Design, Synthesis, and Characterization of Small-Molecule Reagents That Cooperatively Provide Dual Readouts for Triaging and, When Necessary, Quantifying Point-of-Need Enzyme Assays

Article abstract of DOI:10.1021/acs.joc.5b02013

A newly designed small molecule reagent provides both qualitative and quantitative readouts in assays that detect enzyme biomarkers. The qualitative readout enables rapid triaging of samples so that only samples that contain relevant concentrations of the target analyte must be quantified. The reagent is accessible in essentially three steps and 34% overall yield, is stable as a solid when heated to 44 °C for >1 month, and does not produce background signal when used in an assay. This paper describes the design and synthesis of the reagent, characterizes its response properties, and establishes the scope of its reactivity.

Full text of DOI:10.1021/acs.joc.5b02013

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pubs.acs.org/joc
Design, Synthesis, and Characterization of Small-Molecule Reagents
That Cooperatively Provide Dual Readouts for Triaging and, When
Necessary, Quantifying Point-of-Need Enzyme Assays
Adam D. Brooks, Hemakesh Mohapatra, and Scott T. Phillips*
Department of Chemistry, Pennsylvania State University, University Park, Pennsylvania 16802, United States
S
* Supporting Information
ABSTRACT: A newly designed small molecule reagent provides both qualitative
and quantitative readouts in assays that detect enzyme biomarkers. The qualitative
readout enables rapid triaging of samples so that only samples that contain relevant
concentrations of the target analyte must be quantified. The reagent is accessible in
essentially three steps and 34% overall yield, is stable as a solid when heated to 44
°C for >1 month, and does not produce background signal when used in an assay.
This paper describes the design and synthesis of the reagent, characterizes its
response properties, and establishes the scope of its reactivity.
including those with minimal training. While no current assay
satisfies all of the requirements for an effective high-throughput,
low-cost, quantitative point-of-need assay, new approaches are
being developed to address this deficiency. These strategies
include assays based on cell phones,5−17 repurposing glucose
meters to detect analytes other than glucose,18−25 using a
voltage meter to measure the results of an electrochemical
assay,26−35 assays based on relative measurements of time,36−41
detection based on the release of odorous compounds,42−45 as
well as assays based on the distance that a sample travels on an
assay platform.46−48
These approaches, however, all require that every sample be
analyzed quantitatively. Performing quantitative assays on every
sample typically is slow, expensive, and operationally involved,
which is not an ideal scenario when many of the analytical
samples to be tested may lack relevant levels of the analyte of
interest and therefore do not require a quantitative measure-
ment (a qualitative assay would suffice for these samples).
An alternative approach to running quantitative assays on all
samples is to use a triaging assay first, followed by quantitative
assays on samples that test positive for the target analyte. Of
course, this approach requires two separate assays, likely two
aliquots of a sample, and twice as many manipulations by the
user. An approach that provides both quantitative and
qualitative readouts simultaneously, so that only one test is
performed, offers a practical alternative. We demonstrated an
example of a dual readout approach (Figures 1a and 2a) in a
recent report in which only samples with relevant levels of the
INTRODUCTION
■
This paper describes a small molecule assay reagent that
provides the dual readouts of fluorescence (a coumarin
derivative) and smell (ethanethiol) (Figure 1), which enables
rapid triaging of samples for the presence of a target analyte.
This assay strategy requires only one aliquot of a sample and
minimizes the frequency of time-consuming quantitative
measurements. This article focuses on the organic chemistry
of this new reagent, whereas future studies will detail the
analytical performance of the reagent.
The design of this new small molecule reagent addresses an
unrealized goal in analytical chemistry, which is the develop-
ment of high-throughput, low-cost, quantitative assays that
operate in low-resource point-of-need settings.1−4 Example
scenarios where such assays are needed include (i) testing
multiple sources of recreational water for the presence of
Escherichia coli (E. coli) derived from fecal contamination, and if
present, then measuring the quantity of the contamination; (ii)
evaluating streams, lakes, and rivers for specific pollutants from
mining or fracking; or (iii) screening fruits and vegetables or
other food products for the presence and quantity of specific
bacterial pathogens. In these scenarios, not only is information
needed about the presence (and if present, the abundance) of a
contaminant to make an informed decision, but the tests often
must be rapid so that tens or hundreds of samples can be
measured easily and quickly without the resources of a
laboratory.
Designing tests that meet these criteria has proven difficult,
particularly since the assay reagents must be stable, the assay
components must be portable and inexpensive, and the assay
itself must proceed smoothly with little input by users,
Received: August 28, 2015
© XXXX American Chemical Society
A
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the assays. Herein, we describe a small-molecule reagent with
substantially improved properties (compared with our initial
design) for this type of dual readout assay (Figure 1b).
Our approach uses reagent 2 (Figure 1), which provides the
dual readoutone qualitative (4) and one quantitative (3)
all in a single-step assay. If the qualitative readout (odor of 4)
indicates that the analyte is in the sample, then a hand-held
fluorescence meter is used to measure the quantity of 3. We
chose ethanethiol (4) for the qualitative readout because
humans have a keen ability to smell this compound.49 In fact,
ethanethiol is in a class of compounds to which the human nose
is most sensitive, thus providing the required sensitivity to
match the sensitivity of the quantitative readout using 3.49
A
match in sensitivity for detecting 4 vs 3 is critical for the
triaging portion of the assay in order to provide information
that is relevant to deciding whether to run the quantitative
assay.
The complete triaging assay requires three reagents that
operate simultaneously in a single-pot reaction (Figure 1b).
These reagents include the following: (i) a detection reagent
(1) that selectively reacts with the target analyte (enzymes are
the targets in this work) and releases glucose; (ii) glucose
oxidase, which converts glucose into D-gluconic acid δ-lactone
and simultaneously generates hydrogen peroxide; and (iii) a
readout reagent (2) that responds to the hydrogen peroxide to
produce the dual readout. We use two reagents (i.e., 1 and 2)
instead of a single reagent for detecting the analyte and
providing readouts because it is easier to prepare two simple
reagents than one complex reagent and because we envisage
assays in which the detection reagent (1) is varied easily to
target different enzymes, while the readout reagent (2) remains
constant.
Figure 1. Design for a triaging assay where odor (ethanethiol) signals
which samples require quantitative measurements. (a) Schematic
representation of the assay. (b) Specific reagents used in the assay.
These reagents include detection reagents (1) that select for a target
enzyme analyte; glucose oxidase, which produces hydrogen peroxide
upon oxidation of glucose that is released during the detection event;
and readout reagent 2 that provides the dual readouts of fluorescence
(3) and smell (4).
RESULTS AND DISCUSSION
■
Advantages of Reagent 2 vs 5. Our first-generation
reagent for the triaging assay (5, Figure 2a) was limited in (i)
sensitivity (particularly matching the sensitivity of the
qualitative and quantitative readouts for the assay), (ii)
reproducibility during an assay, and (iii) stability (which gave
rise to background signal). In contrast, our new design, reagent
2 (Figure 2b), is thermally stable, readily accessible syntheti-
cally, responds to a target analyte faster than previous reagent 5,
eliminates background smell that limited the sensitivity of 5,
and provides a nearly perfect match in sensitivity between the
qualitative and quantitative readouts for triaging assays (which
is the ideal scenario for such an assay). These improvements are
essential for enabling fast, sensitive, and reproducible triaging
assays, and thus represent a critical synthetic step toward
evaluating the feasibility of smell-based triaging assays in
practical applications.
