J = 7, 1H),
7.66 (d, J = 5 Hz, 1H), 7.72 (m, 2H), 8.54 (d, J = 15 Hz, 1H). 13C
NMR (125 MHz, DMSO-d6): 11.09, 20.09, 20.89, 23.60, 27.57, 27.62,
28.50, 40.43, 45.91, 50.20, 50.24, 53.71, 57.10, 101.27, 103.55, 113.07,
113.32, 113.52, 113.69, 113.85, 115.69, 116.11, 118.06, 120.42, 122.76,
125.63, 126.78, 128.29, 128.44, 128.93, 130.06, 130.37, 131.40, 141.64,
141.85, 144.41, 154.66, 158.02, 158.04, 158.51, 158.81, 160.80, 161.29,
3
3
4699
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
a
Scheme 2. Synthesis of 3 and DHX1
a
Reagents and conditions: (a) Et3N, DMF, 110 °C, 20 min, 78%; (b) SOCl2, pyridine, CH2Cl2, DMF, 0 °C, 30 min; (c) Cyclam, DIPEA, CH3CN,
reflux, 30 min, 18% (over steps b and c); (d) Benzyl bromide, DIPEA, CH3CN, 25 °C, 2 h, 28%. Counterions are omitted for clarity. DIPEA =
diisopropylethylamine.
the fluorescence emission spectra of a 2 μM solution of CuDHX1 in
aqueous buffer at pH = 7, before and after treatment with 100 equiv of
CaCl2, MgCl2, NaCl, ZnCl2, KNO3, NaNO2, KO2, H2O2, NaClO,
sodium ascorbate, NaONOO, L-(+)-cysteine hydrochloride, glutha-
thione, methionine, Na2S, or Angeli’s salt. NO gas, 5000 equiv, and
100 μL of 10 mM NaOH (solvent of Angeli’s salt solutions) were also
tested. In each case, the response was quantified by integrating the
emission intensity from 660 to 900 nm and normalized to that of 2 μM
CuDHX1 in aqueous buffer. For the NO and HNO selectivity studies,
the samples were prepared anaerobically. For the NO, HNO, and Na2S
studies, fluorescence spectra were acquired every 1 min for 10 min. For
the remaining analytes, the fluorescence spectra were recorded at 0, 5,
and 10 min. In addition, the fluorescence turn-on of 2 μM CuDHX1 in
the presence of 100 equiv of KO2 and Angeli’s salt was measured in
CH3CN. To determine the effect of pH on the fluorescence emission
of CuDHX1, 2 μM solutions of CuDHX1 were prepared anaerobically
in aqueous buffer (either 50 mM MES, 100 mM KCl; pH 4 and 5 or
50 mM PIPES, 100 mM KCl; pH 6, 7, and 8), and fluorescence
spectra were recorded before and after addition of 100 equiv of
Angeli’s salt.
Cell Culture and Staining Procedures. HeLa cells were cultured
in Dulbecco’s modified Eagle medium (DMEM; Cellgro, MediaTec,
Inc.), supplemented with 10% fetal bovine serum (FBS; HyClone), 1%
penicillin-streptomycin, 1% sodium pyruvate, and 1% L-glutamine. The
cells were grown to 90% confluence at 37 °C with 5% CO2 before
being passed and plated onto poly-D-lysine-coated plates 48 h before
imaging. All cells were used between passage number 5 and 15.
Imaging was conducted when plates reached 50−70% confluence. The
growth medium was replaced with phosphate-buffered saline (PBS)
containing 5 μM CuDHX1 and 3 μM Hoechst 33528 dye, and the
cells were incubated for 15 min. Cells were rinsed with PBS (2 × 2
mL) followed by addition of fresh PBS (2 mL) and mounted on the
microscope. All cell imaging experiments involving addition of HNO
were carried out in PBS because addition of Angeli’s salt to plates
containing DMEM did not lead to any turn-on response (Figure S23),
presumably because of reaction of Angeli’s salt with cysteine or other
components of DMEM. For cell imaging experiments with ZP1, the
growth medium was replaced with dye-free DMEM containing 5 μM
ZP1, 3 μM Hoechst 33528 dye, and the cells were incubated for 1 h.
