Electronic and Steric Optimization of Fluorogenic Probes for Biomolecular Imaging

Article abstract of DOI:10.1021/acs.joc.7b00285

Fluorogenic probes are invaluable tools for spatiotemporal investigations within live cells. In common fluorogenic probes, the intrinsic fluorescence of a small-molecule fluorophore is masked by esterification until entry into a cell, where endogenous esterases catalyze the hydrolysis of the masking groups, generating fluorescence. The susceptibility of masking groups to spontaneous hydrolysis is a major limitation of these probes. Previous attempts to address this problem have incorporated auto-immolative linkers at the cost of atom economy and synthetic adversity. Here, we report on a linker-free strategy that employs adventitious electronic and steric interactions in easy-to-synthesize probes. We find that X···C = O n→π? interactions and acyl group size are optimized in 2′,7′-dichlorofluorescein diisobutyrate. This probe is relatively stable to spontaneous hydrolysis but is a highly reactive substrate for esterases both in vitro and in cellulo, yielding a bright, photostable fluorophore with utility in biomolecular imaging.

Full text of DOI:10.1021/acs.joc.7b00285

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Article
Electronic and Steric Optimization of
Fluorogenic Probes for Biomolecular Imaging
Wen Chyan, Henry R. Kilgore, Brian Gold, and Ronald T Raines
J. Org. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.joc.7b00285 • Publication Date (Web): 27 Mar 2017
Downloaded from http://pubs.acs.org on March 29, 2017
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Electronic and Steric Optimization of Fluorogenic Probes
for Biomolecular Imaging
Wen Chyan,† Henry R. Kilgore,‡ Brian Gold,† and Ronald T. Raines*,†,§
†Department of Chemistry, ‡Graduate Program in Biophysics, and §Department of Biochemistry,
University of Wisconsin–Madison, Madison, Wisconsin 53706, United States
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ABSTRACT: Fluorogenic probes are invaluable t
http://www.sbnation.com/2017/3/18/14968936/nigel-hayes-game-winner-wisconsin-villanova-
michael-jordan ools for spatiotemporal investigations within live cells. In common fluorogenic
probes, the intrinsic fluorescence of a small-molecule fluorophore is masked by esterification
until entry into a cell, where endogenous esterases catalyze the hydrolysis of the masking groups,
generating fluorescence. The susceptibility of masking groups to spontaneous hydrolysis is a
major limitation of these probes. Previous attempts to address this problem have incorporated
auto-immolative linkers at the cost of atom economy and synthetic adversity. Here, we report on
a linker-free strategy that employs adventitious electronic and steric interactions in easy-to-
synthesize probes. We find that X···C=O n→π* interactions and acyl group size are optimized in
2′,7′-dichlorofluorescein diisobutyrate. This probe is relatively stable to spontaneous hydrolysis
but is a highly reactive substrate for esterases both in vitro and in cellulo, yielding a bright,
photostable fluorophore with utility in biomolecular imaging.
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INTRODUCTION
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Fluorogenic probes with specific responses to physiological events or environmental conditions
are invaluable for deciphering complex biological processes.1 Masked probes are a class of
fluorogenic probes in which a pendant functional group attenuates the fluorescence of a
fluorophore.2 Fluorescence is restored upon removal of the masking group by an enzyme-
catalyzed or uncatalyzed chemical reaction. In cell biological applications, masking groups are
frequently designed to serve as substrates for esterases,3 phosphatases,4 azoreductases,5 or
cytochrome P450s.6 Caged fluorophores (which are also known as photoactivatable
fluorophores) are related but are activated instead by illumination at specific wavelengths.7
Fluorogenic probes that are substrates for esterases are of special interest because they can be
activated by an endogenous intracellular enzyme.8 Conjugation of fluorogenic esterase-activated
probes to biomolecules can provide detailed spatiotemporal information about biomolecular
uptake and localization in live cells.3a,9 These biomolecule–probe conjugates are, however,
typically internalized by endocytic vesicles and can be exposed therein to acidity as low as
pH 4.5,10 making insensitivity to low pH essential to probe function.
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Halogenation of xanthene dyes is a reliable strategy for altering spectroscopic properties and
tuning dye pKa to match those desired for biological applications. Oregon green, which is a
fluorescein derivative in common use, is fluorinated at the 2′ and 7′ positions.11 Although
fluorogenic probes based on fluorinated and chlorinated scaffolds are available from commercial
vendors, little is known about the effects of halogenation on probe stability. Prior work with
fluorinated derivatives demonstrated improved photostability, but accompanied by the
accelerated spontaneous loss of masking groups.12 In contrast, a report of an unusually stable
chlorinated probe13 suggested that halogens other than fluorine merit attention. With fluorinated
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Oregon green, destabilization of the masked substrate was thought to stem from inductive effects
resulting in lowered pKa of the conjugate acid of the fluorescein leaving group.8d Accordingly,
design strategies for stable fluorogenic esterase probes have relied heavily on interjecting self-
immolative linkers with a higher pKa between the low pKa fluorophore and the site of enzymatic
cleavage (Scheme 1).12 Platforms for such auto-immolative linkers include the acetoxymethyl
(AM) ether,8d,12 quinone methides,14 and the trimethyl lock.3a,15 The beneficial stability provided
by auto-immolative linkers does, however, come at the expense of a longer synthetic route to add
atoms that are, ultimately, unnecessary.
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Scheme 1. Auto-immolative linkers (red) inserted between fluorophores and esterase targets to
enhance stability.
The AM ether linker has a small size and facile synthetic accessibility compared to other
auto-immolative linkers.8d Still, AM ether masking groups are installed on fluorescein by
O-alkylation, which often yields undesirable ether–ester mixed byproducts from O-alkylation of
the 2-carboxyl group.16 Addition of a 6-amido group for bioconjugation exacerbates the problem
by shifting the equilibrium away from the “closed” lactone form of a fluorescein derivative and
towards the “open” quinoid form. Accordingly, we sought a simple “linker-free” probe.
Here, we combine electronic and steric effects to create linker-free fluorogenic probes with
high hydrolytic stability, enzymatic reactivity, and photostability. We begin by characterizing the
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effects of ortho-halogenation to identify an optimal substitution pattern. Then, we identify an
ideal acyl masking group after searching for a high rate of enzyme-catalyzed unmasking along
with a low rate of spontaneous hydrolysis. The ensuing probe is small and readily accessible, and
has superior photostability and enzymatic unmasking kinetics in vitro and in cellulo relative to
auto-immolative probes.
