Near-Infrared Photoactivatable Nitric Oxide Donors with Integrated Photoacoustic Monitoring

Article abstract of DOI:10.1021/jacs.8b05514

Photoacoustic (PA) tomography is a noninvasive technology that utilizes near-infrared (NIR) excitation and ultrasonic detection to image biological tissue at centimeter depths. While several activatable small-molecule PA sensors have been developed for various analytes, the use of PA molecules for deep-tissue analyte delivery and monitoring remains an underexplored area of research. Herein, we describe the synthesis, characterization, and in vivo validation of photoNOD-1 and photoNOD-2, the first organic, NIR-photocontrolled nitric oxide (NO) donors that incorporate a PA readout of analyte release. These molecules consist of an aza-BODIPY dye appended with an aryl N-nitrosamine NO-donating moiety. The photoNODs exhibit chemostability to various biological stimuli, including redox-active metals and CYP450 enzymes, and demonstrate negligible cytotoxicity in the absence of irradiation. Upon single-photon NIR irradiation, photoNOD-1 and photoNOD-2 release NO as well as rNOD-1 or rNOD-2, PA-active products that enable ratiometric monitoring of NO release. Our in vitro studies show that, upon irradiation, photoNOD-1 and photoNOD-2 exhibit 46.6-fold and 21.5-fold ratiometric turn-ons, respectively. Moreover, unlike existing NIR NO donors, the photoNODs do not require encapsulation or multiphoton activation for use in live animals. In this study, we use PA tomography to monitor the local, irradiation-dependent release of NO from photoNOD-1 and photoNOD-2 in mice after subcutaneous treatment. In addition, we use a murine model for breast cancer to show that photoNOD-1 can selectively affect tumor growth rates in the presence of NIR light stimulation following systemic administration.

Full text of DOI:10.1021/jacs.8b05514

Article
pubs.acs.org/JACS
Cite This: J. Am. Chem. Soc. XXXX, XXX, XXX−XXX
Near-Infrared Photoactivatable Nitric Oxide Donors with Integrated
Photoacoustic Monitoring
Effie Y. Zhou,†,§ Hailey J. Knox,†,§ Christopher J. Reinhardt,† Gina Partipilo,† Mark J. Nilges,‡
and Jefferson Chan*,†
‡
†Department of Chemistry and Beckman Institute for Advanced Science and Technology, and Illinois EPR Research Center,
University of Illinois at Urbana−Champaign, Urbana, Illinois 61801, United States
S
* Supporting Information
ABSTRACT: Photoacoustic (PA) tomography is a non-
invasive technology that utilizes near-infrared (NIR) ex-
citation and ultrasonic detection to image biological tissue at
centimeter depths. While several activatable small-molecule
PA sensors have been developed for various analytes, the use
of PA molecules for deep-tissue analyte delivery and
monitoring remains an underexplored area of research.
Herein, we describe the synthesis, characterization, and in
vivo validation of photoNOD-1 and photoNOD-2, the first
organic, NIR-photocontrolled nitric oxide (NO) donors that
incorporate a PA readout of analyte release. These molecules consist of an aza-BODIPY dye appended with an aryl N-
nitrosamine NO-donating moiety. The photoNODs exhibit chemostability to various biological stimuli, including redox-active
metals and CYP450 enzymes, and demonstrate negligible cytotoxicity in the absence of irradiation. Upon single-photon NIR
irradiation, photoNOD-1 and photoNOD-2 release NO as well as rNOD-1 or rNOD-2, PA-active products that enable
ratiometric monitoring of NO release. Our in vitro studies show that, upon irradiation, photoNOD-1 and photoNOD-2 exhibit
46.6-fold and 21.5-fold ratiometric turn-ons, respectively. Moreover, unlike existing NIR NO donors, the photoNODs do not
require encapsulation or multiphoton activation for use in live animals. In this study, we use PA tomography to monitor the
local, irradiation-dependent release of NO from photoNOD-1 and photoNOD-2 in mice after subcutaneous treatment. In
addition, we use a murine model for breast cancer to show that photoNOD-1 can selectively affect tumor growth rates in the
presence of NIR light stimulation following systemic administration.
molecules (NORMs) that can be activated by NIR light and
generate a PA readout for real-time monitoring of delivery.12
Light-mediated analyte release is uniquely powerful because it
enables spatiotemporally controlled stimulation of a biological
response. In particular, light in the NIR range is useful as an in
vivo bioorthogonal trigger due to its low phototoxicity and
enhanced tissue penetration relative to higher energy wave-
lengths.14 The utility of NIR activation strategies has been
demonstrated by a variety of recent studies in protein15 and
drug delivery,16−22 photodynamic therapy,17,21,23 carbon
monoxide delivery,24,25 antibody−drug conjugate activation,26
gene expression,27−29 and metal ion delivery.30
NO is an endogenously produced gasotransmitter that is
important for vascular tone and neuronal signaling.31 However,
NO also has key roles in various other conditions, including
inflammation,32,33 infection,34 reperfusion injury,35 and
cancer.36,37 The role of NO is often studied in small-animal
models by knocking out endogenous NO synthase (NOS)
enzymes38,39 or by systemic dosing of NOS inhibitors39 or NO
donors.35,39,40 However, these strategies affect NO generation
INTRODUCTION
■
Photoacoustic (PA) tomography is a noninvasive technology
that combines the advantages of tissue-penetrant near-infrared
(NIR) excitation with ultrasonic detection to achieve high-
resolution imaging at depths of up to 10 cm in biological
tissues.1,2 These properties make PA imaging a powerful tool
for live animal imaging relative to purely optical methods such
as fluorescence imaging. While optical strategies have excellent
utility for studies in cell culture and shallow tissues (≤1.5
mm), they suffer from poor resolution at greater depths due to
the scattering of emitted light. Moreover, PA tomography has
great translational potential, as it has already been employed
for noninvasive, label-free studies of human subjects with
breast cancer,3 thyroid cancer,4 inflammatory arthritis,5 and
scleroderma.6 Several recent developments in small-molecule
activatable PA sensors, or acoustogenic probes, have enabled
the detection of metal ions,7−9 hypoxia,10,11 and nitric oxide
(NO)12,13 using PA readouts. However, the photochemical
delivery of bioactive analytes (like NO) in vivo with PA
monitoring is an underexplored area of research with great
potential for the controlled modulation of biological processes.
Our recent progress toward in vivo detection of NO with PA
tomography inspired our interest in developing NO-releasing
Received: May 25, 2018
© XXXX American Chemical Society
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in a nonspecific manner and can alter normal NO signaling,
potentially leading to incorrect conclusions. This has resulted
in a growing interest in NORMs that can be selectively
activated in local regions or in response to specific stimuli to
enable controlled, local modulation of NO levels without
eliciting a global response.
attenuates tumor growth in a murine model of breast
carcinoma without affecting distal tissues.
