Organelle-specific detection of phosphatase activities with two-photon fluorogenic probes in cells and tissues

Article abstract of DOI:10.1021/ja3036256

Two-photon fluorescence microscopy (TPFM) provides key advantages over conventional fluorescence imaging techniques, namely, increased penetration depth, lower tissue autofluorescence and self-absorption, and reduced photodamage and photobleaching and therefore is particularly useful for imaging deep tissues and animals. Enzyme-detecting, small molecule probes provide powerful alternatives over conventional fluorescent protein (FP)-based methods in bioimaging, primarily due to their favorable photophysical properties, cell permeability, and chemical tractability. In this article, we report the first fluorogenic, small molecule reporter system (Y2/Y1) capable of imaging endogenous phosphatase activities in both live mammalian cells and Drosophila brains. The one- and two-photon excited photophysical properties of the system were thoroughly investigated, thus confirming the system was indeed a suitable Turn-ON fluorescence pair for TPFM. To our knowledge, this is the first enzyme reporting two-photon fluorescence bioimaging system which was designed exclusively from a centrosymmetric dye possessing desirable two-photon properties. By conjugation of our reporter system to different cell-penetrating peptides (CPPs), we were able to achieve organelle- and tumor cell-specific imaging of phosphatase activities with good spatial and temporal resolution. The diffusion problem typically associated with most small molecule imaging probes was effectively abrogated. We further demonstrated this novel two-photon system could be used for imaging endogenous phosphatase activities in Drosophila brains with a detection depth of >100 μm.

Full text of DOI:10.1021/ja3036256

Article
pubs.acs.org/JACS
Organelle-Specific Detection of Phosphatase Activities with Two-
Photon Fluorogenic Probes in Cells and Tissues
Lin Li, Jingyan Ge, Hao Wu, Qing-Hua Xu, and Shao Q. Yao*
Department of Chemistry, National University of Singapore, Singapore 117543
S
* Supporting Information
ABSTRACT: Two-photon fluorescence microscopy (TPFM)
provides key advantages over conventional fluorescence
imaging techniques, namely, increased penetration depth,
lower tissue autofluorescence and self-absorption, and reduced
photodamage and photobleaching and therefore is particularly
useful for imaging deep tissues and animals. Enzyme-detecting,
small molecule probes provide powerful alternatives over
conventional fluorescent protein (FP)-based methods in
bioimaging, primarily due to their favorable photophysical
properties, cell permeability, and chemical tractability. In this
article, we report the first fluorogenic, small molecule reporter
system (Y2/Y1) capable of imaging endogenous phosphatase
activities in both live mammalian cells and Drosophila brains.
The one- and two-photon excited photophysical properties of the system were thoroughly investigated, thus confirming the
system was indeed a suitable Turn-ON fluorescence pair for TPFM. To our knowledge, this is the first enzyme reporting two-
photon fluorescence bioimaging system which was designed exclusively from a centrosymmetric dye possessing desirable two-
photon properties. By conjugation of our reporter system to different cell-penetrating peptides (CPPs), we were able to achieve
organelle- and tumor cell-specific imaging of phosphatase activities with good spatial and temporal resolution. The diffusion
problem typically associated with most small molecule imaging probes was effectively abrogated. We further demonstrated this
novel two-photon system could be used for imaging endogenous phosphatase activities in Drosophila brains with a detection
depth of >100 μm.
polycythemia vera, primary myelofibrosis, and chronic/accute
chronic myelogenous leukemia.4b Consequently, biological and
chemical approaches capable of reporting phosphatase
expression, and more importantly their activities, would provide
invaluable insights into how these enzymes work under
physiological settings and how they could be modulated
therapeutically.5−8 Of particular significance are probes suitable
for live-cell and deep-tissue imaging.9,10 Existing methods in
phosphatase biology, such as those based on gene knockout or
overexpression techniques, do not provide an adequate view of
the temporal, spatial, and dynamic properties of phosphatases
in cells.5 Emerging tools have begun to address some of these
issues by focusing on the detection of phosphatase activities at
the proteome level or in living cells.6−8 Among them, imaging-
based approaches are highly desirable due to their high
sensitivity and good resolution.9 Bastiaen and co-workers
INTRODUCTION
■
Phosphatases are a large and structurally diverse class of
signaling enzymes that remove the phosphate group from their
protein and nonprotein substrates. Defective regulation of these
enzymes has significant implications in human diseases.1 Of the
various types of phosphatases, protein phosphatases (PPs) are
the most common and important as they control the reversible
phosphorylation/dephosphorylation process in up to 30% of all
human proteins. There are two major classes of PPs, namely
protein tyrosine phosphatases (PTPs) and serine/threonine
phosphatases. It is well documented that elevated levels of
endogenous PP activities (especially those of PTPs) are
associated with tumorigenesis in numerous cells and tissues.2
Acid and alkaline phosphatases (abbreviated as ACPs and
ALPs, respectively) are the other two types of phosphatases
that are present in most human organs and tissues, and their
levels of expression have been routinely used for disease
diagnosis.3,4 For example, the serum level of certain ACPs is
used to evaluate the success of the surgical treatment of
prostate cancer.3 Since ALP is a byproduct of osteoblast
activity, elevated ALP has been used to diagnose bond-forming
diseases such as Paget’s disease.4a The ALP found within white
blood cells, commonly referred to as Leukocyte alkaline
phosphatase (LAP), is used in diagnosis of conditions such as
developed the first PTP imaging approach based on Forster
̈
resonance energy transfer (FRET).7 The method relies on
spatial resolution of the enzyme−substrate (ES) interaction
between a PTP genetically fused to a fluorescent protein (FP)
donor and a dye-labeled, synthetic phosphopeptide acceptor,
Received: April 16, 2012
Published: June 26, 2012
© 2012 American Chemical Society
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Figure 1. Two practical strategies for generation of two-photon, enzyme-detecting small molecule probes. (A) The FRET-based strategy recently
developed in our previously work.17 (B) The two-photon fluorogenic approach presented in the current study, which necessitates the discovery of a
centrosymmetric dye with detectable two-photon excited fluorescence. (C) Commercially available 1P fluorogenic phosphatase reporters (left), all of
which have undesirable 2P properties, and the 2P fluorogenic phosphatase reporter developed in the current study (Y2; right). Upon removal of the
phosphate groups in Y2 by a phosphatase, the highly fluorescent Y1, which possesses good 2P photophysical properties, is generated. Y1 was
designed based on a previously known 2P centrosymmetric dye, Y1*.19
and therefore was able to indirectly monitor PTP activities in
cells. The need to deliver the peptide into cells by
microinjection, however, makes this strategy technically
demanding. A genetically encoded calcineurin (a serine/
threonine phosphatase) biosensor was recently reported,8 but
similarly to other macromolecular imaging probes based on
FRET and protein conformation changes, it offers limited
utilities as a general phosphatase imaging probe.11 Small
molecule-based probes provide a powerful tool for live-cell and
tissue imaging, primarily due to their desirable properties,
including permeability (due to their small sizes), photophysical
properties (e.g., good photo stability), chemical tractability
(e.g., different molecular structures/designs can be installed),
and amenability to a wide variety of biological events.10 Of the
various types of small molecule probes, those capable of
detecting enzymatic activities (either in vitro or in situ) are
typically designed based on either the FRET (Figure 1A) or the
fluorogenic approach (Figure 1B), but most of them can readily
diffuse away from the site of the enzymatic reaction in situ, thus
providing limited utilities for subcellular imaging.12 Several
fluorogenic substrates of phosphatases, which are based on
well-known small organic fluorophores such as coumarin,
fluorescein, and their analogues, are available (Figure 1C; left),
and have been widely used in vitro for general phosphatase
detection.13 These phosphatase probes, due the presence of the
negatively charged phosphate group in the reporters, are not
cell-permeable, but may be made so by “caging” the phosphate
with a removable 2-nitrobenzyl protecting group.14 Some of
them have even been developed into magnetic resonance
Imaging (MRI)-based agents for detection of ALP activities in
bone tissues.15 Notwithstanding, the diffusible nature of these