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RESULTS AND DISCUSSION
Tuning Acyl Probe Stability with Halogenation. Previous reports have hinted at a role for
halogenation in probe stability.8d,13 We pursued this strategy, synthesizing halogenated
fluorescein diacetate probes 1a–e to characterize ortho-halogen effects (Scheme 2).17
Scheme 2. Halogenated fluorescein diacetate probes.
We began by assessing the spectrophotometric properties of the unmasked probes. The
product of the extinction coefficient and the quantum yield (ε × Φ) accounts for both the amount
of light absorbed by a fluorophore and its quantum efficiency. This product is directly
proportional to the brightness of the dye. By this measure, the hydrolysis of chlorinated probe 1c
provides the brightest fluorophore.
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Table 1. Spectroscopic attributes and pKa values of unmasked halogenated fluorescein
diacetate probes.
ε (M–1cm–1)
9.3 × 104
Φ
ε × Φ (M–1cm–1)
8.4 × 104
pKa
6.418
4.711
4.6
λabs (nm)
Probe
490
492
503
525
521
1a
1b
1c
1d
1e
0.9218
0.92
8.6 × 104
7.9 × 104
1.01 × 105
1.12 × 105
8.25 × 104
0.88
8.9 × 104
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0.2419
0.0219
2.7 × 104
3.820
3.820
1.6 × 103
Next, we assessed spontaneous hydrolysis by incubating each compound in either a simple
buffer or a mammalian cell culture medium. The observed rates of spontaneous hydrolysis for
probes 1a–e varied with ortho substituents in the order F > H > Cl > Br > I (Figure 1). Moreover,
probes 1c–e exhibited increased stability relative to fluorinated probe 1b, despite their low pKa
values (Table 1), suggesting that inductive electron-withdrawal is not the dominant contributor to
probe stability with larger halogen substituents.
We hypothesized that the resistance of probe 1c to hydrolysis was due to stabilization by a
donor–acceptor interaction. Specifically, donation of a lone-pair of electrons (n) from an ortho-
halo group into the antibonding orbital of the adjacent carbonyl group (π*)—an n→π*
interaction21—could decrease the electrophilicity of the carbonyl group by raising the energy of
its π* orbital.22 In addition, an intimate interaction with a halo group would shield one face of the
acyl group from nucleophilic attack by water.
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Figure 1. Graphs showing the time-course for the spontaneous hydrolysis of probes 1a–e as
measured by the generation of fluorescence. (A) Hydrolysis in 10 mM HEPES–NaOH buffer,
pH 7.3. (B) Hydrolysis in OptiMEM cell culture medium supplemented with FBS (10% v/v).
To characterize the interaction between the halo and acyl groups in 1a–e at higher resolution,
we synthesized o-halophenyl acetates 2a–e (Scheme 3). Infrared carbonyl stretching frequencies
can report on electronic effects on carbonyl groups,23 including n→π* interactions.24 We found
that the introduction of ortho halogens induced modest hypsochromic shifts in the carbonyl
stretching frequencies of 2a–e with magnitudes in the order: Cl > Br > F > I > H, whereas the
hypsochromic shifts observed in 1a–e followed the order: Br > F ~ Cl ~ I > H (Table 2). The
observed hypsochromic shifts in 2a–e follow a pattern similar to that of known rate constants for
the hydrolysis of o-halophenyl acetates.25 Still, hypsochromic shifts of 1a–e and 2a–e did not
follow a pattern based simply on electron-withdrawal. Accordingly, we turned to quantum
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mechanical calculations to evaluate the origin of the anomalous hypsochromic shifts and their
implication in the observed hydrolysis trends.
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Scheme 3. Halogenated model compounds.
Table 2. Carbonyl stretching frequencies, donor–acceptor geometries and interaction energies,
and hydrolytic stabilities of compounds 1a–e and 2a–e.
νC=O
En→π*
∆EX,C=O
Probe
dX···C (Å)b
θX···C=O (°)b
t1/2 (h)c
(cm–1)a
(kcal/mol)b
(kcal/mol)b
1a
1b
1c
1d
1e
2a
2b
2c
2d
2e
1766
1774
1774
1776
1774
1766
1770
1774
1772
1767
3.06
2.97
80.9
77.0
—
0.22
0.20
0.85
4.7
1.8
11
3.25
84.7
0.54
0.98
0.46d, 0.54e
0.81d, 1.01e >2,000
3.50d, 3.55e
3.62d, 3.65e
3.41
91.0d, 92.1e
91.0d, 92.1e
85.1
0.25d, 0.33e
0.40d, 0.44e
—
33
0.11
0.19
1.10
1.17
1.03
ND
ND
ND
ND
ND
3.27
85.9
0.22
3.00
85.8
0.51
3.13
87.6
0.52
3.59
91.1
0.51
aMeasured with FT-IR spectroscopy. bCalculated for each X···C=O interaction in the optimized
geometry. cExperimental half-life for spontaneous hydrolysis in OptiMEM containing FBS (10%
v/v). dData for 4′,5′-halo substituents. eData for 2′,7′-halo substituents. ND, not determined.
We used second-order perturbation theory calculations provided by Natural Bond Orbital
(NBO) analysis26 to assess possible n→π* interactions in compounds 2a–e. The stabilizing
effects of n→π* interactions were pivotal for each compound but could not provide the sole
explanation for the observed trends. The zenith in n→π* interaction energies (En→π*) occurs
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when X = Cl (Figure 2), suggesting that increasing the size of the halogen plays dichotomous
roles. Every favorable n→π* interaction is counteracted, at least partially, by unfavorable Pauli
repulsion between the lone pair and π bonding orbital—a factor that is of increasing importance
for larger halogen atoms. The steric exchange energy (∆EX,C=O), which is the energetic penalty
associated with the overlap of the lone pair and π bonding orbital, is substantial only in
compounds bearing larger halo substituents: Cl (1c and 2c), Br (1d and 2d), and I (1e and 2e).
Hence, we proceeded to assess in greater detail how n)(π Pauli repulsion contributes to the
reactivity of the acyl masking groups in compounds 2a–e.
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Figure 2. Graph showing the strength of n→π* interactions in compounds 2a–e as calculated
with second-order perturbation theory. Data are listed in Table 1. Insert: NBO orbital rendering
of n→π* interactions in 2-chlorophenyl acetate (2c).