RESULTS AND DISCUSSION
■
Design and Synthesis. We hypothesized that the aza-
BODIPY dye platform would be an ideal antenna for
harvesting NIR light due to its large extinction coefficient
(>104 M−1 cm−1) at NIR wavelengths, which could provide
sufficient energy to mediate N-nitroso bond cleavage.54,56,66
This property also facilitates generation of strong PA signals
upon excitation.2 Moreover, the photophysical properties of
aza-BODIPYs are known to be modulated by changes in
chemical substitution, making this an ideal scaffold for
ratiometric probe generation.10−12,67 photoNOD-1 and its
water-soluble congener photoNOD-2 were designed with a
photolabile alkylated N-nitroso moiety, which releases NO and
an aza-BODIPY with a free aryl amine (rNOD-1 or rNOD-2)
upon irradiation (Scheme 1). We hypothesized that the
conversion of the electron-withdrawing N-nitroso bond to a
more electron-rich N−H bond would cause a bathochromic
shift in the absorbance maximum. Irradiation at both
wavelengths would then generate two corresponding PA
signals, enabling the simultaneous identification of photoNOD
and rNOD.10−12,67
To date, several classes of activatable NORMs have been
developed with various release mechanisms and kinetics. NO
release from these molecules can be triggered by pH, thermal
activation, enzymatic activity, thiol-based transnitrosylation, or
irradiation with light.41 Photoactivatable NO-releasing mole-
cules (photoNORMs) that incorporate iron, manganese, or
ruthenium-nitrosyl moieties have been shown to release NO
upon irradiation at wavelengths ranging from UV to NIR.
However, these metal-nitrosyl donors commonly require
encapsulation in nanodelivery systems to mitigate off-target
NO release and toxicity.42,43 Select examples of metal-nitrosyl
complexes have been used in isolated tissues without
encapsulation, but their toxicity has yet to be evaluated.44−47
On the other hand, organic photoNORMs that typically offer
greater stability in biological samples have also been developed.
The first examples required the use of damaging UV irradiation
to release NO from caged diazeniumdiolates,48 aryl N-
nitrosamines,49 and nitrobenzenes.50,51 Molecules stimulated
by visible52−56 and two-photon NIR light57,58 were soon
realized to improve tissue penetration and biocompatibility.
It was not until recently that visible light59,60 and two-
photon NIR-activated61,62 photoNORMs were developed to
incorporate a fluorescent readout following NO delivery. This
enables calibration of the release based on the fluorescence
signal, without the need for an additional NO detection
method (e.g., NO-specific fluorescent dye or electrode). These
have excellent potential as tools to study NO biology in cell
culture due to their ability to elicit NO release in precise
femtoliter volumes.63 However, two-photon methods are
limited to shallow tissue depths (1.5 mm),64 and studies in
cell culture do not accurately mimic the complex, three-
dimensional network of cell types and biochemical signals
found in vivo. This can generate discrepancies between cellular
and animal studies. Therefore, it is critical to develop
photoNORMs that can be used in native, deep-tissue
contexts.65
photoNOD-1 and photoNOD-2 were synthesized from 4′-
aminoacetophenone (Scheme 2). Carbamate protection
yielded 1, which was alkylated with methyl iodide to afford
the N-methylaniline trigger precursor. Next, this intermediate
was subjected to Claisen−Schmidt condensation conditions
with benzaldehyde, followed by Michael addition of nitro-
methane to yield 2a. Dimerization with 3a in the presence of
ammonium acetate yielded aza-dipyrromethene 4a. Alterna-
tively, deprotection of 2a with trifluoroacetic acid to afford 2b
followed by dimerization with 3b12 yielded compound 4b.
Boron chelation in the presence of triethylamine and boron
trifluoride diethyl etherate yielded rNOD-1 and aza-BODIPY
5. Copper-catalyzed click chemistry was utilized to install the
tetramethylammonium PEG solubilizing group 612 onto aza-
BODIPY 5, forming rNOD-2. Nitrosation of rNOD-1 and
rNOD-2 with sodium nitrite in acetic acid yielded photoNOD-
1 and photoNOD-2, respectively.
In Vitro Characterization of photoNODs. Because the
PA signal is proportional to molar absorptivity, the relative PA
intensity of a molecule can be conveniently approximated by
its absorbance spectrum at any given wavelength.68 Absorb-
ance spectra were acquired for each compound in chloroform
and aqueous buffer with additives to enable solubilization
(50% ethanol for photoNOD-1, Figure 1a; 0.1% Cremophor
EL (CrEL) for photoNOD-2, Figure 1b; full spectra may be
found in Figure S1). As hypothesized, the photoNODs exhibit
absorbance maxima near 680 nm (λblue), while the rNODs are
red-shifted to wavelengths greater than 730 nm (λred). All four
species maintain absorptions in the first biological imaging
window with extinction coefficients in the expected range
(Tables 1 and S1). Importantly, these absorbance maxima lie
within the wavelength range of commercial PA tomographers
(typically 680−950 nm).11
Herein, we describe the synthesis and validation of NIR
photoactivatable NO donors (photoNOD-1 and photoNOD-
2) whose irradiation-based NO release can be monitored with
PA tomography (Scheme 1). We characterize the NO release
from these molecules and validate their chemostability and
biocompatibility. Moreover, we illustrate the applicability of
these first-in-class donors for NO release in vivo and
demonstrate that selective irradiation of photoNOD-1
Scheme 1. Irradiation of photoNOD Generates NO with
Concomitant Release of rNOD, Which Can Be Monitored
by Ratiometric PA Imaging
To assess the ability of NIR irradiation to penetrate through
tissue and activate the photoNODs, a solution of each
compound was placed within a tissue-mimicking phantom
(depth of ∼1 cm) that can scatter incident light to evenly
illuminate the sample. These solutions were irradiated for up to
40 min using a PA tomographer, and the change in ratiometric
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Scheme 2. Synthesis of rNOD-1, rNOD-2, photoNOD-1, and photoNOD-2
a
Table 1. λred and λblue (in nm) in Various Solvent Systems
species
λblue/λred (a)
λblue/λred (b)
photoNOD/rNOD-1
photoNOD/rNOD-2
681/733
678/745
677/746
691/762
a
Chloroform (a) and aqueous buffer (b): photoNOD-1: 50% EtOH/
HEPES, pH 7.4; photoNOD-2: 0.1% CrEL/HEPES, pH 7.4.
PA signal was monitored over time (Figure 1c,d). Formation of
the rNODs was apparent after 5 min of irradiation, and
maximal release was noted after 30−40 min of irradiation
(Figure 1c−e). Absorbance spectra of the irradiated solutions
were then acquired, and spectral analysis revealed that
approximately 54% of photoNOD-1 and 88% of photoNOD-
2 were consumed, corresponding to 5.4 and 8.8 μM NO,
respectively (Figure S2; calculations based on measured
extinction coefficients, Table S1). Further irradiation beyond
this point did not increase the yield of NO. The identity of the
rNODs was further confirmed with high-resolution mass
spectrometry (Figures S3−S8).
We then sought to confirm that NO is the species released
by electron paramagnetic resonance (EPR) spectroscopy.
Deoxygenated solutions of each photoNOD (200 μM in
water containing 50% DMF) were irradiated at λblue (Figure
S9) in the presence of Fe(MGD)2, a known NO spin-trap.