Considerations for Reagent 2. The advantages of reagent
2 vs 5 result from modifications to reagent 5, such as replacing
the phenol in 5 with a methyl ether, which we believe limits
nonspecific release of ethanethiol via ketene 6 (Figure 2a).50
Background release of thiol was further minimized by installing
a cis-α,β-unsaturated thioester in 2 (Figure 2b) versus the
dithioester of 5 (Figure 2a). This modification reduces the
number of thioesters that are available for background
hydrolysis, either upon storage or during the assay. Finally,
we installed the vinyl boronate with the goal of increasing the
rate of the oxidative cleavage reaction of the boronate with
hydrogen peroxide that is generated during the assay (Figure
1).51 We reasoned that the aryl boronate in 5 likely suffers from
Figure 2. (a) Structure and proposed mechanism for the desired
response and possible degradation pathway for first-generation triaging
reagent 5.43 (b) Structure of 2 as well as a possible mechanism by
which 2 releases ethanethiol.
analyte (as determined from the qualitative readout) required
quantitative measurements of the second readout.43 The
advantage of this dual readout approach over other methods
is that time-consuming quantitative measurements are saved for
those samples that contain relevant concentrations of the target
analyte, thus reducing consumption of time when conducting
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(i) unfavorable nonbonding interactions that slow the reaction
of hydrogen peroxide with boron, (ii) a conformation around
the aryl carbon−boron bond that slows the bond rearrange-
ment steps during the oxidative cleavage reaction, and (iii)
dative bonding between the lone pairs on the thioester and the
empty p-orbital on boron that could hinder the ability of
hydrogen peroxide to bind to the empty p-orbital on boron to
initiate the oxidative cleavage reaction.52 This type of dative
bonding also might contribute to hydrolysis of the thioester via
Lewis acid-mediated activation of the thioester.52,53 Moving the
boronate away from the aromatic ring (i.e., installing a vinyl
boronate instead of an aryl boronate) in 2 minimizes the
likelihood of these unfavorable interactions.
Synthesis of reagent 2. We prepared reagent 2 in three
primary steps and 34% overall yield (Scheme 1) using a scalable
synthetic strategy that makes available substantial quantities of
2.
a
Scheme 1. Synthesis of Reagent 2
a
Reagents and conditions: (a) BBr3, DCM, (94%); (b) vinyl boronate
allyl chloride, TBAI, Cs2CO3, acetone (53%); (c) porcine liver
esterase, phosphate buffer, acetone (99%); (d) EtSH, EDCI, DMAP,
DCM (88%); (e) K2CO3, THF, −40 °C (68%).
Figure 3. Response of 2 to hydrogen peroxide. (a) Liquid
chromatograms (LC) (obtained from LCMS experiments) before
and 5 min after exposure of 2 to hydrogen peroxide. (b, c) Time-
dependent UV−vis and fluorescent spectra, respectively, after exposure
of 2 (2.3 mM) to hydrogen peroxide (6.0 mM).
Response of Reagent 2 to Hydrogen Peroxide. As
expected, treatment of 2 with hydrogen peroxide in aqueous
solutions quickly and cleanly (i.e., no observable intermediates)
produces fluorescent product 3 and ethanethiol (4) (Figure 3).
(The presence of 3 was confirmed using LCMS (Figure 3a),
and 4 was detected by smell.) Time-dependent UV−vis spectra
of the same reaction reveal isosbestic points at 310 and 358 nm
(Figure 3b), which further supports conversion of 2 to 3
without proceeding through observable intermediates. More-
over, the intensity of the fluorescent signal for 3 increases over
time when 2 is exposed to hydrogen peroxide (Figure 3c), thus
indicating that fluorescence measurements can be used
effectively to quantify assays.
Demonstration of Reagent 2 in Model Assays for
Specific Enzyme Biomarkers. The ability of reagent 2 to
operate in simulated assays is depicted in Figure 4 for detecting
two model enzyme analytes: β-D-galactosidase (a general
marker for fecal coliforms in water) (1a, Figure 1, was used
as its substrate during the assay) and alkaline phosphatase (a
reporter of liver function) (1b was used as the substrate). The
assays involved dissolving 2 (final concentration = 230 μM)
and either 1a or 1b (final concentration = 57 mM) in a sample
containing the desired enzyme target, followed by incubation at
20 °C for 30 min. The linear calibration curves in Figure 4 were
Figure 4. Calibration curves based on the fluorescent signal of 3 when
either 1a (for β-D-galactosidase) or 1b (for alkaline phosphatase)
combined with 2 were exposed to samples of enzyme at different initial
concentrations. The data points are the average of three measure-
ments, and the error bars represent the standard deviation from the
averages.
C
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obtained using a hand-held, portable fluorescence meter that
contains an LED that emits at 365 nm and measures
absorbance from 440−470 nm (for reference, coumarin
product 3 absorbs maximally at 345 nm and emits maximally
at 430 nm). The limits-of-detection54 for 30 min assays are 3.5
nM (0.4 U·mL−1) and 8.5 nM (8 U·mL−1) for the two model
enzymes, respectively.
Comparison of Reagent 2 to Reagent 5. Reagent 5, in
comparison, provides a limit-of-detection for β-D-galactosidase
that is 6× worse than 2.43 This improvement in limit-of-
detection provided by 2 is impressive when considering that (i)
the sensitivity for 2 was determined using a simple, fixed-
intensity, fixed-gain, low-cost hand-held fluorescence meter,
whereas the value for 5 was determined using a high-powered
benchtop UV/vis spectrometer with perfectly tuned emission
and excitation wavelengths to maximize the sensitivity of the
assay.43 This improvement in sensitivity for 2 compared to 5
likely is the result of the resistance of 2 to undergo nonspecific
background reactions.
When compared to standard fluorescent reagents (such as 4-
methylumbelliferyl β-D-galactopyranoside (4-MU); λex = 365
nm; λem = 455 nm) for detecting β-D-galactosidase, 2 enables
assays that are only 3.5× less sensitive (using the hand-held
fluorescence meter and 30 min assay time), but it also enables
rapid triaging based on smell, which is a capability that 4-MU
and other standard reagents do not provide.55
Evaluation of the Practical Benefits of Reagent 2. Even
more important than the absolute sensitivity of the fluorescence
assay, however, is the match in sensitivity between the
qualitative (smell) and quantitative (fluorescent) readouts.
The limits-of-detection for the qualitative assays for β-D-
galactosidase and alkaline phosphatase were determined using
the recommended protocol provided by the American Society
for Testing and Measurements (ASTM) (i.e., “forced-choice
ascending concentration series method of limits”).56
Figure 5. LC chromatographs that reveal the stability of 2. (a)
Solution-phase stability study (1.2 mM in 1:1 MeOH−HEPES buffer
(pH 7.9)) at 31 °C. Naphthalene was used as an internal standard. (b)
Solid-state stability study at 44 3 °C, open to the air. The LC solvent
gradient was different in (b) vs (a) to separate the cis and trans
isomers of 2 that develop after heating solid 2 for days at 44 °C (b).
Briefly, this method involves panelists correctly identifying
the most odorous sample in a set of three samples, where only
one sample contains the analyte, while the other two are
control samples that lack the analyte.56 This procedure was
repeated using six sets of samples that increase sequentially in
the concentration of the analyte. Based on this protocol (and
the responses of 10 panelists), the limits of detection for the
qualitative assays using 2 were calculated as 6.2 nM (0.6 U·
mL−1) for β-D-galactosidase and 15 nM (14 U·mL−1) for
alkaline phosphatase. These values both are only 1.8× higher
than the limits-of-detection of the fluorescence assays for the
same enzymes. This close match in limits-of-detection between
the qualitative and quantitative readouts demonstrates that 2 is
a highly effective reagent for enabling down-selection assays.