Potential energy surfaces can provide valuable insight into the interplay between n→π*
interactions and n)(π Pauli repulsion.22,27 Favorable interactions dominate when the dX···C
distance and θX···C=O′ angle between the halo and carbonyl groups provide sufficient orbital
overlap for n→π* donation, generating a trough in the potential energy surface. This surface is
manicured further by unfavorable steric interactions [e.g., n)(π Pauli repulsion] when the value of
d is too small for a particular value of θ (Figure S1).
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To provide additional information, we calculated potential energy surfaces for compounds
2a–d by scanning the Ca–Cb–O–Cc dihedral angle (Figure 3). The dominant feature in these
surfaces is a trough in which n→π* interactions are favorable and steric repulsion is minimal.
The most productive angle formed between an attacking nucleophile and carbonyl group for the
formation of a tetrahedral intermediate is the Bürgi–Dunitz trajectory.28 Due to the covalent
nature of n→π* interactions, energies are minimized near the Bürgi–Dunitz trajectory. Moving
along the Bürgi–Dunitz trajectory (~107°), the unfavorable n)(π interaction dominates until d ≈
3.6 and 3.8 Å for X = Cl and Br, respectively (Figures 3C and 3D), which recapitulates the
length of the C–X bond. When X = F (Figure 3B), the small van der Waals radii and weak
overlap of 2p orbitals restrict favorable conformations to relatively small values of d and θ. In
the absence of halo substituents, the surface has a singular trough (Figure 3A), unaltered by
significant changes in steric repulsion. Thus, the observed trend in carbonyl stretching
frequencies (Table 1) is a balance between n→π* interactions, n)(π Pauli repulsion, and through-
bond inductive and resonance effects. These findings, in conjunction with spectroscopic
attributes (Table 1), anoint 2′,7′-dichlorofluorescein-based probes as having an optimal
combination of stability and brightness.
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Optimization of the Acyl Masking Group. Encouraged by the attributes endowed by
2′,7′-dichlorination, we suspected that tuning the sterics of the acyl mask group could enhance
probe stability. In particular, an ideal acyl masking group could provide steric hindrance to
spontaneous hydrolysis without slowing the rate of enzymatic cleavage.29 Accordingly, we
synthesized a small library of fluorogenic probes with various acyl masking groups (Scheme 4).
Then, we assessed their susceptibility to spontaneous hydrolysis and enzymatic unmasking in
vitro.
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Figure 3. Calculated potential energy surfaces generated by scanning the Ca–Cb–O–Cc dihedral
angle of compounds 2a–d. Minimal energies (blue) follow a trough that correlates with favorable
n→π* interactions for a given X···Cc distance (d) and X···Cc=O′ angle (θ).
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Scheme 4. Acylated 2′,7′-dichlorofluorescein probes.
As expected, the combination of steric and n→π* stabilization in probes 3a–g reduced the
rate of spontaneous hydrolysis significantly (Figure 4A). Nevertheless, the bulkier acyl groups in
probes 3c–g also diminished the rate of enzymatic hydrolysis in vitro (Figure 4B). The isobutyryl
masking groups in probe 3b provided the best combination of increased stability (~10-fold
greater than that of fluorescein diacetate) and rapid enzymatic unmasking.
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Figure 4. Graphs showing the effect of acyl groups on the hydrolytic stability of
2′,7′-dichlorofluorescein probes in vitro and in cellulo. (A) Spontaneous hydrolysis after a 24-h
incubation in OptiMEM containing FBS (10% v/v). Raw data are shown in Figure S2B. Inset:
structure of compound AM. (B) Hydrolysis by pig liver esterase in 1 h. Raw data are shown in
Figure S2C. (C) Hydrolysis by intracellular esterases in live HeLa cells. Data were quantified
from images in Figure 5. RFU: relative fluorescence units.
The resistance of probe 3b to spontaneous hydrolysis likely arises from a combination of
electronic and steric effects. In its optimized geometry, one face of its ester carbonyl group is
shielded from solvent water by an n→π* interaction with the ortho-chloro group (Scheme 5).
The other face is shielded by a methyl group. These effects might be less detrimental to enzyme-
catalyzed hydrolysis.
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Scheme 5. Optimized geometry of the butyryl ester moiety in probe 3b.
We evaluated the steady-state kinetic parameters for esterase-catalyzed hydrolysis of the
probes (Figure S3). Although the probes undergo two step hydrolysis, full fluorescence is
generated only after release of the second ester group. Accordingly, hydrolysis data can be
reliably fit to the Michaelis–Menten equation to obtain apparent kinetic parameters. Chlorinated
probes tended to interact more strongly with the enzymic active site than did unmodified or
fluorinated probes (Table S1). The additional steric bulk in butyryl probe 3b had only a modest
effect on the rate of enzymatic hydrolysis compared to acetyl probe 1c. Optimized acyl probe 3b,
with apparent kcat/KM = 1.8 × 106 M–1s–1 and KM = 4.6 µM, outperformed probes with auto-
immolative linkers (Table S1).
Human esterases often exhibit higher substrate specificity than does pig liver esterase.30
Accordingly, we sought to corroborate in vitro kinetic data with in cellulo data to ensure probe-
applicability in human cells. Confocal images of live HeLa cells incubated with probes 3a–g
confirmed the trends observed in vitro (Figures 4C and 5). Additionally, probe 3b showed
enhanced rates of enzyme-catalyzed unmasking compared to the analogous AM ether probe
(AM). Finally, we monitored the fluorescence of the 2′,7′-dichlorofluorescein scaffold in live
cells under constant illumination and found that probes based on this scaffold have superior
photostability (Figure S4).
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CONCLUSIONS
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We have described a new strategy for stabilizing esterase-activated fluorogenic probes. An
n→π* interaction between an ortho-halogen and pendant acyl group in 2′,7′-fluorescein diacetate
deters spontaneous hydrolysis. n)(π Pauli repulsion from larger halo groups limits the benefit that
can be gained from this n→π* interaction, and an optimum is achieved with chloro-substitution.
Spontaneous hydrolysis is deterred further with little effect on esterase-catalyzed cleavage when
the esters derive from isobutyric acid rather than acetic acid. Thus, 2′,7′-dichlorofluorescein
diisobutyrate is a simple linker-free probe derived by optimizing electronic and steric effects.