Cells were rinsed with PBS (2 × 2 mL) before addition of fresh PBS
containing 5 μM CuDHX1 and incubated for 15 min. Cells were
rinsed with PBS (2 × 2 mL) followed by addition of fresh PBS (2 mL)
and mounted on the microscope.
Fluorescence Microscopy. Imaging experiments were performed
using a Zeiss Axiovert 200 M inverted epifluorescence microscope
equipped with an EM-CCD digital camera (Hamamatsu) and a
MS200 XY Piezo Z stage (Applied Scientific Instruments). The light
source was an X-Cite 120 metal-halide lamp (EXFO), and the
fluorescence images were obtained using an oil-immersion objective at
63× or 100× magnification. The filters sets used are defined as blue:
excitation G 365 nm, beamsplitter FT 395 nm, emission BP 445/50
nm; green: excitation BP 470/40 nm, beamsplitter FT 495 nm,
emission 525/50 nm; NIR: excitation HQ 650/45 nm, beamsplitter Q
680 nm, emission HQ 710/50 nm. The microscope was operated
using Volocity software (Perkin-Elmer).
The exposure times for acquisition of fluorescence images were kept
constant for each series of images at each channel. To measure analyte-
induced fluorescence changes, a solution of Angeli’s salt, Na2S, or S-
nitrosoglutathione was added to the plate on the microscope stage to
reach a concentration of 1.5 mM, and images were taken immediately
after addition and after 5 min. Mobilization of intracellular zinc was
induced by treating the cells with 3 mM Angeli’s salt and the released
Zn2+ was chelated by bathing the cells in a solution of fresh PBS
containing 50 μM N,N,N′,N′-tetrakis(2-pyridylmethyl)-
ethylenediamine (TPEN). Quantification of fluorescence intensity
was performed using ImageJ (version 1.45, NIH). The whole cell was
selected as the region of interest, and the integrated fluorescence from
the background region was subtracted from the cell body region.
4700
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
650 nm gives a fluorescence spectrum with a maximum at 715
nm that extends across the NIR region (Figure 1). The
brightness (εϕ) of DHX1 is 1.1 × 103 M−1 cm−1. This value is
comparable to that of Indocyanine Green (1.1 × 103 M−1
cm−1),40 a benchmark NIR fluorophore for in vivo imaging,41
indicating that the photophysical properties of DHX1 are
suitable for in vivo applications. The quantum yield of this
compound is decreased by more than 1 order of magnitude
upon binding of Cu(II), a consequence of paramagnetic
quenching induced by the transition metal.
Analyte Selectivity Studies. The ability of a molecule to
sense a particular analyte in a complex biological sample
depends on the specificity of its chemical reactions. We tested
the reactivity of Cu-3 and CuDHX1 toward HNO, the analyte
of interest, and other species that are present in live cells and
can potentially interfere with nitroxyl sensing. All reactivity tests
were performed in aqueous buffer (50 mM PIPES, 100 mM
KCl, pH = 7). Samples of Cu-3 and CuDHX1 were prepared
anaerobically to ensure that the sensors react directly with
HNO or NO and not with an oxidation product. Cu-3
displayed no change in fluorescence intensity upon addition of
NO gas and showed only a very small fluorescence increase
upon addition of Angeli’s salt, an HNO donor (Figure S11).
CuDHX1, however, reacted rapidly and selectively with HNO.
Upon addition of 100 equiv of Angeli’s salt, CuDHX1
underwent a ∼5-fold increase in fluorescence intensity (Figure
2) after only 2 min (Figure S13). We determined 50 equiv to
RESULTS AND DISCUSSION
■
Synthesis and Photophysical Properties. Scheme 2
summarizes the synthesis of ligand DHX1. 5-(Hydroxymethyl)-
benzene-1,2-diol was allowed to react with IR780, a
commercially available tricarbocyanine dye, to produce
compound 1.35 Alcohol 1 was transformed into chloride 2
with thionyl chloride. Compound 2 was used immediately in a
nucleophilic substitution reaction with 1,4,8,11-tetraazacyclo-
tetradecane (cyclam) to give compound 3. This product was
obtained after purification by RP-HPLC using a mobile phase
containing 0.1% TFA and isolated as the trifluoroacetate salt, as
ascertained by 13C and 19F NMR spectroscopy. Compound 3
was alkylated with benzyl bromide to produce DHX1, which
was purified by RP-HPLC and isolated as the trifluoroacetate
salt of the trans (1,8-substituted cyclam) isomer. Small amounts
of other isomers were recovered after HPLC but not further
characterized. The purity of both 3 and DHX1 was determined
to be >95% by analytical HPLC analysis (Figure S6).