An additional practical consideration is the stability of 2
(Figure 5). In the assay solution, reagent 2 does hydrolyze to
the boronic acid within ∼5 min, but this hydrolysis product
undergoes the oxidative cleavage reaction with hydrogen
peroxide essentially as readily as the boronate, so the hydrolysis
reaction does not appear to affect the assay. Other than this
hydrolysis reaction, reagent 2 is stable for at least 5 days in the
assay solution (which is far longer than is needed for an assay),
with <8% isomerization of the cis alkene in the α,β-unsaturated
thioester to form the trans alkene in 9 (Figure 6a). Formation
of trans-α,β-unsaturated thioester 9 from 2 over the course of 5
days does not affect the sensitivity of assays using 2, since no
detectable quantity of 9 is present in a solution of 2 that is
stored for 1 h (which is 30 min longer than the duration of the
Figure 6. Structural variants of reagent 2 for studying the mechanism
of coumarin production and ethanethiol release. (a) Possible
cyclization with a ketene intermediate that would allow the trans
alkene to generate 3 and 4. (b) Comparison of the structure of cis
ester 13 to the cis thioester triaging reagent 2.
current assays). Our original reagent (5), in contrast, degraded
to several byproducts in less than 20 min under solution-phase
conditions, which decreased the quantity of 5 that was available
to react with hydrogen peroxide generated during an assay
(Figure 2a). Reagent 5 also generated a noticeable thiol smell
within this short period of time, which negatively affected the
D
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sensitivity of the qualitative down-selection portion of the assay
when 5 was used instead of 2.
mechanism from 2 (pathway in Figure 2b) likely is operative
over an electrocyclization pathway (pathway in Figure 6a).
The second concern involved losing ethanethiol through
oxidation−reduction reactions between ethanethiol and hydro-
gen peroxide that is generated during the detection event. To
test the effects of this scenario, we prepared control compound
13 (Figure 6b) that contains an ethyl ester instead of an ethyl
thioester. Exposure of 13 to hydrogen peroxide leads to
formation of 3 and release of ethanol rather than ethanethiol,
the former of which does not undergo rapid oxidation−
reduction reactions with hydrogen peroxide. Comparison of
emission intensities (Table 1) for 3, when 2 and 13 are exposed
Reagent 2 also is stable when stored as a solid (as it would be
supplied before use in an assay; Figure 1). Solid 2 showed little
decomposition when heated to 40 °C, open to the air for 1
month (this was the duration of our stability study) (Figure
5b). In fact, the only decomposition reaction of 2 involved
isomerization of the α,β-unsaturated thioester to form 12. Most
importantly, no background release of thiol was detected by
smell. Furthermore, when 2 was stored in the solid state under
inert atmosphere at 8 °C, there was no isomerization or
decomposition observed after more than 1 year (Figure S16).
Evaluation of Structural Variants of Reagent 2.
Reagent 2 clearly is effective for use in triaging assays,
particularly since it is readily accessible synthetically, thermally
stable, and provides qualitative readouts that are almost as
sensitive as the quantitative readout. Because of these
promising results, we tested possible limitations of 2, such as
(i) whether the trans α,β-unsaturated thioester 12 could
function in the assay and (ii) if side reactions with the released
thiol reduce the overall sensitivity of the assay.
Table 1. Normalized Fluorescent Signal for 13 vs 2 upon
Treatment of Each Reagent with H2O2
[H2O2]
(μM)
normalized fluorescent signal for relative fluorescent signal for
2 (thioester)
13 (ester)
10
20
50
1
1
1
1.4 0.1
1.8 0.1
2.4 0.2
To address the first topic, we prepared 12 (Scheme 2) and
then tested whether 12 responds to hydrogen peroxide to form
to hydrogen peroxide under identical conditions, reveals that
the impact of the proposed oxidation−reduction on the
sensitivity of the quantitative and qualitative assays is
substantial (i.e., 13 produces 2.5× more fluorescence signal
than 2) when high concentrations of hydrogen peroxide (>0.08
equiv relative to 2 or 13) are present. High concentrations of
hydrogen peroxide, however, are unlikely to form during an
assay. In an assay, the hydrogen peroxide that is generated via
the detection event with 1 likely is consumed immediately via
the oxidative cleavage reaction with 2; therefore, the loss in
signal as a result of this oxidation−reduction reaction between
hydrogen peroxide and ethanethiol is presumably small
(<28%), as suggested by the limited loss in signal when 2
and 13 are treated with only 0.04 equiv (10 μM) of hydrogen
peroxide. Therefore, the down-selection strategy described in
Figure 1 should remain a viable strategy so long as the rate of
oxidative cleavage of reagents like 2 remain competitive with
the combined rates with which hydrogen peroxide is generated
from reaction of the analyte with 1, followed by reaction of the
released glucose with glucose oxidase to generate hydrogen
peroxide (Figure 1).
a
Scheme 2. Synthesis of Reagent 12
a
Reagents and conditions: (a) EtSH, EDCI, DMAP, DCM (34%); (b)
9, Cs2CO3, THF, 50 °C (20%).
3 and 4. Reagent 12 indeed undergoes the oxidative cleavage
reaction when exposed to hydrogen peroxide, but no
cyclization or other subsequent reactions occur within 4 d
under the assay conditions (Figure 7). While the stability of the
oxidative cleavage product of 12 obviates the use of 12 in
down-selection assays, it does suggest that the direct cyclization
CONCLUSIONS
■
In conclusion, we designed a readout reagent that is capable of
providing rapid, sensitive, qualitative, and quantitative results
for triaging a large number of samples for a target analyte. In
principle, this reagent can be paired with a variety of detection
reagents for use in a wide range of enzyme assays. While the
approach is not yet compatible with all classes of analytes, it
does offer the potential to simplify quantitative point-of-need
assays for detecting and measuring enzyme biomarkers, such as
the various extracellular enzymes indicative of bacterial
contamination, including necrotizing enzymes, kinases, hyalur-
onidase, hemolysins, hydrolases, and laccases.
Perhaps more importantly, the design of the readout reagent
minimizes background reaction, enables rapid assays, and
provides matching of the intensities of the qualitative and
quantitative readouts for the assay. The straightforward
synthesis of the readout reagent adds to these useful qualities
and thus should provide a starting point for others to explore
various analytical applications of this reagent.
Figure 7. Exposure of trans alkene 12 (2.3 mM in 1:1 MeOH−HEPES
buffer (pH 7.9)) at 31 °C to H2O2 (6 mM) yields the phenol, but
none of 3. Naphthalene was used as an internal standard.
E
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dehyde (9) as a white solid (305 mg, 0.876 mmol, 53%): mp 115−118
EXPERIMENTAL SECTION
■
1
°C; H NMR (300 MHz, CDCl3): δ 10.4 (1H, s), 7.33 (1H, s), 6.76
General Experimental Methods. All reactions requiring inert
atmosphere were performed in flame-dried glassware under a positive
pressure of argon. Air- and moisture-sensitive liquids were transferred
by syringe or stainless steel cannula. Organic solutions were
concentrated by rotary evaporation (25−40 mmHg) at ambient
temperature, unless otherwise noted. D-Lactose, D-glucose-6-phos-
phate, β-D-galactosidase, alkaline phosphatase, porcine liver esterase,
and all other reagents were purchased commercially and used as
received. Dry solvents were purified by the method developed by
Pangborn et al.57 Flash column chromatography was performed as
described by Still et al.58 Fluorescence assays were performed in UV-
transparent ultramicro UV cuvettes, and smell-based assays were
performed in microcentrifuge tubes (0.6 mL), unless otherwise noted.