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Figure 5. In cellulo hydrolysis of fluorogenic probes 1a, 1c, 3a–g, and AM. HeLa cells were
incubated with a probe (5 µM) for 15 min, counterstained with Hoechst 33392, and imaged by
confocal microscopy. Quantitation shows that cells incubated with 1c or 3b had comparable
levels of fluorescence signal generation (Figure 5). Scale bars: 25 µm.
EXPERIMENTAL SECTION
General Information. Phenyl acetate (2a), 2-iodophenyl acetate (2e), and all other commercial
chemicals were from Sigma–Aldrich (St. Louis, MO), Fischer Scientific (Hampton, NH), or Alfa
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Aesar (Haverhill, MA) and were used without further purification. Porcine liver esterase (PLE)
was from Sigma–Aldrich.
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Chemical reactions were monitored by thin-layer chromatography (TLC) using EMD 250-
µm silica gel 60-F254 plates and visualization with UV illumination or KMnO4-staining. Flash
chromatography was performed with a Biotage Isolera automated purification system using pre-
packed SNAP KP silica gel columns.
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All procedures were performed in air at ambient temperature (~22 °C) and pressure (1.0 atm)
unless specified otherwise. The phrase “concentrated under reduced pressure” refers to the
removal of solvents and other volatile materials using a rotary evaporator at water aspirator
pressure (<20 torr) while maintaining a water-bath temperature below 40 °C. Residual solvent
was removed from samples at high vacuum (<0.1 torr), which refers to the vacuum achieved by
mechanical belt-drive oil pump.
All fluorogenic probes and fluorescent molecules were dissolved in spectroscopic grade
DMSO and stored as frozen stock solutions. For all applications, DMSO stock solutions were
diluted such that the DMSO concentration did not exceed 1% v/v.
Instrumentation. Absorbance data were acquired with an Agilent Cary 60 UV-vis
spectrometer. Hydrolysis kinetics were measured with a Tecan Infinite M1000 plate reader. All
other fluorescence data were acquired with a PTI QuantaMaster spectrofluorometer. 1H and 13C
NMR spectra were acquired on Bruker Spectrometers at the National Magnetic Resonance
Facility at Madison (NMRFAM) operating at 500 MHz for 1H and 125 MHz for 13C. Mass
spectrometry was performed with a Q ExactiveTM Plus electrospray ionization quadrupole-ion
trap (ESI–QIT-MS) mass spectrometer at the Mass Spectrometry Facility in the Department of
Chemistry at the University of Wisconsin–Madison. IR spectra were acquired with a Micro FT-
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IR spectrometer at the Materials Science Center of the University of Wisconsin–Madison.
Microscopy images were acquired with a Nikon Eclipse Ti inverted confocal microscope at the
Biochemistry Optical Core of the University of Wisconsin–Madison.
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Optical Spectroscopy. UV–visible and fluorescence spectra were recorded by using 1-cm
path length, 4-mL quartz cuvettes or 1-cm path length, 1-mL quartz microcuvettes. Analyte
solutions were stirred with a magnetic stir bar. Quantum yields were determined by referencing
probe solutions to fluorescein (λex = 495 nm; Φ = 0.95) in 0.1 M NaOH(aq)
.
FT-IR spectra were recorded on compounds solvated with a minimum quantity of
dichloromethane or dimethyl sulfoxide, and sandwiched between two sodium chloride windows.
FT-IR spectra were collected with 128 scans at 1200–3500 cm–1 and a resolution of 2 cm–1. A
background spectrum was taken of the solvent alone every 20 min. Plates were washed with
acetone (3×) after recording the spectrum of each compound.
Spontaneous Probe Hydrolysis. Probe stocks were diluted to a final concentration of 5 µM
in 300 µL of either 10 mM HEPES–NaOH buffer, pH 7.3, or OptiMEM containing FBS
(10% v/v). Fluorescence was measured with a plate reader (Costar 96 well clear bottom, bottom
measurement mode) at 30-min intervals for 72 h. Hydrolysis data were fitted to single-phase
decay curves with GraphPad Prism software.
PLE-Catalyzed Probe Hydrolysis. PLE (168 kDa, ≥15 units/mg solid) was suspended in 10
mM HEPES–NaOH buffer, pH 7.3, and diluted to appropriate concentrations before use in
protein LoBind tubes from Eppendorf. Initial rate measurements for each probe were acquired
and the resulting data was fit to the Michaelis–Menten equation in GraphPad Prism software to
obtain apparent kinetic parameters for the enzyme-catalyzed unmasking of probes.
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Cell Culture and Live Cell Imaging. HeLa cells were from American Type Culture
Collection (Manassas, VA) and were maintained according to recommended procedures. Gibco
brand Dulbecco’s Modified Eagle Medium (DMEM), fetal bovine serum (FBS), trypsin (0.25%
w/v), OptiMEM, and Dulbecco’s PBS (DPBS) were from Thermo Fisher Scientific (Waltham,
MA). HeLa cells were grown in DMEM supplemented with FBS (10% v/v), penicillin (100
units/mL), and streptomycin (100 µg/mL). For all imaging experiments, 8-well microscopy
slides from Ibidi (Madison, WI) were seeded with HeLa cells (105 cells/mL) 24 h before use. All
imaging experiments were performed in live cells without fixation. ImageJ software from the
National Institutes of Health (Bethesda, MD) was used for all image processing, signal
quantification, and colocalization measurements.31
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In Cellulo Probe Hydrolysis. HeLa cells in 8-well microscopy slides were incubated with
5 µM probe for 15 min, and counterstained with Hoechst 33342 (2 µg/mL) for 10 min at 37 °C.
Cells were washed twice and imaged with a confocal microscope. Background-subtracted cell
fluorescence signal was quantified in each image.
Photobleaching. HeLa cells in 8-well microscopy slides were incubated with 5 µM of 1a,
1b, or 1c for 15 min, and counterstained with Hoechst 33342 (2 µg/mL) for 10 min at 37 °C. The
cells were washed twice and imaged with a confocal microscope using a 488-nm filter set.
Images were acquired every 2 min with continuous, constant-intensity illumination between
acquisitions. Normalized fluorescence signal was quantified with ImageJ software and plotted in
Figure S4.
