A Ratiometric Acoustogenic Probe for in Vivo Imaging of Endogenous Nitric Oxide

Article abstract of DOI:10.1021/jacs.7b10783

Photoacoustic (PA) imaging is an emerging imaging modality that utilizes optical excitation and acoustic detection to enable high resolution at centimeter depths. The development of activatable PA probes can expand the utility of this technology to allow for detection of specific stimuli within live-animal models. Herein, we report the design, development, and evaluation of a series of Acoustogenic Probe(s) for Nitric Oxide (APNO) for the ratiometric, analyte-specific detection of nitric oxide (NO) in vivo. The best probe in the series, APNO-5, rapidly responds to NO to form an N-nitroso product with a concomitant 91 nm hypsochromic shift. This property enables ratiometric PA imaging upon selective irradiation of APNO-5 and the corresponding product, tAPNO-5. Moreover, APNO-5 displays the requisite photophysical characteristics for in vivo PA imaging (e.g., high absorptivity, low quantum yield) as well as high biocompatibility, stability, and selectivity for NO over a variety of biologically relevant analytes. APNO-5 was successfully applied to the detection of endogenous NO in a murine lipopolysaccharide-induced inflammation model. Our studies show a 1.9-fold increase in PA signal at 680 nm and a 1.3-fold ratiometric turn-on relative to a saline control.

Full text of DOI:10.1021/jacs.7b10783

Article
Cite This: J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
pubs.acs.org/JACS
A Ratiometric Acoustogenic Probe for in Vivo Imaging of
Endogenous Nitric Oxide
Christopher J. Reinhardt, Effie Y. Zhou, Michael D. Jorgensen, Gina Partipilo, and Jefferson Chan*
Department of Chemistry and Beckman Institute for Advanced Science and Technology, University of Illinois at Urbana−Champaign,
Urbana, Illinois 61801, United States
*
S Supporting Information
ABSTRACT: Photoacoustic (PA) imaging is an emerging imaging
modality that utilizes optical excitation and acoustic detection to
enable high resolution at centimeter depths. The development of
activatable PA probes can expand the utility of this technology to
allow for detection of specific stimuli within live-animal models.
Herein, we report the design, development, and evaluation of a series
of Acoustogenic Probe(s) for Nitric Oxide (APNO) for the
ratiometric, analyte-specific detection of nitric oxide (NO) in vivo.
The best probe in the series, APNO-5, rapidly responds to NO to
form an N-nitroso product with a concomitant 91 nm hypsochromic
shift. This property enables ratiometric PA imaging upon selective
irradiation of APNO-5 and the corresponding product, tAPNO-5.
Moreover, APNO-5 displays the requisite photophysical character-
istics for in vivo PA imaging (e.g., high absorptivity, low quantum yield) as well as high biocompatibility, stability, and selectivity
for NO over a variety of biologically relevant analytes. APNO-5 was successfully applied to the detection of endogenous NO in a
murine lipopolysaccharide-induced inflammation model. Our studies show a 1.9-fold increase in PA signal at 680 nm and a 1.3-
fold ratiometric turn-on relative to a saline control.
3
,11
INTRODUCTION
progression,
where the location, source, and role of NO
■
12
remain unclear. The current arsenal of NO donors and ex vivo
analyses provides important preliminary information but has
yet to address these outstanding questions.
Current methods for detecting NO are inadequate for in vivo
applications. For example, the colorimetric Griess assay is useful
for analyzing NO in cell lysates; however, this is performed
indirectly via detection of nitrite and nitrate and requires acidic
Nitric oxide (NO) is a metastable free radical with key roles in
1
2
the regulation of vascular tone, neuronal signaling, tumor
3
4,5
progression, and immunology. Under normoxic conditions,
NO is biosynthesized by nitric oxide synthases (NOS), which
generate NO through the oxidation of L-arginine to L-citrulline.
The NOS family includes an inducible, calcium-independent
isoform (iNOS), which is primarily linked to the immune
response, and constitutively expressed, calcium-dependent
isoforms (ncNOS and ecNOS), which are linked to
13
conditions that are not biocompatible. In contrast, techniques
such as electron paramagnetic resonance (EPR) spectroscopy
14
15
6
7
and magnetic resonance imaging (MRI) have been employed
for NO detection in vivo at relevant imaging depths; however,
these approaches are limited by low resolution and sensitivity,
homeostatic maintenance. NO can also be enzymatically or
nonenzymatically generated by the reduction of nitrite under
hypoxic conditions, a process which has been observed in the
1
6
8
respectively. Amperometry displays high sensitivity (pM) but
requires invasive procedures and can only detect NO in direct
ischemic heart and results in NO-mediated tissue damage. Due
to its various roles in physiological and pathological processes,
the identification, detection, and quantification of NO
continues to be an active area of research. Since ex vivo
samples (e.g., cultured cells, tissue slices, etc.) do not
adequately mimic the complexities of in vivo samples (e.g.,
interaction between the cell and its surrounding environment),
it is essential to develop new methods for the real-time
detection of NO. However, noninvasive, in vivo detection of
NO with high spatiotemporal resolution remains a profound
challenge in the field. The ability to perform longitudinal
studies with high-resolution in vivo imaging would provide
exciting avenues for studying the role of NO as a neuro-
1
7−19
contact with the probe.
luminescence and fluorescence imaging are noninvasive
and enable high resolution with high contrast at shallow
imaging depths; as a result, a diverse palette of reaction-based
fluorescent probes has been developed, primarily for cellular
studies. In principle, many of the red-shifted and multiphoton
congeners can be employed for in vivo imaging, but these
studies require invasive surgical procedures, are confined to
peripheral limbs, and/or suffer from limited resolution
Optical methods such as
20
21
16
22
23
23,24
Received: October 10, 2017
9
,10
transmitter
and elucidating its dichotomous role in cancer
©
XXXX American Chemical Society
A
DOI: 10.1021/jacs.7b10783
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Journal of the American Chemical Society
Article
due to the scattering of emitted light within tissue. Several
other fluorescent NO probes have been applied to living
animals; however, in these examples, the specimen is sacrificed
to enable harvesting and ex vivo imaging of the tissue of
began by incorporating the o-phenylenediamine (OPD)
functionality onto the aza-BODIPY dye platform to afford
APNO-1 (Figure 1). This scaffold was selected over the various
39
2
5−29
interest.
Photoacoustic (PA) imaging is a noninvasive imaging
modality with the potential to overcome these challenges. PA
imaging couples the precision of optical imaging with the deep
tissue capabilities of ultrasound imaging. Specifically, PA
imaging relies on the ability of optically excited small molecules
to release heat through nonradiative relaxation. By pulsing the
excitation source, it is possible to generate fluctuating
thermoelastic expansions that propagate through the tissue as
ultrasound and can be measured by transducers. Since sound
scatters 3 orders of magnitude less than light within tissue, PA
imaging can generate high-resolution images at depths
inaccessible to fluorescence imaging. Empirically, it has been
shown that the spatial resolution of PA imaging is ∼1/200 of
the imaging depth. For example, at 8 cm depths, the resolution
is estimated be ∼400 μM, while at 1 cm the resolution would
30
be predicted to be ∼50 μm. Owing to the noninvasive nature
of PA imaging, it is possible to conduct longitudinal
experiments and acquire high-resolution deep tissue images
within live animals. To date, analyte-specific acoustogenic
31
probes have been developed for the detection of copper and
3
2,33
calcium;
however, the utility of these probes has only been
demonstrated in tissue-mimicking systems and in translucent
larval zebrafish. More recently, a stimulus-specific acoustogenic
probe for hypoxia has been reported and applied for imaging
Figure 1. Structures, triggers, and sensing mechanisms for the panel of
APNO (red) and tAPNO (blue).
34
hypoxia within murine tumor and ischemia models.
Herein, we report the design, synthesis, characterization, and
application of a series of five reaction-based PA probes for NO.