Upon nitrosylation, Fe(MGD)2 generates a diagnostic triplet
(g = 2.04) corresponding to the MGD-Fe-NO complex
(Figure 2a,b) as well as an apparent color change consistent
with the generation of rNODs (Figure 2c). Importantly,
because only NO (and not NO+) can react with Fe(MGD)2 to
generate an EPR-active species,51,69,70 this confirmed that
irradiation induces denitrosylation of both photoNODs. NO
release was then quantified by EPR after 5 or 40 min of
irradiation to confirm that release can be controlled by
irradiation time. photoNOD-1 released 20.4 μM (10%) NO
after 5 min and 30.1 μM (15%) after 40 min, and photoNOD-
2 released 38.7 μM (20%) NO after 5 min and 73.9 μM (37%)
Figure 1. (a) Absorbance spectra of photoNOD-1 and rNOD-1 and
(b) photoNOD-2 and rNOD-2 in CHCl3 (solid line) and aqueous
buffer (dashed line; photoNOD-1: 50% EtOH/HEPES, pH 7.4;
photoNOD-2: 0.1% CrEL/HEPES, pH 7.4). (c) In vitro PA
monitoring of 10 μM photoNOD-1 (PA730 nm/PA680 nm) and (d)
photoNOD-2 (PA 750 nm/PA680 nm) turnover upon irradiation at 680
nm in CHCl3 in tissue-mimicking phantoms. Data presented as mean
SD (n = 3). (e) Representative PA images of photoNOD-2 (10
μM) without irradiation (left), after 5 min of irradiation (center), and
an authentic sample of rNOD-2 shown for comparison (right) in
tissue-mimicking phantoms.
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was observed. Finally, we sought to ensure that the
photoNODs are stable to cytochrome P450 (CYP450)
enzymes. CYP450s are known to oxidize some N-alkyl-N-
nitrosamines to α-hydroxy-N-nitrosamine intermediates, which
can form DNA-alkylating diazonium species.71 We incubated
the photoNODs with NADPH-supplemented rat liver micro-
somes, a source of CYP450s, and verified that these conditions
did not generate a ratiometric turn-on, suggesting that the
photoNODs are not substrates for these enzymes. This is an
especially important finding because N-nitrosamines are often
considered untenable for biological studies based upon the
assumption that they are carcinogenic.71−74 In contrast,
solutions of photoNOD-1 subjected to 5 min of irradiation
at λblue yielded a 46.6-fold turn-on (theoretical maximum of
115.4-fold calculated for rNOD-1, 40.4%, Figure 3a), while
photoNOD-2 yielded a 21.5-fold ratiometric turn-on (the-
oretical maximum of 45.4-fold for rNOD-2, 47.4%, Figure 3b),
indicating robust and selective turnover of the photoNODs
following irradiation. Additionally, the absorbance properties
of the photoNODs proved to be independent of pH in the
range of 4.0−10.0, indicating that NO could be reliably
released by irradiation at λblue regardless of pH (Figure S12).
Collectively, these data indicate that irradiation at λblue
generates the rNODs with concomitant release of NO, and
this process can be conveniently monitored by ratiometric
absorbance or PA imaging. The apparent stability of the
photoNODs, in conjunction with the turnover response
following irradiation, suggests that the photoNODs should
be applicable for controlled NO release in living systems.
We next surveyed the effect of the photoNODs and rNODs
on the viability of live cells to evaluate their biocompatibility
for in vivo applications. HEK293T cells were treated with
photoNODs and rNODs (30 μM) for 24 h. The toxicity was
evaluated by fluorescence microscopy using Hoechst and
propidium iodide counterstaining.75,76 All compounds that
were tested were noncytotoxic under these conditions,
maintaining greater than 95% cell viability (Figure S13).
Characterization of photoNODs in Vivo. Having
established stability and cellular compatibility, we sought to
characterize the photoNODs and rNODs in live animals. Each
compound was subcutaneously administered into the flank of a
BALB/c mouse, and PA spectra from 680 to 850 nm were
acquired (Figure 4a,d). From these data, the optimal
Figure 2. EPR spectra collected after 0, 5, and 40 min of irradiation of
(a) photoNOD-1 (690 nm) or (b) photoNOD-2 (680 nm) (200 μM
in water containing 50% DMF) in the presence of Fe(MGD)2. (c)
Representative images showing colorimetric change of the EPR
samples at each time point. (d) Quantification of NO release. Data
presented as mean SD (n = 3).
NO after 40 min (Figures 2d and S10). We attribute the
difference in release efficiencies to solubility-dependent
changes in the photophysical properties rather than intrinsic
differences.
To evaluate chemostability, each photoNOD was incubated
with reduced glutathione, copper(I), copper(II), iron(II), and
iron(III) to survey the effect of these species toward
transnitrosylation, denitrosylation, or denitrosation. The
absorbance spectra of these solutions were then acquired,
and the ratiometric fold turn-on was calculated. However, no
significant ratiometric turn-on was detected in the presence of
any of these species (Figures 3a,b and S11). Next, the
photoNODs were incubated in human plasma to evaluate
stability during circulation, and again no ratiometric turn-on
wavelengths for in vivo irradiation/excitation, λPAblue and λPAred
,
were determined (700 and 830 nm for photoNOD-1 and
rNOD-1; 710 and 810 nm for photoNOD-2 and rNOD-2).
These wavelengths were selected to maximize the ratiometric
signal (Figure 4a,d).
To validate the irradiation-mediated release of NO in vivo,
each photoNOD was administered subcutaneously into both
flanks of BALB/c mice, and the ratiometric turn-on was
monitored with and without 5 min of irradiation at λPAblue
(Figure 4c,f). This time point was selected to enable fast and
convenient experiments while also sustaining significant NO
release (as described in our in vitro characterization). In vivo
irradiation of photoNOD-1 yielded a 1.37-fold ratiometric
turn-on (theoretical maximum of 2.09-fold, 65.6% release),
with a 1.12-fold turn-on in the absence of irradiation (Figure
4b). photoNOD-2 yielded a 2.29-fold ratiometric turn-on
(theoretical maximum turn-on of 2.60-fold, 88.1% release),
while only a 1.27-fold turn-on was observed in the absence of
irradiation (Figure 4e). Of note, the values obtained for the
controls without irradiation are not statistically different from
Figure 3. Ratiometric fold turn-on of (a) photoNOD-1 and (b)
photoNOD-2 (5 μM) upon exposure to redox-active metals (20 μM),
glutathione (1 mM), rat liver microsomes (10 μL, 200 μg/mL final
concentration with 5 μM NADPH), plasma, and irradiation at λblue at
37 °C in aqueous buffers (photoNOD-1:50% EtOH/HEPES, pH 7.4;
photoNOD-2:0.1% CrEL/HEPES, pH 7.4). Data presented as mean
SD (n = 3).
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Figure 4. (a) In vivo PA spectra of photoNOD-1 and rNOD-1 after injection into the flank. Dashed lines indicate wavelengths selected for
ratiometric imaging. (b) Ratiometric fold turn-on of vehicle control without irradiation and photoNOD-1 after injection and a 5 min period with/
without irradiation at λPAblue (700 nm) and (c) representative PA images at λPAred (830 nm). (d) In vivo PA spectra of photoNOD-2 and rNOD-2
after injection into the flank. Dashed lines indicate wavelengths selected for ratiometric imaging. (e) Ratiometric fold turn-on of vehicle control
without irradiation and photoNOD-2 after injection and a 5 min period with/without irradiation at λPAblue (710 nm) and (f) representative PA
images at λPAred (n = 3). photoNODs were administered via subcutaneous injection (25 μL, 30 μM in sterile saline with 2% DMSO). Scale bars
represent 2.0 mm. Each set of images has been adjusted to the same contrast for comparison. Data presented as mean SD (n = 3 for (b),(e)). *p
< 0.05, **p < 0.01.
values obtained for a vehicle control. These results encouraged
the application of photoNODs modulate biology within small-
animal models.
control, corresponding to uptake (Figure S14). For this reason,
we chose to employ photoNOD-1 for experiments requiring
systemic administration. We posit that photoNOD-2 would be
superior for experiments in which local administration with
temporal control of NO release is required.
Modulation of Tumor Progression with photoNOD-1.