Computational Procedures. Geometry optimization calculations were performed with
Gaussian 09, revision D.01 with the functional M06-2X and 6-311+g(2d,p) basis set.32
Frequency calculations were performed to ensure that the optimized structure was at a true
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minimum. All calculations were performed in water with the integral formalism polarizable
continuum model (IEFPCM implicit solvent model as implemented in Gaussian 09). Potential-
energy surfaces were generated by varying the Ca–Cb–O–Cc dihedral using a relaxed scan. All
calculations performed on systems containing iodine were done by using the Stuttgart relativistic
electron core potential for treating iodine, whereas the 6-311+g(2d,p) basis set was used for all
other atoms. NBO calculations were performed with NBO 6.0.26 The pairwise steric exchange
energy (∆EX,C=O) between the lone pair of a halo group and the π orbital of a carbonyl group was
calculated using natural steric analysis as implemented in NBO 6.0. All energies include zero-
point corrections.
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2′,7′-Dichlorofluorescein. 4-Chlororesorcinol (14.4 g, 99.6 mmol) and phthalic anhydride
(7.3 g, 49.3 mmol) were dissolved in MeSO3H (50 mL), and the resulting solution was heated at
90 °C for 24 h. After cooling to room temperature, the reaction mixture was added slowly to 1 L
of stirred ice water. The resulting suspension was filtered and triturated with cold water to afford
2′,7′-dichlorofluorescein as a yellow solid (18.1 g, 90.7% yield). 1H NMR (500 MHz, (CD3)2SO,
δ): 11.08 (s, 2H), 8.01 (d, J = 7.7 Hz, 1H), 7.82 (t, J = 7.7 Hz, 1H), 7.75 (t, J = 7.6 Hz, 1H), 7.34
(d, J = 7.7 Hz, 1H), 6.91 (s, 2H), 6.66 (s, 2H). 13C NMR (125 MHz, (CD3)2SO, δ): 168.3, 155.1,
151.5, 150.1, 135.9, 130.5, 128.2, 125.9, 125.1, 124.0, 116.3, 110.5, 103.7, 81.5. HRMS (ESI–
QIT) m/z: [M + H]+ Calcd for C20H11Cl2O5 400.9979; Found 400.9982.
Diesters 1a–e and 3a–h, and Esters 2b–d. To a suspension of phenol or fluorescein
derivative (0.15 mmol, 1 equiv) in DCM (2.0 mL) were added 4-dimethylaminopyridine
(15 µmol, 0.1 equiv) and pyridine (0.33 mmol, 2.2 equiv). Acyl chloride (0.33 mmol, 2.2 equiv)
was added dropwise, and the resulting solution was stirred for 1 h or until completion of the
reaction. After dilution with water and extraction with DCM, the combined organic extracts were
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washed with saturated aqueous NH4Cl and brine, dried with MgSO4(s), and concentrated under
reduced pressure. Purification by column chromatography on silica gel (0–40% v/v EtOAc in
hexanes) afforded the title compounds as white solids or clear oils.
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Characterization data for fluorescein diacetate (1a) and 2′,7′-difluorofluorescein diacetate (1b)
were in accord with those reported previously.12
2′,7′-Dichlorofluorescein Diacetate (1c). Off-white solid (68.6 mg, 94.2% yield). 1H NMR
(500 MHz, CDCl3, δ): 8.08 (d, J = 7.6 Hz, 1H), 7.73 (dd, J = 26.5, 1.0 Hz, 2H), 7.22 (d, J = 7.6
Hz, 1H), 7.16 (s, 2H), 6.87 (s, 2H), 2.38 (s, 6H). 13C NMR (125 MHz, CDCl3, δ): 168.7, 168.1,
151.9, 149.8, 148.6, 136.0, 130.8, 129.1, 125.8, 125.8, 124.2, 122.8, 117.8, 112.9, 80.6, 20.8.
HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C24H15Cl2O7 485.0190; Found 485.0196.
Eosin Y diacetate (1d). White solid (40.6 mg, 37% yield). 1H NMR (500 MHz, CDCl3, δ):
8.13 (dd, J = 7.5, 1.0 Hz, 1H), 7.82 (t, J = 7.2 Hz, 1H), 7.77 (dt, J = 7.5, 3.8 Hz, 1H), 7.28 (d, J =
7.3 Hz, 1H), 7.07 (s, 2H), 2.48 (s, 6H). 13C NMR (125 MHz, CDCl3, δ): 171.0, 169.5, 151.1,
150.6, 138.8, 133.7, 132.6, 128.7, 127.9, 126.7, 121.9, 115.4, 111.1, 82.8, 23.3. HRMS (ESI–
QIT) m/z: [M + H]+ Calcd for C24H13Br4O7 728.7389; Found 728.7390.
Erythrosin B Diacetate (1e). White solid (44.2 mg, 32% yield). 1H NMR (500 MHz, CDCl3,
δ): 8.09 (d, J = 7.7 Hz, 1H), 7.78 (m, 1H), 7.72 (t, J = 7.4 Hz, 1H), 7.22 (m, 3H), 2.46 (s, 4H).
13C NMR (125 MHz, CDCl3, δ): 168.4, 166.9, 154.0, 152.0, 137.4, 137.3, 136.1, 130.9, 125.9,
125.2, 124.1, 119.5, 84.4, 83.1, 21.4. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C24H13I4O7,
920.6835; Found 920.6837.
2-Fluorophenyl Acetate (2b). Clear oil (9 mg, 39% yield). 1H NMR (500 MHz, CDCl3, δ):
7.22– 7.07 (m, 1H), 2.35 (s, 1H). 13C NMR (125 MHz, CDCl3, δ): 168.5 (s), 154.2 (d, JC–F = 249
Hz), 138.2 (d, JC–F = 13.0 Hz), 127.2 (d, JC–F = 7.2 Hz), 124.6 (d, JC–F = 3.9 Hz), 123.8 (d, JC–F
=
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0.8 Hz), 116.8 (d, JC–F = 18.6 Hz), 20.6 (s). HRMS (ESI–QIT) m/z: [M + H]+ Calcd for
C8H8FO2, 155.0503; Found, 155.0502.