While all members of this panel exhibited excellent in vitro
selectivity for NO over other reactive nitrogen species, oxidants,
and reactive carbonyl species (Figure S1), we employed our
best probe, APNO-5, to noninvasively detect endogenous NO
in a murine lipopolysaccharide (LPS)-mediated inflammation
model using PA imaging. This work highlights the utility of
small-molecule, reaction-based photoacoustic probes for in vivo
analyte-specific detection.
other NIR absorbing dye platforms (e.g., cyanines) because of
its well-documented photo- and chemostability. We rational-
ized that strategic positioning of the trigger would allow for
NO-mediated disruption of the push−pull system, resulting in a
concomitant hypsochromic (blue) shift in the wavelength of
maximum absorbance (λmax). Empirically, a shift ≥50 nm is
desirable for ratiometric imaging such that each form can be
selectively irradiated to produce a corresponding PA signal.
Treatment of APNO-1 with NO resulted in a 55 nm λmax
blueshift from 731 to 676 nm (Figure S2). When APNO-1 and
turned-over APNO-1 (tAPNO-1) were excited at 740 and 680
nm, respectively, a ratiometric turn-on PA response of 7.4-fold
was observed (Figure S3). Experimental excitation wavelengths
differ from the reported λmax values due to a combination of
instrumental range (680−900 nm) and experimentally larger
PA ratio. Although the observed ratiometric response would be
sufficient to detect most analyte targets, we deemed it essential
to improve this property further due to the low biological
abundance of NO. Moreover, the preparation of APNO-1 was
hampered by decomposition of key synthetic intermediates
which resulted in low overall yields.
RESULTS AND DISCUSSION
■
Acoustogenic Probe Design. When developing acousto-
genic probes for NO, it is essential to satisfy six design criteria
to optimize in vivo performance. First, a probe must exhibit
maximal absorption in the NIR-I (620−950 nm) or NIR-II
3
5
(
1000−1700 nm) window to ensure incident light penetrates
deep within tissue without significant attenuation. Second, it
4
−1
−1
should possess a large extinction coefficient (>10 M cm )
and low quantum yield, since absorptivity and nonemissive
relaxation are proportional to PA signal intensity. Third, it must
react with NO in a rapid and highly selective manner, since NO
is present in low abundance (nM to low μM range) and has a
In order to address these limitations, we turned our attention
to developing probes that rely on NO-mediated deamina-
36
short biological lifetime (seconds). Fourth, a probe should be
optimized for ratiometric imaging, which is necessary to
account for imaging artifacts (e.g., dye loading, heterogeneous
40,41
tion.
Specifically, APNO-2 features an electron-rich o-
aminophenol moiety that undergoes NO-mediated diazonium
formation and subsequent homolytic bond cleavage to afford
tAPNO-2 (Figure 1). It was rationalized that the removal of the
strongly electron-donating aniline would have a larger effect on
the λmax than triazole formation. Treatment of APNO-2 with
NO resulted in a 59 nm blueshift from 735 to 676 nm (Figure
S2) and 13.1-fold ratiometric turn-on response when irradiated
at 750 and 680 nm (Figure S3). While this improvement in the
37
distribution, and variability in imaging depth). Fifth, the
probe must display favorable pharmacokinetic and pharmaco-
dynamic properties (e.g., rapid uptake by the tissue of interest).
Last, the probe should be biocompatible and produce minimal
38
effects on the native biological system.
To this end, we have rationally developed and synthesized a
panel of Acoustogenic Probe(s) for Nitric Oxide (APNO). We
B
DOI: 10.1021/jacs.7b10783
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Journal of the American Chemical Society
Article
dynamic range may have been sufficient, it was limited by
insufficient reaction kinetics. Specifically, the reaction proceeds
via an N-nitroso intermediate (which was detected via
fluorescence and UV-vis) followed by slow diazonium
Scheme 1. Synthesis of APNO-5
40
formation, and decomposition to yield the product. These
properties significantly complicate the interpretation of imaging
results. Of note, we also prepared APNO-3, a constitutional
isomer in which the aniline is not directly in conjugation with
the π-system. While this probe also underwent a blueshift upon
reaction with NO, the change in λmax was small and the probe
suffered from pH dependence in the biological range (Figure
S4). Taken together, the two deamination-based probes do not
satisfy the necessary criteria for in vivo detection of NO.
In light of the rapid formation of N-nitroso intermediates, we
reasoned that monoalkylation of a reactive aniline could both
enhance the reactivity and block the necessary rearrangements
for diazonium formation and deamination. This led to the
development of APNO-4 (Figure 1) which features an N-
methylaniline nucleophile trigger. Additionally, the hydroxyl
group was replaced with a methoxy substituent for synthetic
ease and to prevent possible pH dependence. While there are
examples of photoinduced electron transfer (PET)-based
fluorogenic probes relying on NO-responsive secondary
28,29,42
anilines in the literature,
to the best of our knowledge,
APNO-4 represents the first probe (for any imaging modality)
that employs the N-nitrosation for modulation of a push−pull
system for ratiometric imaging. We observed rapid formation of
the anticipated N-nitroso product (tAPNO-4) and a significant
86 nm blueshift of the λmax from 764 to 678 nm upon treatment
with NO (Figure S2); however, tAPNO-4 was poorly soluble in
aqueous solution. This resulted in inconsistent results due to
aggregation and precipitation. To overcome this limitation, we
developed APNO-5, a positively charged analogue featuring a
PEGylated tetraalkylammonium group to improve solubility,
prevent dye aggregation, and facilitate uptake by cells in vivo.
Synthesis. Each APNO was synthesized via Paal−Knorr
cyclization and coupling of two 4-nitro-1,3- diphenylbutan-1-
one analogues, followed by boron chelation to afford the aza-
Figure 2. (a) Normalized absorbance spectra of 2 μM APNO-5 (red)
and tAPNO-5 (blue). (b) APNO-5 (2 μM) fluorescence enhancement
following 1 h treatment with excess (100 equiv) reactive metal (red),
carbonyl (green), oxygen (blue), or nitrogen (purple) species. (c) PA
spectra of 10 μM APNO-5 (red) and tAPNO-5 (blue). (d) PA images
of 10 μM APNO-5 and tAPNO-5 at 680 and 770 nm. All spectra and
assays were performed in ethanolic 20 mM potassium phosphate
buffer (pH 7.4, 50% v/v). Data presented as mean ± standard
deviation (n = 3) for parts b and c.
4
3
BODIPY dye.
The synthesis of APNO-5 began with o-aminophenol.
Carbamate protection with carbonyldiimidazole followed by
Friedel−Crafts acylation afforded acetophenone 2. Clai-
sen−Schmidt condensation between 2 and benzaldehyde
afforded chalcone 3 in good yields. Sequential Boc-protection
and carbamate deprotection were performed in one pot under
basic conditions, followed by methylation of the chalcone
intermediate 4 with sodium hydride and methyl iodide to yield
5 at 770 and 680 nm, respectively, resulted in a 18.6-fold
ratiometric PA turn-on response in vitro (Figure 2c). Addi-
tionally, the improved aqueous solubility due to the appendd
tetraalkylammonium PEG group enhanced its properties for
ratiometric PA imaging. Insoluble aza-BODIPY dyes are known
to form J-aggregates and nanoparticles, where the λ is red-
max
44
shifted relative to the nonaggregated form. Since our strategy
relies heavily on selective excitation at specific wavelengths, it is
essential that formation of wavelength-shifted aggregates is
minimized. To verify this, we monitored the degree of
aggregation as a function of surfactant concentration at a
fixed dye concentration. At 2 μM, aggregation was attenuated at
concentrations as low as 0.1% surfactant for APNO-5, whereas
APNO-4 required ≥10% (Figure S5). Moreover, the
tetraalkylammonium PEG appendage increased the water
solubility of APNO-5 and tAPNO-5, as indicated by clogP
values of 3.8 and 3.8, relative to 7.6 and 7.3 for APNO-4 and
tAPNO-4 (Table S2). Since the calculation methods are unable
to account for charge, APNO-5 and tAPNO-5’s predicted
5
. Michael addition with the nitromethane anion under basic
conditions provided the 4-nitro-1,3- diphenylbutan-1-one 6 in
6% yield. The asymmetric dipyrromethane core was formed in
reasonable yields followed by boron chelation with boron
trifluoride dietherate in the presence of Hunig’s base to afford 8
8
̈
in 33% yield. Copper(I)-catalyzed alkyne−azide cycloaddition
with trimethylammonium-PEG-N gave APNO-5 in quantita-
3
tive yields (Scheme 1).