small molecule probes still poses a serious challenge for cell-
based imaging where sufficient spatial resolutions are needed.12
The commercially available product ELF 97 provides one such
exception (Figure 1C; left). Upon enzymatic removal of the
phosphate group, the compound forms a bright yellow-green
fluorescent precipitate at the site of reaction and thus is able to
detect phosphatase activities in cells with sufficient spatial
resolution. ELF 97, however, is not cell-permeable and
therefore can only be used in vitro or in fixed cells. Innovative
designs in several newly emerging, enzyme-detecting small
molecule probes have begun to address this diffusion
problem.16−18 Bogyo and co-workers recently developed
quenched activity-based probes (qABPs) to monitor real-time
protease activities in live human cells. By using acyloxymethyl
ketone (AOMK)-based probes which emit a strong fluorescent
signal only after proteolysis by a cysteine protease target, with
the simultaneous formation of a covalent probe enzyme adduct,
this strategy couples imaging and self-immobilization within the
same design to achieve the intended “Turn-ON” imaging
capability for cysteine proteases in vivo with good spatial
resolution.16 The design, however, was not easily amenable to
imaging most other types of enzymatic activities (e.g.,
phosphatases). On the basis of a similar concept, we recently
developed the first small molecule probe capable of detecting
endogenous phosphatase activities with acceptable spatial
resolution in live cells (Figure 1A);17 our method incorporated
a quinone methide system into the FRET-based approach
(Figure 1A) with a two-photon (2P) dye, 2-hydroxy-4,6-bis(4-
(diethylamino)styryl)pyrimidine (Y1*; Figure 1C; right),19 and
a quencher, 4-(4-dimethylaminophenyl)diazenylbenzoic acid
(Dabcyl). Upon enzymatic phosphate removal, the quencher
was released from the probe, and at the same time a highly
reactive quinone methide intermediate was generated which
subsequently “fixed” the fluorescent adduct near the site of the
reaction, thus achieving good spatial resolution. The need for
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Figure 2. Overall strategy of the four CPP-conjugated 2P fluorogenic substrates, YP1 to YP4. They were used for subcellular bioimaging of
endogenous phosphatase activities in mammalian cells and Drosophila brains. The 2P reporter in all four conjugates was in the “caged” form (similar
to Y3, where R = H). Sequences of the four CPPs (P1 to P4) were shown by their single-letter amino acid alphabets. Unnatural AAs were
represented as Fx = L-cyclohexylalanine; r = D-Arg; e = D-Glu; U = succinic acid. The full structure of YP4 is shown, with the negatively charged D-
Glu/succinic acid and the positively charged D-Arg highlighted in red and blue, respectively. MMP cleavage site in YP4 is indicated. In the tumor
cell-specific approach, membrane-bound MMPs first cleaved the masking “red” peptide fragment (Step 1), exposing the “blue” CPP and delivering
the probe into the cells (Step 2). Subsequent UV irradiation (Step 3) released the phosphatase-responsive probe Y2, which was dephosphorylated by
endogenous PPs, generating highly fluorescent and localized Y1 (Step 4), possibly through the monophosphorylated intermediate Y5.
introducing the Dabcyl group, however, made the resulting
probe significantly bulkier than needed and rendered it cell-
impermeant.
previously reported 2P dye, Y1*), upon attachment of an
electron-withdrawing phosphate group (giving Y2; Figure 1C,
right), was effectively turned OFF in aqueous buffers. This thus
makes Y1, to our knowledge, the first fluorogenic reporter
developed from a centrosymmetric dye possessing desirable 2P
photophysical properties. Herein, we report the successful
demonstration of Y1, and the resulting probes based on it, as
efficient Turn-ON 2P imaging reporters for detection of
endogenous phosphatase activities with good spatial resolution
in both cells and tissues.
Two-photon (2P) fluorescence microscopy (TPFM) pro-
vides key advantages over conventional one-photon (1P)
imaging techniques, namely, increased penetration depth, lower
tissue autofluorescence and self-absorption, reduced photo-
damage and photobleaching, and therefore is particularly useful
for imaging deep tissues and animals.9,20,21 Various reactive 2P
probes have been developed for metal cations, organic and
inorganic anions, DNA and other targets.21 Practical 2P small
molecule probes (that is, probes made of fluorescent dyes that
possess desirable 2P photophysical properties) capable of
detecting endogenous enzyme activities, to our knowledge, are
unavailable at present, apart from our most recent report.17 In
principle, a “piggyback” approach may be taken by using some
of the existing 1P probes (e.g., the coumarin and fluorescein
probes shown in Figure 1C) in a 2P experiment.20 This,
however, has not been very successful, primarily due to
undesirable 2P photochemical and photophysical properties of
coumarin and fluorescein dyes. For TPFM to be widely
adopted for bioimaging of enzymatic activities, it is essential
that fluorogenic enzyme substrates based on novel 2P dyes are
developed.10,20 Inspired by our recent work of the FRET-based
2P probe, we reasoned that Y1* and other similarly designed
2P dyes having the same core structure (e.g., 2-hydroxy-4,6-
bis(4-hydroxystyryl)pyrimidine, or Y1, in Figure 1C, right)
might indeed be made into fluorogenic dyes/reporters. A well-
known strategy to turn ON/OFF a dye is to change the pull−
push conjugated π-electron system within the dye. This
principle had previously been successfully applied in the
development of fluorogenic probes based on coumarin and
fluorescein dyes (Figure 1C; left).22 As part of the current
study, we found that fluorescence of the newly developed 2P
dye, Y1 (Figure 1C right; via NR2→OH conversion from the
RESULTS AND DISCUSSION
■
Design of the Two-Photon Fluorogenic Probes. In our
approach (Figure 2), Y1 serves as the 2P fluorescence reporter.
Its phosphorylated version Y2, where the two phenolic groups
were masked by an electron-withdrawing, phosphatase-
responsive phosphate warhead, has a significantly reduced
conjugated π-electron system, and thereby represents the Turn-
OFF state of Y1. Y3, the “caged” form of Y2, which releases the
active phosphatase reporter upon UV irradiation, increases the
cell permeability of the reporter and at the same time provides
additional spatial and temporal controls over imaging
endogenous phosphatase activities.14,17 In the current study,
we chose a simple photolabile 2-nitrobenzyloxy group as the
caging molecule for synthetic convenience, but it may be readily
replaced, if necessary, with other known two-photon caging
molecules.21d,e To minimize the diffusion problem of our
probes while imaging endogenous phosphatase activities and to
make them suitable for subcellular detection of localized
enzymatic activities in live cells, we conjugated Y4, which is an
alkyne-containing analog of Y3 (Supporting Information,
Scheme S1), to different azide-modified, cell-penetrating
peptides (CPPs; P1 to P4 in Figure 2 and Scheme S1), giving
YP1/YP2/YP3/YP4 (Figure 2 and Scheme S1). We chose the
Cu(I)-catalyzed cycloaddition, or click chemistry, for highly
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modular and rapid assembly of these small molecule probe−
peptide conjugates, which would have otherwise been
challenging to obtain synthetically.23 Of the four CPPs chosen,
P1 is a peptide derived from Simian virus 40 (SV40) T-
antigen.24a The resulting P1-conjugated probe, YP1, was
expected to be delivered to the endoplasmic reticulum (ER)
of mammalian cells and detect ER-localized phosphatase
activities therein. YP2, conjugated to the N-palmitoylated
plasma membrane localization peptide P2 which is commonly
found on membrane-bound proteins,24b would detect mem-
brane-localized phosphatase activities. YP3 (conjugated to a
well-known mitochondria-localizing peptide, P3, developed by
Kelley et al.24c) would detect mitochondria-localized phospha-
tase activities. To investigate whether our strategy could also be
used to image endogenous phosphatase activities in a tumor
cell-specific manner, we designed YP4, which is conjugated to
the P4 peptide. The P4 peptide contains a polycationic CPP
peptide (in blue) fused to a polyanionic masking peptide (in
red) through a linker cleavable by matrix metalloproteases
(MMPs) which are overexpressed on the surface of numerous
cancer cells including HeLa cells. Previously, Tsien and co-
workers had successfully used this peptide as a vehicle for
tumor cell-specific delivery.25 As shown in Figure 2, cellular
uptake of YP4 would normally be blocked due to the charge
interaction between the two peptide fragments, but upon
cleavage by MMPs present on tumor cells (Step 1), the CPP
peptide would be locally unleashed, causing the cleaved YP4 to
be internalized to the intended subcellular organelle (i.e., ER;
Step 2). Subsequent UV irradiation (Step 3) would release the
phosphatase-responsive probe Y2 (still conjugated to the
peptide), which would then be dephosphorylated by
endogenous phosphatases, generating the highly fluorescent
and localized Y1 (Step 4).