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2-Chlorophenyl Acetate (2c). Clear oil (7.9 mg, 31% yield). 1H NMR (500 MHz, CDCl3, δ):
7.46 (dd, J = 8.0, 1.6 Hz, 1H), 7.33–7.27 (m, 1H), 7.20 (td, J = 7.7, 1.6 Hz, 1H), 7.15 (dd, J =
8.0, 1.6 Hz, 1H), 2.37 (s, 3H). 13C NMR (125 MHz, CDCl3, δ): 168.5, 146.9, 130.3, 127.7, 127.0,
126.8, 123.7, 20.6. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C8H8ClO2 171.0207; Found
171.0206
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2-Bromophenyl Acetate (2d). Clear oil (9.4 mg, 29% yield). 1H NMR (500 MHz, CDCl3, δ):
7.46 (dd, J = 8.0, 1.6 Hz, 1H), 7.33–7.27 (m, 1H), 7.20 (td, J = 7.7, 1.6 Hz, 1H), 7.15 (dd, J =
8.0, 1.6 Hz, 1H), 2.37 (s, 3H). 13C NMR (125 MHz, CDCl3, δ): 168.7, 148.4, 133.5, 128.7, 127.6,
123.9, 116.4, 21.0. HRMS (ESI–QIT) m/z: [M + NH4] + Calcd for C8H11NBrO2 231.9968; Found
231.9965.
2′,7′-Dichlorofluorescein Dipropionate (3a). White solid (66.2 mg, 86% yield). 1H NMR (500
MHz, CDCl3, δ): 8.08 (d, J = 7.6 Hz, 1H), 7.73 (dtd, J = 31.0, 7.5, 0.9 Hz, 2H), 7.22 (d, J = 7.6
Hz, 1H), 7.16 (s, 2H), 6.87 (s, 2H), 2.67 (q, J = 7.5 Hz, 4H), 1.31 (t, J = 7.6 Hz, 6H). 13C NMR
(125 MHz, CDCl3, δ): 171.6, 168.7, 152.0, 149.9, 148.7, 135.9, 130.8, 129.0, 125.8, 125.8,
124.1, 122.8, 117.7, 112.9, 80.6, 27.6, 9.1. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for
C26H19Cl2O7 513.0502; Found 513.0503.
2′,7′-Dichlorofluorescein Diisobutyrate (3b). White solid (60.1 mg, 74% yield). 1H NMR (500
MHz, CDCl3, δ): 8.08 (d, J = 7.6 Hz, 1H), 7.74 (tdd, J = 15.0, 11.0, 4.1 Hz, 2H), 7.22 (d, J = 7.6
Hz, 1H), 7.14 (s, 2H), 6.87 (s, 2H), 2.96–2.79 (m, 2H), 1.36 (dd, J = 7.0, 4.0 Hz, 12H). 13C NMR
(125 MHz, CDCl3, δ): 174.1, 168.6, 152.0, 149.8, 148.6, 135.8, 130.7, 128.9, 125.7, 125.7,
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124.0, 122.7, 117.5, 112.7, 80.5, 34.2, 18.9, 18.9. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for
C28H23Cl2O7 541.0815; Found 541.0815.
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2′,7′-Dichlorofluorescein Dipivalate (3c). White solid (73.5 mg, 86% yield). 1H NMR (500
MHz, CDCl3, δ): 8.08 (d, J = 7.5 Hz, 1H), 7.72 (dtd, J = 29.1, 7.4, 1.0 Hz, 2H), 7.20 (d, J = 7.6
Hz, 1H), 7.13 (s, 2H), 6.87 (s, 2H), 1.40 (s, 18H). 13C NMR (125 MHz, CDCl3, δ): 175.6, 168.6,
152.0, 149.8, 148.8, 135.8, 130.6, 128.9, 125.7, 125.7, 124.0, 122.8, 117.4, 112.7, 80.6, 39.4,
27.1. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C30H27Cl2O7 569.1128; Found 569.1128.
2′,7′-Dichlorofluorescein Di-tert-butylacetate (3d). White solid (76.2 mg, 85% yield). 1H
NMR (500 MHz, CDCl3, δ): 8.08 (d, J = 7.6 Hz, 1H), 7.73 (dtd, J = 23.7, 7.4, 1.0 Hz, 2H), 7.20
(d, J = 7.5 Hz, 1H), 7.14 (s, 2H), 6.88 (s, 2H), 2.53 (s, 4H), 1.16 (s, 16H). 13C NMR (125 MHz,
CDCl3, δ): 169.3, 168.6, 152.0, 149.7, 148.5, 135.8, 130.6, 129.0, 125.7, 125.6, 124.0, 122.7,
117.5, 112.7, 80.5, 47.3, 31.1, 29.6. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C32H31Cl2O7
597.1441; Found 597.1438.
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2′,7′-Dichlorofluorescein Dicyclobutyrate (3e). White solid (78 mg, 92% yield). 1H NMR (500
MHz, CDCl3, δ): 8.08 (d, J = 7.6 Hz, 1H), 7.79–7.68 (m, 2H), 7.22 (d, J = 7.6 Hz, 1H), 7.15 (s,
2H), 6.87 (s, 2H), 3.52–3.41 (m, 2H), 2.55–2.45 (m, 4H), 2.43–2.32 (m, 4H), 2.15–1.97 (m, 4H).
13C NMR (125 MHz, CDCl3, δ): 172.5, 168.7, 152.1, 149.9, 148.7, 135.9, 130.8, 129.0, 125.8,
125.8, 124.2, 122.9, 117.6, 112.9, 80.7, 38.0, 25.5, 25.4, 18.6. HRMS (ESI–QIT) m/z: [M + H]+
Calcd for C30H23Cl2O7 565.0815; Found 565.0817.
2′,7′-Dichlorofluorescein Dihexanoate (3f). White solid (56.5 mg, 63% yield) 1H NMR (500
MHz, CDCl3, δ): 8.08 (d, J = 7.5 Hz, 1H), 7.78–7.68 (m, 2H), 7.22 (d, J = 7.6 Hz, 1H), 7.15 (s,
2H), 6.87 (s, 2H), 2.63 (t, J = 7.5 Hz, 4H), 1.78 (dd, J = 14.9, 7.4 Hz, 4H), 1.46–1.34 (m, 8H),
0.93 (t, J = 7.0 Hz, 6H). 13C NMR (125 MHz, CDCl3, δ): 171.0, 168.7, 152.0, 149.9, 148.7,
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135.9, 130.8, 129.1, 125.8, 125.8, 124.2, 122.8, 117.6, 112.90, 80.6, 34.1, 31.3, 24.6, 22.4, 14.0.
HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C32H31Cl2O7 597.1441; Found 597.1441.