Compounds 3 and DHX1 were treated with CuCl2 to form
their Cu(II) complexes. ESI-MS analysis revealed that under
these conditions the complexes Cu-3 and CuDHX1 were
formed with a coordinated trifluoroacetate ligand with m/z =
799.33 and 889.38, respectively.
The photophysical properties of 3 (Figure S12) and DHX1,
as well as their Cu(II) complexes, as measured in buffered
aqueous solution (50 mM PIPES, 100 mM KCl, pH = 7) are
summarized in Table 1. The absorption spectrum of DHX1 is
Table 1. Summary of Photophysical Properties of
Compounds 3 and DHX1
absorption:
emission:
λmax (nm); φ
λmax (nm); ε (cm−1 M−1
)
ligand
Cu complex
ligand
Cu complex
3
693;
693;
715;
0.059(2)
715;
0.0050(4)
2.7(1) × 104
3.3(2) × 104
DHX1
693;
693;
715;
0.048(3)
715;
0.0027(1)
2.3(1) × 104
2.9(1) × 104
broad with a maximum at 693 nm (Figure 1). The absorbance
of the Cu(II) complex is also broad and, in addition to a
maximum at 693 nm, contains a peak at 650 nm. Excitation at
Figure 2. Fluorescence spectra of 2 μM CuDHX1 (black dashed line)
in aqueous buffer (50 mM PIPES, 100 mM KCl, pH = 7) and 2 min
after the addition of 100 equiv of Angeli’s salt (red solid line). The
black solid line is the spectrum of ligand DHX1. λex: 650 nm.
be the minimum amount of Angeli’s salt needed to induce turn-
on of 2 μM CuDHX1 in aqueous buffer at pH = 7. It is difficult
to estimate the effective concentration of HNO in a solution of
Angeli’s salt because of the high reactivity of nitroxyl.42 We
therefore can only conclude that the detection limit of HNO by
CuDHX1 is ≤50 equiv.
Addition of 5000 equiv of NO resulted in no detectable
change in fluorescence intensity, demonstrating that CuDHX1
detects HNO selectively over NO (Figure 3). The reaction of
CuDHX1 with HNO did not restore the fluorescence to that of
the free ligand (Figure 2). Moreover, after 2 min, the
fluorescence intensity decreased slowly (Figure S13). Upon
reaction of ligand DHX1 with Angeli’s salt, there was a decrease
Figure 1. Fluorescence (dotted lines) and absorbance (solid lines)
spectra of DHX1 (black lines) and CuDHX1 (red lines) in aqueous
buffer (50 mM PIPES, 100 mM KCl, pH = 7).
4701
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
able to reduce CuDHX1. Reduction of the nitrosonium cation
(NO+) to NO occurs at 1.52 V (vs NHE),44 and consequently
NO cannot reduce Cu(II) in CuDHX1. These data explain the
selectivity of CuDHX1 for HNO over NO.
ESI-MS studies in buffer (50 mM PIPES, 100 mM KCl, pH =
7) revealed that both CuDHX1 and Cu-3 exist as mixtures of
complexes with Cu(II) bound to trifluoroacetate or chloride as
fifth ligand. Addition of 100 equiv of Angeli’s salt to CuDHX1
or Cu-3 resulted in loss of the copper ion to give primarily the
m/z of ligands DHX1 and 3, respectively (Figures S9 and S10).
In the case of Cu-3, the m/z of the copper complex without the
fifth ligand was also detected (Figure S10).
