Lanthanide(III) complexes of rhodamine-DO3A conjugates as agents for dual-modal imaging

Article abstract of DOI:10.1021/ic402233g

Two novel dual-modal MRI/optical probes based on a rhodamine-DO3A conjugate have been prepared. The bis(aqua)gadolinium(III) complex Gd.L1 and mono(aqua)gadolinium(III) complex Gd.L2 behave as dual-modal imaging probes (r₁ = 8.5 and 3.8 mM^-1 s^-1 for Gd.L1 and Gd.L2, respectively; λ_ex = 560 nm and λ_em = 580 nm for both complexes). The rhodamine fragment is pH-sensitive, and upon lowering of the pH, an increase in fluorescence intensity is observed as the spirolactam ring opens to give the highly fluorescent form of the molecule. The ligands are bimodal when coordinated to Tb(III) ions, inducing fluorescence from both the lanthanide center and the rhodamine fluorophore, on two independent time frames. Confocal imaging experiments were carried out to establish the localization of Gd.L2 in HEK293 cells and primary mouse islet cells (～70% insulin-containing β cells). Colocalization with MitoTracker Green demonstrated Gd.L2's ability to distinguish between tumor and healthy cells, with compartmentalization believed to be in the mitochondria. Gd.L2 was also evaluated as an MRI probe for imaging of tumors in BALB/c nude mice bearing M21 xenografts. A 36.5% decrease in T₁ within the tumor was observed 30 min post injection, showing that Gd.L2 is preferentially up taken in the tumor. Gd.L2 is the first small-molecule MR/fluorescent dual-modal imaging agent to display an off-on pH switch upon its preferential uptake within the more acidic microenvironment of tumor cells.

Full text of DOI:10.1021/ic402233g

Article
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Lanthanide(III) Complexes of Rhodamine−DO3A Conjugates as
Agents for Dual-Modal Imaging
Charlotte Rivas,† Graeme J. Stasiuk,† Juan Gallo,† Florencia Minuzzi,‡ Guy A. Rutter,‡
and Nicholas J. Long*,†
†Department of Chemistry, Imperial College London, South Kensington, London SW7 2AZ, U.K.
‡Section of Cell Biology, Division of Diabetes, Endocrinology and Metabolism, Department of Medicine, Imperial College London,
South Kensington, London SW7 2AZ, U.K.
S
* Supporting Information
ABSTRACT: Two novel dual-modal MRI/optical probes based on a rhodamine−DO3A conjugate have been prepared. The
bis(aqua)gadolinium(III) complex Gd.L1 and mono(aqua)gadolinium(III) complex Gd.L2 behave as dual-modal imaging
probes (r1 = 8.5 and 3.8 mM−1 s−1 for Gd.L1 and Gd.L2, respectively; λex = 560 nm and λem = 580 nm for both complexes). The
rhodamine fragment is pH-sensitive, and upon lowering of the pH, an increase in fluorescence intensity is observed as the
spirolactam ring opens to give the highly fluorescent form of the molecule. The ligands are bimodal when coordinated to Tb(III)
ions, inducing fluorescence from both the lanthanide center and the rhodamine fluorophore, on two independent time frames.
Confocal imaging experiments were carried out to establish the localization of Gd.L2 in HEK293 cells and primary mouse islet
cells (∼70% insulin-containing β cells). Colocalization with MitoTracker Green demonstrated Gd.L2’s ability to distinguish
between tumor and healthy cells, with compartmentalization believed to be in the mitochondria. Gd.L2 was also evaluated as an
MRI probe for imaging of tumors in BALB/c nude mice bearing M21 xenografts. A 36.5% decrease in T1 within the tumor was
observed 30 min post injection, showing that Gd.L2 is preferentially up taken in the tumor. Gd.L2 is the first small-molecule
MR/fluorescent dual-modal imaging agent to display an off−on pH switch upon its preferential uptake within the more acidic
microenvironment of tumor cells.
disease and to establish the efficacy of the concurrent treatment
in a noninvasive manner. This can be achieved by the in vivo
use of a dual-modal MRI/fluorescent probe that responds to a
characteristic trait of tumor cells.7−9 It is known that the
mitochondrial potential in cancer cells is greater than that of
healthy cells,10 and the design of a probe that can accumulate in
these energized mitochondria will lead to a tumor-targeting
agent. The combination of MRI and fluorescence will allow for
improved diagnostic accuracy by tumor localization via MRI
imaging and guided surgery via fluorescence imaging.
INTRODUCTION
■
Molecular imaging is a rapidly growing area of chemistry that
aims to visualize cellular function and structure in a noninvasive
manner with the in vivo use of specially designed imaging
agents. The combination of magnetic resonance imaging (MRI)
and optical imaging offers synergistic advantages over either
modality alone. MRI has high spatial resolution and good soft-
tissue contrast1,2 while optical imaging displays high sensitivity
and can give information on the local chemical environment.3
Lanthanide-based optical probes have superior fluorescence
properties in comparison to organic fluorophores, with
luminescence lifetimes stretching to the millisecond region,
and time-gated techniques can be employed in order to
eliminate interfering background autofluorescence.4−6
MRI is a noninvasive diagnostic technique that relies on the
enhancement of local water proton relaxation.11,12 Different
tissues have different relaxation times, resulting in endogenous
With the constant advances in cancer treatment, it is
becoming increasingly important to detect the early signs of the
Received: September 2, 2013
Published: December 4, 2013
© 2013 American Chemical Society
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a
Scheme 1. Synthesis of L1 and Ln.L1 (Ln = Gd, Tb)
a
Reagents and conditions: (a) (i) POCl3, reflux, 18 h; (ii) bromoethylamine, CH3CN, room temperature, 24 h. (b) Tri-t-Bu-DO3A, K2CO3,
CH3CN, reflux, 24 h. (c) TFA, DCM, room temperature, 16 h. (d) LnCl3·6H2O, H2O, NaOH(aq), pH 5.5, room temperature, 24 h.
magnetic resonance contrast. Exogenous agents can be used to
enhance this contrast by selectively shortening the T1
(longitudinal) or T2 (transverse) relaxation time; these agents
include Gd(III) chelates, manganese complexes, and super-
paramagnetic nanoparticles.13,14 The most successful and
commonly used of these exogenous agents are complexes
containing the Gd(III) ion because of its high magnetic
moment (7.9μB) given by its seven unpaired electrons and its
totally symmetric electronic ground state (8S7/2).15 According
to Solomon−Bloembergen−Morgan theory, the relaxivity of
Gd(III) complexes is governed by a number of factors,
including the number of bound water molecules (q), the
rotational tumbling time (rotational correlation time) of the
complex (τR), and the mean residence lifetime of Gd(III)-
bound water molecules (τM).2 An effective way to increase the
relaxivity of a Gd(III) complex is to increase the number of
bound water molecules. Herein we report two novel probes
possessing different hydration states. We envisioned that the
relaxivity could be improved by decreasing the number of
coordination sites on the Gd(III) chelate from eight to seven,
allowing for a change from one bound water molecule to two. It
is reported however that heptadentate ligands, when bound to
Gd(III), have unfavorable properties for use as MRI probes in
terms of thermodynamic and kinetic stability because of the
coordinatively unsaturated nature of the complex.16
Eu(III) and Tb(III) chelates based on a DO3A core have
received a lot of interest as luminescent lanthanide
systems.17−20 These lanthanides are characterized by their
long-lived luminescent excited states (ms time scale) and
linelike emission spectra, and their complexes have been shown
to be useful in biomedicine as luminescent probes.21
Lanthanide metals or ions however, have low absorption
coefficients due to Laporte-forbidden f−f transitions.22 This can
be overcome by attaching an aromatic chromophore to the
macrocyclic core to act as an “antenna” by transferring
absorbed excitation energy from its triplet excited state to the
excited state of the coordinated lanthanide ion.23 Rhodamine
derivatives have received significant attention as fluorescent
labeling agents because of their long absorption and emission
wavelengths, large absorption coefficients, and high quantum
yields.24 They have been used in a wide range of applications,
having great success as chemosensors (both in vitro and in
vivo),25,26 protein labels,27 and dual-modal imaging agents.28,29
Rhodamine derivatives have had particular success as
mitochondria targets,30−32 as their delocalized organic cationic
forms tend to accumulate in the mitochondria of tumor cells as
a result of the increased negative mitochondrial potential.33−35
The spiro ring-opening mechanism of rhodamine has also led
to its extensive use as a chemosensor for metal detection as well
as a pH-sensitive probe.36−38 Under acidic conditions, the
rhodamine moiety exists in its highly fluorescent pink-colored
open form. Upon addition of OH−, the structure exists in its
spirocyclic form, which is colorless and nonfluorescent.