Chemical Synthesis. Synthesis of the newly discovered 2P
fluorogenic dye, 2-hydroxy-4,6-bis(4-hydroxystyryl)pyrimidine,
Y1, its phosphorylated Y2 and the “caged” version Y3 is shown
in Scheme S1 (Supporting Information). The compounds were
synthesized according to previously published protocols with
some modifications.19 Briefly, 2-hydroxy-4,6-dimethylpyrimi-
dine (1) was condensed with 2.2 equivalents of 4-
hydroxybenzaldehyde in the presence of hydrochloric acid
under reflux conditions to give Y1 in a single step (80% yield).
Similarly, Y2 was prepared from diethyl 4-formylphenyl
phosphate (2) followed by TMSI deprotection in 15% yield
(in two steps). The nitrobenzyl protected Y3 was prepared by
condensation between (1) and 4-aldehydephenyl bis(2-nitro-
benzyl) phosphate (3) in 21% yield. The propargyl group-
containing Y4 was prepared by coupling propargyl bromide to
Y3 in the presence of K2CO3 (37.5% yield). Y5 (used as a
control in our studies), the monophosphorylated version of Y2,
was synthesized by the coupling of (1) with 1.1 equiv of 4-
hydroxybenzaldehyde, giving (E)-4-(4-hydroxystyryl)-6-meth-
ylpyrimidin-2-ol (4) (72% yield). The subsequent reaction
between (4) and (2) followed by TMSI deprotection gave the
desired product Y5 in 11% yield (in two steps). The four azide-
functionalized localization peptides were synthesized on rink
amide resin using protocols modified from standard solid phase
peptide synthesis (SPPS; Supporting Information, Scheme
S1).26 For P1 and P3, the corresponding amino acids were
assembled on the resin. At the end of the synthesis, 4-
azidobutanoic acid was coupled to the resin using HOBt/
HBTU/DIEA activation strategy. For P2, the palmitoyl group
was first attached to Ω-amino group of the first Lys residue by
HOBt/HBTU/DIEA coupling. Subsequent SPPS and the
coupling of 4-azidobutanoic acid were similarly followed. For
P4, since a succinic acid moiety was needed as the N-terminal
cap of the peptide, 4-azidobutanoic acid was added to the Ω-
amino group of the first Lys residue instead (Supporting
Information, Scheme S1). The peptides were cleaved from the
resin and purified by reverse-phase preparative HPLC to
homogeneity and characterized by LC−MS. To obtain the
peptide-reporter conjugates, YP1−YP4, a variety of ‘‘click’’
conditions were tested for the coupling between Y4 and P1-P4;
at the end, it was found a cosolvent system of DMSO/H2O
(1:1) with copper catalysts and additives (CuSO4/NaAsc/
TBTA) gave the best results for YP1/YP2/YP3, and for YP4, a
cosolvent system of DMSO/CH3CN/H2O (4:4:1) with CuI/
DIEA worked the best. All four conjugates were obtained in
good coupling efficiency and product quality. They were
similarly purified and characterized.
Photophysical and Enzymatic Properties of Probes.
To unequivocally establish that our newly developed Y1/Y2
pair was indeed a good 2P fluorogenic system capable of
emitting strong fluorescence during Y2→Y1 conversion and
thus reporting phosphatase activities in real time, we next
investigated their photophysical (1P and 2P) and enzymatic
properties (Table 1, Figures 3 and 4; Figures S1−S3 in the
Table 1. Photophysical Properties of Y1/Y2/Y3/Y5
a
(ab)
max
b
(em)
max
c
d
e
λ
ε/103
λ
εΦ
τ
δΦ
Y1
Y2
Y3
Y5
421
406
404
405
49
28
18
33
512
532
481
522
5390
1960
1620
3300
0.340
0.315
0.320
0.325
82
21
25
39
a
b
Peak position of the longest absorption band. Peak position of
c
emission, exited at the absorption maxima in Hepes. Brightness of the
d
e
relative compound. Life time of the emission in ns. The maximum
two-photon action cross-section values upon excitation from 750 to
860 nm in GM (1 GM = 10−50 cm4 s photon−1).
Supporting Information). Y3 (the “caged” version of Y2), Y5
(the monophosphorylated version of Y1) as well as two
common 1P dyes, coumarin and fluorescein, were included in
the experiments as references. As shown in Table 1, under
physiological conditions (Hepes buffer at pH = 7.5), Y1/Y2/
Y3/Y5 had maximum absorption spectra at 400−420 nm,
Figure 3. (A) pH Effect on the one-photon excited fluorescence
intensity of Y1 at room temperature in Hepes buffer (supplemented
with 50 mM NaCl, 2.5 mM EDTA, 2 mM DTT and 0.02% Triton X-
100; probe concentration: 1.0 μM; λex/em = 405/550 nm). (B)
Photostability profiles of Y1 and three commercial dyes (coumarin,
HPQ, and fluorescein) over the course of 60 min in Hepes buffer
(λex/em = 405/550 nm for Y1, 350/450 nm for courmarin, 350/520 nm
for HPQ, and 488/520 nm for fluorescein).
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bovine intestinal mucosa, two human serine/threonine
phosphatases, PP1 and PP2B, which are involved in cell
division and T cell activation, respectively27b,c) gave rise to
almost complete Y2→Y1 conversion in 60 min, with a
concomitant increase in fluorescence (Figure 4C). With the
presence of two phosphate groups in the molecule, Y2 might
have been first dephosphorylated to Y5 on route to Y1 (Figures
2 and 4B), and if so, it might potentially complicate the real-
time reporting of phosphatase activities. We were pleased to
find, however, that the formation of Y5 was not detected in our
LC−MS experiments throughout the duration of Y2→Y1
process (Figure 4A). This indicates a faster rate of Y5→Y1 (k2)
than that of Y2→Y5 (k1) as illustrated in Figure 4B. As such,
with Y2→Y5 being the rate-determining step, readouts from
the fluorescence increase of Y2 upon phosphatase treatment
should accurately reflect the overall rate of Y2→Y1 (k).
Subsequently, time-dependent fluorescence measurements
were carried out with PTP1B-treated reactions (Y2 in Figure
4C and Y5 in Figure 4D), and the results were extrapolated to
provide the corresponding kinetic parameters. On the basis of
the reaction model shown in Figure 4B, we obtained reaction
rate constants of k = 4.00 0.27 × 103 M−1 s−1 and k2 = 1.83
0.25 × 104 M−1 s−1. These results thus corroborated well with
our earlier LC−MS results (Figure 4A).
Figure 4. (A) Y3→Y2 conversion by UV irradiation (5 min), followed
by Y2→Y1 dephosphorylation by PTP1B (60 min). Reactions were
carried out at room temperature in Hepes buffer (supplemented with
50 mM NaCl, 2.5 mM EDTA, 2 mM DTT and 0.02% Triton X-100,
pH = 7.5; probe concentration: 1.6 μM). Experiments were monitored
by LC−MS, and peaks were unambiguously assigned based on their
molecular weights. PTP1B/probe concentration ratio = 1:30. Y5 was
not detected throughout the entire Y2→Y1 process. (B) Overall
pathway of Y3→Y2→Y1. (C,D) Time-dependent one-photon excited
fluorescence emission spectra of PTP1B-treated Y2 and Y5 reactions,
respectively, over the course of 60 min.
Subcellular Bioimaging of Endogenous Phosphatase
Activities with YP1/YP2/YP3. We first established that our
newly developed Y1/Y2/Y3 fluorogenic system was indeed
useful for general bioimaging applications to detect endogenous
phosphatase activities. Since Y3 (the “caged”version of Y2) is
cell-permeable, it was used in our preliminary imaging
experiments (Figure 5). The ELF 97 detection kit was used
making them suitable to be excited by a common two-photon
laser. Y1 had a fluorescence maximum at λem = 512 nm (ε =
49000 M−1 cm−1 and ε·Φ = 5390) which was largely
independent of pH changes (Figure 3A), and a two-photon
action cross-section (δ·Φ = 82 GM) comparable to other
common 2P dyes.21 On the contrary, Y2 and Y3 were only
weakly fluorescent (ε·Φ = 1960 and 1620, respectively) and
had much smaller two-photon action cross sections (δ·Φ = 21
and 25 GM, respectively). Y5, with an absorption maximum at
λab = 405 nm and fluorescence emission maximum at λem = 522
nm (ε = 33000 M−1cm−1, ε·Φ = 3300, δ·Φ = 39 GM), was
moderately fluorescent as well, due to removal of one of the
phosphate groups in Y2. Y1 showed excellent photostability
which was better than courmarin, fluorescein, and 2-(2′-
hydroxy-5′-chlorophenyl)-6-chloro-4-(3H)-quinazolinone
(HPQ) (Figure 3B). The consistent fluorescence lifetime (τ) in
the Y1/Y2 system indicates the fluorogenicity of the system
was based on the increase in the absorbance during Y2→Y1
conversion, which is similar to other well-known fluorogenic
phosphatase reporter systems (e.g., DiFMUP, MUP, ELF-97,
and FDP in Figure 1C).22 Unlike Y2, both Y1 and Y3 were
shown to be highly cell-permeable (Papp = 223 and 233 nm·s−1,
respectively). We therefore concluded that the Y1/Y2/Y3
system was suitable for endogenous phosphatase imaging using
TPFM.