The X-band EPR spectrum of CuDHX1, measured at 77 K in
CH3OH, revealed a rhombic signal (Figure 4), which
Figure 3. Normalized integrated fluorescence intensity (660−900 nm)
of 2 μM CuDHX1 in aqueous buffer (50 mM PIPES, 100 mM KCl,
pH = 7) and 10 min after addition of 100 equiv of the analyte or 2 min
after addition of 100 equiv of HNO. λex: 650 nm.
in fluorescence intensity (Figure S14), indicating that one of
the products of the decomposition of the salt reacts with DHX1
to diminish its emission in buffered solution.
Angeli’s salt, which decomposes readily (t1/2 = 3 min)38 at
pH = 7 to give HNO and NaNO2, is usually handled as an
aqueous solution at pH = 12. To verify that HNO is
responsible for the observed fluorescence increase, the
reactivity of CuDHX1 toward NaNO2 (100 equiv) and 10
mM NaOH (100 μL) was evaluated. No increase in
fluorescence intensity was observed upon reaction of
CuDHX1 with either 100 equiv of NaNO2 or 10 mM NaOH
in aqueous buffer (Figure 3). Additionally, no turn-on was
induced when 100 equiv of CaCl2, MgCl2, NaCl, ZnCl2, KNO3,
H2O2, NaClO, sodium ascorbate, NaONOO, L-(+)-cysteine,
GSH, or methionine were added to a solution of CuDHX1
(Figure 3). CuDHX1 is selective for HNO over superoxide
(KO2) both in buffer (Figure 3) and in CH3CN (Figure S16).
The only analyte that promoted some detectable fluorescence
enhancement after 10 min was H2S (Figure 3), which elicited a
much slower turn-on compared to that induced by HNO. This
selectivity is remarkable in view of the susceptibility of some
Cu-based sensors toward reducing thiols26−28 and makes
CuDHX1 a much more valuable sensor for use in biological
samples where thiols are abundant. In addition, the
fluorescence of CuDHX1 was measured at different pH values.
A blue shift and decrease in intensity was observed at pH < 6,
consistent with protonation of the hydroxyl group on the
fluorophore (pKa ∼5.6).35 At low pH values, HNO-induced
turn-on was considerably smaller than at neutral pH (Figure
S17).
Figure 4. X-band EPR spectra of 400 μM CuDHX1 in CH3OH. Top:
CuDHX1 before (black line) and after addition of 5000 equiv of NO
(red line). Middle: CuDHX1 before (black line) and after (red line)
addition of 100 equiv of Angeli’s salt. Bottom: CuDHX1 after
reduction by HNO (black line) and after reoxidation by air (red line).
Collection parameters: temperature, 77 K; modulation amplitude, 20
G; microwave power, 0.2 mW at 9.23 GHz.
disappeared upon treatment with 100 equiv of Angeli’s salt,
as expected for reduction of Cu(II) to Cu(I). In contrast,
addition of 5000 equiv of NO gas to a solution of CuDHX1 in
CH3OH did not evoke a noticeable change in the EPR
spectrum (Figure 4). The reduced form of CuDHX1 could be
reoxidized by allowing air into the EPR tube, giving a more
axially symmetric signal; simulated traces are provided in Figure
S20. Notably, we could not observe an EPR-silent species after
reaction of Cu-3 with Angeli’s salt (Figure S22). This
Mechanistic Investigation. Cyclic voltammetric studies of
CuDHX1 and Cu-3 revealed quasi-reversible Cu(II)/Cu(I)
reductions at 370 and 325 mV (vs Fc/Fc+), respectively
(Figures S18 and S19). The reduction of NO to give HNO at
pH = 7 occurs at −0.11 V (vs NHE,)43 and the reduction
potential of the Cu(II)/Cu(I) couple in CuDHX1 is 1.01 V (vs
NHE). These values indicate that HNO is thermodynamically
4702
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
observation suggests that, if Cu(I)-3 is formed, it converts
immediately to Cu(II), even under anaerobic conditions. The
nature of the species responsible for this reoxidized copper
material was not further investigated.
channels (Figure 6D,G). Treatment of these cells with 3 mM
Angeli’s salt led to an increase in fluorescence in both the NIR
Live Cell Imaging. We assessed the ability of CuDHX1 to
detect HNO in live cells. Human cervical cancer (HeLa) cells
were incubated with 5 μM CuDHX1 and 3 μM Hoechst 33528
nuclear stain in PBS for 15 min at 37 °C. Fluorescence
microscopy images revealed only faint fluorescence in the NIR
channel (Figure 5C), consistent with the low brightness of
Figure 6. Multicolor/multianalyte imaging of HNO and mobile zinc.