In Vitro Characterization of APNO-5. As anticipated,
APNO-5 reacted rapidly with NO to afford the N-nitroso
product (tAPNO-5) along with a significant 91 nm blueshift of
the λmax from 764 to 673 nm, the largest in the APNO series
45
(Figure 2a).
values should serve as overapproximations of the true values.
The extinction coefficient of APNO-5 was sufficiently large
3.2 × 10 M cm ) to ensure generation of strong PA signals
in vivo (Table 1). Selective irradiation of APNO-5 and tAPNO-
This key structural feature was also required for APNO-5 to
permeate deep into tissue after subcutaneous administration
into live mice. Other congeners pooled at the injection site and
4
−1
−1
(
C
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Journal of the American Chemical Society
Article
Table 1. Photophysical Characterization of APNO and tAPNO in Ethanolic 20 mM Potassium Phosphate Buffer (pH 7.4, 50%
v/v)
1
a
ε (M−1 cm−1
3.7 × 104
a
APNO
ƛmax (nm)
ε (M− cm−1
)
ϕ
tAPNO
ƛmax (nm)
)
ϕ
4
1
2
3
4
5
731
735
679
764
764
3.3 × 10
7.4 × 10
2.2 × 10
3.7 × 10
0.0065
0.0006
0.0708
0.0003
0.0003
1
2
3
4
5
676
676
678
678
673
0.14
0.12
4
4
6.9 × 10
4
b
b
n.d.
n.d.
n.d.
4
c
c
n.d.
4
3.3 × 104
3.2 × 10
0.23
a
b
Measured relative to dimethoxy aza-BODIPY (ϕ = 0.32, CHCl ). Not determined: tAPNO-3 displays pH dependence under these conditions.
3
c
Not determined: tAPNO-4 precipitated out of solution.
showed minimal response owing to poor tissue uptake (data
not shown).
Green, MitoTracker Green, and ER-Tracker Green) within
RAW 264.7 macrophages. On the basis of the overlap between
APNO-5 and each of the trackers, it was clear that APNO-5
broadly localizes throughout the cell (except the nucleus) with
slight preference for the lysosome and mitochondria (Figure
S8).
Next, we determined that the fluorescence quantum yields of
APNO-5 and tAPNO-5 were sufficiently low (Table 1). This is
ideal for PA imaging, since the quantum yield is inversely
proportional to the intensity of the PA signal. Nevertheless, we
conducted selectivity assays using fluorimetry due to its
sensitivity and dynamic range. Treatment of APNO-5 with
excess NO (100 equiv) resulted in a significant 174.4-fold
fluorescence enhancement, whereas other reactive nitrogen
species (RNS) (e.g., nitroxyl and peroxynitrite) did not elicit an
appreciable response (Figure 2d). High selectivity is important
because NO can rapidly convert into other RNS under
biological conditions (besides N O , the hypothesized nitro-
Noninvasive Photoacoustic Imaging of NO in a
Murine Inflammation Model. As mentioned previously,
the noninvasive detection of NO in vivo remains a significant
challenge despite the general availability of NO-specific
fluorescent probes. To evaluate the utility of APNO-5 for the
detection of NO in vivo, we turned to a LPS-induced murine
inflammation model. This system was selected because mice
show similar inflammatory responses to LPS as humans,
2
3
2
8
46
sating species). Importantly, most reported NO triggers react
with N O rather than directly with NO. While this decreases
making it a useful model for inflammation.
Mice were treated with either LPS or a saline vehicle control
via subcutaneous (s.c.) injection into their flank. Following
background scans, the dye was administered in the same
location as the treatment. APNO-5 and tAPNO-5 were
irradiated at 770 and 680 nm, respectively, and the change in
ratiometric signal (PAI680/PAI770) was monitored over time.
Following administration, we observed significant pooling of
the probe at the injection site. The initial broad absorbance
band of APNO-5 in the injection solution resulted in a ratio of
2
3
sensitivity, it is suggested to prevent perturbation of NO signal
3
9
transduction. When APNO-5 was treated with a panel of
oxidants, no oxidation of the aza-BODIPY core nor the N-
methylaniline trigger was observed. Finally, the reactivity of
APNO-5 was assessed against a series of key biological carbonyl
species including formaldehyde (FA), ascorbic acid (AA),
dehydroascorbic acid (DHA), and glyoxal to test for side-
reactivity and false turn-on via Schiff base formation (Figure
∼
1.07 at 1 h. Diffusion of the PA signal shortly thereafter (2 h)
indicated uptake into the surrounding tissue (Figure 3). At this
2b). Even with large stoichiometric excesses of each carbonyl,
no cross-reactivity was observed.
We then determined the chemo- and photostability of
APNO-5 in the interest of supporting in vivo imaging over
extended periods (hours to days). First, the absorption spectra
of a solution of APNO-5 were monitored over the course of 5
days. During this time, no significant decomposition of the
probe was observed. Next, we subjected a solution of APNO-5
to continuous irradiation with an OPO laser at 770 nm for 60
min. Under these conditions, minimal photobleaching was
observed (Figure S6). Interestingly, it was discovered that
tAPNO-5 undergoes photoinduced denitrosylation when
continuously irradiated; however, only moderate signal loss is
observed (<30%) under standard imaging conditions (con-
firmed by UV−vis; data not shown). Moreover, since APNO-5
is released as the product during prolonged irradiation, it is
reasonable to predict minimal decrease in sensitivity.
Figure 3. Representative PA images of APNO-5 responding to
endogenous NO within a murine LPS-induced inflammation model.
Images of APNO-5 (68 μg/kg) in a mouse flank following
subcutaneous LPS or saline treatment. APNO-5 and tAPNO-5 were
selectively imaged at 770 and 680 nm, respectively.
After confirming the requisite selectivity, solubility, and
stability, the probe’s biocompatibility and subcellular local-
ization were assessed. First, RAW 264.7 macrophages were
incubated with APNO-5 (1 and 5 μM) for 3, 6, and 24 h.
Biocompatibility was assessed for both membrane integrity
time, we observed robust conversion of APNO-5 to afford
tAPNO-5 in LPS-treated animals, as determined by an increase
in the ratiometric PA signal to ∼1.2, which remained constant
for the remainder of the experiment. In contrast, the PA ratio
for the saline-treated mice was ∼0.9 and did not change
throughout the experiment. The initial decrease in PA ratio for
the control animal can be attributed to sharpening of APNO-5’s
(trypan blue assay) and cell metabolism (MTT assay). Both
assays indicated that APNO-5 was minimally cytotoxic at a
concentration of 5 μM, with greater than 79% viability observed
after 24 h (Figure S7). Next the localization was determined via
colocalization studies with organelle-specific dyes (LysoTracker
D
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Journal of the American Chemical Society
Article
absorbance profile following uptake and solubilization within
the tissue. This was confirmed by monitoring the absorbance
profile in the absence and presence of rat liver microsomes, a
source of membrane lipids and proteins for association, and the
PA spectrum of APNO-5 in a mouse as a function of time
the reaction with NO is rapid and gives rise to a single product
with predictable chemostability.
12
The best probe in our series, APNO-5, exceeds all six of the
essential criteria for optimal in vivo performance. This
performance was evaluated in a well-established murine LPS-
induced inflammation model: rapid conversion of APNO-5 to
tAPNO-5 was observed exclusively following LPS treatment
and not in the saline-treated control. Importantly, since PA
imaging is noninvasive, it is possible to monitor a single animal
throughout the entire time course of the experiment. This is in
contrast to most other imaging modalities, which provide
information at only a single time point because the animal must
be sacrificed to harvest the tissue for ex vivo imaging.