Figure 5. (A) One-photon excited fluorescence microscopy of live
HepG2 cells treated with Y3 (10 μM). Following 2-min UV
irradiation, the cells were imaged over the course of 120 min with
60-s intervals (λex = 405 nm; PMT range: 460−550 nm). The image
shown was taken at the 80-min time point. Scale bar = 5 μm. (B)
Graphical representation of time-dependent changes in fluorescence
intensity at the circled position in panel A.
as a control to detect endogenous phosphatase activities in fixed
mammalian cells (Supporting Information, Figure S4);13a
fluorescence signals were detected mostly throughout the
whole cell (except nucleus), indicating the ubiquitous presence
of phosphatase activities across most intracellular space and
organelles. Real-time imaging of live HepG2 cells treated with
the cell-permeable Y3 probe, upon UV irradiation (to give
intracellular Y2), showed a gradual increase in fluorescence
signals in the whole cell (similar to ELF 97) over the course of
60 min. After 60 min, the fluorescence reached saturation
Next, it was established that a completely uncaged product
Y2 could be obtained from Y3 (through its monoprotected
intermediate Y3* which was detectable by HPLC) under UV
exposure of 500 μJ/cm2 for 5 min (Figure 4A). Further
incubation of the uncaged Y3 with PTP1B (a key human PTP
involved in diabetes and obesity),27a and other types of
phosphatases (Figure S3; including alkaline phosphatase from
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(Figure 5). This kinetic profile was similar to what was
observed in the in vitro experiment with recombinant PTP1B
(Figure 4C). We thus concluded Y3 alone could be used to
detect endogenous phosphatase activities with similar kinetic
profiles as those from in vitro experiments. We further
confirmed that the short UV irradiation conditions used in
our imaging experiments did not cause any apparent changes in
cell morphology and viability, as well as endogenous
phosphatase activities (Supporting Information, Figure S5).
We next investigated whether the probes could be
successfully delivered to subcellular organelles in live
mammalian cells, for the maging of phosphatase activities
localized therein. Direct administration of Y3 to cells followed
by UV irradiation/imaging, as shown in Figure 5, was not
expected to provide sufficient spatial resolutions needed for
subcellular imaging due to the diffusible nature of this
compound. We therefore conjugated the probe to various
cell-penetrating peptides (CPPs), giving YP1/YP2/YP3
(Supporting Information, Scheme 1 Figure 2). Subcellular
targeting was previously demonstrated for inhibitors and probes
in cultured cells, using delivery vehicles including small
molecules, localized CPPs and SNAP-tag fusion proteins.26a,28
The use of localization peptides as delivery tools offers the
advantages of better controlled administration, ease of synthesis
and organelle specificity.28c Subcellular imaging of endogenous
metabolites aided by small molecules or proteins has also been
demonstrated recently.24c,28b Our aim in the current study was
for the CPPs to achieve both organelle-specific delivery and
subcellular retention of the probes, thus ensuring the released
fluorescence be spatially imaged with sufficient resolutions.
HeLa cells incubated with control dye-CPP conjugates
indicated expected organelle-specific localization for all CPPs
tested (Supporting Information, Figure S6). Subsequently, the
imaging of endogenous phosphatase activities was carried out
with both one- and two-photon fluorescence microscopy in live
HeLa and HepG2 cancer cells (Supporting Information, Figure
S7 and Figure 6). To image subcellular phosphatase activities
using the CPP-conjugated probes, live cells were first incubated
with the corresponding compound, irradiated by UV then
imaged using TPFM (excitation at 800 nm; Figure 6). All three
probes (YP1, YP2, and YP3) showed strong fluorescence
signals ONLY in their intended organelles (ER, membrane and
mitochondria, respectively), but not in other subcellular
organelles and intracellular space (see image panels 3/7/11 in
Figure 6). The same cells were simultaneously treated with
commercially available organelle-specific imaging markers to
independently verify the results (see image panels 2/6/10, for
ER-Tracker, Cell-Mask, and Mito-Tracker, respectively). A
reconstructed two-photon fluorescence 3-D image of HepG2
cells treated with YP2 provided further evidence that our
strategy was indeed organelle-specific, enabling high-resolution
imaging of phosphatase activities only in predetermined
intracellular locations, and thereby minimizing interference
fluorescence from other sources (Supporting Information,
Figure S9). The diffusion problem encountered by most
other small molecule imaging probes,12 was effectively
abrogated in the current study through the organelle-specific
“trapping” of CPPs by their cellular interacting counterparts.28c
Probe-treated cells, if not UV-irradiated, showed no fluo-
rescence, indicating the temporal control of our “caging”
strategy (Supporting Information, Figure S8). Cells treated
with Na3VO5 (a general phosphatase inhibitor) showed
undetectable fluorescence signals under the same imaging
Figure 6. Two-photon fluorescence microscopy (800 nm) of
endogenous phosphatase activities in live mammalian cells (A,
HeLa; B, HepG2) by YP1/YP2/YP3 probes. Live cells were incubated
with the probe (8−10 μM) for 1−2 h, followed by UV irradiation (2
min) and the addition of a tracker (0.25 μg/mL), then further
incubated for 30 min, before being imaged. Probes used: YP1 in panels
1−4; YP2 in panels 5−8; YP3 in panels 9−12. All images were
acquired in the same way. Scale bar = 5 μm.
conditions (Figure S8). Both HeLa and HepG2 cells showed
similar organelle-specific imaging results (compare Figure 6A
and 6B), indicating the strategy is amenable to other
mammalian cells. Both one- and two-photon fluorescence
microscopy provided similar imaging results (compare
Supporting Information, Figure S7 and Figure 6).21a
Tumor Cell-Specific Imaging Using YP4. We next
investigated whether this newly developed strategy could be
extended to imaging phosphatase activities in a tumor cell-
specific manner. Elevated levels of phosphatase activities are
found in many cancerous cells and tissues.2 Imaging agents that
are highly responsive toward diseased cells/tissues hold great
promises in diagnosis and therapeutics. Previously, Tsien and
co-workers developed a tumor-targeting strategy based on
selective local unleashing of CPPs.25 Herein, we showed the
successful application of this method to our newly developed
imaging probes (Figure 2). Cellular uptake of YP4, which was
conjugated to the azide-modified P4 peptide, by normal
mammalian cells was not expected due to a lack of
overexpressed MMPs on their cell surface. In our imaging
experiments with YP4 and live HeLa cells (known to
overexpress MMPs on the cell surface), we observed both
MMP and time-dependent cellular uptake of the probe and
imaging of endogenous phosphatase activities (Figure 7). A
control experiment carried out in the presence of GM6001 (a
general MMP inhibitor29) showed only moderate fluorescence
signals at cell periphery after 120 min of incubation (Figure
7A), indicating nonspecific association between the uncleaved
YP4 and cell membranes, but no cellular internalization. On the
contrary, HeLa cells treated with YP4 started to develop strong
fluorescence signals on the plasma membrane after 30 min
(Figure 7B), indicating membrane association. After 60 min,
strong fluorescence was observed both on the plasma
membrane and in the cytoplasm. After 120 min, most
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Figure 8. 2P imaging of endogenous phosphatase activities in fresh
brains of one day old live female Drosophila using Y2 and uncaged
YP2. The images were taken at 800 nm with 40× magnification.
Drosophila brains were incubated with the probes (20 μM) for 6 h.