(A) DIC image; (B) blue channel showing nuclei; (C) quantification
of the fluorescence intensity in the green and NIR channels; (D) green
channel before addition of Angeli’s salt; (E) green channel after
addition of Angeli’s salt; (F) green channel after addition of TPEN;
(G) NIR channel before addition of Angeli’s salt; (H) NIR channel
after addition of Angeli’s salt; and (I) NIR channel after addition of
TPEN. Scale bar =10 μm.
Figure 5. Fluorescence microscopy images of cells incubated with
CuDHX1 in PBS before and after addition of Angeli’s salt: (A)
Differential interference contrast (DIC) image; (B) blue channel
showing nuclei; (C) NIR channel before addition of Angeli’s salt; and
(D) NIR channel 5 min after treatment with 1.5 mM Angeli’s salt.
Scale bar =25 μm.
CuDHX1. Treatment with 1.5 mM Angeli’s salt resulted in a
∼3-fold increase in fluorescence intensity after only 5 min to
restore a bright signal in the NIR channel (Figure 5D).
Time-lapsed microscopy experiments indicated that, in the
absence of HNO, CuDHX1 displays only a slight turn-on over
time in live cells (Figure S24), significantly less than that
induced by HNO. Treatment of HeLa cells with GSNO, an NO
donor, did not elicit any detectable fluorescence turn-on
(Figure S25), demonstrating that the sensor is selective for
HNO over NO and S-nitrosothiols in live cells. In addition,
incubation of live cells with Na2S resulted in only a very small
increase in fluorescence, and subsequent treatment of the same
cells with Angeli’s salt still induced a strong fluorescence turn-
on (Figure S26). This experiment proves that the presence of
H2S does not interfere with intracellular HNO sensing by
CuDHX1.
An advantage of NIR probes is that they can be used in
combination with visible-color sensors to image two or more
analytes simultaneously. In this kind of experiment, it is
possible to investigate in real time the interplay between
signaling molecules in live organisms. An example of such a
relationship is the release of mobile zinc upon nitrosation of
metallothionein.45,46 Mobilization of endogenous zinc induced
by NO donors can be detected in live cells by fluorescent
sensors,47 but NO is unable to nitrosate thiolates directly.48
HNO, however, reacts directly with thiols16 and may be capable
of releasing chelatable zinc. To investigate this possibility, we
incubated HeLa cells with CuDHX1 and the green-fluorescent,
zinc-selective, sensor ZP1.36 Before any treatment, the cells
showed only very faint fluorescence in both the NIR and green
channel and the green channel (Figure 6C,E,H). Addition of 50
μM TPEN reduced the fluorescence only in the green channel
(Figure 6C,F), demonstrating that the observed turn-on
corresponded to an increase in the intracellular levels of mobile
zinc. Addition of TPEN did not significantly change the
fluorescence of CuDHX1 in the NIR channel (Figure 6C,I).
As discussed above, Angeli’s salt was handled as a solution in
aqueous 10 mM NaOH, which produces HNO and NaNO2.
To prove that the mobilization of Zn2+ was induced by HNO,
HeLa cells were incubated with 3 mM NaNO2 in aqueous 10
mM NaOH. No significant change in fluorescence intensity was
detected in the green channel under these conditions (Figure
S27).
These experiments reveal the value of CuDHX1 as a versatile
sensor for multicolor imaging, allowing for simultaneous
observation of HNO and mobile Zn2+. They also prove that
release of mobile Zn2+ is a downstream effect of addition of
HNO. To the best of our knowledge, our results provide the
first application of a multicolor microscopy experiment to study
the cellular chemistry of HNO.