2′,7′-Dichlorofluorescein Di-2-propylvalerate (3g). White solid (68.5 mg, 70% yield). 1H
NMR (500 MHz, CDCl3, δ): 8.09 (d, J = 7.5 Hz, 1H), 7.74 (tdd, J = 14.9, 10.8, 4.2 Hz, 2H), 7.21
(d, J = 7.6 Hz, 1H), 7.11 (s, 2H), 6.88 (s, 2H), 2.73–2.64 (m, 2H), 1.84–1.76 (m, 4H), 1.63–1.56
(m, 5H), 1.51–1.43 (m, 8H), 0.97 (td, J = 7.3, 2.2 Hz, 12H). 13C NMR (125 MHz, CDCl3, δ):
173.6, 168.8, 152.1, 149.8, 148.6, 135.9, 130.8, 129.1, 125.8, 125.7, 124.1, 122.8, 117.6, 112.9,
80.6, 45.4, 34.6, 34.5, 20.8, 20.8, 14.1 HRMS (ESI–QIT) m/z: [M + H]+ Calcd for C36H39Cl2O7
653.2067; Found 653.2067.
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2′,7′-Dichlorofluorescein diacetoxymethyl ether (AM). Ag2O (145 mg, 0.63 mmol), 2′,7′-
dichlorofluorescein (0.25 mmol), and powdered activated 4-Å molecular sieves (208 mg) were
added to an oven-dried round-bottom flask. Anhydrous CH3CN (4.0 mL) was added, and the
resulting suspension was stirred under N2(g) for 5 min. To this mixture was added bromomethyl
acetate (0.1 mL, 1.0 mmol) dropwise, and the resulting mixture was stirred under N2(g) for 48 h.
The reaction mixture was then diluted with DCM and filtered through a pad of Celite®.
Purification by column chromatography on silica gel (0–40% v/v EtOAc in hexanes with
constant 40% v/v DCM as cosolvent) afforded the title compound as a white solid (53.2 mg, 39%
yield). 1H NMR (500 MHz, CDCl3, δ): 8.08 (d, J = 7.4 Hz, 1H), 7.72 (td, J = 7.4, 1.1 Hz, 2H),
7.21–7.14 (m, 1H), 7.09 (s, 2H), 6.79 (s, 2H), 5.86 (dd, J = 8.4, 6.5 Hz, 4H), 2.19 (s, 6H). 13C
NMR (125 MHz, CDCl3, δ): 169.6, 168.8, 154.1, 151.9, 150.4, 135.7, 130.6, 129.4, 126.3, 125.7,
123.9, 119.7, 113.9, 104.1, 85.4, 81.2, 21.0. HRMS (ESI–QIT) m/z: [M + H]+ Calcd for
C26H19Cl2O9 545.0401; Found 545.0400.
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Supporting Information
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The Supporting Information is available free of charge on the ACS Publications website at DOI:
10.1021/acs.joc.xxxxxxx.
Table S1, Figures S1–S4, 1H NMR, 13C NMR, and IR spectra, and computational data.
AUTHOR INFORMATION
Corresponding Author
*E-mail: rtraines@wisc.edu
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
We are grateful to Dr. C. L. Jenkins for her critical review of the manuscript. This work was
supported by grant R01 GM044783 to R.T.R. (NIH). W.C. was supported by an NSF Graduate
Research Fellowship. B.G. was supported by an Arnold O. Beckman Postdoctoral Fellowship.
This work used data acquired at the National Magnetic Resonance Facility at Madison, which is
supported by Grant P41 GM103399 (NIH). The work also made use of a Thermo Q Exactive™
Plus mass spectrometer (NIH grant S10 OD020022), Phoenix Cluster at the UW–Madison HPC
center (NSF grant CHE-0840494), and Micro FT-IR spectrometer (NSF grant DMR-1121288).
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REFERENCES
5
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8
9
(1) (a) Lavis, L. D.; Raines, R. T. ACS Chem. Biol. 2008, 3, 142-155. (b) Chan, J.; Dodani, S.
C.; Chang, C. J. Nat. Chem. 2012, 4, 973-984.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
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39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(2) (a) Grimm, J. B.; Heckman, L. M.; Lavis, L. D. Prog. Mol. Biol. Transl. Sci. 2013, 113,
1-34. (b) Lavis, L. D.; Raines, R. T. ACS Chem. Biol. 2014, 9, 855-866.
(3) (a) Lavis, L. D.; Chao, T.-Y.; Raines, R. T. ACS Chem. Biol. 2006, 1, 252-260. (b) Lavis,
L. D.; Chao, T.-Y.; Raines, R. T. ChemBioChem 2006, 7, 1151-1154.
(4) (a) Levine, M. N.; Raines, R. T. Anal. Biochem. 2011, 418, 247-252. (b) Park, J.; Kim, Y.
Bioorg. Med. Chem. Lett. 2013, 23, 2332-2335.
(5) Chevalier, A.; Mercier, C.; Saurel, L.; Orenga, S.; Renard, P.-Y.; Romieu, A. Chem.
Commun. 2013, 49, 8815-8818.
(6) Yatzeck, M. M.; Lavis, L. D.; Chao, T.-Y.; Chandran, S. S.; Raines, R. T. Bioorg. Med.
Chem. Lett. 2008, 18, 5864-5866.
(7) Heckman, L. M.; Grimm, J. B.; Schreiter, E. R.; Kim, C.; Verdecia, M. A.; Shields, B. C.;
Lavis, L. D. Angew. Chem., Int. Ed. 2016, 55, 8363-8366.
(8) (a) Cohen, J. D.; Husic, H. D. FASEB J. 1991, 5, A1511-A1511. (b) Karlsson, H. J.;
Westman, G. Tetrahedron 2000, 56, 8939-8944. (c) Shan, G. M.; Hammock, B. D. Anal.
Biochem. 2001, 299, 54-62. (d) Tallman, K. R.; Beatty, K. E. ChemBioChem 2015, 16, 70-75. (e)
Tallman, K. R.; Levine, S. R.; Beatty, K. E. ACS Chem. Biol. 2016, 11, 1810-1815.
(9) Levine, M. N.; Hoang, T. T.; Raines, R. T. Chem. Biol. 2013, 20, 614-618.
(10) Sorkin, A.; von Zastrow, M. Nat. Rev. Mol. Cell Biol. 2002, 3, 600-614.
(11) Sun, W. C.; Gee, K. R.; Klaubert, D. H.; Haugland, R. P. J. Org. Chem. 1997, 62, 6469-
6475.