The key advantage of using a photocontrollable NO donor
over a standard NORM is the ability to release NO with high
spatiotemporal control. This enables systemic administration
of photoNORMs while preventing off-target side effects
outside of the tissue area of interest. After establishing that
the photoNODs were capable of selectively releasing NO upon
irradiation, we sought to demonstrate the capability of this
system for local release following systemic administration in a
proof-of-principle study. NO has been implicated for its
toxicity to cancer cells at high concentrations, and various NO
donors have been demonstrated to be effective for inhibiting
tumor growth in vivo.77−80 Moreover, it has been shown that
the administration of NO can sensitize hypoxic tumors to both
chemo- and radiotherapy.81−91 Thus, we hypothesized that an
in vivo tumor model could serve as an effective system to
demonstrate controlled NO release using the photoNODs.
For our initial studies, we selected photoNOD-2 for in vivo
administration based on its water solubility and resulting
superior PA signal. Unfortunately, when photoNOD-2 was
administered systemically via retroorbital injection,92 no
apparent signs of tumor uptake were observed via PA imaging.
We hypothesized that this charged molecule was likely
undergoing rapid clearance,93,94 and therefore we repeated
the study with the more lipophilic photoNOD-1. Indeed, PA
imaging at 4 h following administration revealed a substantial
increase in the ratiometric PA signal (λPAblue/λPAred) for
photoNOD-1 when compared to photoNOD-2 and a vehicle
Inspired by various reports suggesting the ability of NO to
manipulate tumor growth, and our own preliminary studies
suggesting that NO can decrease proliferation of cultured 4T1
cells (Figure S15), we sought to assess the physiological impact
of selective light-mediated NO release. To this end, 4T1
tumors were implanted in both flanks of BALB/c mice. At 5
days post-implantation, mice were randomly divided into
control and treatment groups. Either photoNOD-1 or vehicle
was administered systemically. Four hours post-injection, the
left tumor was irradiated at λPAblue (700 nm) for 5 min while
the right tumor was not (Figure 5a). Release was monitored by
PA imaging before and after irradiation (left tumor) or a 5 min
period without irradiation (right tumor) (Figure 5b,c). A
ratiometric turn-on response was observed selectively in the
irradiated tumors of the treatment group, corresponding to the
formation of rNOD-1 and NO release (Figures 5c and S16).
This process was repeated every other day for a total of 7 days
(4 doses). Over the course of the experiment, tumor volumes
were measured using calipers, the mice were weighed, and their
behavior was monitored (e.g., signs of pain and distress).95
One week following the initial treatment, we observed that the
average tumor volumes for vehicle-treated (with and without
irradiation) and photoNOD-1-treated (without irradiation)
tumors were approximately 90 mm3. In contrast, the average
volume for irradiated tumors in photoNOD-1-treated mice was
only 42 mm3 (Figure 5d). The difference in tumor volume
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Figure 5. (a) Schematic illustration of photoNOD-1 administration and NO release with PA monitoring. (b) Photograph of mouse in the imaging
tray of the PA tomographer. Dashed line indicates region of PA imaging. (c) PA images (λPAred) acquired before/after a 5 min period with/without
irradiation (λPAblue) 4 h following systemic administration of photoNOD-1 (1.2 mg/kg, 150 μL, 20% DMSO in sterile saline). Both sets of images
are adjusted to identical contrast for comparison. (d) Tumor volumes measured by calipers on each day of treatment. (e) Tumor volumes
measured by calipers on day 7 of treatment. Scale bar represents 2.0 mm. Data presented as mean SD (n ≥ 3). *p < 0.05.
between the treated mice and each control group was greater
than 50% (Figure 5e), indicating that NO release was
modulating tumor progression via NO-induced cytotoxicity
or suppression of proliferation.96−98 To further confirm these
results, we excised tumor tissue following treatment and
performed histological staining for apoptotic cells. We found
that the number of apoptotic cells was greater following
irradiation as compared to the control, indicating that tumor
growth suppression was likely occurring via NO-induced
apoptosis (Figure S17). Of note, we also confirmed that
photoNOD-1 and rNOD-1 are nontoxic to 4T1 cells even at
elevated concentrations (Figure S18).
Importantly, mice treated with photoNOD-1 experienced no
significant weight loss (Figure 6b) or behavioral changes
during this experiment. To determine the localization of
photoNOD-1 in various tissue types, tumor-bearing mice were
sacrificed 4 h following administration, and ex vivo PA images
of various organs were acquired. A trend of increased PA signal
at λPAblue was observed in the heart, liver, kidneys, spleen, and
tumor tissue of treated mice as compared to the vehicle control
(Figure 6a and c), possibly corresponding to photoNOD-1
uptake. The uptake of photoNOD-1 in a variety of tissues is
advantageous, because it suggests that NO release from
photoNOD-1 could be selectively targeted to different organs
by local irradiation for a breadth of applications. Overall, these
experiments confirmed that NO release from photoNOD-1 is
limited to the area of irradiation, demonstrating that it can be
used for NO delivery with high spatiotemporal control.
CONCLUSION
■
The photoNODs represent the first small-molecule analyte
donors that incorporate a direct PA readout to enable
noninvasive monitoring of analyte release in live animals.
Existing PA-compatible analyte carriers (e.g., carbon nano-
tubes, gold nanoparticles, and porphysomes)99 that simply
track analyte distribution do not undergo an intrinsic PA signal
change upon release and cannot provide reliable confirmation
of delivery.100−105 On the other hand, organic nanoparticles,
microsomes, and liposomes that are coloaded with an analyte
donor and responsive PA sensor can detect analyte flux;
however, these constructs are not ideal because they indirectly
couple analyte release to the PA response.106,107 Small-
molecule-based donors such as the photoNODs overcome
these limitations because each state (pre- and post-release) has
a different PA signature, allowing for facile identification of
each. This unique feature, coupled with the noninvasiveness
and high resolution of PA imaging, mitigates the need for
invasive procedures (e.g., analyte-specific electrode), additional
contrast agents, or ex vivo analyses to confirm analyte delivery.
Furthermore, the PA readouts of the photoNODs make them
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Figure 6. (a) PA signal (λPAblue) of various mouse tissues. Mice were sacrificed, and tissues were removed for imaging 4 h following retroorbital
injection of photoNOD-1 (1.2 mg/kg, 150 μL, 20% DMSO in sterile saline) or vehicle control (n ≥ 3). Signals are normalized to average control
values for comparison. (b) Mouse weight for photoNOD-treated and control animals measured on each treatment day (n ≥ 3). (c) Representative
PA images (λPAblue) of various tissues of mice treated with photoNOD-1 or vehicle control. Scale bar represents 2.0 mm.
superior for in vivo applications as compared to existing
fluorescence-based photoNORMs,62,66,108 which are optimal
for applications in cells and in shallow tissues due to the depth-
dependent attenuation of light in tissue.
NO, this design strategy may be generalizable to other
analytes, broadening the scope of synergistic NIR photorelease
and PA imaging.