In recent years there have been an increasing number of
reports of multimodal imaging probes wherein two or more
imaging modalities are combined.39,40 The combination of
modalities leads to probes that give enhanced visualization and
improved reliability of data by synergistically combining
imaging techniques to overcome their inherent disadvantages.
MRI gives high anatomical resolution and deep tissue
penetration but lacks sensitivity. Optical imaging, however,
boasts high sensitivity but has limited tissue penetration. As
discussed earlier, the combination of the two techniques yields
a probe that is able to provide a more complete picture of the
biological area of interest. There are few reports of dual-modal
MRI/optical imaging agents in the literature that are not of
nanoparticle nature. These include Gou et al.’s description of
the binding of a cyanine−DTPA conjugate to BSA to afford
relaxivity rate constants of 15 mM−1 s−1.41 The aliphatic nature
of the chelate is unfavorable, however, as release of Gd(III) ions
into the body from these types of complexes has been related to
nephrogenic systemic fibrosis (NSF).42 In 1998, it was reported
that a rhodamine−Gd(III) complex conjugate showed no MRI
contrast enhancement when introduced to Xenopus laevis
embryos.43 The same complex conjugate, Gd(Rhoda-DOTA),
was studied alongside a series of hydrophobic fluorescent dye−
Gd(III) complex conjugates in 2011 by Yamane et al.,44 who
found that incorporation of the hydrophobic fluorescent moiety
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a
Scheme 2. Synthesis of L2 and Ln.L2 (Ln = Gd, Tb)
a
Reagents and conditions: (a) Ethylenediamine, EtOH, reflux, 24 h. (b) Chloroacetyl chloride, NEt3, DCM, room temperature, 2 h. (c) Tri-t-Bu-
DO3A, K2CO3, CH3CN, reflux, 24 h. (d) TFA, DCM, room temperature, 16 h. (e) LnCl3·6H2O, H2O, NaOH(aq), pH 5.5, room temperature, 24 h.
improves cell permeability compared to Gd(DOTA). They
reported that Gd(Rhoda-DOTA) showed a large increase in r1
relaxivity when bound to albumin and efficient cell permeation
when incubated with HeLa cells. Again, however, no significant
change in the MR signal of HeLa cells was observed in the
presence of the complex, even though its fluorescence was
observed by microscopy. Similarly, a rhodamine moiety has
previously been conjugated to DO3A and consequently labeled
with 64Cu to arrive at a dual-modal PET/optical probe.29 It was
found that 64Cu(DOTA-LRB) had a surprisingly high tumor
uptake compared with other 64Cu-labeled organic cations. The
probe was able to selectively localize to the tumor mitochondria
with long tumor retention times.45 The probe and its
derivatives have been the subjects of extensive in vitro and in
vivo studies showing that there is promise for a Gd(III) MRI/
optical analogue to be as successful in vivo.
In this paper, we report two novel methods for conjugating
fluorescent rhodamine derivatives to DO3A. We evaluated the
two rhodamine−DO3A Gd(III) derivatives as multimodal
MRI/optical imaging agents by measuring their relaxivity and
fluorescence properties as well as their in vitro tumor cellular
localization. The Tb(III) analogues were synthesized to assess
their dual-luminescence properties, and it was found that the
organic and metal-based luminescence can be separated on
different time scales.
both commencing from commercially available rhodamine B
and involving five straightforward steps. Reaction of the
carboxylic acid moiety on rhodamine B with bromoethylamine
resulted in amide bond formation, giving product 2. The ligand
precursor 3 was prepared by N-alkylation of the tri-tert-butyl
ester derivative of cyclen (tri-t-Bu-DO3A). Removal of the tert-
butyl ester groups was then carried out using trifluoroacetic acid
(TFA) to obtain L1. The successful isolation of the product
1
was confirmed through H and 13C NMR spectroscopy along
with high-resolution mass spectrometry. L1 was coordinated to
a series of lanthanide metal chloride salts [Ln = Gd(III),
Tb(III)] in water in a mildly acidic environment (pH ∼5.5).
After purification, the absence of free lanthanide ions was
verified by the use of xylenol orange indicator solution.46 For
each complex, a peak corresponding to the molecular ion with
the correct isotopic pattern could be observed by high-
resolution electrospray ionization mass spectrometry. The
Ln.L1 complexes were found to be only partially water-soluble
and therefore unsuitable for in vivo applications. Thus, the
second ligand L2 was developed to overcome the solubility
issues of L1 in water. The synthesis of L2 was very similar to
that of L1. Two subsequent amide bond formation reactions on
rhodamine B were carried out, first with ethylenediamine to
form 4 and then with chloroacetyl chloride resulting in product
5. This was then conjugated to tri-t-Bu-DO3A to give 6 in 39%
yield, and subsequent deprotection with TFA gave L2 in the
same manner as for L1. The second amide bond was
introduced to increase the hydrogen-bond donor and acceptor
ability and thereby to increase the water solubility. L2 was
RESULTS AND DISCUSSION
■
Synthesis. Two new synthetic routes were developed to
obtain L1 and L2 (outlined in Schemes 1 and 2, respectively),
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Figure 1. Structures of Ln.L1 and Ln.L2 [Ln = Gd(III), Tb(III)].
Figure 2. Fluorescence emission pathways in Tb.L1 and Tb.L2.
Figure 3. Left panel: excitation spectrum (left) and emission spectrum (right) of Gd.L2 with λem = 580 nm and λex = 560 nm (0 ms delay). These
are also representative spectra of Gd.L1. Right panel: absorption spectrum (left), terbium emission (center), and rhodamine fluorescence (right) of
Tb.L2 with λem = 545 nm and λex = 310 and 560 nm, respectively (0.1 ms delay for terbium emission). These are also representative spectra of
Tb.L1, although no delay was needed to observe the terbium emission. All of the Ln.L1 complex measurements were carried out in methanol, and all
of the Ln.L2 complex measurements were carried out in H2O (pH 7.4, 298 K, 100 μM concentration).
complexed with Gd(III) and Tb(III) as above, and the
complexes Ln.L2 were found to be water-soluble.
were found to be very similar, as they exhibited fluorescence
emission at 580 nm upon excitation at 560 nm. This short
Stokes shift and orange fluorescence was expected, as the
fluorescence properties of rhodamine and its derivatives have
been extensively studied.47 Modification of the carboxylic acid
moiety of rhodamine B does not lead to significant changes in
the photophysical properties of the chromophore, as no
Photophysical Properties. Complexes Ln.L1 and Ln.L2
(Figure 1) behave as dual-modal MRI/optical imaging probes
with Gd(III) and dual-luminescent probes with Tb(III). In this
section, we discuss the photophysical properties of both
systems. The photophysics of the two ligands with Gd(III)
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electronic or structural changes occur on the xanthene core, so
a very small bathochromic shift is observed (rhodamine B: λex =
553 nm, λem = 572 nm in ethanol).