SUMMARY AND CONCLUSIONS
■
CuDHX1 provides a fast, selective, NIR fluorescent sensor for
the detection of HNO. CuDHX1 is selective against a variety of
analytes present in live cells, including thiols and H2S, which is
an advantage over previous Cu(II)-based HNO sensors. EPR
spectroscopic and ESI-MS studies showed that the sensing
mechanism relies on reduction of Cu(II) by HNO, with
concomitant dissociation of Cu(I) from the cyclam-based
4703
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
(12) Paolocci, N.; Saavedra, W. F.; Miranda, K. M.; Martignani, C.;
Isoda, T.; Hare, J. M.; Espey, M. G.; Fukuto, J. M.; Feelisch, M.; Wink,
D. A.; Kass, D. A. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 10463−10468.
(13) Dautov, R. F.; Ngo, D. T. M.; Licari, G.; Liu, S.; Sverdlov, A. L.;
Ritchie, R. H.; Kemp-Harper, B. K.; Horowitz, J. D.; Chirkov, Y. Y.
Nitric Oxide 2013, 35, 72−78.
ligand. The sensor detects HNO in live cells, even in the
presence of H2S. The narrow emission of CuDHX1 in the NIR
region makes it useful for multicolor imaging experiments,
exemplified here by its use in combination with ZP1 to track
the increase in intracellular levels of mobile zinc elicited by
exogenously applied Angeli’s salt. These results represent the
first direct observation of a relationship between HNO and
mobile zinc in a biological environment. We anticipate that
CuDHX1 will be useful for investigating the biology of HNO
and its interplay with other signaling molecules by multicolor
fluorescence microscopy in the NIR.
(14) Choe, C.-u.; Lewerenz, J.; Fischer, G.; Uliasz, T. F.; Espey, M.
G.; Hummel, F. C.; King, S. B.; Schwedhelm, E.; Boger, R. H.; Gerloff,
̈
C.; Hewett, S. J.; Magnus, T.; Donzelli, S. J. Neurochem. 2009, 110,
1766−1773.
(15) Choe, C.-U.; Lewerenz, J.; Gerloff, C.; Magnus, T.; Donzelli, S.
Antioxid. Redox Signal. 2011, 14, 1699−1711.
(16) Fukuto, J. M.; Carrington, S. J. Antioxid. Redox Signal. 2011, 14,
1649−1657.
ASSOCIATED CONTENT
■
(17) Kroncke, K. D.; Fehsel, K.; Schmidt, T.; Zenke, F. T.; Dasting,
̈
S
* Supporting Information
I.; Wesener, J. R.; Bettermann, H.; Breunig, K. D.; Kolb-Bachofen, V.
Biochem. Biophys. Res. Commun. 1994, 200, 1105−1110.
(18) Maret, W. Adv. Nutr. 2013, 4, 82−91.
NMR spectroscopic data, analytical HPLC traces, high-
resolution ESI-MS, fluorescence and absorbance spectra of 3,
time-dependent fluorescence experiments, cyclic voltammo-
grams, simulated EPR spectra, time-lapsed microscopy
measurements, and control imaging experiments. This material
is available free of charge via the Internet at http://pubs.acs.org
(19) Switzer, C. H.; Flores-Santana, W.; Mancardi, D.; Donzelli, S.;
Basudhar, D.; Ridnour, L. A.; Miranda, K. M.; Fukuto, J. M.; Paolocci,
N.; Wink, D. A. Biochim. Biophys. Acta Bioenerg. 2009, 1787, 835−840.
(20) Sakai, N.; Kaufman, S.; Milstien, S. Mol. Pharmacol. 1993, 43,
6−10.
(21) Adak, S.; Wang, Q.; Stuehr, D. J. J. Biol. Chem. 2000, 275,
33554−33561.
AUTHOR INFORMATION
■
(22) Murphy, M. E.; Sies, H. Proc. Natl. Acad. Sci. U.S.A. 1991, 88,
10860−10864.