(
Figure S9). Taken together, this represents a 1.31-fold
ratiometric turn on (PAI680/PAI770) and 1.89-fold signal
enhancement at 680 nm (Figure 4) after 5 h.
In conclusion, this work represents one of very few studies
that have successfully accomplished in vivo imaging of NO and
the first using PA imaging. Compared to previous reports, our
strategy does not require the surgical generation of an intravital
window and extends the detection of NO to significantly
deeper regions of the animal. Owing to the noninvasive nature
of PA imaging and excellent performance of APNO-5, we
anticipate our work will provide a means to study other NO-
mediated biological processes. Current efforts are focused on
applying these photoacoustic probes for NO to study the role
of NO in tumor progression.
Figure 4. Ratiometric imaging (PAI680/PAI770) of endogenously
produced NO within an LPS-induced murine inflammation model
over 5 h. Data presented as mean ± standard deviation (n = 8).
Statistical analysis between LPS and saline treated mice was performed
using 2-way ANOVA (α = 0.05). LPS and saline ratiometric signals
were compared at each time point using Sidak’s multiple comparison
test (α = 0.05). *, p < 0.05; **, p < 0.01.
EXPERIMENTAL DETAILS
■
Materials and Synthetic Procedures. All materials were
purchased from Acros Organic, Airgas, AK Scientific, Alfa Aesar,
Amersco, Cambridge Isotope Laboratories, Cayman Chemicals, Click
Chemistry Tools, Combi-Blocks, Fischer Scientific, Gold Biotechnol-
ogy, Life Technologies, Macron Fine Chemicals, McMaster-Carr,
Oakwood Chemical, Sigma-Aldrich, or Thermo Scientific and used
without further purification. Specific purchasing information can be
found in the Supporting Information. Thin-layer chromatography
To confirm that the signal was a result of APNO-5
nitrosation and not LPS-induced changes in endogenous
chromophores (e.g., oxy-hemoglobin, deoxy-hemoglobin), PA
spectra were acquired following LPS or saline treatment both in
the presence and in the absence of dye. As expected, LPS
induces a small increase in ratiometric signal in the absence of
dye; however, it does not account for the total signal
enhancement (Figure S10). This data, together with our in
vitro results, provides compelling evidence that APNO-5 is
capable of detecting endogenously generated NO in vivo. We
propose that APNO-5 may be effective at even greater depths
with the use of general administration (e.g., intravenous) and/
or alternative instrumentation, since our specific PA tomogra-
pher's imaging window is limited to 2.5 cm.
(TLC) was performed on glass-backed TLC plates precoated with
silica gel containing an UV254 fluorescent indicator (Macherey-
Nagel). Compounds were visualized with a 254/365 nm hand-held UV
lamp (UVP). Flash chromatography was performed using 230−400
mesh silica gel P60 (SiliCycle Inc.). Preparative TLC purification was
performed on 1.00 mm thick TLC plates precoated with silica gel
containing a UV254 fluorescent indicator (Macherey-Nagel). Non-
commercially available anhydrous solvents were dried over 3 Å
molecular sieves activated via heating under a vacuum at 300 °C. All
glassware used in anhydrous reactions was flame-dried under a vacuum
and cooled immediately prior to use. Additional detailed synthetic
procedures can be found in the Supporting Information.
CONCLUSION
Benzo[d]oxazol-2(3H)-one (1). A solution of 2-aminophenol
10.0 g, 92.0 mmol, 1 equiv) and di(1H-imidazol-1-yl)methanone
CDI) (37.0 g, 228.0 mmol, 2.5 equiv) in anhydrous MeCN (250.0
■
(
(
PA imaging is an emerging imaging technology that has the
potential to perform real-time, noninvasive molecular imaging
in deep tissue. In this work, we rationally developed a series of
NO-responsive probes for PA imaging. Since we installed the
most common aerobic NO triggers on a single dye platform, it
was possible to systematically compare OPD-, deamination-,
and N-nitrosation-based sensing mechanisms. It is clear that the
OPD-based trigger is a reasonable approach if triazole
formation can be coupled to a robust readout. This is in
contrast to some reports implicating cross-reactivity with
mL) was stirred at 70 °C for 4 h. When judged to be complete by
TLC, the reaction was concentrated and suspended in EtOAc. Excess
CDI was quenched with 2 M aq HCl, and the organic layer was
washed sequentially with a 2 M solution of aq HCl (3×), sat.
NaHCO , and H O (2×). The organic fraction was dried over Na SO
4
3
2
2
and concentrated to afford the product as an orange solid (10.5 g, 78.0
mmol, 85%), which was used without further purification. R = 0.39
f
1
(3:7 v/v EtOAc/hexanes). H NMR (500 MHz, DMSO-d
) δ 11.56
6
(
bs, 1H), 7.29−7.24 (m, 1H), 7.17−7.11 (m, 1H), 7.10−7.04 (m, 2H).
1
3
47
C NMR (125 MHz, DMSO-d ) δ 154.43, 143.35, 130.40, 123.75,
ascorbic acid and dehydroascorbic acid. This suggests that
the side reaction is probe dependent, as we did not observe any
response to these two carbonyl species, nor with glyoxal. While
our work demonstrates that NO-mediated deamination can be
expanded, it is limited by the potential formation of metastable
intermediates that can complicate interpretation. We found the
N-nitrosation trigger to be the optimal of the three triggers
examined. When the N-alkyl amine is sufficiently nucleophilic,
6
1
21.80, 109.76, 109.48.
6
-Acetylbenzo[d]oxazol-2(3H)-one (2). The product was
48
synthesized according to previously reported protocols. A flame-
dried two-neck round-bottom flask was charged with AlCl (156 g,
3
1
.16 mol, 10 equiv). The solid was heated to 45 °C under a nitrogen
atmosphere and treated with dropwise addition of anhydrous DMF
(26.0 mL). Compound 1 (15.7 g, 116.2 mmol, 1 equiv) was added
portion-wise to the slurry. Acetyl choride (50.0 mL, 697.0 mmol, 6
E
DOI: 10.1021/jacs.7b10783
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
equiv) was added dropwise via addition funnel then heated to 75 °C
for 2 h. The reaction was slowly poured into ice, and the resultant
δ 194.87, 146.29, 144.41, 139.75, 129.11, 127.81, 127.60, 124.86,
124.62, 107.80, 106.78, 79.88, 55.69, 40.82, 40.00, 29.81.
precipitate was filtered and rinsed with H O to afford the product as
(Z)-2-Methoxy-N-methyl-4-(4-phenyl-5-((3-phenyl-5-(4-
(prop-2-yn-1-yloxy)phenyl)-2H-pyrrol-2-ylidene)amino)-1H-
pyrrol-2-yl)aniline (7). A suspension of 6 (0.40 g, 1.2 mmol, 1 equiv)
and 35 (0.80 g, 2.5 mmol, 2 equiv) in n-butanol (25 mL) was heated
2
an orange solid (15.6 g, 88.0 mmol, 76%), which was used without
1
further purification. R = 023 (1:1 v/v EtOAc/hexanes) H NMR (500
f
MHz, DMSO-d ) δ 12.04 (s, 1H), 7.83, (m, 2H), 7.20−7.17 (m, 1H),
6
.56 (s, 3H). 1 C NMR (125 MHz, DMSO-d ) δ 196.30, 154.41,
3
4
to 110 °C to dissolve all solids. NH OAc (1.4 g, 19 mmol, 15 equiv)
2
1
6
was added in one portion and the reaction stirred at the same
temperature for 5 h. The reaction was azeotroped with PhMe and
dried to a blue-green solid. The product was extracted from brine with
43.28, 134.81, 131.21, 125.33, 109.33, 109.03, 26.60.