Table 1. Selected Photophysical Data for Complexes Tb.L1
and Tb.L2
complex
τH O/ms
τD O/ms
q
ϕem/%
The photophysical properties of the Tb(III) analogues were
investigated (Figure 2). Under excitation at λex = 310 nm,
complex Tb.L1 displays exclusively green emission arising from
5D4−7FJ transitions of terbium. The rhodamine-centered
emission is of a very low intensity in comparison with the
Tb(III) emission, indicating efficient ligand-to-lanthanide
energy transfer. At λex = 560 nm, emission is seen at 580 nm,
featuring fluorescence from the xanthene core. Therefore, two
fluorescence pathways are available to Tb.L1: emission via the
lanthanide with energy transfer from rhodamine and direct
excitation of the xanthene core. Conversely, under excitation at
λex = 310 nm, the emission spectrum of complex Tb.L2 is
dominated by rhodamine emission at 580 nm. Applying a 0.1
ms delay to the fluorescence measurement allowed for isolation
of the short-lived organic fluorescence of the rhodamine moiety
from the long-lived lanthanide luminescence in both complexes.
The extra amide bond in L2 renders the complex water-soluble
but also introduces a quenching pathway. The additional N−H
bond is close to the lanthanide center, and fluorescence
deactivation may occur via the N−H vibrational energy level
oscillator.48
2
2
Tb.L1
Tb.L2
1.23
1.55
2.53
2.05
1.8
0.8
18.6
0.8
hydration states of two and one respectively. These hydration
states were confirmed by the lifetime measurements of the
terbium analogues.
It is well-known that the presence of hydrophobic moieties
within a probe is one of the basic structural requirements for
ligand binding to human serum albumin (HSA).51 HSA is the
most abundant protein in blood, and binding to it results in an
increase in intravascular retention and relaxivity (due to a
decrease in τR of the complex).52 In view of the fact that the
rhodamine moiety has extended aromatic rings and hence
hydrophobicity, a study was carried out to investigate possible
interactions between Gd.L2 and HSA. The relaxation rates of a
series of solutions of Gd.L2 with increasing concentrations of
HSA were measured. The relaxation rate was found to remain
constant across the series (Figure 4), therefore indicating that
Therefore, when complexed with a luminescent lanthanide
ion, both ligands behave as dual-luminescent probes. In both
Tb.L1 and Tb.L2, excitation at 560 nm leads to emission from
the organic chromophore at 580 nm, with the fluorescence
originating from the xanthene core. Using time-gating
techniques, Tb(III) emission is induced in both complexes
upon excitation of the isolated phenyl ring of the rhodamine
moiety at 310 nm by energy transfer to the Tb(III) metal
center (Figures 2 and 3). In Tb.L1 only, Tb(III) emission can
be seen upon excititation at 310 nm without the use of a time
delay. The quantum yields of both terbium complexes were
determined, and the results were in agreement with the
observed spectra, confirming the efficient energy transfer to the
terbium metal center in Tb.L1 (Φ = 18.6%) and the inefficient
energy transfer in Tb.L2 (Φ = 0.8%).
The luminescence decays of the Tb(III) complexes in both
H2O and D2O were also measured (see the Supporting
Information). The luminescence lifetimes in D2O were
increased in comparison to H2O because less nonradiative
deactivation was induced by O−D vibrations than by O−H
vibrations.49 From these differences in the rates of energy
transfer to H2O and D2O oscillators, the hydration state can be
calculated using eq 1, in which the term 0.06 reflects the
Figure 4. Titration of Gd.L2 with HSA.
there is no significant binding of the probe to the protein. The
binding mechanisms of rhodamine B with bovine serum
albumin (BSA) were previously investigated, and site-selective
binding was observed. However, these results were obtained
using high molar ratios of rhodamine B to BSA (>50).53 Such
high concentrations are not practical for use of MRI probes in
vivo, and indeed, large changes in relaxivity have been observed
in cases where the molar ratio of probe to serum albumin have
been much lower.54,55 It is hypothesized that the steric bulk of
the complex is unsuitable for selective binding to HSA.
pH Titration. As previously discussed (vide supra), the
lactam ring of rhodamine can be intramolecularly opened and
closed via changes in pH.36 The pH sensitivity of Gd.L2 was
investigated, and to show the sensitivity of the proton-triggered
ring opening, a pH titration was carried out wherein the
emission intensity was monitored at various pH values (Figure
5). The pH titration showed that Gd.L2, which was
nonfluorescent at pH 6.5 or above, rearranged into its
fluorescent form at acidic pH. The fluorescence of Gd.L2 at
pH 4.3 was 50-fold brighter than that at pH 6.5 and 15-fold
brighter than that at pH 6.0, demonstrating that Gd.L2 is a
sensitive, acid-responsive probe capable of sensing small
changes in pH in the range 4−7.
qTb = 5(Δk − 0.06 − 0.01n)
(1)
quenching effect of unbound water molecules and n is the
number of coordinated secondary amide groups, reflecting the
quenching effect of the proximate NH oscillators.50 The
emission decays were fitted to monoexponential decays, and
the corresponding lifetime measurements of both terbium
complexes confirmed the expected coordination numbers of
seven for Tb.L1 and eight for Tb.L2 (Table 1).
Relaxometric Studies. T1 measurements of the Gd(III)
complexes were performed at 400 MHz (9.4 T, 25 °C), and
their r1 values were determined. The r1 values of 8.5 and 3.8
mM−1 s−1 obtained for complexes Gd.L1 and Gd.L2,
respectively, were within the range for Gd(III) chelates with
A titration curve was produced by plotting the emission
intensity at λem = 580 nm versus pH, which yielded a pKa value
of 5.11. It was hypothesized that the change in the charge of
Gd.L2 on going from pH ∼ 7.0 to 4.0 would aid the probe to
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Figure 5. Upper panel: pH-mediated ring opening of the intramolecular sprirolactam of Gd.L2, yielding a highly fluorescent species. Lower panels:
(a) pH-dependent emission spectra of Gd.L2 (λex = 560 nm, λem = 580 nm); (b) pH titration curve used to determine pKa.
localize in the more acidic microenvironment of tumor cells.56
It has been shown that the microenvironment of tumor cells is
more acidic than that of healthy tissues (Warburg effect). Upon
entering an acidic environment, the probe can undergo
rearrangement to its ring-opened form, which is fluorescent.
This cationic acid form of the probe would then be expected to
accumulate in the mitochondria.57
Images B, C and E, F in Figure 6 show clear accumulation of
Gd.L2 within each cell type. However, a clear colocalization of
Gd.L2 and MitoTracker was not evident (Figure 6C, box).