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5
6
7
8
9
(12) Lavis, L. D.; Chao, T.-Y.; Raines, R. T. Chem. Sci. 2011, 2, 521-530.
(13) Chyan, W.; Zhang, D. Y.; Lippard, S. J.; Radford, R. J. Proc. Natl. Acad. Sci. USA
2014, 111, 143-148.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(14) Hakamata, W.; Tamura, S.; Hirano, T.; Nishio, T. ACS Med. Chem. Lett. 2014, 5, 321-
325.
(15) Levine, M. N.; Raines, R. T. Chem. Sci. 2012, 3, 2412-2420.
(16) (a) Ouyang, H.; Vander Velde, D. G.; Borchardt, R. T.; Siahaan, T. J. J. Peptide Res.
2002, 59, 183-195. (b) Yang, Y. Z.; Babiak, P.; Reymond, J. L. Helv. Chim. Acta 2006, 89, 404-
415.
(17) (a) Hammett, L. P. Trans. Faraday Soc. 1938, 34, 156-166. (b) McDaniel, D. H. B.,
Herbert C. J. Am. Chem. Soc. 1954, 77, 3756-3764. (c) Hansch, C.; Leo, A.; Hoekman, D. H.,
Exploring QSAR: Fundamentals and Applications in Chemistry and Biology. American
Chemical Society: Washington, DC, 1995. (d) Santiago, C. B.; Milo, A.; Sigman, M. S. J. Am.
Chem. Soc. 2016, 138, 13424-13430.
(18) Smith, S. A.; Pretorius, W. A. Water SA 2002, 28, 395-402.
(19) Zhang, X. F.; Zhang, J. L.; Liu, L. M. J. Fluoresc. 2014, 24, 819-826.
(20) Batistela, V. R.; Pellosi, D. S.; de Souza, F. D.; da Costa, W. F.; Santin, S. M. D.; de
Souza, V. R.; Caetano, W.; de Oliveira, H. P. M.; Scarminio, I. S.; Hioka, N. Spectrochim. Acta,
Part A 2011, 79, 889-897.
(21) Choudhary, A.; Kamer, K. J.; Raines, R. T. J. Org. Chem. 2011, 76, 7933-7937.
(22) Choudhary, A.; Fry, C. G.; Kamer, K. J.; Raines, R. T. Chem. Commun. 2013, 49, 8166-
8168.
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4
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6
7
8
9
(23) (a) Jones, R. N.; Spinner, E. Can. J. Chem. 1958, 36, 1020-1027. (b) Bistoni, G.;
Rampino, S.; Scafuri, N.; Ciancaleoni, G.; Zuccaccia, D.; Belpassi, L.; Tarantelli, F. Chem. Sci.
2016, 7, 1174-1184.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
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33
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35
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40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(24) Singh, S. K.; Mishra, K. K.; Sharma, N.; Das, A. Angew. Chem., Int. Ed. 2016, 55,
7801-7805.
(25) Nishioka, T.; Fujita, T.; Kitamura, K.; Nakajima, M. J. Org. Chem. 1975, 40, 2520-
2525.
(26) Glendening, E. D.; Badenhoop, J. K.; Reed, A. E.; Carpenter, J. E.; Bohmann, J. A.;
Morales, C. M.; Weinhold, F. NBO 6.0, Madison, WI, 2012.
(27) Jakobsche, C. E.; Choudhary, A.; Raines, R. T.; Miller, S. J. J. Am. Chem. Soc. 2010,
132, 6651-6653.
(28) Bürgi, H. B.; Dunitz, J. D.; Lehn, J. M.; Wipff, G. Tetrahedron 1974, 30, 1563-1572.
(29) (a) Calloway, N. T.; Choob, M.; Sanz, A.; Sheetz, M. P.; Miller, L. W.; Cornish, V. W.
ChemBioChem 2007, 8, 767-774. (b) Bender, D. M.; Peterson, J. A.; McCarthy, J. R.; Gunaydin,
H.; Takano, Y.; Houk, K. N. Org. Lett. 2008, 10, 509-511.
(30) Tian, L.; Yang, Y. L.; Wysocki, L. M.; Arnold, A. C.; Hu, A.; Ravichandran, B.;
Sternson, S. M.; Looger, L. L.; Lavis, L. D. Proc. Natl. Acad. Sci. USA 2012, 109, 4756-4761.
(31) French, A. P.; Mills, S.; Swarup, R.; Bennett, M. J.; Pridmore, T. P. Nat. Protoc. 2008,
3, 619-628.
(32) Frisch, M. J.; Trucks, G. W.; Schlegel, H. B.; Scuseria, G. E.; Robb, M. A.; Cheeseman,
J. R.; Scalmani, G.; Barone, V.; Mennucci, B.; Petersson, G. A.; Nakatsuji, H.; Caricato, M.; Li,
X.; Hratchian, H. P.; Izmaylov, A. F.; Bloino, J.; Zheng, G.; Sonnenberg, J. L.; Hada, M.; Ehara,
M.; Toyota, K.; Fukuda, R.; Hasegawa, J.; Ishida, M.; Nakajima, T.; Honda, Y.; Kitao, O.;
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Nakai, H.; Vreven, T.; Montgomery, J. A., Jr.; Peralta, J. E.; Ogliaro, F.; Bearpark, M.; Heyd, J.
J.; Brothers, E.; Kudin, K. N.; Staroverov, V. N.; Kobayashi, R.; Normand, J.; Raghavachari, K.;
Rendell, A.; Burant, J. C.; Iyengar, S. S.; Tomasi, J.; Cossi, M.; Rega, N.; Millam, J. M.; Klene,
M.; Knox, J. E.; Cross, J. B.; Bakken, V.; Adamo, C.; Jaramillo, J.; Gomperts, R.; Stratmann, R.
E.; Yazyev, O.; Austin, A. J.; Cammi, R.; Pomelli, C.; Ochterski, J. W.; Martin, R. L.;
Morokuma, K.; Zakrzewski, V. G.; Voth, G. A.; Salvador, P.; Dannenberg, J. J.; Dapprich, S.;
Daniels, A. D.; Farkas, Ö.; Foresman, J. B.; Ortiz, J. V.; Cioslowski, J.; Fox, D. J. Gaussian 09,
Revision D.01, Gaussian, Inc.: Wallingford, CT, 2009.
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