When designing the photoNODs, it was essential to strike a
balance between chemostability and light-based lability of the
N-nitroso bond. Our in vitro analysis demonstrates that both
photoNODs are stable unless activated by light, including in
the presence of CYP450 enzymes, alleviating potential
concerns of formation of potent alkylating agents. Additionally,
the rNODs generated by irradiation were designed to be
weakly nucleophilic and hence less susceptible to back-capture
of released NO, unlike nitrosation-based probes for NO that
feature electron-rich aniline triggers.12,109 PA imaging follow-
ing systemic injection demonstrated that photoNOD-1 was
superior to photoNOD-2 in terms of distribution and tissue
accumulation. However, photoNOD-2 presents good PA
properties following local administration. This phenomenon
highlights the utility of noninvasive PA monitoring, because
evaluation of compound accumulation would otherwise require
the sacrifice of multiple animals and subsequent tissue analysis
at various time points. Finally, photoNOD-1 can be selectively
activated in a tissue of interest following systemic admin-
istration without causing the global effects that would be
expected for standard NORMs.
EXPERIMENTAL DETAILS
■
Synthetic Methods. tert-Butyl (4-Acetylphenyl)carbamate (1).
To a 250 mL round-bottomed flask (RBF) were added 4′-
aminoacetophenone (6.7 g, 50 mmol, 1.0 equiv) and di-tert-butyl
dicarbonate (13.1 g, 60 mmol, 1.2 equiv). The reaction vessel was
capped and flushed with nitrogen. 1,4-Dioxane (60 mL) was added,
and the solution was heated to 100 °C for 8.5 h. The reaction was
cooled and concentrated to an oil that crystallized to a solid. The solid
was washed with 1:3 v/v EtOAc/hexanes to give one batch of pure
product. The filtrate was then concentrated to a solid and washed
with 1:4 EtOAc/hexanes to afford a second batch of pure product
1
(total: 10.1 g, 43 mmol, 86% yield). H NMR (400 MHz, CDCl3) δ
7.89 (d, J = 8.4 Hz, 2H), 7.46 (d, J = 8.3 Hz, 2H), 2.55 (s, 3H), 1.50
(s, 9H). 13C NMR (125 MHz, CDCl3) δ 197.13, 152.35, 143.15,
131.84, 129.94, 117.53, 81.35, 28.37, 26.49.
tert-Butyl Methyl (4-(4-Nitro-3-phenylbutanoyl)phenyl)-
carbamate (2a). To a 250 mL RBF were added 1 (3.0 g, 12.8
mmol, 1.0 equiv) and anhydrous THF (40 mL) under nitrogen. The
reaction mixture was cooled to 0 °C, and sodium hydride (60 wt %
dispersion in mineral oil) (0.61 g, 15.3 mmol, 1.2 equiv) was added
portion-wise over the course of 10 min. After 20 min, methyl iodide
(1.57 mL, 31.9 mmol, 2.5 equiv) was added, and the white suspension
was warmed to room temperature and stirred until it formed an amber
solution. After 40 min at this temperature, completion was noted by
TLC (3:17 v/v EtOAc/hexanes); the reaction mixture was
concentrated, and the crude residue was taken up in EtOAc and
washed with brine (3×). The organic layer was dried over sodium
sulfate and concentrated to a yellow oil (quantitative yield). A portion
was transferred to a 50 mL RBF (0.59 g, 2.38 mmol, 1.0 equiv) and
stirred with benzaldehyde (0.27 mL, 2.62 mmol, 1.1 equiv),
potassium hydroxide (10 M, 0.72 mL, 7.15 mmol, 3.0 equiv), and
ethanol (10 mL) for 2.5 h. Nitromethane (1.90 mL, 35.7 mmol, 15
We envision that slight modifications to the photoNOD
scaffold, such as incorporating tumor- or tissue-targeting
motifs, could significantly increase uptake, dosage, and
specificity. More broadly, the development of small molecule
donors that can be activated to modulate biological processes
with a deep-tissue compatible PA readout provides an exciting
avenue for validating cellular findings in the native biological
context. While our work in this study focuses on the delivery of
G
DOI: 10.1021/jacs.8b05514
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Journal of the American Chemical Society
Article
equiv) was added, and the solution was stirred at room temperature
for 10.5 h, fully converting to a brown solution. The reaction was
concentrated, diluted with brine, and extracted with dichloromethane.
The organic layers were dried over sodium sulfate, filtered, and
concentrated. The product was purified via silica column chromatog-
raphy in 1:4 v/v EtOAc/hexanes to afford a yellow oil (0.47 g, 1.4
gradient from 1:2 v/v EtOAc/hexanes to 1:1 v/v EtOAc/hexanes to
afford a red-brown solid (72.7 mg, 0.131 mmol, 79% yield). 1H NMR
(500 MHz, CDCl3) δ 8.14 (d, J = 8.9 Hz, 2H), 8.11−8.02 (m, 6H),
7.50−7.34 (m, 6H), 7.18 (s, 1H), 7.01 (d, J = 8.8 Hz, 2H), 6.95 (s,
1H), 6.70−6.60 (m, 2H), 3.88 (s, 3H), 2.93 (s, 3H). 13C NMR (125
MHz, CDCl3) δ 161.19, 160.12, 154.34, 152.40, 146.61, 144.02,
143.70, 140.21, 133.35, 132.79 (t, J = 5.5 Hz), 132.52, 131.26 (t, J =
4.6 Hz), 129.45, 129.36, 129.18, 128.63, 128.61, 128.58, 125.25,
119.67, 119.38, 117.30, 114.18, 112.38, 55.51, 30.23. 19F NMR (470
MHz, CDCl3) δ −132.68 (dd, J = 64.8, 32.3 Hz). 11B NMR (161
MHz, CDCl3) δ 1.27 (t, J = 32.4 Hz).
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)-N-
methylaniline (5). To a 50 mL RBF were added 4b (0.15 g, 0.28
mmol, 1 equiv), dichloromethane (11.3 mL), Et3N (4.9 mL, 28
mmol, 100 equiv), and boron trifluoride diethyl etherate (5.2 mL, 43
mmol, 150 equiv) under nitrogen at 0 °C, and this was stirred for 2.5
h at room temperature. Additional boron trifluoride diethyl etherate
(5.2 mL, 43 mmol, 150 equiv) was added at 0 °C, and the reaction
was stirred at room temperature for 21 h. The reaction mixture was
diluted with dichloromethane, washed with a saturated solution of
sodium bicarbonate, and dried over sodium sulfate. The crude
product was purified via silica column chromatography in 1:2 v/v
EtOAc/hexanes to afford a red-brown solid (0.14 g, 0.23 mmol, 82%
yield). 1H NMR (500 MHz, CDCl3) δ 8.15−8.10 (m, 2H), 8.06 (dt, J
= 8.3, 6.0 Hz, 6H), 7.50−7.39 (m, 5H), 7.37 (t, J = 7.3 Hz, 1H), 7.18
(d, J = 3.6 Hz, 1H), 7.10−7.05 (m, 2H), 6.93 (s, 1H), 6.63 (t, J = 8.0
Hz, 2H), 4.75 (t, J = 2.2 Hz, 2H), 2.93−2.82 (m, 3H), 2.56 (t, J = 2.4
Hz, 1H). 13C NMR (125 MHz, CDCl3) δ 160.68, 159.05, 153.28,
152.74, 146.99, 143.92, 143.75, 139.59, 133.52, 133.11, 132.46,
131.23, 129.58, 129.54, 129.23, 128.72, 128.70, 128.63, 126.36,
120.15, 119.01, 117.12, 115.05, 112.49, 78.57, 76.14, 56.08, 30.18. 19F
NMR (471 MHz, CDCl3) δ −132.29 (dd, J = 61.2, 32.6 Hz). 11B
NMR (128 MHz, CDCl3) δ 1.31 (t, J = 32.3 Hz).