Moreover, we noted that in cells where significant uptake of
Gd.L2 was apparent (as indicated in the primary cells in D−F),
MitoTracker accumulation was lower and more diffuse than in
neighboring cells, and little, if any, colocalization of red and
green staining was evident (Figure 6F). These findings are
probably best explained by the fact that the Gd.L2 fluorescence,
which is strongly suppressed at pH >6.5 (Figure 5B), is only
readily detectable in cells in which mitochondria are relatively
depolarized, such that the mitochondrial matrix pH falls to
levels permissive of Gd.L2 fluorescence. Such mitochondria,
however, are expected to be poorly able to retain MitoTracker,
whose accumulation depends on the inner mitochondrial
membrane potential and the pH gradient (i.e., high intra-
mitochondrial pH >7). These results are thus consistent with
the known properties of both MitoTracker and Gd.L2 but are
unable to definitively establish a mitochondrial localization of
the latter.
BALB/c nude mice with neck M21 (human melanoma cell)
xenograft implantations were injected intravenously with 0.1
mmol kg−1 Gd.L2. The mice were imaged in a 4.7 T MRI
instrument at different times (preinjection, 30 min post
injection, and 60 min post injection) to evaluate the effect of
the contrast agent. Thirty minutes after the injection of Gd.L2,
a 36.5% decrease in tumor average T1 was observed compared
with the baseline of the preinjection image (Figure 7a,b). This
decrease in longitudinal relaxation time was retained at least up
In Vitro/in Vivo Studies. To test the viability of Gd.L2 as a
dual-modal agent for cancer imaging, in vivo and in vitro
studies were carried out. Because of a lack of water solubility,
no in vivo studies could be carried out on L1 complexes. The
addition of an extra amide functionality within L2 introduced
water solubility, and this allowed for the investigation of its
biological properties via in vitro and in vivo studies. Cell
penetration is crucial for the use of imaging probes in the
monitoring of biological mechanisms. To this end, preliminary
in vitro cell studies were carried out using Gd.L2. Rhodamine
has been widely used to determine the mitochondrial potential
in tumor cells.9 The negative mitochondrial potential is
reported to be greater in tumor cells than in normal cells,
and organic cations such as rhodamine have been shown to
accumulate in the energized mitochondria.57,58 With this in
mind, colocalization experiments were undertaken wherein a
known mitochondrial stain (MitoTracker Green FM) was
tested against Gd.L2. Confocal microscopy images of human
embryonic kidney (HEK293) cells and primary mouse islet
cells (∼70% insulin-containing β cells) were recorded after a 30
min incubation with Gd.L2 and MitoTracker Green (Figure 6).
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Figure 6. (A−C) Confocal microscopy images of HEK cells. Cells were incubated with 215 mM Gd.L2 and 100 nM MitoTracker Green for 30 min,
washed twice with KREBS buffer, and visualized under a fluorescence microscope. (A) MitoTracker Green signal. (B) Gd.L2 signal. (C) Merged
signals. Scale bar: 50 μm. (D−F) Confocal microscopy images of dissociated primary mouse islet cells. Cells were incubated with 215 mM Gd.L2
and 100 nM MitoTracker Green for 30 min, washed twice with KREBS buffer, and visualized under a fluorescence microscope. (D) MitoTracker
Green signal. (E) Gd.L2 signal. (F) Merged signals. Scale bar: 50 μm.
Figure 7. Parametric T1 maps of a BALB/c nude mouse tumor: (a) preinjection; (b) 30 min post injection with Gd.L2; (c) 60 min post injection
with Gd.L2. Scale bars in ms.
to 60 min (31.3% decrease; Figure 7c). These preliminary
images demonstrate that Gd.L2 functions successfully in vivo
by shortening the T1 of protons in the tumor sufficiently so as
to give a more detailed image. The observed accumulation of
Gd.L2 in the tumor area is consistent with the predicted effect
of a lower pH environment on the probe; the rearrangement of
the lactam ring at acidic pH introduces a positive charge in the
molecule that can promote the intracellular uptake of the
molecule responsible of the average T1 decrease observed in the
tumor.
of the sensitizer and linker design from the lanthanide metal
center on the efficiency of energy transfer. When complexed
with Gd(III), they form dual-modal MRI/fluorescent probes.
Ln.L2 is water-soluble and suitable for in vitro/in vivo studies,
and Gd.L2 has r1 = 3.84 mM−1 s−1 (9.4 T, 25 °C) and has been
shown to be viable both in vitro and in vivo. The pH sensitivity
of the probe means that it functions as an “off−on” luminescent
probe that is sensitive to the small pH changes within cells. By
coexpression with a known mitochondrial stain, the probe has
been hypothesized to localize in the mitochondria of HEK cells.
These studies demonstrate that the probe is cell-penetrating,
has a pH sensitivity to acidic microenvironments, and is a
tumor specific, dual-modal MRI/fluorescent contrast agent.
CONCLUSIONS
■
We have reported the preparation and full characterization of
two rhodamine−DO3A conjugate derivatives. The DO3A units
have been coordinated to paramagnetic Gd(III) and
luminescent Tb(III) centers. Complexation with Tb(III) gives
dual-luminescent probes with two different time scales, and
photophysical data collected also show the impact of distance
EXPERIMENTAL SECTION
■
Materials and Conditions. No special precautions were taken to
exclude air or moisture during reactions or workups, unless otherwise
stated. Products 259 and 460 were prepared via literature methods from
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commercially available starting materials. All other materials were
purchased from commercial suppliers and used without further
purification.
continuously using calipers, and tumor volumes (V) were calculated
using the equation V = πabc/6, where a, b, and c represent the lengths
of the three orthogonal axes of the tumor.
Instrumentation. 1H and 13C{1H} NMR spectra were recorded at
ambient temperature on a Bruker 400 MHz spectrometer and
internally referenced to the residual solvent peaks of CDCl3 at 7.26
ppm (1H) and 77.16 ppm (13C{1H}) or MeOD at 3.31 ppm (1H) and
49.15 ppm (13C{1H}). 13C{1H} spectra were fully assigned where
possible using 2D correlation spectroscopy. Mass spectrometry
analyses were conducted by the Mass Spectrometry Service, Imperial
College London. Microanalyses were carried out by Stephen Boyer of
the Science Centre, London Metropolitan University. ICP-MS analysis
was carried out by Stanislav Strekopytov of the Natural History
Museum.
In Vivo MRI. In vivo MRI was performed in a 4.7 T horizontal-bore
DirectDrive MRI system (Varian, Palo Alto, CA, USA) equipped with
40 G/cm actively shielded gradients (VnmrJ 3.1). For imaging of the
tumors and generation of the T1 parametric maps, a 45 mm diameter
saddle coil was used in transmit mode and a separate 20 mm diameter
stripline resonator was used as a surface coil to accommodate the
tumors.65 Imaging was performed with an inversion recovery (IR)
sequence and the following parameters: axial images; FOV = 35 × 35
mm; No slices = 14; slice thickness = 1 mm (zero slice gap); image
bandwidth = 156 kHz; TR = 4500 ms; TI = 6, 14.6, 35.4, 86.1, 209,
508, 1230, or 3000 ms; NA = 2; image matrix = 192 × 192; TA = 12
min. During imaging, animals were anesthetized with 2% isofluorane
and respiration and body temperature were monitored via SA
physiological monitoring systems (SA Instruments, Stony Brook,
NY, USA). Body temperature was monitored by a rectal probe and
Fluorescence Measurements. Absorption and fluorescence
spectra of all complexes in either aqueous or methanolic solutions
depending on solubility were obtained at room temperature on a
PerkinElmer Lambda 25 spectrometer and a Cary Varian luminescence
spectrometer (using SCAN for Windows), respectively. Samples were
held in 10 mm × 4 mm quartz Hellma cuvettes. Lifetimes were
measured by direct excitation of the sample at 325 nm with a 40 ms
pulse of light (50 pulses per point) followed by monitoring of the
integrated intensity of light emitted at 545 nm during a fixed gate time
of 0.1 ms at a delay time later. Delay times were set at 0.1 ms, covering
four or more lifetimes. Excitation and emission slits were set to 10:10
nm bandpass respectively. The obtained decay curves were fitted to a
simple monoexponential first-order decay curve using Microsoft Excel.