Instrumentation. Assay-related fluorescence data was obtained
using a Promega QuantiFluor-P hand-held fluorometer (λex = 365 nm,
λem = 440−470 nm). Proton nuclear magnetic resonance (1H NMR)
spectra were recorded at 25 °C. Proton chemical shifts are expressed in
parts per million (ppm, δ scale) and are referenced to residual protium
in the NMR solvent (CDCl3, δ 7.26 ppm). Data are represented as
follows: integration, chemical shift, multiplicity (s = singlet, d =
doublet, t = triplet, q = quartet, m = multiplet and/or multiple
resonances, br s = broad singlet, dd = doublet of doublet), and
coupling constant (J) in hertz. Carbon nuclear magnetic resonance
spectra (13C NMR) were recorded at 25 °C. Carbon chemical shifts
are expressed in parts per million (ppm, δ scale) and are referenced to
the carbon resonance of the NMR solvent (CDCl3, δ 77.16 ppm).
LC−MS data were obtained on an analytical reversed-phase HPLC
coupled to a quadrupole mass spectrometer using a phenyl hexyl
column (150 mm × 5 mm, 5 μm particle size) for separation. The
column was equilibrated with the initial acetonitrile−water ratio of the
solvent gradient selected for column separation at 1.0 mL min−1 flow
rate. Solvent gradients after injection are reported with each
experiment. A portion of the HPLC stream was automatically injected
into the mass spectrometer. The mass spectrometer (APCI) settings
were as follows: gas temperature = 350 °C, drying gas flow = 11 L
min−1, nebulizer pressure = 35 psi, and voltage = 3000 V.
(1H, d of t, J = 18, 4.5 Hz), 6.45 (1H, s), 5.85 (1H, d, J = 17 Hz), 4.71
(2H, d of d, J = 4.3, 1.7 Hz), 3.93 (3H, s), 3.87 (3H, s), 1.28 (12H, s);
13C NMR (75 MHz, CDCl3): δ 187.9, 157.6, 155.7, 146.2, 143.9, 120.7
(br.), 117.9, 109.0, 97.4, 83.7, 70.7, 56.3, 24.9; IR (neat) 3216, 2984,
2536, 2161, 2027, 1657, 1603 cm−1; (TOF MS APCI+, m/z): 349.3
(M + H+); HRMS (TOF MS ES+, m/z) calcd for C18H26O6B (M +
H+) 349.1822, found 349.1818.
(Bis(2,2,2-trifluoroethoxy)phosphinyl)acetic Acid (17). This
procedure was adapted from the work of Sano et al.60 Porcine liver
esterase (1000 units/mm0l) was added to a solution of 9:1 1 M
phosphate buffer (pH 7.8) and acetone (50 mL). Ethyl [bis(2,2,2-
trifluoroethoxy)phosphinyl]acetate (10) (0.62 mL, 2.62 mmol, 1
equiv) was added to the solution in one portion and stirred in a round-
bottom flask for 16 h. Concentrated HCl was added to reduce the pH
to 1. The solution was saturated with NaCl, and 20 mL of EtOAc was
added with vigorous stirring. The excess NaCl was removed by
vacuum filtration and the organic elutant separated from the aqueous.
The aqueous layer was extracted with EtOAc (3 × 75 mL), and the
combined organic layers were washed once with an equal volume of
saturated brine. The organic layer was then dried over MgSO4, filtered,
and concentrated under reduced pressure to a viscous brown oil. The
oil was dried overnight to yield [bis(2,2,2-trifluoroethoxy)phosphinyl]-
acetic acid (17) as brown solid that was used without further
purification (795 mg, 2.61 mmol, 99%): 1H NMR (300 MHz, CDCl3)
δ 9.95 (1H, broad s), 4.48 (4H, quin, J = 7.9 Hz), 3.22 (2H, d, JP−H
=
21 Hz). This spectral data is consistent with data for the known
compound.60
S-Ethyl (Bis(2,2,2-trifluoroethoxy)phosphinyl)ethanethioate
(11). DMAP (54 mg, 0.442 mmol, 0.11 equiv) and 17 (1.26 g, 4.14
mmol, 1.0 equiv) were mixed in CH2Cl2 (16 mL) under argon and
cooled to 0 °C. To this solution was added ethanethiol (3.2 mL, 43.2
mmol, 10.4 equiv) followed by EDCI (940 mg, 4.90 mmol, 1.2 equiv).
The reaction mixture was stirred for 5 min, at which point the argon
inlet was removed and the reaction stirred for 6 h at 0 °C in a tightly
sealed vessel. The mixture was concentrated under reduced pressure to
an oil and redissolved in Et2O (50 mL). The organic layer was washed
with NaHCO3 (2 × 50 mL), 0.5 M HCl (2 × 50 mL), DI H2O (1 ×
50 mL), and brine (3 × 50 mL), dried over MgSO4, filtered, and
concentrated in vacuo to a brown oil. This oil was passed through a
silica plug with 25% EtOAc/hexanes to yield S-ethyl (bis(2,2,2-
trifluoroethoxy)phosphinyl)ethanthioate (11) as a golden oil (1.27 g,
3.66 mmol, 88%): 1H NMR (300 MHz, CDCl3) δ 4.48 (4H, quin, J =
8.1 Hz), 3.44 (2H, d, JP−H = 21 Hz), 2.95 (2H, q, J = 7.4 Hz), 1.28
(3H, t, J = 7.4 Hz); 13C NMR (75 MHz, CDCl3) δ 189.4, 128.0 (q),
124.3 (q), 120.6 (q), 116.9 (q), 63.1 (q), 62.6 (q), 62.1 (q), 61.6 (q),
42.8 (d), 41.0 (d), 24.2, 13.9; IR 2975, 2937, 1679, 1455, 1419 cm−1;
MS (TOF MS APCI+, m/z) 348.9 (M + H+); HRMS (TOF MS ES+,
m/z) calcd for C8H12F6O4PS (M + H+) 349.0098, found 349.0092.
(2Z)-S-Ethyl 3-(2-((2E)-3-(4,4,5,5-Tetramethyl-1,3,2-dioxa-
borolanyl)-2-propenoxy)-4,5-dimethoxyphenyl)-
propenethioate (2). This procedure was modified from a similar
method reported by Touchard.61 18-Crown-6 ether (115 mg, 0.435
mmol, 2.02 equiv) and anhydrous K2CO3 (115 mg, 0.871 mmol, 4.05
equiv) were dissolved in 5 mL of THF under argon and allowed to
equilibrate overnight. The cloudy solution was cooled to −40 °C, and
then 9 (88 mg, 0.253 mmol, 1.18 equiv) was added in one portion.