Probes used: Y2 in panels 1−3; uncaged YP2 in panels 4−6. (1,4)
Bright-field images of half brain. (2,5) TPEF images of medulla region;
insets are images obtained from the brain treated with Na3VO5 (100
μM) 30 min prior to incubation with Y2; (3,6) Enlarged images of the
red boxes in panels 2 and 5 at a depth of ∼110 μm with 40×
magnification. Scale bar: (1,4, and insets) 150 μm; (3,6) 20 μm.
Figure 7. (A,B) TPFM (800 nm) of endogenous phosphatase
activities in live HeLa cells treated with the tumor-specific probe, YP4.
(A) YP4 + HeLa cells + GM6001 (500 nM). Images were taken after
120 min of incubation. (B) YP4 + HeLa cells. Images were taken after
incubation for 30 min (panels 1−4), 60 min (panels 5−8), and 120
min (panels 9−12). Tracker channel: ER-Tracker (0.25 μg/mL). Scale
bar = 5 μm. (C) Flow cytometry analysis of HeLa cells incubated with
YP4 (120 min), followed by UV irradiation and FACS analysis: (1)
untreated cells; (2) cells incubated with YP4 in the presence of
GM6001 (500 nM); (3) cells incubated with YP4 only.
97 detection kit, which was shown to detect fluorescence
signals of the tissues only at a depth of up to 35 μm
(Supporting Information, Figure S10). This much deeper
penetration, coupled with the significantly lower background
fluorescence and better clarity observed in our TPFM images,
clearly demonstrated the potential application of these newly
developed, two-photon probes for tissue-based imaging experi-
ments in future.
CONCLUSION
■
fluorescence signals were detected in the ER, indicating
complete internalization of the probe and successful detection
of subcellular phosphatase activities. Similar results were
obtained with a flow cytometry experiment (Figure 7C),
where at least 2- to 3-fold increase in fluorescence was detected
from HeLa cells treated with YP4 versus those treated with
YP4 plus GM6001 (Figure 7C).
A novel two-photon dye, 2-hydroxy-4,6-bis(4-hydroxystyryl)-
pyrimidine (Y1), based on a previously reported dye, 2-
hydroxy-4,6-bis(4-(diethylamino)styryl)pyrimidine,19 was dis-
closed for the first time. A significant advance in our current
study is the finding that the highly fluorescent Y1, upon
attachment of electron-withdrawing phosphate groups to its
phenolic moieties (to make Y2), could be rendered only weakly
fluorescent. This thus makes the Y1/Y2 system the first
fluorogenic, two-photon pair capable of real-time detection of
phosphatase activities. Other existing small molecule phospha-
tase reporters based on common one-photon dyes including
DiFMUP, FDP, and ELF 97 are not ideal for robust two-
photon imaging applications, as they are made of one-photon
dyes such as coumarin and fluorescein which have undesirable
2P photophysical properties. Despite having two phosphate
groups in Y2, we found the monodephosphorylation step (k1)
was the rate-determining step, with the second dephosphor-
ylation step (k2) calculated to be ∼4× faster. In fact, we were
not able to detect the presence of the monodephosphorylated
intermediate Y5 throughout the Y2→Y1 process. This indicates
the overall fluorescence release from the Y2→Y1 conversion in
our reporter system could be used to directly follow
endogenous phosphatase activities in real time. We have
subsequently conjugated Y3 (the caged version of Y2) to
different cell-penetrating peptides. The resulting probes YP1−
YP4 were successfully used as imaging probes to detect
endogenous phosphatase activities in both live mammalian cells
and Drosophila brains. We showed that with the aid of CPPs,
Two-Photon Imaging of Animal Tissues. As earlier
discussed, two-photon fluorescence microscopy is particularly
useful for imaging deep tissues and animals, owing to its
increased penetration depth and other key advantages.9,20,21
Despite recent advances in the development of 2P small
molecule probes, few of them have been successfully designed
to image endogenous enzymatic activities.17,20,21 Our long-
standing research interests in Catalomics has prompted us to
start looking into enzyme-detecting, small molecule probes that
can work in live tissues and animals.30 In the current study, we
have successfully imaged endogenous phosphatase activities
deep inside animal tissues using our probes (Figure 8). The
fresh brain of one day-old female Drosophila was dissected and
incubated with 20 μM of either Y2 or uncaged YP2 for 6 h at
room temperature. Subsequently, TPFM images of the tissues
were taken at a depth of ∼110 μm (Figure 8); the results
revealed distribution of endogenous phosphatase activities
mostly in the medulla region of the tissue (panels 2 and 5).
Na3VO5-treated tissues shows undetectable fluorescence signals
under similar imaging conditions (insets in panel 2). Control
experiments were carried out with tissues treated with the ELF
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PMT detector ranged from 420 to 700 nm for steady state
fluorescence and nondescanned detectors (NDD) for the two-photon
excited fluorescence, immediately. Images were processed with Leica
Application Suite Advanced Fluorescence (LAS AF). Flow cytometry
analysis was performed on a BD LSR II bioanalyzer equipped with a
350 nm UV air-cooled laser. The data was analyzed using BD
FACSDiva software.
our probes could be effectively delivered to cells in an
organelle- and tumor cell-specific manner. Diffusion problems
normally associated with small molecule imaging probes were
abrogated. We found our probes could image endogenous
phosphatase activities in Drosophila brains at a depth of >100
μm. Further work will focus on the extension of this novel
strategy to the imaging of other classes of enzymes.31 One of
the most obvious drawbacks about our current strategy is that
the probes at present are not able to distinguish different classes
of phosphatases. In fact, imaging results obtained from this
study, including those in mammalian cells and Drosophila
brains, likely came from cumulative enzymatic activities of
different endogenous phosphatases, e.g. protein phosphatases,
acidic and alkaline phosphatases that were present at the
specific organelle during the time of the experiment. We are
however enthusiastic that, in future, class- and substrate-specific
small molecule probes of phosphatases may be discovered.6,32
Chemical Synthesis. The overall synthetic scheme was provided
as Scheme S1 in the Supporting Information.
Diethyl 4-Formylphenyl Phosphate (2). 4-Hydroxybenzaldehyde
(0.24 g, 2 mmol) was dissolved in dry DCM (10 mL), followed by the
addition of DMAP (0.03 g, 0.25 mmol), and the mixture was stirred at
room temperature for 10 min, then cooled on an ice bath.
Subsequently, diethyl phosphorochloridate (0.52 g, 3 mmol) in 10
mL of dry DCM was added dropwise to the mixture, followed by
addition of triethylamine (0.7 mL, 5 mmol). The reaction was stirred
on the ice bath for 2 h, then at room temperature for another 8 h. The
reaction mixture was concentrated and purified by flash chromatog-
raphy to afford 2 as light yellow oil (0.31 g; 60% yield). 1H NMR (300
MHz, CDCl3) δ 1.37 (dt, J1 = 6.9 Hz, J2 = 1.2 Hz, 6H), 4.27 (m, 4H),
7.41 (d, J = 8.7 Hz, 2H), 7.90 (d, J = 8.7 Hz, 2H), 9.97 (s, 1H). 13C
NMR (75 MHz, CDCl3) δ 15.64, 15.73, 64.58, 64.66, 120.16, 120.23,
131.27, 133.02, 155.04, 155.12, 190.34. 31P NMR (121 MHz, CDCl3)
δ −6.67.
EXPERIMENTAL SECTION
■
General Information. All chemicals were purchased from
commercial vendors and used without further purification, unless
otherwise noted. N,N-Dimethylformamide (DMF) and dichloro-
methane (DCM) were distilled over CaH2. All nonaqueous reactions
were carried out under a nitrogen/argon atmosphere in oven-dried
4-Aldehydephenyl Bis(2-nitrobenzyl) Phosphate (3). The com-
pound bis(2-nitrobenzyl)hydrogen phosphate was synthesized accord-
ing to reported procedures (63% yield).34 1H NMR (300 MHz,
MeOD) δ 8.08 (d, 2H), 7.69−7.78 (m, 4H), 7.54 (t, J = 9.0 Hz, 2H),
5.43 (d, J = 3.8 Hz, 4H). 13C NMR (75 MHz, MeOD) δ 166.60,
148.50, 135.11, 130.13, 129.76, 125.87. 31P NMR (121 MHz, MeOD)
δ −0.71. After adding bis(2-nitrobenzyl)hydrogen phosphate (5.5 g,
15 mmol) into dry DCM, oxalyl chloride (6.5 mL, 75 mmol) was
added with catalytic amount of DMF to initiate the reaction. The
reaction solution was stirred at room temperature for 2 h. The solvent
and oxalyl chloride were removed in vacuo to give a light yellow oil.