Fluoresence quantum yields of terbium complexes were determined
from the integrated fluorescence intensities of the complexes following
a previously reported method61 using the reference compound62
Na3[Tb(dpa)3] in water or methanol at 279 nm (quantum yield =
26.5% or 18.2%, respectively).
pH Titration. A 1.49 mM solution of Gd.L2 in 0.1 M NaCl (1 mL)
was prepared, from which 15 × 67 μL aliquots (microlitre pipet,
Eppendorf) were taken. Each aliquot was adjusted to the required pH
using a digital pH meter (Jenway 3510) equipped with a glass
electrode (Jenway 924005). The pH was monitored and adjusted to
acidic or basic conditions using small aliquots of a 1 M HCl or 1 M
NaOH solution, respectively. The pH was allowed to stabilize. The
solution was then added to a quartz cuvette for fluorescence analysis.
The fluorimeter was operated using an excitation slit width of 10 nm
and an emission slit width of 2.5 nm.
Relaxivity Measurements. Gd.L1 was dissolved in a methanol/
water (50:50) solvent mixture to give a solution of concentration 5
mM. Gd.L2 was dissolved in water to give a solution concentration of
5 mM. The resulting solutions were placed in 1.7 mm diameter
capillaries, which were sealed. The 1/T1 measurements were
performed on a Bruker Avance 400 spectrometer (400 MHz). These
experiments were carried out a minimum of three times, and the
relaxivity results given are averages of all experiments. The
concentration of Gd(III) was checked by the chemical shift
measurement of HOD induced by magnetic susceptibility.63
HSA Relaxivity Titration. A 1.49 mM solution of Gd.L2 in 0.1 M
NaCl (pH 7.4) was prepared, from which 10 × 67 μL aliquots were
taken. To each aliquot was added an aliquot of a 2 mM solution of
HSA (0.1 M NaCl, pH 7.4) to give 10 different sample concentrations
(0, 0.1, 0.25, 0.35, 0.5, 0.65, 0.85, 1, 1.5, and 2 equiv of HSA). The
solutions were made up to a volume of 100 μL by addition of water
where required. The resulting solutions were placed in 1.7 mm
diameter capillaries, which were sealed. The 1/T1 measurements were
performed on a Bruker Avance 400 spectrometer (400 MHz).
In Vivo Tumor Models. All of the animal experiments were
performed by licensed investigators in accordance with the United
Kingdom Home Office Guidance on the Operation of the Animal
(Scientific Procedures) Act 1986 and in keeping with the newly
published guidelines for the welfare and use of animals in cancer
research.64 Female BALB/c nude mice (aged 6−8 weeks; Harlan)
were used. M21 cells were injected subcutaneously on the back of mice
(5 × 106 cells in 100 μL of sterile PBS). Animals were used when the
xenografts reached ∼100 mm3. Tumor dimensions were measured
maintained at 35
1 °C. T1 maps were generated in ImageJ66 by
fitting the acquired data to the equation S = S0[1 − 2 exp(−TI/T1)],
where TI is the inversion time and T1 is the longitudinal relaxation
time. In order to assign the signal with the appropriate sign for the
fitting process, the χ2 value of the fit was evaluated during the fitting.
ROIs were then drawn on the T1 maps to delineate the whole tumor
with consideration to avoid the edges of the tumor and thus
contamination from neighboring tissues and partial volume effects.
In Vitro Cell Culture and Imaging. HEK293 cells were cultured
in DMEM (Lonza) supplemented with 10% fetal bovine serum (FBS)
(Lonza), 1% glutamine (Lonza), and 1% penicillin/streptomycin
(Gibco). On the day of visualization, cells were washed twice with
KREBS buffer (140 mM NaCl, 3.6 mM KCl, 0.5 mM NaH2PO4, 0.2
mM MgSO4, 1.5 mM CaCl2, 10 mM HEPES, pH 7.4, 2 mM
NaHCO3) pre-equilibrated with 95:5 O2/CO2 and containing 25 mM
glucose. Cells were then incubated with KREBS buffer containing 215
mM Gd.L2 and 100 nM MitoTracker Green for 30 min and washed
twice with KREBS buffer before visualization. Images were captured
using a Zeiss Axiovert confocal microscope coupled to a Nipkow
spinning-disk head (Yokogawa CSU-10) using a 63× oil objective. A
solid-state laser (CrystaLaser) controlled by a laser-merge module
(Spectral Applied Physics) provided wavelengths of 491 nm to excite
MitoTracker Green (emission filtered at 525 nm) and 561 nm to
excite Gd.L2 (emission filtered at 620 nm). Images were captured by a
highly sensitive 16-bit, 512 × 512 pixel back-illuminated EM-CCD
camera (ImageEM 9100-13, Hamamatsu). Volocity software (Perki-
nElmer) provided the user interface.
In Vivo Tissue Isolation and Imaging. Islet isolation was carried
out as shown in ref 67. In short, a female CD1 mouse (8−12 weeks
old) was sacrificed by cervical dislocation. The pancreas was injected
with 1 mg/mL collagenase solution through the bile duct and excised.
The pancreas was digested by heating with collagenase solution, and
islets were isolated through the use of ficoll gradient centrifugation.
Islets were hand-picked and kept at 37 °C in Islet medium (RPMI
supplemented with 10% FBS and 1% penicillin/streptomycin). Islets
were dispersed into clusters and single cells according to the procedure
in ref 67. Briefly, islets were incubated with Hank’s based enzyme-free
dissociation buffer (Invitrogen, Paisley, UK) and centrifuged. All but 1
mL of the supernatant was removed, and the pellet was resuspended
vigorously via pipetting. The cells were washed and resuspended with
RPMI 1640 and then loaded onto a poly-L-lysine-treated coverslip. On
the day of visualization, cells were washed twice with KREBS buffer
(140 mM NaCl, 3.6 mM KCl, 0.5 mM NaH2PO4, 0.2 mM MgSO4, 1.5
mM CaCl2, 10 mM HEPES, pH 7.4, 2 mM NaHCO3) pre-equilibrated
with 95:5 O2/CO2 and containing 25 mM glucose. Cells were then
incubated with KREBS buffer containing 215 mM Gd.L2 and 100 nM
MitoTracker Green for 30 min and washed twice with KREBS buffer
before visualization. Images were captured using a Zeiss Axiovert
confocal microscope coupled to a Nipkow spinning-disk head
(Yokogawa CSU-10) using a 63× oil objective. A solid-state laser
(CrystaLaser) controlled by a laser-merge module (Spectral Applied
Physics) provided wavelengths of 491 nm to excite MitoTracker Green
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Inorganic Chemistry
Article
(emission filtered at 525 nm) and 561 nm to excite Gd.L2 (emission
filtered at 620 nm). Images were captured by a highly sensitive 16-bit,
512 × 512 pixel back-illuminated EM-CCD camera (ImageEM 9100-
13, Hamamatsu). Volocity software (PerkinElmer) provided the user
interface.
solid (0.12 g, 21%). 1H NMR (400 MHz, CDCl3): δ 1.18 (t, 12H, 3JHH
= 7.0 Hz), 1.40 (s, 27H), 1.98−2.92 (m, 28H), 3.28 (q, 8H, 3JHH = 7.1
Hz), 6.24−6.41 (m, 6H), 7.01−7.04 (m, 1H), 7.35−7.45 (m, 2H),
7.76−7.83 (m, 1H). 13C NMR (100 MHz, CDCl3): δ 12.53, 27.86,
30.94, 40.00, 40.55, 44.31, 49.01, 50.27, 51.13, 55.66, 56.22, 65.43,
81.52, 97.68, 104.84, 108.17, 122.73, 123.84, 128.38, 130.05, 132.84,
148.8, 153.18, 169.63, 170.58, 171.01, 172.31. ESI-HRMS: calcd for
C58H87N8O9 1039.6596, found m/z 1039.6676.