Compound 11 (75 mg, 0.215 mmol, 1 equiv) in 1 mL of THF was
then added, and additional 1 and 0.5 mL portions of THF were used
for quantitative transfer of 11. The vibrant yellow solution was stirred
in the dark under argon at −40 °C for 8 h (or until reaction
completion). This mixture was concentrated in vacuo in the dark,
redissolved in the Et2O (50 mL), and washed with 1 M phosphate
buffer pH 6.8 (2 × 50 mL), DI H2O (1 × 50 mL), and brine (2 × 50
mL). The organic layer was dried over MgSO4, filtered, and
concentrated to an oil that was purified by column chromatography
(30% EtOAc/hexanes). The product was dried in vacuo to a canary
yellow solid, (2Z)-S-ethyl 3-(2-((2E)-3-(4,4,5,5-tetramethyl-1,3,2-
dioxaborolanyl)-2-propenoxy)-4,5-dimethoxyphenyl)propenethioate
2-Hydroxy-4,5-dimethoxybenzaldehyde (16). This procedure
was modified from a method reported by Gunduz et al.59 2,4,5-
̈
̈
Trimethoxybenzaldehyde (8) (1.00 g, 5.09 mmol, 1 equiv) was
dissolved in CH2Cl2 (50 mL) under argon and cooled to −78 °C. To
this solution was added 1 M BBr3 in CH2Cl2 (8 mL) and the mixture
stirred for 30 min before the cooling bath was removed and the
reaction mixture stirred overnight. The brown reaction solution was
then cooled to 0 °C, acidified with concd HCl (∼5 mL), diluted with
water, and extracted. The organic layer was washed with 1 M HCl (3 ×
50 mL), water (3 × 50 mL), and brine (3 × 50 mL). The aqueous
layers were combined and back-extracted with 100 mL of EtOAc. The
combined organic layers were dried over MgSO4, filtered, and
concentrated to give 2-hydroxy-4,5-dimethoxybenzaldehyde (16) as
an off-white solid that was used without further purification (873 mg,
1
4.79 mmol, 94%): mp 101−103 °C; H NMR (360 MHz, CDCl3) δ
11.4 (1H, s), 9.70 (1H, s), 6.90 (1H, s), 6.47 (1H, s), 3.93 (3H, s),
3.88 (3H, s). This spectral data is consistent with data for the known
compound.59
2-((2E)-3-(4,4,5,5-Tetramethyl-1,3,2-dioxaborolanyl)-2-pro-
penoxy)-4,5-dimethoxybenzaldehyde (9). (E)-2-Chloromethylvi-
nylboronic acid pinacol ester (515 mg, 2.54 mmol, 1.55 equiv), 16
(298 mg, 1.64 mmol, 1 equiv), and tetrabutylammonium iodide (61
mg, 0.17 mmol, 0.10 equiv) were added to a round-bottom flask and
dissolved in DMA (6 mL) under argon. Anhydrous K2CO3 (374 mg,
2.71 mmol, 1.65 equiv) was added and the solution heated at 80 °C for
8 h. This solution was diluted with 50 mL of Et2O and 50 mL of water.
The aqueous layer was removed and extracted once more with 50 mL
of Et2O. The organic layers were combined, washed with water (2 ×
100 mL) and brine (3 × 100 mL), dried with MgSO4, filtered, and
concentrated in vacuo. The yellow oil was purified using column
chromatography (30% EtOAc/hexanes) to obtain 2-((2E)-3-(4,4,5,5-
tetramethyl-1,3,2-dioxaborolanyl)-2-propenoxy)-4,5-dimethoxybenzal-
F
DOI: 10.1021/acs.joc.5b02013
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1
(2) (64 mg, 0.146 mmol, 68%): H NMR (360 MHz, CDCl3) δ 7.79
(400 MHz, CDCl3) δ 7.68 (1H, s), 7.23 (1H, d, J = 12.8 Hz), 6.74
(1H, d of t, J = 18.2, 4.3 Hz), 6.43 (1H, s), 5.84 (1H, d, J = 12.8 Hz),
5.81 (1H, s), 4.59 (2H, d of d, J = 4.2, 1.7 Hz), 4.16 (2H, q, 3.88, J =
7.12 Hz), 3.87 (3H, s), 3.83 (3H, s), 1.29 (15H, m); 13C NMR (75
MHz, CDCl3) δ 166.5, 152.0, 151.2, 147.3, 142.7, 138.2, 120 (br.),
117.3, 116.1, 114.5, 98.10, 83.6, 70.9, 60.2, 56.6, 56.2, 25.0, 14.4; IR
(neat) 2973, 2921, 2852, 2531, 2162, 2030, 1670, 1647 cm−1; (TOF
MS APCI+, m/z) 419.2 (M + H+); HRMS (TOF MS ES+, m/z) calcd
for C22H32O7B (M + H+) 419.2241, found 419.2213.
Experimental Procedure for LC/MS Monitoring of Reagent 2
Response to Hydrogen Peroxide (Figure 3b). A hydrogen
peroxide solution (3 μL, 1.0 M in water) was added to a solution of
2 (500 μL, 2.3 mM in 1:1 MeOH−40 mM HEPES buffer, pH 7.9) and
naphthalene (0.5 mM). The mixture was agitated by vortex for ∼5 s.
An aliquot of the resulting mixture was injected into an analytical
reversed-phase HPLC coupled to a mass spectrometer, and additional
aliquots were injected at regular intervals. After injection of the sample,
an isocratic solvent gradient was run as 3:2 acetonitrile−water for 20
min.
Procedure for Monitoring the Time-Dependent UV−vis
Response of 2 to Hydrogen Peroxide (Figure 3b). A hydrogen
peroxide solution (690 μL, 0 and 3 μM in 40 mM phosphate buffer,
pH 7.0) was added to a solution of 2 (20 μL, 4 mM in MeOH) in a 1.5
mL centrifuge tube, agitated with a vortex mixer for ∼7 s, and then
transferred to a quartz cuvette. The UV−vis spectrum was then
monitored until no further changes were detected (∼10 min).
Procedure for Monitoring the Time-Dependent Fluores-
cence Response of 2 to Hydrogen Peroxide (Figure 3c). A
hydrogen peroxide solution (330 μL, 0 and 3 μM in 40 mM phosphate
buffer, pH 7.0) was added to a solution of 2 (20 μL, 4 mM in MeOH)
in a 1.5 mL centrifuge tube, agitated with a vortex mixer for ∼7 s, and
then transferred to a quartz cuvette. The fluorescence spectrum was
then monitored until no further changes were detected (∼10 min).
General Experimental for Hydrogen Peroxide Fluorescence
Emission Calibration Curve. A hydrogen peroxide solution (330
μL, 0−100 μM in 40 mM HEPES buffer, pH 7.9) was added to a
solution of 2 (20 μL, 4 mM in MeOH) in an ultramicro cuvette,
capped, and agitated with a vortex mixer for ∼7 s. After 30 min, the
fluorescence emission (I) of the mixture was measured, and the
average fluorescence of the 0 μM hydrogen peroxide sample (I0) was
subtracted (I − I0); this difference was then plotted to obtain a
calibration curve. (I = fluorescence intensity after 30 min from the
initial mixing of solutions; I0 = fluorescence intensity after 30 min from
the initial mixing of the solutions with 0 μM hydrogen peroxide.)
General Procedure for Enzyme Assays (Figure 4). To a
premixed solution of 2 (20 μL, 4 mM in MeOH), glucose oxidase
(100 μL, 15 U·mL−1 in pH 7.01 20 mM phosphate buffer and 400 mM
NaCl), and glucose-6-phosphate or lactose (100 μL, 200 mM in DI
H2O) was added a solution of alkaline phosphatase or β-D-
galactosidase (130 μL, 0−100 nM, in pH 7.01 20 mM phosphate
buffer and 400 mM NaCl) respective of enzyme substrate. This
solution was vortex mixed for ∼10 s and the fluorescence measured at
30 min and 1 h. The fluorescence emission (I) of the mixture was
measured, and the average fluorescence of the 0 μM enzyme sample
(I0) was subtracted (I − I0), this difference was plotted to obtain a
calibration curve. (I = fluorescence intensity after 30 min or 1 h from
the initial mixing of solutions; I0 = fluorescence intensity after 30 min
from the initial mixing of the solutions with 0 μM enzyme.)