Next, 4-hydroxybenzaldehyde (0.61 g, 5 mmol) was dissolved in dry
DCM (20 mL), followed by the addition of DMAP (0.06 g, 0.5
mmol), and the mixture was stirred at room temperature for 10 min.
Then the light yellow oil (now redissolved in 10 mL dry DCM) was
added dropwise to the mixture on an ice bath, followed by addition of
triethylamine (1.4 mL, 10 mmol). The reaction was left on ice for 2 h,
and then warmed to room temperature for another 8 h. The reaction
mixture was concentrated and purified by flash chromatography to
1
glasswares. H NMR, 31P NMR, and 13C NMR spectra were recorded
on a Bruker model Avance 300 MHz or DPX-500 MHz NMR
spectrometer. Chemical shifts are reported in parts per million relative
to internal standard tetramethylsilane (Si(CH3)4 = 0.00 ppm) or
residual solvent peaks (CDCl3 = 7.26 ppm, DMSO-d6 = 2.50 ppm,
1
MeOD = 3.31 ppm). H NMR coupling constants (J) are reported in
Hertz (Hz) and multiplicity is indicated as follows: s (singlet), d
(doublet), t (triplet), q (quartet), m (multiplet), br s (broad singlet),
br d (broad doublet), dd (doublet of doublet), dt (doublet of triplet),
dq (doublet of quartet), tq (triplet of quartet). The chemical shift of
31P NMR is reported relative to H3PO4 = 0.00 ppm. Analytical HPLC
and mass spectra were recorded on a Shimadzu LC-IT-TOF or LC-
ESI system equipped with an autosampler, using reverse-phase
Phenomenex Luna 5 μm C18 100 Å 50 × 3.0 mm columns.
Preparative HPLC was carried out on a Gilson preparative HPLC
system using Trilution software and a reverse-phase Phenomenex
Luna 5 μm C18(2) 100 Å 50 × 30.00 mm column. 0.1% TFA/H2O and
0.1% TFA/acetonitrile were used as eluents. The flow rate was 0.6
mL/min for analytical HPLC and 10 mL/min for preparative HPLC.
UV−vis absorption and fluorescence spectra were measured by using a
Shimadzu UV−vis spectrophotometer and a Perkin-Elmer LS50
spectrofluorometer, respectively. The two-photon excited fluorescence
measurements were performed by using a Spectra Physics femto-
second Ti:sapphire oscillator (Tsunami) as the excitation source. The
output laser pulses have a tunable center wavelength from 750 to 860
nm with pulse duration of 40 fs and a repetition rate of 76 MHz. The
laser beam was focused onto the samples that were contained in a
cuvette with a path length of 1 cm. The emission from the samples was
collected at 90° angle by a pair of lenses and an optical fiber that was
connected to a monochromator (Acton, Spectra Pro 2300i) coupled
with CCD (Princeton Instruments, Pixis 100B) system. A short pass
filter with a cutoff wavelength at 700 nm was placed before the
spectrometer to minimize the scattering from the pump beam. All the
measurements were performed at room temperature. Cell-permeability
assay was carried out, as previously described,33 with MDCK (Madin-
Darby canine kidney) cells and with caffeine and lucifer yellow as
controls. All images were acquired on Leica TCS SP5X confocal
microscope system equipped with Leica HCX PL APO 63x/1.20 W
CORR CS, 405 nm diode laser, argon ion laser, white laser (470 to
670 nm, with 1 nm increments, with 8 channels AOTF for
simultaneous control of 8 laser lines, each excitation wavelength
provides 1.5 mV), an PMT detector ranging from 420 to 700 nm for
steady state fluorescence, and Ti-Sapphire laser (∼4 W at 800 nm), the
1
afford 3 as a light yellow oil (1.2 g; 51% yield). H NMR (300 MHz,
CDCl3) δ 5.21 (d, J = 7.8 Hz, 4H), 7.44 (d, J = 8.4 Hz, 2H), 7.45 (dt,
J1 = 8.4 Hz and J2 = 1.8 Hz, 2H), 7.65 (m, 4H), 7.82 (d, J = 8.7 Hz,
2H), 8.06 (d, J = 7.8 Hz, 2H), 9.90 (s, 1H). 13C NMR (75 MHz,
CDCl3) δ 67.07, 67.13, 120.55, 120.62, 125.16, 128.62, 129.35, 131.36,
131.47, 131.75, 133.67, 14.23, 146.75, 154.70, 190.68. 31P NMR (121
MHz, CDCl3) δ −6.98. IT-TOF-MS: m/z [M+Na]+ calcd, 495.07;
found, 495.05.
2-Hydroxy-4,6-bis(4-hydroxystyryl)pyrimidine (Y1). Y1 was syn-
thesized according to reported methods.19 A mixture of 2-hydroxy-4,6-
dimethylpyrimidine hydrochloride (1; 0.8 g, 5 mmol) and 4-
hydroxybenzaldehyde (1.34 g, 11 mmol) in 20 mL of ethanol was
stirred under an N2 atmosphere at room temperature for 1 h.
Hydrochloric acid (2 mL, 3 M) was added dropwise to the reaction
mixture which was then refluxed for 30 h. The solvent was removed,
and the residue was washed with DCM three times. The resulting solid
was purified by flash chromatography (10:1 DCM/methanol) to give
Y1 as a red powder (1.33 g; 80% yield). 1H NMR (300 MHz, DMSO-
d6) δ 6.80 (d, J = 16.2 Hz, 2H), 6.82 (s, 1H), 6.85 (d, J = 8.4 Hz, 4H),
7.51 (d, J = 8.4 Hz, 4H), 7.78 (d, J = 16.2 Hz, 2H), 10.06 (s, 1H),
11.74 (s, 1H). 13C NMR (75 MHz, DMSO-d6) δ 98.90, 115.97,
119.05, 126.12, 129.70, 139.13, 156.76, 159.59, 162.01. IT-TOF-MS:
m/z [M+H]+ calcd, 333.12; found, 333.12.
2-Hydroxy-4,6-bis(4-dihydrogen phosphate)pyrimidine (Y2). A
mixture of 2-hydroxy-4,6-dimethylpyrimidine hydrochloride (1; 0.08 g,
0.5 mmol) and diethyl 4-formylphenyl phosphate (0.39 g, 1.5 mmol)
in 10 mL of ethanol was stirred under an Ar atmosphere at room
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temperature for 1 h. Hydrochloric acid (0.2 mL, 3 M) was added
dropwise to the reaction mixture which was then refluxed for 48 h.
The solvent was removed, and the resulting residue was washed with
ethanol (3×) then concentrated in vacuo. Twenty milliliters of DCM
was subsequently added, and the resulting mixture was stirred at room
temperature for 10 min before being cooled on an ice bath. TMSI
(0.46 g, 2.2 mmol) was added dropwise. The reaction was stirred
further for 4 h before being quenched by the addition of 10 mL of
saturated ammonium chloride solution. Upon concentration in vacuo,
the residue was purified by semipreparative HPLC on a C18 column to
Solid-Phase Peptide Synthesis (SPPS). The peptides were
synthesized on rink amide resin using Irori microkan reactors (150
mg/reactor; loading level =0.6 mmol/g), as previously described.26
Briefly, Fmoc deprotection was carried out with piperidine (20% v/v
in DMF; 1 h). The washing cycles were done with DMF (3×), DCM
(3×), MeOH (3×) and DMF (1×). The resins were then added to a
preactivated DMF solution containing Fmoc-protected amino acid (4
equiv), HOBt (4 equiv), HBTU (4 equiv), and DIEA (4 equiv). The
coupling reaction was carried out for 12 h at room temperature with
constant shaking. Complete coupling was confirmed by the Ninhydrin
test. For the coupling of palmitoyl group to the first Lys residue in P2,
Fmoc-Lys(Mtt)−OH was first coupled to the resin. Subsequently, the
Mtt group on the Ω-amino group was removed with 1% TFA ((v/v)
in DCM for 1 h). Upon neutralization with DIEA, to the resin was
added a preactivated DMF solution containing palmitic acid (4 equiv),
HOBt (4 equiv), HBTU (4 equiv), and DIEA (4 equiv). The reaction
was carried out for 12 h at room temperature with constant shaking.