Synthesis of 3. DO3A-t-Bu-ester (0.16 g, 0.28 mmol) and K2CO3
(0.05 g, 0.34 mmol) were dissolved in MeCN (5 mL), and the mixture
was stirred for 5 min. 2 (0.15 g, 0.28 mmol) in MeCN (15 mL) was
added dropwise. The reaction mixture was heated to reflux
temperatures for 24 h. The resulting solution was filtered, and the
filtrate was concentrated under reduced pressure. The crude product
was purified by silica gel chromatography with 1−20% MeOH/DCM,
1% NH3 solution (28%), affording 3 as a beige solid (0.21 g, 37%). 1H
NMR (400 MHz, CDCl3): δ 1.18 (t, 12H, 3JHH = 7.1 Hz), 1.43 (s, 27
H), 2.09−2.15 (m, 2H), 2.29−3.06 (m, 16H), 3.13−3.19 (m, 2H),
Synthesis of L2. 6 (0.30 g, 0.3 mmol) was stirred in CH2Cl2 (1
mL), and trifluoroacetic acid (1 mL, excess) was added dropwise. The
solution was stirred at room temperature, open to air, for 24 h. The
solvents were then removed in vacuo, and the residue was redissolved
in CH2Cl2. This was again removed, and diethyl ether was added. After
removal of diethyl ether, this process was repeated, affording L2 as a
1
pink solid (0.23 g, 92%). H NMR (400 MHz, MeOD): δ 1.15 (t,
12H, 3JHH = 6.0 Hz), 2.84 (t, 2H, 3JHH = 7.0 Hz), 2.97−3.95 (m, 26H),
3
3.36 (q, 8H, JHH = 7.4 Hz), 3.50 (s, 6H), 6.23−6.43 (m, 6H), 7.04−
3
7.08 (m, 1H), 7.41−7.47 (m, 2H), 7.81−7.84 (m, 1H). 13C NMR
(100 MHz, CDCl3): δ 12.6, 28.0, 30.9, 36.0, 44.3, 46.2, 49.6, 51.0,
53.5, 55.8, 56.4, 65.2, 82.0, 82.4, 97.9, 105.1, 107.8, 122.6, 123.8, 128.0,
128.7, 131.0, 132.4, 148.8, 153.4, 153.5, 168.4, 172.5. ESI-HRMS:
calcd for C56H83N7O8 982.6381, found m/z 982.6398.
3.54 (q, 8H, JHH = 6.0 Hz), 6.60−6.66 (m, 6H), 7.11 (d, 1H), 7.59
(m, 2H), 7.90 (d, 1H). 13C NMR (100 MHz, CDCl3): δ 10.88, 38.29,
39.01, 45.44, 46.52, 50.41, 53.10, 110.98, 115.25, 122.42, 123.66,
128.55, 128.91, 133.06, 153.05, 161.17, 161.51, 169.20. ESI-HRMS:
calcd for C46H63N8O9 871.4702, found m/z 871.4718.
Synthesis of Gd.L2. L2 (0.19 g, 0.22 mmol) and GdCl3·6H2O
(0.04 g, 0.11 mmol) were dissolved in water (2 mL), and the pH was
adjusted to 5.5 using 1 M NaOH(aq). The reaction was left stirring at
room temperature overnight. The solution was removed in vacuo until
a small volume was left. This crude solution was purified using
Sephadex G10 chromotography, affording Gd.L2 as a pale-beige solid
(0.03 g, 27%). ESI-HRMS: calcd for C46H60GdN8O9 1026.3724, found
m/z 1026.3834.
Synthesis of Tb.L2. The complex was prepared by an method
analogous to that for Gd.L2 using L2 (0.19 g, 0.21 mmol) and TbCl3·
6H2O (0.04 g, 0.11 mmol) to yield Tb.L2 as a pale-beige solid (0.03 g,
27%). ESI-HRMS (MeOH): calcd for C46H60N8O9Tb 1027.3737,
found m/z 1027.3805.
Synthesis of L1. 3 (0.20 g, 1 mmol) was dissolved in CH2Cl2 (2
mL), and trifluoroacetic acid (2 mL) was added dropwise. The
solution was stirred at room temperature, open to air, for 24 h. The
solvents were removed in vacuo, and the residue was redissolved in
CH2Cl2. This was repeated with diethyl ether and again with CH2Cl2
1
until L1 was obtained as a bright-pink solid (0.16 g, 96%). H NMR
3
(400 MHz, CDCl3): δ 1.17 (t, 12H, JHH = 7.2 Hz), 3.06−3.71 (m,
24H), 4.20 (m, 2H), 6.54−6.75 (m, 6H), 7.14 (d, 1H), 7.60−7.64 (m,
2H), 7.80 (d, 1H). 13C NMR (100 MHz, CDCl3): δ 11.06, 34.14,
45.42, 45.63, 48.18, 49.33, 51.97, 52.26, 54.41, 110.36, 122.90, 124.22,
126.14, 128.62, 128.82, 129.83, 133.57, 153.00, 153.35, 160.78, 169.12.
ESI-HRMS: calcd for C44H60N7O8 814.4503, found m/z 814.4503.
Synthesis of Gd.L1. L1 (0.04 g, 0.05 mmol) and GdCl3·6H2O
(0.01 g, 0.03 mmol) were dissolved in water (2 mL), and the pH was
adjusted to 5.5 using 1 M NaOH solution. The mixture was stirred at
room temperature for 24 h. The reaction precipitate was filtered and
then dissolved in DCM. The organic layer was washed with water and
then dried over MgSO4. Addition of ether to this solution precipitated
Gd.L1 as a pale-beige powder (0.022 g, 84%). ESI-HRMS: calcd for
C44H57N7O8Gd 969.3510, found m/z 969.3569.
ASSOCIATED CONTENT
■
S
* Supporting Information
Luminescence decay data for Tb.L1 and Tb.L2. This material is
available free of charge via the Internet at http://pubs.acs.org.
Synthesis of Tb.L1. The complex was prepared by an method
analogous to that for Gd.L1 using L1 (0.26 g, 0.32 mmol) and TbCl3·
6H2O (0.06 g, 0.16 mmol) to yield Tb.L1 as a pale-beige solid (0.12 g,
80%). ESI-HRMS: calcd for C44H57N7O8Tb 970.3522, found m/z
970.3555.
AUTHOR INFORMATION
■
Corresponding Author
*E-mail: n.long@imperial.ac.uk.