6
-Cinnamoylbenzo[d]oxazol-2(3H)-one (3). A solution of 2
(
8.0 g, 45.3 mmol, 1 equiv) and benzaldehyde (4.8 g, 45.3 mmol, 1
CH Cl2 (3×). The combined organic fractions were dried over
equiv) in EtOH (100 mL) was treated with dropwise addition of a 10
M solution of aq KOH (13.6 mL, 136.0 mmol, 3 equiv). The reaction
was allowed to stir at room temperature for 4 h. The slurry was
concentrated to an orange solid and recrystallized in EtOH to yield the
2
Na SO and purified twice via silica gel column chromatography (3:7
2
4
v/v EtOAc/hexanes and then 4:1 CH Cl /hexanes) to afford the
2
2
product as a dark green solid (340 mg, 0.60 mmol, 48%). R = 0.50
f
1
(
(
7
3:7 v/v EtOAc/hexanes). H NMR (500 MHz, CDCl ) δ 8.07−8.03
product as a yellow solid (8.3 g, 48.4 mmol, 68%). R = 0.48 (2:1:7 v/v
3
f
1
m, 4H), 7.67−7.63 (m, 2H), 7.52−7.48 (m, 2H), 7.43−7.33 (m, 5H),
acetone/MeOH/hexanes). H NMR (400 MHz, DMSO-d ) δ 12.11
6
.32−7.27 (m, 1H), 7.24 (s, 1H), 6.91 (d, J = 2.0 Hz, 1H), 6.89 (s,
(
(
s, 1H), 8.10 (d, J = 1.4 Hz, 1H), 8.06 (dd, J = 8.2, 1.6 Hz, 1H), 7.98
d, J = 15.6 Hz, 1H), 7.89 (dd, J = 6.7, 2.9 Hz, 2H), 7.73 (d, J = 15.5
1H), 6.54 (d, J = 8.1 Hz, 1H), 4.66 (d, J = 2.5 Hz, 2H), 3.93 (s, 3H),
13
13
Hz, 1H), 7.45 (m, 3H), 7.23 (d, J = 8.1 Hz, 1H). C NMR (100 MHz,
2.90 (s, 3H), 2.56 (t, J = 2.4 Hz, 1H). C NMR (125 MHz, CDCl ) δ
3
DMSO-d ) δ 187.30, 154.49, 143.78, 143.50, 134.98, 134.76, 131.85,
165.10, 157.81, 157.16, 146.83, 146.36, 142.93, 142.52, 141.97, 135.43,
134.75, 133.79, 129.30, 128.78, 128.26, 128.22, 128.12, 128.12, 127.09,
126.44, 125.66, 124.00, 120.73, 119.45, 115.39, 108.94, 108.27, 106.90,
78.46, 75.98, 55.98, 55.30, 29.91.
6
1
30.58, 128.93, 128.91, 125.78, 125.74, 121.77, 109.60, 109.56, 109.54.
tert-Butyl (4-Cinnamoyl-2-hydroxyphenyl) Carbamate (4). A
suspension of 3 (0.9 g, 3.4 mmol, 1 equiv), K CO (1.4 g, 10 mmol, 3
2
3
equiv), and di-tert-butyl dicarbonate (1.5 g, 7 mmol, 2 equiv) in
MeOH (16 mL) was stirred at 45 °C for 3 h. The reaction was diluted
4-(5,5-Difluoro-1,9-diphenyl-7-(4-(prop-2-yn-1-yloxy)-
phenyl)-5H-5l4,6l4-dipyrrolo[1,2-c:2′,1′-f][1,3,5,2]-
triazaborinin-3-yl)-2-methoxy-N-methylaniline (8). A solution of
with sat. NaHCO and extracted with EtOAc (3×). The combined
3
7
(0.077 g, 0.14 mmol, 1 equiv) and N,N-diisopropylethylamine (0.36
organic layers were dried over Na SO , concentrated, and purified via
2
4
mL, 2.1 mmol, 15 equiv) in anhydrous CH Cl (5.5 mL) was cooled
silica gel column chromatography (3:7 v/v EtOAc/hexanes) to afford
2
2
to 0 °C in an ice bath and treated portion-wise with boron trifluoride
diethyl etherate (0.18 mL × 3, 4.3 mmol, 45 equiv) over 24 h. After
each addition, the reaction was allowed to warm to room temperature.
When complete, the reaction was diluted with sat. NaHCO3 and
the product as a yellow solid (0.8 g, 2.3 mmol, 69%). R = 0.44 (3:7 v/
f
1
v EtOAc/hexanes). H NMR (500 MHz, CDCl ) δ 8.51 (bs, 1H),
3
7
7
9
1
1
.93−7.88 (m, 2H), 7.81 (d, J = 15.6 Hz, 1H), 7.67−7.59 (m, 3H),
.56 (d, J = 15.7 Hz, 1H), 7.43−7.39 (m, 3H), 7.30 (s, 1H), 1.55 (s,
H). 13C NMR (125 MHz, CDCl ) δ 189.94, 153.28, 145.71, 145.04,
extracted with CH Cl (3×). The combined organic fractions were
2 2
3
dried over Na SO , concentrated, and purified via silica gel column
34.97, 132.88, 132.26, 130.76, 129.09, 128.70, 122.66, 121.71, 118.18,
16.10, 81.77, 28.42.
tert-Butyl (4-Cinnamoyl-2-methoxyphenyl)(methyl) Carba-
2
4
chromatography (3:7 v/v EtOAc/hexanes) to afford the product as a
dark purple solid (0.027 g, 0.045 mmol, 33%). R = 0.29 (3:7 v/v
f
1
EtOAc/hexanes). H NMR (500 MHz, CDCl ) δ 8.11−8.03 (m, 7H),
mate (5). A 38 mL pressure flask under nitrogen was charged with 4
3
7
(
.96 (s, 1H), 7.78 (dd, J = 8.6, 2.0 Hz, 1H), 7.49−7.39 (m, 6H), 7.36
(
0.6 g, 1.8 mmol, 1 equiv), anhydrous DMF (8 mL), MeI (1.2 g, 8.8
t, J = 7.3 Hz, 1H), 7.07−7.02 (m, 2H), 6.93 (s, 1H), 6.64−6.57 (m,
mmol, 5 equiv), and NaH (60 wt %, 2 g, 5.5 mmol, 3 equiv) at 0 °C.
The reaction was allowed to warm to room temperature and was
stirred for 1 h. The reaction was then heated to 70 °C for 3 h. The
reaction was quenched with water and extracted with EtOAc (3×).
The organic layers were combined, concentrated, and purified via silica
gel column chromatography (3:17 v/v EtOAc/hexanes) to afford the
1
2
1
1
1
7
H), 4.76 (d, J = 2.4 Hz, 2H), 3.94 (s, 3H), 2.97 (s, 3H), 2.57 (q, J =
.6 Hz, 1H). 13C NMR (125 MHz, CDCl ) δ 160.63, 158.78, 152.53,
3
47.06, 146.35, 143.84, 143.69, 143.52, 139.05, 133.54, 132.43, 131.07,
29.58, 129.44, 129.39, 129.08, 128.81, 128.77, 128.61, 128.55, 128.39,
27.42, 126.40, 120.09, 118.38, 116.69, 115.27, 114.76, 110.96, 108.45,
11
8.45, 75.96, 55.97, 55.78, 29.85. B NMR (161 MHz, CDCl ) δ 1.34
product as a yellow solid (5.9 g, 1.6 mmol, 91%). R = 0.24 (1:9 v/v
3
f
(
6
t, J = 32.8 Hz). 19F NMR (471 MHz, CDCl ) δ −131.51 (dd, J =
1
EtOAc/hexanes). H NMR (500 MHz, CDCl ) δ 7.85 (d, J = 15.7 Hz,
3
3
6.0, 32.9 Hz).
1
H), 7.70−7.63 (m, 4H), 7.58 (d, J = 15.6 Hz, 1H), 7.43 (qd, J = 3.8,
APNO-5. Compound 8 (0.046 g, 0.076 mmol, 1 equiv), tris(3-
hydroxypropyltriazolylmethyl)amine (0.007 g, 0.015, 0.2 equiv), 39
1
.7 Hz, 3H), 7.32 (d, J = 8.4 Hz, 1H), 3.94 (s, 3H), 3.21 (s, 3H), 1.45
(
s, 9H). 13C NMR (125 MHz, CDCl ) δ 189.15, 155.05, 154.77,
3
(0.099 g, 0.25 mmol, 3.5 equiv), and copper sulfate pentahydrate
1
1
44.64, 137.50, 136.91, 134.71, 130.45, 128.84, 128.36, 128.32, 121.65,
21.20, 111.14, 79.86, 55.50, 36.65, 28.13.