(1H, s), 7.05 (1H, d, J = 12.7 Hz), 6.74 (1H, d of t, J = 18.3, 4.1 Hz),
6.41 (1H, s), 6.07 (1H, d, J = 12.7 Hz), 5.83 (1H, d, J = 18.2 Hz), 4.60
(2H, d of d, J = 3.9, 1.4 Hz), 3.88 (6H, s), 2.92 (2H, q, J = 7.4 Hz),
1.26 (15H, m); 13C NMR (75 MHz, CDCl3) δ 190.4, 153.0, 152.1,
147.6, 143.0, 135.3, 123.9, 120 (br.), 116.4, 114.5, 98.1, 84.1, 71.3,
56.9, 56.6, 25.4, 24.3, 15.4; IR (neat) 2974, 2932, 2839, 2531, 2161,
2029, 1655 cm−1; (TOF MS APCI+, m/z) 435.2 (M + H+); HRMS
(TOF MS ES+, m/z) calcd for C22H32O6BS (M + H+) 435.2013,
found 435.1994.
S-Ethyl (Diethoxyphosphinyl)ethanethioate (15). A solution
of diethylphosphonoacetic acid (14) (420 μL, 2.61 mmol, 1 equiv)
and DMAP (31 mg, 0.253 mmol, 0.097 equiv) were mixed in CH2Cl2
(9 mL) under argon and cooled to 0 °C. To this solution were added
ethanethiol (1 mL, 13.5 mmol, 5.17 equiv) and EDCI (600 mg, 3.13
mmol, 1.20 equiv). The reaction was stirred for 10 min, at which point
the argon inlet was removed, and the reaction was stirred for 8 h at 0
°C in a tightly sealed vessel. The mixture was concentrated under
reduced pressure to a clear oil and redissolved in Et2O (50 mL). The
organic layer was washed with NaHCO3 (2 × 50 mL), 0.5 M HCl (2 ×
50 mL), DI H2O (1 × 50 mL), and brine (3 × 50 mL), dried over
MgSO4, filtered, and concentrated in vacuo to yield S-ethyl
(diethoxyphosphinyl)ethanethioate (15) as a clear oil (0.216 g, 0.90
1
mmol, 34%): H NMR (360 MHz, CDCl3) δ 4.17 (4H, quin., J = 7.2
Hz), 3.21 (2H, d, JP−H = 21.3 Hz), 2.92 (2H, q, J = 7.5 Hz), 1.34 (5H,
t, J = 7.1 Hz), 1.27 (3H, t, J = 7.5 Hz). This spectral data is consistent
with data for the known compound.62
(2E)-S-Ethyl 3-(2-((2E)-3-(4,4,5,5-Tetramethyl-1,3,2-dioxa-
borolanyl)-2-propenoxy)-4,5-dimethoxyphenyl)-
propenethioate (12). This procedure was adapted from the work of
Mandal et al.63 Cs2CO3 (29 mg, 0.135 mmol, 1.62 equiv) and 9 (29
mg, 83 μmol, 1 equiv) were dissolved in 2.5 mL of THF under argon.
To this solution was added 15 (22 mg, 0.112 mmol, 1.35 equiv) in 2.5
mL of THF. The solution was heated at 50 °C for 16 h and then
cooled to room temperature. This mixture was concentrated in vacuo,
redissolved in the Et2O (50 mL), and washed with 1 M phosphate
buffer pH 6.8 (2 × 50 mL), DI H2O (1 × 50 mL), and brine (2 × 50
mL). The organic layer was dried over MgSO4, filtered, and
concentrated to an oil that was purified by column chromatography
(30% EtOAc/hexanes). The product was dried in vacuo overnight to
give a vibrant yellow oil, (2E)-S-ethyl-3-(2-((2E)-3-(4,4,5,5-tetrameth-
yl-1,3,2-dioxaborolanyl)-2-propenoxy)-4,5-dimethoxyphenyl)-
1
propenethioate (12) (7 mg, 17 μmol, 20%): H NMR (360 MHz,
CDCl3) δ 7.95 (1H, d, J = 15.9 Hz), 6.99 (1H, s), 6.75 (1H, d of t, J =
18.3, 4.1 Hz), 6.63 (1H, d, J = 15.9 Hz), 6.43 (1H, s), 5.79 (1H, d, J =
18.2 Hz), 4.66 (2H, d, J = 2.3 Hz), 3.87 (6H, s), 2.99 (2H, q, 7.2 Hz),
1.30 (15H, m); 13C NMR (75 MHz, CDCl3) δ 190.2, 153.4, 152.4,
147.1, 143.6, 135.3, 123.1, 120.6 (br), 115.1, 110.5, 98.5, 84.8, 71.0,
56.5, 56.2, 24.9, 23.3, 15.0; IR (neat) 2975, 2932, 2543, 2163, 2030,
1649, 1593 cm−1; (TOF MS APCI+, m/z) 435.2 (M + H+); HRMS
(TOF MS ES+, m/z) calcd for C22H32O6BS (M + H+) 435.2013,
found 435.2007.
(2Z)-Ethyl 3-(2-((2E)-3-(4,4,5,5-Tetramethyl-1,3,2-dioxaboro-
lanyl)-2-propenoxy)-4,5-dimethoxyphenyl)acrylate (13). This
procedure was modified from the procedure reported by Touchard.61
18-Crown-6 ether (70 mg, 0.265 mmol, 1.91 equiv) and anhydrous
K2CO3 (75 mg, 0.543 mmol, 3.91 equiv) were dissolved in 5 mL of
THF under argon and allowed to equilibrate overnight. The cloudy
solution was cooled −40 °C, and 9 (50 mg, 0.144 mmol, 1.04 equiv)
was added in one portion. Compound 10 (33 μL, 0.139 mmol, 1
equiv) was then added, and the pale yellow solution was stirred in the
dark under argon at −40 °C for 5 h. This mixture concentrated in
vacuo in the dark, redissolved in the Et2O (50 mL), and washed with 1
M phosphate buffer pH 7.1 (2 × 50 mL), water (1 × 50 mL), and
brine (2 × 50 mL). The organic layer was dried over MgSO4, filtered,
and concentrated to an oil that was purified by column
chromatography (20% EtOAc/hexanes). The product was concen-
trated dried in vacuo overnight to a white solid, (2Z)-ethyl 3-(2-((2E)-
3-(4,4,5,5-tetramethyl-1,3,2-dioxaborolanyl)-2-propenoxy)-4,5-
Procedure for Forced-Choice Ascending Concentration
Series Method for Limit of Detection by Smell Using ASTM
Protocol. Six sample sets of three mixed solutions of 2 (20 μL, 4 mM
in MeOH), glucose oxidase (100 μL, 15 U·mL−1 in pH 7.01, 20 mM
phosphate buffer and 400 mM NaCl), and glucose-6-phosphate or
lactose (100, μL, 200 mM in DI H2O) were prepared in 600 μL
microcentrifuge tubes. Out of each set, two of the samples were blanks
and had buffer (130 μL, pH 7.01 20 mM phosphate buffer and 400
mM NaCl) added. The remaining sample contained increasing
concentrations of alkaline phosphatase or β-D-galactosidase (130 μL,
0−100 nM, in pH 7.01 20 mM phosphate buffer and 400 mM NaCl).