1
mmol, 59% yield over three steps). H NMR (500 MHz, CDCl3) δ
7.88 (d, J = 8.8 Hz, 2H), 7.36 (d, J = 8.8 Hz, 2H), 7.34−7.31 (m,
2H), 7.30−7.26 (m, 3H), 4.83 (dd, J = 12.5, 6.5 Hz, 1H), 4.68 (dd, J
= 12.5, 8.1 Hz, 1H), 4.22 (p, J = 7.1 Hz, 1H), 3.48−3.35 (m, 3H),
3.30 (s, 3H), 1.48 (s, 9H). 13C NMR (125 MHz, CDCl3) δ 195.85,
154.10, 148.59, 139.24, 132.57, 129.19, 129.16, 128.68, 127.99,
127.57, 124.44, 81.44, 79.69, 41.48, 39.44, 36.90, 28.40, 14.33.
tert-Butyl (Z)-(4-(5-((5-(4-Methoxyphenyl)-3-phenyl-2H-pyrrol-2-
ylidene)amino)-4-phenyl-1H-pyrrol-2-yl)phenyl)(methyl)-
carbamate (4a). To a 50 mL RBF were added 2a (475 mg, 1.2 mmol,
1.0 equiv), 3a12 (718 mg, 2.4 mmol, 2.0 equiv), and ethanol (36 mL),
and the mixture was stirred at 60 °C until completely dissolved.
Ammonium acetate (1.38 g, 18 mmol, 15 equiv) was added, and the
reaction was refluxed for 15 h. The solution was then cooled to room
temperature; the solids were isolated by filtration and purified by
gradient silica column chromatography (dichloromethane, then 1:199
v/v methanol/dichloromethane) to yield a blue-green solid (185 mg,
1
0.30 mmol, 25%). H NMR (500 MHz, CDCl3) δ 8.02−7.92 (m,
4H), 7.79 (d, J = 8.3 Hz, 2H), 7.67 (d, J = 8.1 Hz, 2H), 7.40−7.29
(m, 9H), 7.06 (s, 1H), 6.96 (s, 1H), 6.91 (d, J = 8.3 Hz, 2H), 3.81 (s,
3H), 3.30 (s, 3H), 1.54 (s, 9H). 13C NMR (125 MHz, CDCl3) δ
161.74, 159.90, 154.46, 153.35, 148.00, 145.76, 144.53, 144.45,
139.09, 134.10, 133.60, 130.34, 129.15, 129.05, 128.94, 128.74,
128.38, 128.18, 128.15, 127.98, 127.81, 127.50, 125.98, 125.32,
125.07, 116.92, 114.53, 113.86, 112.02, 80.85, 53.54, 37.07, 28.45.
(Z)-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 (4b). To
a 25 mL RBF were added 2a (1.13g, 2.84 mmol, 1.0 equiv),
trifluoroacetic acid (2 mL), and dichloromethane (10 mL), and this
was stirred for 1 h at room temperature. Upon completion by TLC,
the reaction mixture was cooled to 0 °C and quenched with saturated
sodium bicarbonate (5 mL) followed by slow addition of solid sodium
carbonate until gas evolution ceased. The reaction mixture was
extracted with dichloromethane, dried over sodium sulfate, filtered,
and concentrated to afford a clear oil, 2b, which was used without
purification (0.60 g, 2.52 mmol, 89% yield). A portion (0.2 g, 0.84
mmol, 1.0 equiv) was added to a 50 mL RBF; 3b12 (0.54 g, 1.69
mmol, 2.0 equiv) and n-butanol (20 mL) were added and warmed at
110 °C until solids completely dissolved. Ammonium acetate (0.97 g,
12.6 mmol, 15 equiv) was added, and the reaction was stirred at this
temperature for 8 h. The solution was concentrated via rotary
evaporation, diluted with brine, and extracted with EtOAc. The
organic layer was dried over sodium sulfate, filtered, and concentrated.
The product was purified via silica column chromatography (4:1 v/v
dichloromethane/hexanes with 0.1% Et3N) to afford a blue-green
solid (0.150 g, 0.28 mmol, 33% yield). 1H NMR (400 MHz, CDCl3)
δ 8.07 (dt, J = 8.0, 1.8 Hz, 5H), 7.92 (d, J = 8.6 Hz, 2H), 7.82−7.77
(m, 2H), 7.46−7.27 (m, 6H), 7.13−7.09 (m, 2H), 7.00 (s, 1H), 6.72
(d, J = 8.7 Hz, 2H), 4.79 (d, J = 2.4 Hz, 2H), 4.28 (d, J = 5.4 Hz, 1H),
2.97 (d, J = 5.2 Hz, 3H), 2.59 (t, J = 2.4 Hz, 1H). 13C NMR (126
MHz, CDCl3) δ 162.43, 158.12, 155.00, 151.68, 145.52, 145.29,
144.14, 137.30, 134.45, 133.73, 129.32, 129.16, 128.78, 128.16,
128.14, 128.00, 127.21, 126.99, 125.78, 121.39, 118.05, 115.57,
112.41, 110.40, 78.28, 75.88, 55.95, 29.70.
rNOD-2. To a two-neck 25 mL RBF were added 5 (0.05 g, 0.086
mmol, 1 equiv), 612 (0.12 g, 0.3 mmol, 3.5 equiv), copper sulfate
pentahydrate (0.1 g, 0.43 mmol, 5 equiv), and tris-hydroxypropyl-
triazolylmethylamine (7 mg, 0.02 mmol, 0.2 equiv), and this was
flushed under nitrogen for 1 h. (+)-Sodium L-ascorbate (0.025 g,
0.129 mmol, 1.5 equiv) was added along with degassed THF (4.5
mL) and degassed water (1.5 mL). The reaction was stirred for 21 h
at room temperature, diluted with dichloromethane, and washed with
brine. The aqueous layer was further extracted with a mixture of
dichloromethane and isopropanol (2:1 v/v). The combined organic
layers were dried over sodium sulfate, filtered, and concentrated. The
crude product was loaded onto Celite and purified via neutral alumina
column chromatography in 1:19 v/v MeOH/dichloromethane (2×)
1
to afford a red-brown solid (0.025 g, 0.028 mmol, 33% yield). H
NMR (500 MHz, CD2Cl2) δ 8.14 (d, J = 8.7 Hz, 2H), 8.09 (d, J = 7.4
Hz, 2H), 8.05 (d, J = 7.7 Hz, 2H), 8.02−7.95 (m, 3H), 7.45 (dt, J =
21.2, 7.5 Hz, 5H), 7.38−7.30 (m, 2H), 7.29−7.24 (m, 1H), 7.12 (d, J
= 8.4 Hz, 2H), 6.91−6.83 (m, 3H), 5.25 (s, 2H), 4.54 (t, J = 4.8 Hz,
2H), 3.85 (t, J = 4.9 Hz, 2H), 3.68 (bs, 2H), 3.58−3.45 (m, 10H),
3.12 (s, 9H), 2.91 (d, J = 5.0 Hz, 3H). 13C NMR (125 MHz, CD2Cl2)
δ 161.42, 159.85, 154.97, 151.13, 147.83, 144.40, 143.82, 143.14,
138.04, 133.99, 133.84, 132.57, 131.42, 130.03, 129.79, 129.29,
129.07, 129.00, 128.70, 126.85, 125.05, 121.14, 117.67, 116.63,
115.20, 113.00, 70.97, 70.75, 70.74, 70.56, 69.66, 65.93, 65.37, 62.49,
54.75, 50.86, 29.99. 19F NMR (471 MHz, CD2Cl2) δ −131.65 (dd, J
= 65.5, 31.7 Hz). 11B NMR (128 MHz, CD2Cl2) δ 1.25 (t, J = 32.5
Hz).