For the capping of succinoyl group to P4, the N-terminus of P4 was
coupled with a DMF solution containing succinic anhydride (4 equiv)
and DIEA (4 equiv), for 4 h at room temperature. For coupling of 4-
azidobutanoic acid, the N-termini of P1, P2, and P3 resin-bound
peptides were treated with a preactivated DMF solution containing 4-
azidobutanoic acid (4 equiv), HOBt (4 equiv), HBTU (4 equiv), and
DIEA (4 equiv), for 4 h at room temperature. P4 needed the
attachment of 4-azidobutanoic acid at the Ω-amino group of the first
Lys residue. Upon removal of the Mtt group on the resin-bound
Fmoc-Lys(Mtt)−OH, the resin was then coupled to 4-azidobutanoic
acid similarly. The peptides were cleaved from the solid support by
adding a cleavage cocktail of TFA/TIS/H2O (95/2.5/2.5) to the
resins and shaking for 2 h at room temperature. Following ether
precipitation and preparative HPLC, the peptides (P1−P4) were
obtained as white solids. The peptides were characterized using IT-
TOF LC-MS. The m/z values obtained were fractions of the mass of
target peptides due to positive charges present in the peptides. P1:
(N3−KKKRKV−NH2), C39H77N17O7, [M]+: m/z = 895.62; LC−MS
found for [(M+2H)/2]+ = 448.80. P2: N3−KK(palmitoyl)−NH2),
1
give Y2 as a dark-red solid (37 mg; 15% yield). H NMR (300 MHz,
DMSO-d6) δ 6.98 (d, J = 16.2 Hz, 2H), 7.17 (s, 1H), 7.27 (d, J = 8.4
Hz, 2H), 7.67 (d, J = 8.7 Hz, 4H), 7.97 (d, J = 16.2 Hz, 2H), 8.62 (bs,
1H). 31P NMR (121 MHz, DMSO-d6) δ −5.79. IT-TOF-MS: m/z [M
+H]+ calcd, 493.05; found, 493.06.
2-Hydroxy-4,6-bis(4-bisnitrobenzyl phosphate)pyrimidine (Y3). A
mixture of 2-hydroxy-4,6-dimethylpyrimidine hydrochloride (1; 0.08 g,
0.5 mmol) and 3 (0.71 g, 1.5 mmol; prepared according to ref 34) in
10 mL of ethanol was stirred under an Ar atmosphere at room
temperature for 1 h. Hydrochloric acid (0.2 mL, 3 M) was added
dropwise. The reaction mixture was refluxed for 48 h. The solvent was
removed in vacuo and the resulting residue was washed with DCM
(3×). Next the solid was purified by semipreparative HPLC on a C18
1
column to give Y3 as a dark-red solid (80 mg; 21% yield). H NMR
(300 MHz, DMSO-d6) δ 5.32 (d, J = 7.5 Hz, 4H), 6.98 (d, J = 16.2 Hz,
2H), 7.12 (m, 2H), 7.15 (m, 3H), 7.59 (m, 6H), 7.76 (m, 4H), 8.10
(d, J = 7.8 Hz, 2H), 8.16 (d, J = 16.2 Hz, 2H). 31P NMR (121 MHz,
DMSO-d6) δ −6.23. IT-TOF-MS: m/z [M+H]+ calcd, 763.11; found,
763.09.
2-Propargyloxy-4,6-bis(4-bisnitrobenzyl phosphate) Pyrimidine
(Y4). Y3 (38 mg, 0.05 mmol), propargyl bromide (18 mg, 0.15 mmol)
and K2CO3 (14 mg, 0.1 mmol) were mixed in 5 mL of dry DMF. The
reaction mixture was stirred under an Ar atmosphere at 80 °C for 48 h.
Upon removal of the solvent in vacuo, the resulting residue was
purified by semipreparative HPLC on a C18 column to give Y4 as a
dark-red solid (15 mg; 37.5% yield). 1H NMR (300 MHz, DMSO-d6)
δ 3.53 (s, 1H), 5.11 (s, 2H), 5.32 (d, J = 4.2 Hz, 4H), 7.11 (d, J = 8.4
Hz, 2H), 7.29 (m, 5H), 7.57 (br d, 6H), 7.77 (br s, 4H), 7.87 (d, J =
9.6 Hz, 2H), 8.09 (d, J = 4.5 Hz, 2H). 31P NMR (121 MHz, DMSO-
d6) δ −7.72. IT-TOF-MS: m/z [M+H]+ calcd, 801.13; found, 801.11.
(E)-4-(4-Hydroxystyryl)-6-methylpyrimidin-2-ol (4). A mixture of
2-hydroxy-4,6-dimethylpyrimidine hydrochloride (1; 0.8 g, 5 mmol)
and 4-hydroxybenzaldehyde (0.67 g, 5.5 mmol) in 20 mL of ethanol
was stirred under an N2 atmosphere at room temperature for 1 h.
Hydrochloric acid (2 mL, 3 M) was added dropwise to the reaction
mixture which was then refluxed for 24 h. The solvent was removed,
and the residue was washed with DCM three times. The resulting solid
was purified by flash chromatography (10:1 DCM/methanol) to give 4
as a yellow powder (0.82 g; 72% yield). 1H NMR (300 MHz, DMSO-
d6) δ 2.44 (s, 3H), 6.91 (d, J = 16.0 Hz, 1H), 6.93 (d, J = 8.5 Hz, 2H),
7.16 (s, 1H), 7.59 (d, J = 9.0 Hz, 2H), 8.19 (d, J = 16.0 Hz, 1H). ESI-
MS: m/z [M+H]+ calcd, 229.1; found, 229.2.
C32H62N8O4, [M]+: m/z = 622.49; LC−MS found for [M+H]+
623.50. P3: N3−FxrFxK−NH2, C34H62N12O5, [M]+: m/z = 718.50;
=
L C − M S f o u n d f o r [ M + H ] +
=
7 1 9 . 4 7 . P 4 :
UeeeeeeeeGGPLGLAGrrrrrrrrrK(N3)-NH2, C136H234N58O46, [M]+:
m/z = 3415.78; LC-MS found for [(M+3H)/3]+ = 1139.92, [(M
+4H)/4]+ = 855.44.
Click Assembly of YP1−YP4. For the synthesis of YP1/YP2/YP3,
4 μL of each peptide (20 mM stock), 8 μL of Y4, 8 μL of
CuSO4·5H2O (4 mM stock) and 8 μL of tris-[benzyltriazolylmethyl]-
amine (TBTA) (10 mM stock) were added to 48 μL of DMSO/H2O
(1:1). The resulting solution was mixed, followed by addition of
sodium ascorbate (4 μL in 50 mM stock). The tubes were capped and
shaken at room temperature for 2 days. For YP4 synthesis, 32 μL of
P4 (20 mM stock), 8 μL of Y4 (20 mM stock), 16 μL of CuI (4 mM
stock), and DIEA (10 mM stock) were added to 48 μL of DMSO/
CH3CN/H2O (4:4:1). The tubes were capped and shaken for 24 h.
The products were purified by semipreparatory HPLC and
characterized by LC-MS. YP1: C76H107N21O20P2, [M]+: m/z =
1695.75; LC−MS found for [(M+2H)/2]+ = 848.85. YP2:
C69H92N12O17P2, [M]+: m/z = 1422.62; LC−MS found for [(M
+2H)/2]+ = 712.29. YP3: C71H92N16O18P2, [M]+: m/z = 1518.63;
LC−MS found for [(M+2H)/2]+ = 760.26. YP4: C173H264N62O59P2,
[M]+: m/z = 4215.90; LC−MS found for [(M+3H)/3]+ = 1406.93,
[(M+4H)/4]+ = 1055.45.
Photo Cleavage Assay. The caged phosphate groups of Y3 were
introduced to temporally control the detection of endogenous
phosphatase activities via an UV-mediated uncaging reaction. It was
established that completely uncaged product (Y2) could be obtained
from Y3 (C = 1.6 × 10−6 mol L−1) under conditions of 500 μJ/cm2
UV exposure (at 400 nm) for 2−5 min at room temperature.