These solutions were mixed by vortex for ∼10 s and then timed for 30
1
dimethoxyphenyl)acrylate (13) (35 mg, 83 μmol, 59%): H NMR
G
DOI: 10.1021/acs.joc.5b02013
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The Journal of Organic Chemistry
Featured Article
min. When a set reached 30 min of incubation, a panelist was asked to
identify which of the samples had the strongest odor. The geometric
mean for each panelist was calculated between the last incorrectly and
first correctly identified sample.
Almeida, Anthony DiLauro, and Inanllely Gonzalez in
providing independent evaluation of the qualitative portions
of the assay.
Procedure for Solution-Phase Stability of 2 by LC/MS
(Figure 5a). A solution of 2 (500 μL, 1.2 mM in 1:1 MeOH−40
mM HEPES buffer, pH 7.9) and naphthalene (1.5 mM) was prepared
in a brown glass vial. An aliquot of the resulting mixture was injected
into an analytical reversed-phase HPLC coupled to a mass
spectrometer, and additional aliquots were injected at regular intervals.
The temperature of the autosampler chamber remained between 30
and 32 °C. After injection of the sample, a solvent gradient was run as
follows: time, 0 min, 1:1 acetonitrile−water; time, 10 min, 1:1
acetonitrile−water; time, 15 min, 4:1 acetonitrile−water; time, 20 min,
4:1 acetonitrile−water.
Procedure for Solid-Phase Stability of 2 by LC/MS (Figure
5b). A 50 μL aliquot of 2 (10.2 mM) and naphthalene (10.4 mM) in
acetonitrile was added to a 1.5 mL centrifuge tube. The samples were
allowed to dry under a flow of air at room temperature, after which the
lids were closed and placed in heating block at 44 3 °C. At regular
intervals, a sample was removed from heat, cooled to room
temperature, and diluted with acetonitrile (125 μL). An aliquot of
the resulting mixture was injected into an analytical reversed-phase
HPLC coupled to a mass spectrometer. After injection of the sample,
an isocratic solvent gradient was run as 3:2 acetonitrile−water for 20
min.
Experimental Procedure for LC/MS Monitoring of Reagent
12 Response to Hydrogen Peroxide (Figure 7). A hydrogen
peroxide solution (3 μL, 1.0 M in water) was added to a solution of 12
(500 μL, 2.3 mM in 1:1 MeOH−40 mM HEPES buffer, pH 7.9) and
naphthalene (0.5 mM). The mixture was agitated by vortex ∼5 s. An
aliquot of the resulting mixture was injected into an analytical
reversed-phase HPLC coupled to a mass spectrometer, and additional
aliquots were injected at regular 25 min intervals and then 1 d later.
After injection of the sample, a solvent gradient was run as follows:
time, 0 min, 1:1 acetonitrile−water; time, 10 min, 1:1 acetonitrile−
water; time, 15 min, 4:1 acetonitrile−water; time, 20 min, 4:1
acetonitrile−water.
Procedure for Comparing Fluorescent Output of Thioester
(2) vs ester 13 (Table 1). A hydrogen peroxide solution (330 μL, 0,
10, 20, or 50 μM in 40 mM HEPES buffer, pH 7.9) was added to a
solution of 2 or 13 (20 μL, 4 mM in MeOH) in an ultramicro cuvette,
capped, and agitated with a vortex mixer for ∼7 s. After 30 min, the
fluorescence emission (I) of the mixture was measured, and the
average fluorescence of the 0 μM hydrogen peroxide sample (I0) was
subtracted (I − I0); this difference was plotted to obtain a graph (I =
fluorescence intensity after 30 min from the initial mixing of solutions;
I0 = fluorescence intensity after 30 min from the initial mixing of the
solutions with 0 μM hydrogen peroxide).
REFERENCES
■
(1) Martinez, A. W.; Phillips, S. T.; Whitesides, G. M.; Carrilho, E.
Anal. Chem. 2010, 82, 3−10.
(2) Phillips, S. T.; Lewis, G. G. MRS Bull. 2013, 38, 315−319.
(3) Phillips, S. T.; Lewis, G. G. Expert Rev. Mol. Diagn. 2014, 14,
123−125.
(4) Yager, P.; Edwards, T.; Fu, E.; Helton, K.; Nelson, K.; Tam, M.
R.; Weigl, B. H. Nature 2006, 442, 412−418.
(5) Berg, B.; Cortazar, B.; Tseng, D.; Ozkan, H.; Feng, S.; Wei, Q.;
Chan, R. Y.-L.; Burbano, J.; Farooqui, Q.; Lewinski, M.; Di Carlo, D.;
Garner, O. B.; Ozcan, A. ACS Nano 2015, 9, 7857−7866.
(6) Pohanka, M. Sensors 2015, 15, 13752−13762.
(7) Wargocki, P.; Deng, W.; Anwer, A.; Goldys, E. Sensors 2015, 15,
11653−11664.
(8) Thom, N. K.; Lewis, G. G.; Yeung, K.; Phillips, S. T. RSC Adv.
2014, 4, 1334−1340.
(9) Thongprajukaew, K.; Choodum, A.; Sa-E, B.; Hayee, U. Food
Chem. 2014, 163, 87−91.
(10) Chin, C. D.; Cheung, Y. K.; Laksanasopin, T.; Modena, M. M.;
Chin, S. Y.; Sridhara, A. A.; Steinmiller, D.; Linder, V.; Mushingantahe,
J.; Umviligihozo, G.; Karita, E.; Mwambarangwe, L.; Braunstein, S. L.;
van de Wijgert, J.; Sahabo, R.; Justman, J. E.; El-Sadr, W.; Sia, S. K.
Clin. Chem. 2013, 59, 629−640.
(11) Coskun, A. F.; Wong, J.; Khodadadi, D.; Nagi, R.; Tey, A.;
Ozcan, A. Lab Chip 2013, 13, 636−640.
(12) You, D. J.; Park, T. S.; Yoon, J.-Y. Biosens. Bioelectron. 2013, 40,
180−185.
(13) Wang, S.; Zhao, X.; Khimji, I.; Akbas, R.; Qiu, W.; Edwards, D.;
Cramer, D. W.; Ye, B.; Demirci, U. Lab Chip 2011, 11, 3411−3418.
(14) Zhu, H.; Sikora, U.; Ozcan, A. Analyst 2012, 137, 2541−2544.
(15) Iqbal, Z.; Bjorklund, R. B. Talanta 2011, 84, 1118−1123.
(16) Zhu, H.; Yaglidere, O.; Su, T.-W.; Tseng, D.; Ozcan, A. Lab
Chip 2011, 11, 315−322.
(17) Oncescu, V.; Mancuso, M.; Erickson, D. Lab Chip 2014, 14,
759−763.
(18) Wang, Q.; Wang, H.; Yang, X.; Wang, K.; Liu, R.; Li, Q.; Ou, J.
Analyst 2015, 140, 1161−1165.
(19) Wang, Q.; Liu, F.; Yang, X.; Wang, K.; Wang, H.; Deng, X.
Biosens. Bioelectron. 2015, 64, 161−164.
(20) Chavali, R.; Kumar Gunda, N. S.; Naicker, S.; Mitra, S. K. Anal.
Methods 2014, 6, 6223−6227.
(21) Chen, J.; Wu, W.; Zeng, L. Anal. Methods 2014, 6, 4840−4844.