rNOD-1. To a 100 mL RBF were added 4a (100 mg, 0.166 mmol,
1.0 equiv), dichloromethane (10 mL), and Et3N (0.33 mL, 2.48
mmol, 15 equiv). Boron trifluoride diethyl etherate (0.30 mL, 2.48
mmol, 15 equiv) was added dropwise. The reaction was stirred for 6 h
before additional Et3N (0.33 mL, 2.48 mmol, 15 equiv) and boron
trifluoride diethyl etherate (0.30 mL, 2.48 mmol, 15 equiv) were
added. The reaction was stirred for another 15 h. The reaction
mixture was diluted with dichloromethane, washed with a saturated
solution of sodium bicarbonate, and dried over sodium sulfate. The
crude product was purified via silica column chromatography using a
photoNOD-1. To a 25 mL RBF were added rNOD-1 (72.7 mg,
0.130 mmol, 1.0 equiv), THF (8.7 mL), dichloromethane (4.35 mL),
acetic acid (4.35 mL), and sodium nitrite (44.8 mg, 0.653 mmol, 5.0
equiv). The reaction mixture was stirred for 2 h at room temperature.
Upon completion by TLC, the reaction was diluted in dichloro-
methane and quenched with a saturated solution of sodium
bicarbonate. The organic layer was dried, concentrated, and purified
by silica column chromatography in 4:1 v/v dichloromethane/
1
hexanes to yield a green solid (36.9 mg, 0.63 mmol, 48% yield). H
H
DOI: 10.1021/jacs.8b05514
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Journal of the American Chemical Society
Article
NMR (500 MHz, CDCl3) δ 8.17 (d, J = 8.5 Hz, 2H), 8.14 (d, J = 8.7
Hz, 2H), 8.10−8.04 (m, 4H), 7.69 (d, J = 8.4 Hz, 2H), 7.45 (h, J =
7.0, 6.5 Hz, 6H), 7.12 (s, 1H), 7.03 (s, 2H), 7.01 (s, 1H), 3.89 (s,
3H), 3.48 (s, 3H). 13C NMR (125 MHz CDCl3) δ 162.62, 160.68,
155.17, 146.45, 144.84, 144.71, 143.33, 142.44, 132.57, 132.10 (t, J =
4.9 Hz), 132.03, 130.75 (t, J = 4.5 Hz), 130.68, 129.71, 129.42,
129.24, 129.20, 128.64, 128.61, 123.52, 119.66 (d, J = 3.6 Hz),
118.30, 118.01 (d, J = 4.1 Hz), 114.44, 55.51, 30.65. 19F NMR (471
MHz, CDCl3) δ −131.97 (dd, J = 63.5, 31.7 Hz). 11B NMR (161
MHz, CDCl3) δ 1.08 (t, J = 31.7 Hz).
buffer with 0.1% CrEL (pH 7.4). CYP450 activity was validated prior
to this analysis with N-nitroso-N-methylaniline (NMA), a known
substrate of CYP450s found in rat liver microsomes. Rat liver
microsomes (10 μM) were incubated with 100 μM NMA and 500
μM NADPH in 100 mM potassium phosphate buffer, pH 7.4. After
1.5 h of vigorous shaking at 30 °C and extraction with ethyl acetate,
full consumption of NMA was observed by TLC.
For comparison, solutions of photoNOD-1 (5 μM) in 20 mM
HEPES buffer with 50% EtOH and photoNOD-2 (5 μM) in 20 mM
HEPES buffer with 0.1% CrEL were prepared. Solutions were
irradiated (photoNOD-1, 680 nm; photoNOD-2, 690 nm) in FEP
tubes in tissue-mimicking phantoms as previously described (see In
Vitro Irradiation). Absorbance spectra were obtained before and after
irradiation to determine ratiometric turn-on.
EPR Analysis of photoNOD NO Release. The Fe(II) complex
with N-(dithiocarbamoyl)-N-methyl-D-glucamine (MGD) [(Fe-
MGD2)] was prepared by mixing FeSO4·7H2O with MGD in
degassed aqueous DMF (1:1) to final concentrations of 0.8 and 4
mM, respectively. Samples were maintained under a nitrogen
atmosphere and in the absence of light until immediately before
measurement. Samples were transferred to a hematocrit capillary,
sealed, and placed in a quartz EPR tube to acquire the spectra. EPR
spectra of photoNOD-1 or -2 (200 μM) were acquired either with or
without 5 or 40 min irradiation at 680 or 690 nm in step-and-shoot
mode, respectively (Figure S13). MAHMA-NONOate (a commer-
cially available NO donor) was used as a positive control and for the
preparation of the calibration curve. Varying concentrations were
prepared in degassed 10 mM potassium hydroxide and quickly added
to the Fe(II) complex (Figure S8). EPR spectra were recorded with a
Varian E-line 12″ Century Series X-band CW EPR (VarianXBand)
spectrometer at room temperature. Spectrometer settings: modulation
frequency, 100 kHz; modulation amplitude, 2.5 G; gain, 4000; sweep
width, 100 G; scan time, 0.5 min per scan, 6 scans, 3 min total;
microwave power, 12 dB; and microwave frequency, 9.30 GHz.
Additional information regarding the EPR materials can be found in
the Supporting Information.
photoNOD-2. To a 25 mL RBF were added rNOD-2 (0.014 g,
0.015 mmol, 1 equiv), acetic acid (0.51 mL), THF (1 mL), and
dichloromethane (1 mL), and this was cooled to 0 °C. Sodium nitrite
(5 mg, 0.0772 mmol, 5 equiv) was added and stirred for 1 h. The
reaction was diluted in dichloromethane and washed with saturated
sodium bicarbonate. The compound was extracted from the aqueous
layer using dichloromethane/isopropanol (2:1 v/v) and washed with
brine. The organic layer was dried over sodium sulfate and
concentrated. The product was purified via neutral alumina column
chromatography in 1:19 v/v MeOH/dichloromethane to afford a blue
solid (0.009 g, 0.01 mmol, 66% yield). 1H NMR (500 MHz, CD2Cl2)
δ 8.18 (d, J = 8.9 Hz, 2H), 8.16 (d, J = 9.0 Hz, 2H), 8.09 (ddt, J = 9.6,
6.4, 1.5 Hz, 4H), 8.02 (s, 1H), 7.74−7.71 (m, 2H), 7.52−7.42 (m,
6H), 7.20 (d, J = 1.1 Hz, 1H), 7.19−7.16 (m, 2H), 7.09 (d, J = 1.2
Hz, 1H), 5.30 (s, 2H), 4.58 (dd, J = 5.6, 4.6 Hz, 2H), 3.89 (dd, J =
5.6, 4.7 Hz, 2H), 3.85 (td, J = 5.0, 2.8 Hz, 2H), 3.82−3.77 (m, 2H),
3.61−3.55 (m, 4H), 3.55−3.51 (m, 4H), 3.48 (s, 3H), 3.31 (s, 9H).