Synthesis of 5. 4 (0.50 g, 1.03 mmol) was dissolved in DCM (10
mL), and NEt3 (0.1 mL, 1.24 mmol) was added. The reaction mixture
was cooled to 0 °C using an ice bath, and chloroacetyl chloride (0.1
mL, 1.24 mmol) in DCM (5 mL) was then added dropwise. The
reaction mixture was allowed to come to room temperature. The
solution was stirred at room temperature for 5 h, and then the solvent
was removed under reduced pressure. The resultant solid was
dissolved in DCM (20 mL) and washed with water (10 mL × 2)
and brine. The organic extract was dried over MgSO4 and evaporated
to give 5 as a purple solid (0.36 g, 62%). 1H NMR (400 MHz,
CDCl3): δ 1.18 (t, 12H, JHH = 7.0 Hz), 3.10 (t, 2H, JHH = 7.1 Hz),
3.34 (q, 10H, 3JHH = 7.2 Hz), 3.95 (s, 2H), 6.31−6.47 (m, 6H), 7.08−
7.11 (m, 1H), 7.46−7.48 (m, 2H), 7.93−7.96 (m, 1H). 13C NMR
(100 MHz, CDCl3): δ 12.42, 39.50, 40.39, 41.43, 42.42, 45.01, 65.65,
98.87, 108.96, 123.08, 123.88, 128.41, 128.53, 130.22, 133.03, 148.26,
153.17, 153.55, 166.80, 169.46, 169.85. ESI-HRMS: calcd for
C32H38ClN4O3 561.2632, found m/z 561.2646.
Synthesis of 6. Tri-tert-butyl-DO3A (0.32 g, 0.59 mmol) and
K2CO3 (0.08 g, 0.59 mmol) were dissolved in MeCN (20 mL) and left
to stir for 5 min. 5 in MeCN (20 mL) was then added dropwise. The
reaction mixture was heated to reflux for 24 h. The resulting solution
was filtered, and the filtrate was concentrated under reduced pressure.
The crude product was purified by silica gel chromatography using 1−
20% MeOH/DCM, 1% NH3 solution (33%), affording 6 as a beige
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
N.J.L. and G.A.R. were supported by a grant from the
Wellcome Trust Institutional Strategic Support Fund. G.A.R.
was supported by Wellcome Trust Senior Investigator
(WT098424AIA), MRC Programme (MR/J0003042/1), and
Diabetes UK Project (11/0004210) grants and is the recipient
of a Royal Society Wolfson Research Merit Award. The work
leading to this publication received support from the Innovative
Medicines Initiative Joint Undertaking under Grant Agreement
155005 (IMIDIA), resources of which are composed of
financial contributions from the European Union’s Seventh
Framework Programme (FP7/2007-2013) and EFPIA compa-
nies’ in-kind contribution (G.A.R.). We also thank Imperial
College London for funding, Dr. Pauline Chabosseau (Cell
Biology) for kind assistance with single cell imaging, Dr. Israt
Alam (Surgery and Cancer) for assistance with mice work, and
Dr. Ioannis Lavdas (Surgery and Cancer) for MRI coil setup
and expertise.
3
3
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(37) Best, Q. A.; Xu, R.; McCarroll, M. E.; Wang, L.; Dyer, D. J. Org.
Lett. 2010, 12, 3219−3221.
REFERENCES
■
(1) Aime, S.; Crich, S. G.; Gianolino, E.; Giovenzana, G. B.; Tei, L.;
(38) Lee, M. H.; Han, J. H.; Lee, J. H.; Park, N.; Kumar, R.; Kang, C.;
Kim, J. S. Angew. Chem., Int. Ed. 2013, 52, 6206−6209.
(39) Jennings, L. E.; Long, N. J. Chem. Commun. 2009, 3511−3524.
(40) Stasiuk, G. J.; Long, N. J. Chem. Commun. 2013, 49, 2732−2746.
(41) Guo, K.; Berezin, M. Y.; Zheng, J.; Akers, W.; Lin, F.; Teng, B.;
Vasalatiy, O.; Gandjbakhche, A.; Griffiths, G. L.; Achilefu, S. Chem.
Commun. 2010, 46, 3705−3707.
Terreno, E. Coord. Chem. Rev. 2006, 250, 1562−1579.
́
́
h, E.; Helm, L.; Merbach, A. E. Top. Curr. Chem. 2002, 221,
(2) Tot
61−101.
(3) Frullano, L.; Meade, T. J. J. Biol. Inorg. Chem. 2007, 12, 939−949.
(4) Bunzli, J.-C. G.; Piguet, C. Chem. Soc. Rev. 2005, 34, 1048−1077.
̈
(5) Montgomery, C. P.; Murray, B. S.; New, E. J.; Pal, R.; Parker, D.
Acc. Chem. Res. 2009, 42, 925−937.
(42) Hao, D.; Ai, T.; Goerner, F.; Hu, X.; Runge, V. M.; Tweedle, M.
J. Magn. Reson. Imaging 2012, 36, 1060−1071.
(6) Faulkner, S.; Burton-Pye, B. P.; Pope, S. J. A. Appl. Spectrosc. Rev.
2005, 40, 1−39.
(43) Huber, M. M.; Staubli, A. B.; Kustedjo, K.; Gray, M. H. B.; Shih,
̈
(7) Manella, C. Biochim. Biophys. Acta 2006, 1762, 140−147.
(8) David, S.; Weiss, M. J.; Wong, J. R.; Lampidis, T. J.; Chen, L. B. J.
Biol. Chem. 1985, 260, 13844−13850.
J.; Fraser, S. E.; Jacobs, R. E.; Meade, T. J. Bioconjugate Chem. 1998, 9,
242−249.
(44) Yamane, T.; Hanaoka, K.; Muramatsu, Y.; Tamura, K.; Adachi,
Y.; Miyashita, Y.; Hirata, Y.; Nagano, T. Bioconjugate Chem. 2011, 22,
2227−2236.
(9) Dairkee, S. H.; Hackett, A. J. Breast Cancer Res. Treat. 1991, 18,
57−61.
(10) Duchen, M. R. Mol. Aspects Med. 2004, 25, 365−451.
(11) Caravan, P.; Ellison, J. J.; McMurry, T. J.; Lauffer, R. B. Chem.
Rev. 1999, 99, 2293−2352.
(45) Yan, X.; Zhou, Y.; Liu, S. Theranostics 2012, 2, 988−998.
(46) Barge, A.; Cravotto, G.; Gianolio, E.; Fedeli, F. Contrast Med.
Mol. Imaging 2006, 1, 184−188.
(12) Caravan, P. Chem. Soc. Rev. 2006, 35, 512−523.
(13) Weissleder, R.; Ross, B.; Rehemtulla, A.; Gambhir, S. Molecular
Imaging: Principles and Practice; PMPH-USA: Shelton, CT, 2010.
(14) Comblin, V.; Gilsoul, D.; Hermann, M.; Humblet, V.; Jacques,
V.; Mesbahi, M.; Sauvage, C.; Desreux, J. Coord. Chem. Rev. 1999, 185,
451−470.
(47) Beija, M.; Afonso, C. A. M.; Martinho, J. M. G. Chem. Soc. Rev.
2009, 38, 2410−2433.
(48) Beeby, A.; Clarkson, I. M.; Dickins, R. S.; Faulkner, S.; Parker,
D.; Royle, L.; de Sousa, A. S.; Williams, J. A. G.; Woods, M. J. Chem.
Soc., Perkin Trans. 2 1999, 493−503.
(49) Bruce, J. A.; Lowe, M. P.; Parker, D. Photophysiscal Aspects of
Lanthanide(III) Complexes: The Chemistry of Contrast Agents in Medical
Magnetic Resonance Imaging; Wiley; West Sussex, U.K., 2001.