(22) Fu, X.; Feng, X.; Xu, K.; Huang, R. Anal. Methods 2014, 6,
2233−2238.
ASSOCIATED CONTENT
* Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.joc.5b02013.
■
(23) Wang, Q.; Wang, H.; Yang, X.; Wang, K.; Liu, F.; Zhao, Q.; Liu,
P.; Liu, R. Chem. Commun. 2014, 50, 3824−3826.
(24) Zhu, X.; Xu, H.; Lin, R.; Yang, G.; Lin, Z.; Chen, G. Chem.
Commun. 2014, 50, 7897−7899.
(25) Mohapatra, H.; Phillips, S. T. Chem. Commun. 2013, 49, 6134−
6136.
S
Tables of data and reproductions of NMR spectra (PDF)
(26) Madasamy, T.; Santschi, C.; Martin, O. J. F. Analyst 2015, 140,
AUTHOR INFORMATION
Corresponding Author
*E-mail: sphillips@psu.edu.
Notes
■
6071−6078.
(27) Ren, K.; Wu, J.; Yan, F.; Zhang, Y.; Ju, H. Biosens. Bioelectron.
2015, 66, 345−349.
(28) Koo, K. M.; Ibn Sina, A. A.; Carrascosa, L. G.; Shiddiky, M. J. A.;
Trau, M. Analyst 2014, 139, 6178−6184.
The authors declare no competing financial interest.
(29) Hu, J.; Wang, T.; Kim, J.; Shannon, C.; Easley, C. J. J. Am. Chem.
Soc. 2012, 134, 7066−7072.
ACKNOWLEDGMENTS
■
(30) Safavieh, M.; Ahmed, M. U.; Tolba, M.; Zourob, M. Biosens.
Bioelectron. 2012, 31, 523−528.
This work was supported by the NIH (R01GM105686). We
appreciate the contributions of Dr. Lu Wang, Dr. Kimy Yeung,
Dr. Greg Lewis, Dr. Jessica Robbins, Dr. Saptarshi Chatterjee,
Dr. Hyungwoo Kim, Travis Cordes, Michael Olah, Karoliny
(31) Cass, A. E. G.; Zhang, Y. Faraday Discuss. 2011, 149, 49−61.
(32) Lee, A.-C.; Liu, G.; Heng, C.-K.; Tan, S.-N.; Lim, T.-M.; Lin, Y.
Electroanalysis 2008, 20, 2040−2046.
H
DOI: 10.1021/acs.joc.5b02013
J. Org. Chem. XXXX, XXX, XXX−XXX

The Journal of Organic Chemistry
Featured Article
(33) Centi, S.; Tombelli, S.; Minunni, M.; Mascini, M. Anal. Chem.
2007, 79, 1466−1473.
(34) Liu, H.; Xiang, Y.; Lu, Y.; Crooks, R. M. Angew. Chem., Int. Ed.
2012, 51, 6925−6928.
(35) Wu, Y.; Xue, P.; Hui, K. M.; Kang, Y. Biosens. Bioelectron. 2014,
52, 180−187.
(36) Noh, H.; Phillips, S. T. Anal. Chem. 2010, 82, 8071−8078.
(37) Schilling, K. M.; Lepore, A. L.; Kurian, J. A.; Martinez, A. W.
Anal. Chem. 2012, 84, 1579−1585.
(38) Lewis, G. G.; DiTucci, M. J.; Baker, M. S.; Phillips, S. T. Lab
Chip 2012, 12, 2630−2633.
(39) Lewis, G. G.; DiTucci, M. J.; Phillips, S. T. Angew. Chem., Int. Ed.
2012, 51, 12707−12710.
(40) Lewis, G. G.; Robbins, J. S.; Phillips, S. T. Anal. Chem. 2013, 85,
10432−10439.
(41) Lewis, G. G.; Robbins, J. S.; Phillips, S. T. Macromolecules 2013,
46, 5177−5183.
(42) Nunez, S. A.; Yeung, K.; Fox, N. S.; Phillips, S. T. J. Org. Chem.
̃
2011, 76, 10099−10113.
(43) Mohapatra, H.; Phillips, S. T. Angew. Chem., Int. Ed. 2012, 51,
11145−11148.
(44) Xu, Y.; Zhang, Z.; Ali, M. M.; Sauder, J.; Deng, X.; Giang, K.;
Aguirre, S. D.; Pelton, R.; Li, Y.; Filipe, C. D. M. Angew. Chem., Int. Ed.
2014, 53, 2620−2622.
(45) Zhang, Z.; Wang, J.; Ng, R.; Li, Y.; Wu, Z.; Leung, V.; Imbrogno,
S.; Pelton, R.; Brennan, J. D.; Filipe, C. D. M. Analyst 2014, 139,
4775−4778.
(46) Cate, D. M.; Dungchai, W.; Cunningham, J. C.; Volckens, J.;
Henry, C. S. Lab Chip 2013, 13, 2397−2404.
(47) Fu, E. Analyst 2014, 139, 4750−4757.
(48) Cate, D. M.; Noblitt, S. D.; Volckens, J.; Henry, C. S. Lab Chip
2015, 15, 2808−2818.
(49) Greenman, J.; El-Maaytah, M.; Duffield, J.; Spencer, P.;
Rosenberg, M.; Corry, D.; Saad, S.; Lenton, P.; Majerus, G.;
Nachnani, S. J. Am. Dent. Assoc., JADA 2005, 136, 749−757.
(50) This hypothesis for the source of background signal in 6
remains unsubstantiated.
(51) Mosey, R. A.; Floreancig, P. E. Org. Biomol. Chem. 2012, 10,
7980−7985.
(52) Hall, D. G. In Boronic Acids; Wiley−VCH, 2006; pp 1−99.
(53) Hughes, M. P.; Smith, B. D. J. Org. Chem. 1997, 62, 4492−4499.
(54) The limits of detection were cacluated using the standard
deviation of the data when no enzyme analyte was present. The
calculation used the following expression: (3 × std[enzyme analyte=0])/
slope obtained from the calibration curve.
(55) A Quantifluor-ST Fluorometer Method for 4-Methylumbelliferone;
Application Note S-0080; Promega Corporation: Madison, WI, Nov
2011.
(56) Determination of Odor and Taste Thresholds By a Forced-
Choice Ascending Concentration Series Methods of Limits. ASTM
Standard E679, 2004; ASTM International: West Conshohocken, PA,
2011, DOI: 10.1520/E0679-04R11, www.astm.org.
(57) Pangborn, A. B.; Giardello, M. A.; Grubbs, R. H.; Rosen, R. K.;
Timmers, F. J. Organometallics 1996, 15, 1518−1520.
(58) Still, W. C.; Kahn, M.; Mitra, A. J. Org. Chem. 1978, 43, 2923−
2925.
̈
(59) Gunduz, C.; Salan, U.; Bulut, M. Supramol. Chem. 2010, 22,
̈
̈
491−498.
(60) Sano, S. T.; Takemoto, Y.; Nagao, Y. ARKIVOC 2003, 8, 93−
101.
(61) Touchard, F. P. Tetrahedron Lett. 2004, 45, 5519−5523.
(62) ter Horst, B.; van Wermeskerken, J.; Feringa, B. L.; Minnaard, A.
J. Eur. J. Org. Chem. 2010, 2010, 38−41.
(63) Mandal, P. K.; Liao, W. S. L.; McMurray, J. S. Org. Lett. 2009,
11, 3394−3397.
I
DOI: 10.1021/acs.joc.5b02013
J. Org. Chem. XXXX, XXX, XXX−XXX