(0.095g, 0.38 mmol, 5 equiv) were dissolved in degassed THF (3.8
mL), and then, sodium ascorbate (0.023 g, 0.11 mmol, 1.5 equiv) in
degassed water (0.8 mL) was added under nitrogen. The reaction was
allowed to stir at room temperature for 3 h. When the reaction was
1
-(3-Methoxy-4-(methylamino)phenyl)-4-nitro-3-phenylbu-
tan-1-one (6). A solution of 5 (0.53 g, 1.4 mmol, 1 equiv) and
nitromethane (0.87 g, 14 mmol, 10 equiv) in EtOH (3 mL) was
treated with a 1 M solution of aq KOH (0.29 mL, 0.2 equiv). The
reaction was stirred at 60 °C for 7 h. The reaction was diluted with
brine, extracted with EtOAc (3×), and concentrated. The crude
intermediate was dissolved in CH Cl (8 mL) and cooled to 0 °C in an
complete, the product was extracted with 3:7 v/v MeOH/CH
(3×) from sat. NaI. The combined organic layers were dried over
Na SO and concentrated under reduced pressure. The crude residue
was purified via preparatory TLC (1:9 v/v H O/MeCN). The product
was recovered in 1:9 v/v MeOH/CH Cl with sonication, filtered, and
Cl
2
2
2
4
2
2
2
ice bath. Trifluoroacetic acid (2 mL) was added dropwise, and the
reaction was allowed to stir at room temperature for 3 h. The reaction
was quenched with sat. NaHCO and extracted with CH Cl (3×).
The combined organic fractions were dried over Na SO , concen-
trated, and purified via silica gel column chromatography (3:7 v/v
EtOAc/hexanes) to afford the product as a brown solid (0.40 g, 1.2
mmol, 86%). R = 0.47 (3:7 v/v EtOAc/hexanes). H NMR (500
MHz, CDCl ) δ 7.45 (dd, J = 8.3, 1.9 Hz, 1H), 7.28 (d, J = 1.9 Hz,
H), 7.27−7.16 (m, 6H), 6.40 (d, J = 8.3 Hz, 1H), 4.82 (s, 1H), 4.78
dd, J = 12.5, 6.3 Hz, 1H), 4.60 (dd, J = 12.5, 8.5 Hz, 1H), 4.13 (tt, J =
.3, 6.1 Hz, 1H), 3.79 (s, 3H), 3.30 (dd, J = 16.9, 6.2 Hz, 1H), 3.23
dd, J = 16.9, 8.0 Hz, 1H), 2.85 (s, 3H). 13C NMR (125 MHz, CDCl3)
2
2
dried to afford the product as a red solid (0.077 g, 0.076 mmol, 100%).
1
R = 0.55 (1:9 v/v H O/MeCN). H NMR (500 MHz, DMSO-d ) δ
3
2
2
f
2
6
8.26 (s, 1H), 8.25−8.22 (m, 2H), 8.16 (dd, J = 8.8, 2.0 Hz, 1H), 8.14−
8.11 (m, 2H), 8.09 (d, J = 9.0 Hz, 2H), 8.06 (s, 1H), 7.99−7.96 (m,
1H), 7.56 (dd, J = 8.2, 6.6 Hz, 2H), 7.49 (td, J = 7.5, 5.7 Hz, 3H),
7.41−7.35 (m, 1H), 7.27 (s, 1H), 7.23−7.16 (m, 3H), 6.75 (d, J = 8.9
Hz, 1H), 5.27 (s, 2H), 4.57 (t, J = 5.2 Hz, 2H), 3.95 (s, 3H), 3.84 (t, J
= 5.2 Hz, 2H), 3.82−3.77 (m, 2H), 3.58−3.51 (m, 5H), 3.50−3.48 (m,
5H), 3.06 (s, 9H), 2.94 (d, J = 5.2 Hz, 3H). 13C NMR (125 MHz,
2
4
1
f
3
1
(
8
(
DMSO-d ) δ 159.42, 149.00, 147.30, 146.52, 145.96, 142.83, 142.61,
6
141.90, 135.72, 133.62, 132.04, 130.88, 130.19, 130.08, 129.55, 129.06,
F
DOI: 10.1021/jacs.7b10783
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
1
7
5
1
28.99, 128.87, 128.53, 125.67, 125.49, 116.42, 116.35, 115.10, 109.53,
Angeli’s salt in degassed 10 mM potassium hydroxide solution. NO
was generated in situ from a solution of MAHMA-NONOate in
degassed 10 mM potassium hydroxide. Hydroxyl radical was prepared
using Mohr’s salt in degassed 1 M aqueous hydrochloric acid and
hydrogen peroxide. Fe(II), nitroxyl, and hydroxyl radical selectivity
assays were performed under nitrogen to prevent oxidation.
Peroxynitrite was prepared according to previously reported
literature. All other analytes were prepared by dilution from
commercially available sources.
PA Spectra. Tissue phantoms were prepared by mixing agarose (4
0.24, 70.04, 70.00, 69.88, 69.80, 69.14, 64.89, 64.49, 61.71, 56.10,
1
1
3.60, 53.57, 53.54, 49.93, 29.81. B NMR (161 MHz, DMSO-d ) δ
6
.33 (t, J = 34.5 Hz). 19F NMR (471 MHz, DMSO-d ) δ −129.98 (dd,
6
+
J = 67.0, 29.9 Hz). HR-MS calcd [M] 902.4258, found 902.4345.
1
13
11
19
Instruments. H, C, B, and F NMR spectra were acquired on
Varian 400, Varian 500, or Carver B500 spectrometers. The following
abbreviations were used to describe coupling constants: singlet (s),
doublet (d), triplet (t), quartet (q), quintet (quint), multiplet (m), and
broad singlet (bs). Spectra were visualized and analyzed using
MestReNova (version 10.0). High-resolution mass spectra were
acquired with a Waters Q-TOF Ultima ESI mass spectrometer and a
Waters Synapt G2-Si ESI/LC-MS mass spectrometer. Ultraviolet−
visible (UV−vis) measurements or spectra were recorded on a Cary 60
spectrometer or SpectraMax M2 plate reader (Molecular Devices).
Fluorescence spectra were acquired on a QuantaMaster-400 scanning
spectrofluorometer with micro fluorescence quartz cuvettes (Science
Outlet). Refractive indices were measured using a RHB-32ATC Brix
Refractometer. Cellular imaging was performed on either a EVOS FL
epifluorescence microscope or a Zeiss LSM 700 confocal microscope.
Photoacoustic imaging was performed using the Endra Nexus 128
photoacoustic tomography system (Ann Arbor, MI, USA), and the
data were analyzed using either OsiriX (version 8.0) or Horos (version
5
2
g) in a solution of 2% reduced fat milk (2 mL) and deionized H O (78
2
mL). The mixture was heated in a microwave oven until a viscous gel
was produced (30 s); the gel was then mixed and reheated for an
additional 15 s to ensure homogeneity. The gel was transferred to a
custom Teflon mold made to fit within the Endra bowl system with
hypodermic steel tubing inserted to prepare channels for placing FEP
tubes (0.06 in. diameter) containing the samples. After cooling the gel
for a minimum of 1 h at 4 °C, the phantom was removed from the
mold. When imaging, sample solutions (200 μL) were pipetted into
FEP tubing (0.08 in. diameter, cut to 10 cm long). The tubing was
then inserted into the phantom and sealed by folding over the ends
and securing with additional tubing (0.12 in. diameter, cut to 5 mm
long). Images were acquired using the Step and Shoot mode with 120
angles and 10 pulses per angle. APNO (10 μM) was dissolved in an
ethanolic 20 mM potassium phosphate buffer (pH 7.4, 50% v/v).
tAPNO was generated by reacting APNO with NO (2.5 mM
MAHMA NONOate, 500 equiv NO) for greater than 1 h at room
temperature (complete conversion was confirmed by UV−vis). PA
spectra were measured in the same solvent at 10 nm intervals, and
signal was integrated over the total range of signal. The mean signal for
each FEP tube was averaged for each sample. Fresh samples were
prepared for each tAPNO-5 wavelength to limit photoinduced
cleavage of the N-nitroso.