13C NMR (125 MHz, CD2Cl2) δ 161.93, 160.87, 156.01, 146.95,
145.35, 144.14, 143.46, 143.18, 133.00, 132.68, 132.51, 131.28,
131.08, 130.36, 129.94, 129.89, 129.76, 129.23, 129.20, 125.10,
124.52, 120.42, 118.92, 115.68, 71.09, 70.87, 70.79, 70.77, 69.82,
66.00, 65.66, 62.67, 54.87, 50.90, 31.25. 19F NMR (471 MHz,
CD2Cl2) δ −131.31 (dd, J = 63.7, 31.8 Hz). 11B NMR (161 MHz,
CD2Cl2) δ 1.23, 1.04, 0.83. 11B NMR (161 MHz, CD2Cl2) δ 1.03 (t, J
= 32.0 Hz).
In Vitro Irradiation. A solution of photoNOD-1 or -2 (10 μM in
chloroform) was prepared, and initial absorbance spectra were
obtained (400 to 900 nm). Solutions (400 μL) were pipetted into
FEP tubing (0.08 in. diameter, cut to 5 cm long), and the tubing was
inserted into the tissue phantom (see Supporting Information) and
sealed by folding over the ends and securing with additional tubing
(0.12 in. diameter). Initial PA measurements were acquired at 680
and 730 nm (photoNOD-1) or 680 and 750 nm (photoNOD-2)
(continuous mode, 6 s rotation time). Following the initial
measurement, subsequent PA irradiations at 680 nm were conducted
for a total scan time of 0.5−40 min (continuous mode with 6 s
rotation time for 0.5 and 1 min; step-and-shoot mode (120 angles, 41
pulses per angle) for longer time points), and PA images were
acquired at both wavelengths after each time point. After the 40 min
time point, the solution was removed from the FEP tubing, and
absorbance spectra were obtained (400−900 nm). Control PA
measurements were acquired after periods of 0.5−40 min of darkness
(using fresh samples of photoNOD-1 or -2 for each time point) to
assess turnover in the absence of irradiation. Each time point was
performed in triplicate.
In Vivo Studies. All in vivo 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. Female BALB/c mice (5−7 weeks old) were acquired from The
Jackson Laboratory. Hair was removed from the lower half of the
body by shaving and applying depilatory cream prior to all
experiments.
In Vivo NO Release and Monitoring. Vehicle control or
photoNODs (25 μL, 30 μM in sterile saline containing 2% DMSO)
were injected subcutaneously into both flanks of BALB/c mice. PA
images were acquired both before and after a 5 min period with
irradiation (right flank) or without irradiation (left flank). Irradiation
of photoNOD-1 and photoNOD-2 was performed at 710 and 700
nm, respectively, using step-and-shoot mode (120 angles, 39 pulses
per angle, 4.8 min irradiation). Images were acquired at 710 and 810
nm for photoNOD-1 and 700 and 830 nm for photoNOD-2 using
continuous rotation mode (6 s rotation time, 0.2 min irradiation).
Modulation of Tumor Progression Using photoNOD-1. A
suspension of 2.5 × 105 4T1 cells in serum-free medium containing
50% v/v Matrigel (50 μL) was injected subcutaneously into both
flanks of BALB/c mice. Five days after tumor implantation, mice were
randomly assigned to experimental and control groups. photoNOD-1
(1.2 mg/kg in 150 μL of sterile saline containing 20% DMSO) was
administered to each mouse in the experimental group via retroorbital
injection, while the control mice were treated with vehicle only. Four
hours following administration, the left tumors of all mice were
irradiated for 5 min using 700 nm light (step-and-shoot mode, 120
angles, 39 pulses per angle). PA images (700 and 830 nm) were
acquired both before and after irradiation, and before and after a 5
min period without irradiation for the right tumors. This dosing
session was repeated every other day for a total of 4 sessions. On
treatment days, mice were weighed, and tumors were measured using
Stability Assays. Reported concentrations for these studies
describe the final concentration after addition of all species.
photoNOD-1 (5 μM) was preincubated for >10 min at 37 °C in
20 mM HEPES buffer with 50% EtOH (pH 7.4) before incubation
with GSH (1 mM), [(CH3CN)4Cu]PF6 (20 μM), CuCl2 (20 μM),
FeSO4·7H2O (20 μM), FeCl3 (20 μM). photoNOD-1 was also
incubated in rat liver microsomes (10 μL/mL) with NADPH (5 μM),
and in human plasma (20% by volume) in PBS. Immediately after
addition, initial absorbance spectra were acquired (400−900 nm).
After 1 h, final absorbance spectra were acquired. Stability was plotted
by determining the ratio of absorbance at λred to absorbance at λblue
before and after incubation. All experiments were performed in
triplicate. Note that Cu(I) and Fe(II) experiments were carried out
using degassed solutions under nitrogen atmosphere. The same
experiments were carried out for photoNOD-2 in 20 mM HEPES
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calipers. Tumor volumes were calculated using the formula V = (w2 ×
l)/2, where w is the tumor width and l is the tumor length.110,111
Ex Vivo Biodistribution. Tumor-bearing mice were treated with
photoNOD-1 (1.2 mg/kg in 150 μL of sterile saline containing 20%
DMSO) or vehicle without photoNOD-1 via retroorbital injection.
After 4 h, mice were sacrificed and dissected to remove tissues of
interest. Uptake of photoNOD-1 was determined via PA imaging
(700 nm, continuous mode, 6 s rotation time) of individual organs in
PBS. Custom regions of interest (ROIs) were drawn around each
organ, and the mean ROI PA signal was determined.
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ASSOCIATED CONTENT
* Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/jacs.8b05514.
Chem. Soc. 2015, 137, 15628−15631.
■
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Detailed materials, instrumentation, data processing
methods and statistical analysis, tissue phantom
preparation, pH stability, NO release quantification
(UV−vis, EPR), mass spectrometry, cell culture
information (including cytotoxicity and proliferation
assays), PA spectra, in vivo PA turn-on data, TUNEL
IHC, NMR data, and supplementary figures and tables
(PDF)
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AUTHOR INFORMATION
Corresponding Author
*jeffchan@illinois.edu
ORCID
Effie Y. Zhou: 0000-0002-2222-7391
Hailey J. Knox: 0000-0003-0608-2855
Christopher J. Reinhardt: 0000-0001-9992-1253
Gina Partipilo: 0000-0001-6353-3419
Jefferson Chan: 0000-0003-4139-4379
■
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Sang, D. K.; Xing, C.; Li, Z.; Dong, B.; et al. Proc. Natl. Acad. Sci. U. S.
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§E.Y.Z. and H.J.K. contributed equally.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
(22) Nagaya, T.; Gorka, A. P.; Nani, R. R.; Okuyama, S.; Ogata, F.;
Maruoka, Y.; Choyke, P. L.; Schnermann, M. J.; Kobayashi, H. Mol.
Cancer Ther. 2018, 17, 661−670.
■
This work was supported by the Chemistry-Biology Interface
Training Grant (T32 GM070421 to E.Y.Z. and C.J.R.), the
Tissue Microenvironment Training Grant (T32 EB019944 to
H.J.K.), the National Science Foundation Graduate Research
Fellowship Program (NGE-1144245 to E.Y.Z.), the Spring-
born Fellowship (to E.Y.Z.), the Beckman Fellowship (to
H.J.K.), and the Alfred P. Sloan fellowship (FG-2017-8964 to
J.C.). Major funding for the 500 MHz Bruker CryoProbeTM
was provided by the Roy J. Carver Charitable Trust
(Muscatine, Iowa; grant no. 15-4521) to the School of
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 also acknowledge the Core
Facilities at the Carl R. Woese Institute for Genomic Biology
for access to Leica CM3050S cryostat, tissue staining facility,
and the Hamamatsu Photonics NanoZoomer Digital Pathol-
ogy System.
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