Corresponding Authors
lippard@mit.edu
(23) Sharpe, M. A.; Cooper, C. E. Biochem. J. 1998, 332, 9−19.
(24) Filipovic, M. R.; Miljkovic, J. L.; Nauser, T.; Royzen, M.; Klos,
priveraf@mit.edu
K.; Shubina, T.; Koppenol, W. H.; Lippard, S. J.; Ivanovic-
I. J. Am. Chem. Soc. 2012, 134, 12016−12027.
́
(25) Miljkovic, J. L.; Kenkel, I.; Ivanovic-Burmazovic, I.; Filipovic, M.
́ ́
Burmazovic,
Notes
The authors declare no competing financial interest.
́
R. Angew. Chem., Int. Ed. 2013, 52, 12061−12064.
ACKNOWLEDGMENTS
(26) Rosenthal, J.; Lippard, S. J. J. Am. Chem. Soc. 2010, 132, 5536−
5537.
(27) Royzen, M.; Wilson, J. J.; Lippard, S. J. J. Inorg. Biochem. 2013,
■
This work was supported by the National Science Foundation
(Grant CHE-1265770 to S.J.L.). Instrumentation in the MIT
DCIF is maintained with funding from NIH Grant
1S10RR13886-01. A.T.W. acknowledges support from the
Paul E. Gray (1954) Endowed Fund for UROP. A.D.L. thanks
the NIH for support under Interdepartmental Biotechnology
Training Grant T32 GM008334. P. R.-F. thanks the Swiss
National Science Foundation for a postdoctoral fellowship. We
thank Dr. Robert Radford for providing a sample of ZP1 and
Dr. Ulf-Peter Apfel for insightful discussions.
118, 162−170.
(28) Zhou, Y.; Liu, K.; Li, J.-Y.; Fang, Y.; Zhao, T.-C.; Yao, C. Org.
Lett. 2011, 13, 1290−1293.
(29) Apfel, U.-P.; Buccella, D.; Wilson, J. J.; Lippard, S. J. Inorg.
Chem. 2013, 52, 3285−3294.
(30) Kawai, K.; Ieda, N.; Aizawa, K.; Suzuki, T.; Miyata, N.;
Nakagawa, H. J. Am. Chem. Soc. 2013, 135, 12690−12696.
(31) Hilderbrand, S. A.; Weissleder, R. Curr. Opin. Chem. Biol. 2010,
14, 71−79.
(32) Egawa, T.; Hirabayashi, K.; Koide, Y.; Kobayashi, C.; Takahashi,
N.; Mineno, T.; Terai, T.; Ueno, T.; Komatsu, T.; Ikegaya, Y.;
Matsuki, N.; Nagano, T.; Hanaoka, K. Angew. Chem., Int. Ed. 2013, 52,
3874−3877.
REFERENCES
■
(1) Cannon, R. O., III Clin. Chem. 1998, 44, 1809−1819.
(2) Bogdan, C. Nat. Immunol. 2001, 2, 907−916.
(3) Bredt, D. S.; Hwang, P. M.; Snyder, S. H. Nature 1990, 347, 768−
770.
(33) Kobayashi, H.; Koyama, Y.; Barrett, T.; Hama, Y.; Regino, C. A.
S.; Shin, I. S.; Jang, B.-S.; Le, N.; Paik, C. H.; Choyke, P. L.; Urano, Y.
ACS Nano 2007, 1, 258−264.
(34) Sasakura, K.; Hanaoka, K.; Shibuya, N.; Mikami, Y.; Kimura, Y.;
Komatsu, T.; Ueno, T.; Terai, T.; Kimura, H.; Nagano, T. J. Am. Chem.
Soc. 2011, 133, 18003−18005.
(4) Palmer, R. M. J.; Ferrige, A. G.; Moncada, S. Nature 1987, 327,
524−526.
(5) Bult, H.; Boeckxstaens, G. E.; Pelckmans, P. A.; Jordaens, F. H.;
Van Maercke, Y. M.; Herman, A. G. Nature 1990, 345, 346−347.
(6) Pieper, G. M. Hypertension 1998, 31, 1047−1060.
(7) Wei, G.; Dawson, V. L.; Zweier, J. L. Biochim. Biophys. Acta, Mol.