3
.0) imaging software. All other data analysis was performed using
GraphPad Prism (version 6.0). A Mettler Toledo SevenCompact pH
meter was used for pH measurements.
Photophysical Characterization. Extinction coefficients and
fluorescence quantum yields were acquired in experimental triplicates.
tAPNO was generated by reacting APNO with NO (500 equiv) for
greater than 1 h at room temperature (complete conversion was
confirmed by UV−vis). Extinction coefficients were acquired by
titrating or diluting compound into chloroform or ethanolic 20 mM
potassium phosphate buffer (pH 7.4, 50% v/v) within the linear range
(
typically absorbance values 0.05−1.50). Fluorescence quantum yields
Photostability Studies. APNO-5 or tAPNO-5 solutions (20 μM)
were prepared in potassium phosphate saline with CrEL (0.1% v/v).
tAPNO-5 was generated according to the same procedure as
previously mentioned. PA images were acquired at the wavelength
corresponding to the maximal ratiometric PA response using 120
angles and 10 pulses per angle for a total of 60 min. The mean signal
was acquired, the average PAI was obtained for the two tubes, and
relative signals were calculated according to the initial PA signal.
PA Imaging of the LPS-Induced Murine Inflammation
Model. All animal experiments were performed with the approval of
the Institutional Animal Care and Use Committee of the University of
Illinois at Urbana−Champaign, following the principles outlined by
the American Physiological Society on research animal use. Six to eight
week old BALB/c mice were obtained from Jackson Laboratory.
Lipopolysaccharide was dissolved in sterile saline at a concentration of
were obtained using a modified method for relative fluorescence
49
quantum yield. Briefly, samples were prepared and the absorbance
and fluorescence were monitored (n = 3). Dye was sequentially
titrated or diluted, and the absorbance was kept low (<0.1) to prevent
secondary absorbance events. Quantum yields were calculated relative
to dimethoxy aza-BODIPY (ϕ = 0.36, chloroform) or indocyananine
green (ϕ = 0.106, DMSO). The refractive index of ethanolic 20 mM
potassium phosphate buffer (pH 7.4, 50% v/v) was measured to be
50
51
1
.3573.
Reaction of APNO with NO. APNO was dissolved in ethanolic 20
mM potassium phosphate buffer (pH 7.4, 50% v/v). UV−vis and
fluorescence measurements were acquired from 400 to 950 nm. The
reactions were initiated by the addition of MAHMA-NONOate
(
prepared in degassed 10 mM aq KOH) at room temperature. The
1
mg/mL. Hair was removed from the flanks using depilatory cream,
reaction with NO(g) was confirmed by treating APNO (20 μM) in
ethanolic water (50% v/v) with NO(g) until a color change was
observed. The reaction products for APNO-1, APNO-2, APNO-4, and
APNO-5 were confirmed by ESI/LC-MS (Figures S11−S14).
and then, mice were challenged subcutaneously with either 4 mg/kg of
LPS or an equivolumetric amount of saline in the flank. After 3.5 h,
mice were anaesthetized using isoflurane, and background PA signals
were acquired at 680 and 770 nm. After an additional 0.5 h, a 17 μM
solution of APNO-5 in sterile saline containing 0.5% DMF was
subcutaneously injected into the same flank at 68 μg/kg (80 μL for 20
g mouse). PA signals were monitored every hour for 5 h in triplicate at
Selectivity Studies. Unless otherwise noted, APNO was
preincubated at 37 °C in an ethanolic 20 mM potassium phosphate
buffer (pH 7.4, 50% v/v) for at least 15 min. The initial fluorescence
was measured followed by the addition of 100 equiv of various reactive
metals, oxygen, nitrogen, and carbonyl species. The reaction was
sealed and incubated at 37 °C for 1 h. Final measurements were
recorded, and relative turn-on was determined by the sum of intensity
over the total range of emission. Due to the rapid oxidation of Fe(II)
in phosphate buffer, Fe(II) selectivity studies were performed in
ethanolic 20 mM HEPES buffer (pH 7.4, 50% v/v). All metal solutions
were prepared from their chloride salt in water with the exception of
Fe(II) which was prepared from FeSO ·(H O) . Formaldehyde
solutions were heated to 100 °C before use to ensure depolymerizing
of any paraformaldehyde. Dehydroascorbic acid solutions were
prepared by heating a solution at 65 °C in water. Perchlorate, nitrite,
and nitrate solutions were prepared for their corresponding sodium
salts in water. Superoxide anion was added as a solution of potassium
superoxide in DMSO. Nitroxyl was generated in situ from a solution of
6
80 and 770 nm. Average PA intensities were acquired over the total
3
volume of interest (1 cm ), and the triplicates were averaged. PA
spectra were acquired after 5 h with excitations every 10 nm under the
same conditions as previously described, either with or without the
administration of APNO-5.
ASSOCIATED CONTENT
Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/jacs.7b10783.
■
4
2
7
*
S
Experimental details, including detailed synthetic proce-
dures, spectral data, and supplemental in vitro and in vivo
characterization (PDF)
G
DOI: 10.1021/jacs.7b10783
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX

Journal of the American Chemical Society
Article
(
22) Kashiwagi, S.; Tsukada, K.; Xu, L.; Miyazaki, J.; Kozin, S. V.;
AUTHOR INFORMATION
Corresponding Author
jeffchan@illinois.edu
■
*
Tyrrell, J. A.; Sessa, W. C.; Gerweck, L. E.; Jain, R. K.; Fukumura, D.
Nat. Med. 2008, 14, 255.
(
23) Li, H.; Zhang, D.; Gao, M.; Huang, L.; Tang, L.; Li, Z.; Chen,
X.; Zhang, X. Chem. Sci. 2017, 8, 2199.
24) Huo, Y.; Miao, J.; Han, L.; Li, Y.; Li, Z.; Shi, Y.; Guo, W. Chem.
Sci. 2017, 8, 6857.
25) Sasaki, E.; Kojima, H.; Nishimatsu, H.; Urano, Y.; Kikuchi, K.;
ORCID
(
Christopher J. Reinhardt: 0000-0001-9992-1253
Jefferson Chan: 0000-0003-4139-4379
Notes
(
Hirata, Y.; Nagano, T. J. Am. Chem. Soc. 2005, 127, 3684.
(26) Dong, X.; Heo, C. H.; Chen, S.; Kim, H. M.; Liu, Z. Anal. Chem.
The authors declare no competing financial interest.
2
014, 86, 308.
(27) Mao, Z.; Feng, W.; Li, Z.; Zeng, L.; Lv, W.; Liu, Z. Chem. Sci.
016, 7, 5230.
28) Mao, Z.; Jiang, H.; Li, Z.; Zhong, C.; Zhang, W.; Liu, Z. Chem.
Sci. 2017, 8, 4533.
29) Mao, Z.; Jiang, H.; Song, X.; Hu, W.; Liu, Z. Anal. Chem. 2017,
9, 9620.
30) Wang, L. V.; Hu, S. Science 2012, 335, 1458.
(31) Li, H.; Zhang, P.; Smaga, L. P.; Hoffman, R. A.; Chan, J. J. Am.
ACKNOWLEDGMENTS
■
2
(
This work was supported by the Chemistry-Biology Interface
Training Grant (T32 GM070421 to C.J.R. and E.Y.Z.), the
National Science Foundation Graduate Research Fellowship
Program (NGE-1144245 to E.Y.Z.), the Springborn Fellowship
(
8
(
(
to E.Y.Z.), the Alfred P. Sloan fellowship (FG-2017-8964 to
J.C.), and the Neurotechnology for Memory and Cognition
Award (to M.D.J.). Major funding for the 500 MHz Bruker
CryoProbe was provided by the Roy J. Carver Charitable Trust
Chem. Soc. 2015, 137, 15628.