In vitro Phosphatase Assay. The expression and purification of
PTP1B were previously reported.35 Alkaline phosphatase was
purchased from Sigma (USA). PP1 and PP2B (2500 U/mL) were
obtained from NEB (USA). PTP1B were prepared to ∼0.2 μg/μL in
4-((E)-2-(2-Hydroxy-6-((E)-4-hydroxystyryl)pyrimidin-4-yl)vinyl)-
phenyl Dihydrogen Phosphate (Y5). A mixture of 4 (0.23 g, 1 mmol)
and diethyl 4-formylphenyl phosphate (2; 0.28 g, 1.5 mmol) in 10 mL
of pyridine and 50 μL of piperidine was refluxed for 24 h. The solvent
was removed, and the resulting residue was washed with ethanol (3×)
and then concentrated in vacuo. Twenty milliliters of DCM was
subsequently added, and the resulting mixture was stirred at room
temperature for 10 min before being cooled on an ice bath. TMSI
(0.46 g, 2.2 mmol) was added dropwise. The reaction was stirred
further for 4 h before being quenched by the addition of 10 mL of
saturated ammonium chloride solution. Upon concentration in vacuo,
the residue was purified by semipreparative HPLC on a C18 column to
1
give Y5 as a dark-red solid (45 mg; 11% yield). H NMR (300 MHz,
DMSO-d6) δ 6.82 (d, J = 16.2 Hz, 1H), 6.0 (d, J = 8.4 Hz, 2H), 6.95
(d, J = 16.2 Hz, 1H), 7.05 (s, 1H), 7.20 (d, J = 8.4 Hz, 2H), 7.55 (d, J
= 8.4 Hz, 2H), 7.60 (d, J = 8.4 Hz, 2H), 7.82 (d, J = 16.2 Hz, 1H), 7.90
(d, J = 16.2 Hz, 1H), 9.35 (bs, 1H). 31P NMR (121 MHz, DMSO-d6)
δ −5.76. ESI-MS: m/z [M+H]+ calcd, 413.1; found, 413.2.
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Journal of the American Chemical Society
Article
Hepes buffer (1×, pH = 7.5, supplemented with 0.05 M NaCl, 2.5 mM
EDTA, 2 mM DTT, and 0.02 or 0.08% Triton X-100). The
experiment for testing PTP1B activities was as the following: PTP1B
(∼2 μg) was incubated with either Y3 or Y2 (1.6 μM final
concentration) in 1 mL of Hepes buffer. For the experiment involving
Y3, the reaction mixture was first irradiated with 500 μJ/cm2 UV
exposure (at 400 nm) for 5 min at room temperature, as above-
described, prior to the addition of PTP1B. The dephosphorylation
reaction was closely monitored by both LC−MS and either a Perkin-
Elmer LS50 spectrofluorometer (Figure 4C) or an in-house two-
photon spectrometer. The reaction rate constant was defined from the
half-life equation for a second-order reaction dependent on one
at 37 °C and washed three times with PBS and imaged. For the
inhibition experiment in tumor cell-specific imaging with YP4, the cells
were first incubated with GM6001 (500 nM, Millipore) for 30 min,
before being treated by the probe and imaged as above-described.
Flow cytometry experiments were done as previously reported.18b
HeLa cells were seeded in Nunc 3.5 cm dishes (cat. no. 153066) and
grown until ∼90% confluency. Two sets of cells were prepared with or
without first being incubated with GM6001 (500 nM, Millipore) for
30 min, before being treated by 10 μM of YP4 (prepared in fresh
media). DMSO was used as negative control. Upon further incubation
for 2 h, cells were UV irradiated (2 min) and further incubated for 0.5
h, then washed with PBS (2×), detached from dishes, washed (with
PBS), fixed with 3.8% paraformaldehyde in PBS for 10 min, and then
analyzed by FACS (Figure 7C).
−1
second-order reactant (k = t1/2 [A]0−1). Experiments with Y5 were
similarly carried out (Figure 4D).
Imaging of Drosophila Brains. Brains were prepared from a one
day-old female Drosophila. Brains were incubated with ELF97 (0.5 h),
Y2 (6 h), and uncaged YP2 (6 h) in DMEM under a humidified
atmosphere of 5:95 (v/v) of CO2/air at 37 °C. For inhibition
experiments, the cells were incubated with Na3VO5 (100 μM) 30 min
prior to incubation with Y2. Treated brains were subsequently washed
three times with PBS and transferred to poly-L-lysine-coated
coverslips, and imaged. The images were taken at ∼35 μm (for ELF
97; see Supporting Information, Figure S10) and ∼110 μm (for all
other experiments) by changing the Z-axis thickness.
For photostability experiments of Y1 and three commercial dyes in
Hepes buffer (Figure 3A), fluorescence intensities were collected at
550 nm for Y1, 450 nm for courmarin, 520 nm for HPQ, and 520 nm
for fluorescein with 60 s intervals using an excitation wavelength 405
nm for Y1, 350 nm for courmarin, 350 nm for HPQ, and 488 nm for
fluorescein, respectively. For the pH effect on the one-photon excited
fluorescence intensity of Y1 at room temperature (Figure 3B), the
fluorescence intensity was collected at 550 nm (λex = 405 nm) with 1.0
μM of the probe prepared in Hepes buffers having different pH values.
General Procedures for 1P and 2P Fluorescence Imaging. Both
HeLa and HepG2 cell lines used in our imaging experiments were
cultured in growth media (DMEM) supplemented with 10% fetal
bovine serum, 100.0 mg/L streptomycin, and 100 IU/mL penicillin.
Cells were maintained in a humidified atmosphere of 5% CO2 at 37 °C
to a confluency of around 80%. Tissues used in our imaging
experiments were fresh brains of one day-old live female Drosophila.
All the trackers were from Invitrogen (Ex = 590 nm for ER-Tracker
Red glibenclamide BODIPY TR (PMT range: 600−650 nm), Ex =
554 nm for MitoTracker Red CMXRos dye (PMT range: 570−650
nm), and Ex = 543 nm for CellMask Plasma Membrane Stains C10045
(PMT range: 555−650 nm)).
For time-dependent one-photon excited fluorescence imaging of
HepG2 cells treated with Y3 (Figure 5), cells were added to Y3 (10
μM) and UV-irradiated for 2 min. Images were collected at 460−550
nm with 60 s intervals using an excitation wavelength at 405 nm. For
control experiments with the four dye-CPP conjugates (Supporting
Information, Figure S6), HeLa cells were added to the tested peptide
followed by further incubation (1 μM for SG-KKKRKV × 2 h, SG-
FxrFxK × 1 h, SG-RRRRRRRRR × 2 h, and 0.5 μM for SG-
KK(palmitoyl) × 30 min). Subsequently, cells were treated with 0.25
μg/mL of the corresponding organelle tracker following protocols
provided by the vendor and washed three times with PBS before being
imaged.
The ELF 97 Endogenous Phosphatase Detection Kit (Invitrogen,
E6601) was used, in control experiments, to endogenous phosphatase
activities in fixed mammalian cells and Drosophila brains. Mammalian
cells were seeded on glass-bottom dishes (Mattek) and grown until
70−80% confluency. The growth medium was removed; the cells were
first fixed with 3.7% paraformaldehyde in PBS for 10 min and washed
three times with PBS. The cells were incubated with 0.25 μg/mL of
the corresponding organelle tracker. Next, the fixed cells were
incubated with 1 × ELF 97 in detection buffer and imaged
(Supporting Information, Figure S4).
Live-Cell Imaging with Organelle-Specific Probes. Four probe-
peptide conjugates (YP1 to YP4) were used to image endogenous
phosphatase activities in live HeLa and HepG2 cells. Cells were seeded
in glass-bottom dishes (Mattek) and grown until 70−80% confluency.
Subsequently, the cells were incubated with the peptide (10 μM for
YP1, YP3, and YP4, 8 μM for YP2; prepared in fresh media). Upon
further incubation (2 h for YP1 and YP4, 1 h for YP2 and YP3), cells
were UV irradiated (2 min) then further incubated with 0.25 μg/mL of
the organelle tracker. Next, cells were washed three times with PBS
before and immediately imaged with Leica TCS SP5X Confocal
Microscope System. Meanwhile, another identical set of samples was
similarly treated as described above and without UV-irradiated for
negative control. All the cells were then treated for further incubation
ASSOCIATED CONTENT
* Supporting Information
■
S
Other relevant experimental sections, spectral characterization
of new compounds, and supplementary biological results. This
material is available free of charge via the Internet at http://
pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
chmyaosq@nus.edu.sg
■
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
Financial support was provided by the Agency for Science,
Technology and Research (R-143-000-391-305) and the
Ministry of Education (R-143-000-394-112). We also acknowl-
edge Madam Tong Yan in the Department of Biological
Sciences (NUS) for her technical support on two-photon
microscopy and Dr Lim Kah Leong (NUS) for the generous
donation of the Drosophila brains.
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