(50) Horrocks, W. D.; Sudnick, D. R. J. Am. Chem. Soc. 1979, 101,
334−340.
(15) Esqueda, A. C.; Lop
́
ez, J. A.; Andreu-de-Riquer, G.; Alvarado-
Monzon, J. C.; Ratnaker, J.; Lubag, A. J. M.; Sherry, A. D.; De Leon-
́
́
Rodríguez, L. M. J. Am. Chem. Soc. 2009, 131, 11387−11391.
(16) Polasek, M.; Caravan, P. Inorg. Chem. 2013, 52, 4084−4096.
(17) Reguerio-Figueroa, M.; Bensenane, B.; Ruscsak
́
, E.; Esteban-
(51) Fasano, M.; Curry, S.; Terreno, E.; Galliano, M.; Fanali, G.;
Narciso, P.; Notari, S.; Ascenzi, P. IUBMB Life 2005, 57, 787−796.
(52) Boros, E.; Caravan, P. J. Med. Chem. 2013, 56, 1782−1786.
(53) Cai, H.-H.; Zhong, X.; Yang, P.-H.; Wei, W.; Chen, J.; Cai, J.
Colloids Surf., A 2010, 372, 35−40.
(54) Caravan, P. Acc. Chem. Res. 2009, 42, 851−862.
(55) Caravan, P.; Zhang, Z. Eur. J. Inorg. Chem. 2012, 1916−1923.
(56) Estrella, V.; Chen, T.; Lloyd, M.; Wotjkowiak, J.; Cornnell, H.
H.; Ibrahim-Hashim, A.; Bailey, K.; Balagurunathan, Y.; Rothberg, J.
M.; Sloane, B. F.; Johnson, J.; Gatenby, R. A.; Gillies, R. J. Cancer Res.
2013, 73, 1524−1535.
(57) Zhao, Z.; Chan, P.-S.; Li, H.; Wong, K.-L.; Wong, R. N. S.; Mak,
N.-K.; Zhang, J.; Tam, H.-L.; Wong, W.-Y.; Kwong, D. W. J.; Wong,
W.-K. Inorg. Chem. 2012, 51, 812−821.
(58) Reungpatthanaphong, P.; Dechsupa, S.; Meesungnoen, J.;
Loetchutinat, C.; Mankhetkorn, S. J. Biochem. Biophys. Methods
2003, 57, 1−16.
(59) Shiraishi, Y.; Sumiya, S.; Kohno, Y.; Hirai, T. J. Org. Chem. 2008,
73, 8571−8574.
(60) Zhang, X.; Shiraishi, Y.; Hirai, T. Org. Lett. 2007, 9, 5039−5042.
(61) Zheng, Y.; Yang, Z.; Ying, J. Y. Adv. Mater. 2007, 19, 1475−
1479.
Gomez, D.; Charbonniere, L. J.; Tircso, G.; Toth, I.; de Blas, A.;
́
̀
́
́
Rodríguez-Blas, T.; Platas-Iglesias, C. Inorg. Chem. 2011, 50, 4125−
4141.
(18) Placidi, M. P.; Engelmann, J.; Natarajan, L. S.; Logothetis, N. K.;
Angelovski, G. Chem. Commun. 2011, 47, 11534−11536.
(19) Faulkner, S.; Pope, S. J. A.; Burton-Pye, B. P. Appl. Spectrosc. Rev.
2005, 40, 1−39.
(20) Parker, D.; Dickins, R. S.; Puschmann, H.; Crossland, C.;
Howard, J. A. K. Chem. Rev. 2002, 102, 1977−2010.
(21) Murray, B. S.; New, E. J.; Pal, R.; Parker, D. Org. Biomol. Chem.
2008, 6, 2085−2094.
(22) Atkins, P.; de Paula, J. Physical Chemistry, 8th ed.; Oxford
University Press: Oxford, U.K., 2006.
(23) Bunzli, J.-C. G. Acc. Chem. Res. 2006, 39, 53−61.
̈
(24) Gonca̧ lves, M. S. T. Chem. Rev. 2009, 109, 190−212.
(25) Yuan, L.; Lin, W.; Zheng, K.; He, L.; Huang, W. Chem. Soc. Rev.
2013, 42, 622−661.
(26) Suksai, C.; Tuntulani, T. Chem. Soc. Rev. 2003, 32, 192−202.
(27) Bandichlov, R.; Petrescu, A. D.; Vespa, A.; Kier, A. B.;
Schroeder, F.; Burgess, K. Bioconjugate Chem. 2006, 17, 1219−1225.
(28) Zhou, Y.; Kim, Y.-S.; Yan, X.; Jacobson, O.; Chen, X.; Liu, S.
Mol. Pharmaceutics 2011, 8, 1198−1209.
(62) Chauvin, A.-S.; Gumy, F.; Imbert, D.; Bunzli, J.-C. G. Spectrosc.
̈
(29) You, Y.; Tomat, E.; Hwang, K.; Atanasijevic, T.; Nam, W.;
Jasanoff, A. P.; Lippard, S. J. Chem. Commun. 2010, 46, 4139−4141.
(30) Scadutro, R. C., Jr.; Grotyohann, L. W. Biophys. J. 1999, 76,
469−477.
Lett. 2004, 37, 517−532.
(63) Corsi, D. M.; Platas-Iglesias, C.; van Bekum, H.; Peters, J. A.
Magn. Reson. Chem. 2001, 39, 723−726.
(64) Workman, P.; Aboagye, E. O.; Balkwill, F.; Balmain, A.; Bruder,
G.; Chaplin, D. J.; Double, J. A.; Everitt, J.; Farningham, D. A. H.;
Glennie, M. J.; Kelland, L. R.; Robinson, V.; Stratford, I. J.; Tozer, G.
M.; Watson, S.; Wedge, S. R.; Eccles, S. A. Br. J. Cancer 2010, 102,
1555−1577.
(65) Lavdas, I.; Seton, H. C.; Harrington, C. R.; Wischik, C. M.
Magn. Reson. Mater. Phys. 2011, 24, 331−337.
(66) Rasband, W. S. ImageJ; U.S. National Institutes of Health:
Bethesda, MD, 1997−2012; http://rsb.info.nih.gov/ij/.
(67) Ravier, M. A.; Rutter, G. A. Methods Mol. Biol. 2010, 633, 171−
184.
(31) Longmuir, M. R.; Ogawa, M.; Hana, Y.; Kosaka, N.; Regino, C.
A. S.; Choyke, P. L.; Kobayashi, H. Bioconjugate Chem. 2008, 19,
1735−1742.
(32) Lavis, L. D.; Chao, T.-Y.; Raines, R. T. ACS Chem. Biol. 2006, 1,
252−260.
(33) Hockings, P. D.; Rogers, P. J. Biochim. Biophys. Acta 1996, 1282,
101−106.
(34) Huang, S. G. J. Biomol. Screening 2002, 7, 383−389.
(35) Lefevre, C.; Kang, H. C.; Haugland, R. P.; Malakzadeh, N.;
Arttamangkul, S.; Haugland, R. P. Bioconjugate Chem. 1996, 7, 482−
489.
(36) Chen, X.; Pradhan, T.; Wang, F.; Kim, J. S.; Yoon, J. Chem. Rev.
2012, 112, 1910−1956.
14293
dx.doi.org/10.1021/ic402233g | Inorg. Chem. 2013, 52, 14284−14293