(32) Mishra, A.; Jiang, Y.; Roberts, S.; Ntziachristos, V.; Westmeyer,
G. G. Anal. Chem. 2016, 88, 10785.
(
Muscatine, Iowa; Grant No. 15-4521) to the School of
(33) Roberts, S.; Seeger, M.; Jiang, Y.; Mishra, A.; Sigmund, F.; Stelzl,
Chemical Sciences NMR Lab. The Q-Tof Ultima mass
spectrometer was purchased in part with a grant from the
National Science Foundation, Division of Biological Infra-
structure (DBI-0100085). We acknowledge Professor Elvira de
Mejia (Food Science and Human Nutrition, UIUC) for
providing RAW 264.7 macrophage cells. We also acknowledge
the Core Facilities at the Carl R. Woese Institute for Genomic
Biology for access to the Zeiss LSM 700 Confocal Microscope
and corresponding software.
A.; Lauri, A.; Symvoulidis, P.; Rolbieski, H.; Preller, M.; Dean Ben, X.
́
L.; Razansky, D.; Orschmann, T.; Desbordes, S.; Vetschera, P.; Bach,
T.; Ntziachristos, V.; Westmeyer, G. G. J. Am. Chem. Soc. 2017,
DOI: 10.1021/jacs.7b03064.
(34) Knox, H. J.; Hedhli, J.; Kim, T. W.; Khalili, K.; Dobrucki, L. W.;
Chan, J. Nat. Commun. 2017, 8, 1794.
(35) Jiang, Y.; Upputuri, P. K.; Xie, C.; Lyu, Y.; Zhang, L.; Xiong, Q.;
Pramanik, M.; Pu, K. Nano Lett. 2017, 17, 4964.
(36) Kelm, M. Biochim. Biophys. Acta, Bioenerg. 1999, 1411, 273.
(
37) Zhuang, Y.; Xu, Q.; Huang, F.; Gao, P.; Zhao, Z.; Lou, X.; Xia,
F. ACS Sens. 2016, 1, 572.
38) Reinhardt, C. J.; Chan, J. Biochemistry 2017, DOI: 10.1021/
acs.biochem.7b00888.
39) Kojima, H.; Nakatsubo, N.; Kikuchi, K.; Kawahara, S.; Kirino,
Y.; Nagoshi, H.; Hirata, Y.; Nagano, T. Anal. Chem. 1998, 70, 2446.
40) Shiue, T. W.; Chen, Y. H.; Wu, C. M.; Singh, G.; Chen, H. Y.;
Hung, C. H.; Liaw, W. F.; Wang, Y. M. Inorg. Chem. 2012, 51, 5400.
41) Beltran, A.; Burguete, M. I.; Abanades, D. R.; Perez-Sala, D.;
Luis, S. V.; Galindo, F. Chem. Commun. 2014, 50, 3579.
42) Miao, J.; Huo, Y.; Lv, X.; Li, Z.; Cao, H.; Shi, H.; Shi, Y.; Guo,
W. Biomaterials 2016, 78, 11.
43) Grossi, M.; Palma, A.; McDonnell, S. O.; Hall, M. J.; Rai, D. K.;
Muldoon, J.; Oshea, D. F. J. Org. Chem. 2012, 77, 9304.
44) Chen, Z.; Liu, Y.; Wagner, W.; Stepanenko, V.; Ren, X.; Ogi, S.;
Wurthner, F. Angew. Chem., Int. Ed. 2017, 56, 5729.
45) Tetko, I. V.; Gasteiger, J.; Todeschini, R.; Mauri, A.;
REFERENCES
■
(
(
1) Zhao, Y.; Vanhoutte, P. M.; Leung, S. W. S. J. Pharmacol. Sci.
015, 129, 83.
2
(
(
2
(
5
(
1
(
(
2) Yun, H. Y.; Dawson, V. L.; Dawson, T. M. Mol. Psychiatry 1997,
, 300.
3) Fukumura, D.; Kashiwagi, S.; Jain, R. K. Nat. Rev. Cancer 2006, 6,
21.
4) MacMicking, J.; Xie, Q. W.; Nathan, C. Annu. Rev. Immunol.
997, 15, 323.
5) Bogdan, C. Nat. Immunol. 2001, 2, 907.
6) Nathan, C.; Xie, Q. Cell 1994, 78, 915.
7) Dungel, P.; Penzenstadler, C.; Ashmwe, M.; Dumitrescu, S.;
Stoegerer, T.; Redl, H.; Bahrami, S.; Kozlov, A. V. Sci. Rep. 2017, 7,
2092.
8) Zweier, J. L.; Wang, P.; Samouilov, A.; Kuppusamy, P. Nat. Med.
995, 1, 804.
9) Zhou, L.; Zhu, D.-Y. Nitric Oxide 2009, 20, 223.
10) Vincent, S. R. Prog. Neurobiol. 2010, 90, 246.
(
(
́
́
́
(
(
(
(
1
(
1
(
(
(
̈
(
Livingstone, D.; Ertl, P.; Palyulin, V. A.; Radchenko, E. V.; Zefirov,
N. S.; Makarenko, A. S.; Tanchuk, V. Y.; Prokopenko, V. V. J. Comput.-
Aided Mol. Des. 2005, 19, 453.
11) Floyd, R. A.; Kotake, Y.; Towner, R. A.; Guo, W.-X.; Nakae, D.;
(
46) Copeland, S.; Warren, H. S.; Lowry, S. F.; Calvano, S. E.;
Remick, D. Clin. Diagn. Lab. Immunol. 2005, 12, 60.
47) Zhang, X.; Kim, W. S.; Hatcher, N.; Potgieter, K.; Moroz, L. L.;
Gillette, R.; Sweedler, J. V. J. Biol. Chem. 2002, 277, 48472.
48) Liacha, M.; Yous, S.; Poupaert, J. H.; Depreux, P.; Aichaoui, H.
Monatsh. Chem. 1999, 130, 1393.
49) Wurth, C.; Grabolle, M.; Pauli, J.; Spieles, M.; Resch-Genger, U.
Nat. Protoc. 2013, 8, 1535.
Konishi, Y. J. Toxicol. Pathol. 2007, 20, 77.
12) Wang, P. G.; Xian, M.; Tang, X.; Wu, X.; Wen, Z.; Cai, T.;
Janczuk, A. J. Chem. Rev. 2002, 102, 1091.
13) Tsikas, D. J. Chromatogr. B: Anal. Technol. Biomed. Life Sci. 2007,
51, 51.
14) Komarov, A.; Mattson, D.; Jones, M. M.; Singh, P. K.; Lai, C. S.
Biochem. Biophys. Res. Commun. 1993, 195, 1191.
15) Sharma, R.; Seo, J.-W.; Kwon, S. J. Nanomater. 2014, 2014,
23646.
(
(
(
8
(
(
(
̈
(
5
(
(
(50) Zhao, W.; Carreira, E. M. Angew. Chem., Int. Ed. 2005, 44, 1677.
(51) Brouwer, A. M. Pure Appl. Chem. 2011, 83, 2213.
(52) Uppu, R. M. Anal. Biochem. 2006, 354, 165.
16) Hong, H.; Sun, J.; Cai, W. Free Radical Biol. Med. 2009, 47, 684.
17) Shim, J. H.; Lee, Y. Anal. Chem. 2009, 81, 8571.
(
18) Jo, A.; Do, H.; Jhon, G. J.; Suh, M.; Lee, Y. Anal. Chem. 2011,
8
(
3, 8314.
19) Li, Y.; Hu, K.; Yu, Y.; Rotenberg, S. A.; Amatore, C.; Mirkin, M.
V. J. Am. Chem. Soc. 2017, 139, 13055.
(
20) Li, H.; Wan, A. Analyst (Cambridge, U. K.) 2015, 140, 7129.
(
21) Nagano, T.; Yoshimura, T. Chem. Rev. 2002, 102, 1235.
H
DOI: 10.1021/jacs.7b10783
J. Am. Chem. Soc. XXXX, XXX, XXX−XXX