Basis Dis. 1999, 1455, 23−34.
(35) Yuan, L.; Lin, W.; Zhao, S.; Gao, W.; Chen, B.; He, L.; Zhu, S. J.
Am. Chem. Soc. 2012, 134, 13510−13523.
(36) Burdette, S. C.; Walkup, G. K.; Spingler, B.; Tsien, R. Y.;
Lippard, S. J. J. Am. Chem. Soc. 2001, 123, 7831−7841.
(37) Chapman, G.; Henary, M.; Patonay, G. Anal. Chem. Insights
2011, 6, 29−36.
(8) Heales, S. J. R.; Bolanos, J. P.; Stewart, V. C.; Brookes, P. S.;
̃
Land, J. M.; Clark, J. B. Biochim. Biophys. Acta, Bioenerg. 1999, 1410,
215−228.
(38) Miranda, K. M.; Nagasawa, H. T.; Toscano, J. P. Curr. Top. Med.
Chem. 2005, 5, 649−664.
(9) Hussain, S. P.; Hofseth, L. J.; Harris, C. C. Nat. Rev. Cancer 2003,
3, 276−285.
(39) Stoll, S.; Schweiger, A. J. Magn. Reson. 2006, 178, 42−55.
(40) Philip, R.; Penzkofer, A.; Baumler, W.; Szeimies, R. M.; Abels, C.
̈
(10) Fukuto, J. M.; Cisneros, C. J.; Kinkade, R. L. J. Inorg. Biochem.
2013, 118, 201−208.
J. Photochem. Photobiol. A 1996, 96, 137−148.
(41) Alander, J. T.; Kaartinen, I.; Laakso, A.; Patila, T.; Spillmann, T.;
(11) Bullen, M. L.; Miller, A. A.; Andrews, K. L.; Irvine, J. C.; Ritchie,
R. H.; Sobey, C. G.; Kemp-Harper, B. K. Antioxid. Redox Signal. 2011,
14, 1675−1686.
̈
̈
Tuchin, V. V.; Venermo, M.; Val
2012, 940585.
isuo, P. Int. J. Biomed. Imaging 2012,
̈
4704
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705

Journal of the American Chemical Society
Article
(42) Miranda, K. M.; Dutton, A. S.; Ridnour, L. A.; Foreman, C. A.;
Ford, E.; Paolocci, N.; Katori, T.; Tocchetti, C. G.; Mancardi, D.;
Thomas, D. D.; Espey, M. G.; Houk, K. N.; Fukuto, J. M.; Wink, D. A.
J. Am. Chem. Soc. 2005, 127, 722−731.
(43) Armstrong, D. A.; Huie, R. E.; Lymar, S.; Koppenol, W. H.;
́
Merenyi, G.; Neta, P.; Stanbury, D. M.; Steenken, S.; Wardman, P.
BioInorg. React. Mech. 2014, 9, 59−61.
(44) Lee, K. Y.; Kuchynka, D. J.; Kochi, J. K. Inorg. Chem. 1990, 29,
4196−4204.
(45) Berendji, D.; Kolb-Bachofen, V.; Meyer, K. L.; Grapenthin, O.;
Weber, H.; Wahn, V.; Kroncke, K.-D. FEBS Lett. 1997, 405, 37−41.
̈
(46) Stamler, J. S.; Simon, D. I.; Osborne, J. A.; Mullins, M. E.; Jaraki,
O.; Michel, T.; Singel, D. J.; Loscalzo, J. Proc. Natl. Acad. Sci. U.S.A.
1992, 89, 444−448.
(47) Spahl, D. U.; Berendji-Grun, D.; Suschek, C. V.; Kolb-Bachofen,
̈
V.; Kroncke, K.-D. Proc. Natl. Acad. Sci. U.S.A. 2003, 100, 13952−
̈
13957.
(48) Kozhukh, J.; Lippard, S. J. Inorg. Chem. 2012, 51, 7346−7353.
4705
dx.doi.org/10.1021/ja500315x | J. Am. Chem. Soc. 2014, 136, 4697−4705