Measuring human carboxylesterase 2 activity in pancreatic cancer patient-derived xenografts using a ratiometric fluorescent chemosensor

Article abstract of DOI:10.1039/c9sc00283a

Irinotecan-based therapy is a common treatment for pancreatic cancer. To elicit its anticancer activity, the drug requires first the hydrolysis action of the enzyme human carboxylesterase 2 (hCES2). It has been established that pancreatic cancer patients have various levels of hCES2, whereby patients having low levels respond poorer to Irinotecan than patients with higher levels, suggesting that hCES2 can be used to predict response. However, current methods that measure hCES2 activity are inaccurate, complex or lengthy, thus being incompatible for use in a clinical setting. Here, we developed a small molecule ratiometric fluorescent chemosensor that accurately measures hCES2 activity in a single-step within complex mixtures. Our chemosensor is highly selective for hCES2 over hCES1, cell permeable and can measure hCES2 activity in pancreatic cancer patient-derived xenografts. Given the simplicity, accuracy and tissue compatibility of our assay, we anticipate our chemosensor can be used to predict patient response to Irinotecan-based therapy.

Full text of DOI:10.1039/c9sc00283a

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Measuring human carboxylesterase 2 activity in
pancreatic cancer patient-derived xenografts using
a ratiometric fluorescent chemosensor†
Cite this: DOI: 10.1039/c9sc00283a
All publication charges for this article
have been paid for by the Royal Society
of Chemistry
Karishma Kailass,a Oleg Sadovski,a Michela Capello,b Ya'an Kang,c Jason B. Fleming,d
a
Samir M. Hanashb and Andrew A. Beharry
*
Irinotecan-based therapy is a common treatment for pancreatic cancer. To elicit its anticancer activity, the
drug requires first the hydrolysis action of the enzyme human carboxylesterase 2 (hCES2). It has been
established that pancreatic cancer patients have various levels of hCES2, whereby patients having low
levels respond poorer to Irinotecan than patients with higher levels, suggesting that hCES2 can be used
to predict response. However, current methods that measure hCES2 activity are inaccurate, complex or
lengthy, thus being incompatible for use in a clinical setting. Here, we developed a small molecule
ratiometric fluorescent chemosensor that accurately measures hCES2 activity in a single-step within
complex mixtures. Our chemosensor is highly selective for hCES2 over hCES1, cell permeable and can
measure hCES2 activity in pancreatic cancer patient-derived xenografts. Given the simplicity, accuracy
and tissue compatibility of our assay, we anticipate our chemosensor can be used to predict patient
response to Irinotecan-based therapy.
Received 17th January 2019
Accepted 28th July 2019
DOI: 10.1039/c9sc00283a
rsc.li/chemical-science
Irinotecan, is converted to its active metabolite SN-38 by hCES2
to inhibit the DNA repair enzyme topoisomerase 1.7
Introduction
Given the importance of hCES2 in facilitating the conversion
of prodrugs to drugs, a major factor governing anticancer drug
efficacy has been the level of hCES2 activity found in cancers
and patients. Most notably, hCES2 has been used as an indi-
cator of response to Irinotecan treatment against neuroblas-
toma, metastatic colorectal cancer, and non-small cell lung
cancer.8–10 In pancreatic ductal adenocarcinoma (PDAC)
patients, hCES2 expression and activity was found to be a major
contributor to Irinotecan sensitivity when administered in the
chemotherapeutic cocktail, FOLFIRINOX.11 Thus, by using
hCES2 as a predictive marker in biopsied pancreatic tissues,
a patient's response towards drugs like Irinotecan can be
determined before therapy to help guide the choice of the most
appropriate therapeutic drug for that individual. Moreover, by
continuously monitoring hCES2 activity over the course of
therapy, any acquired resistance to Irinotecan can be caught
early and the treatment plans can be adjusted accordingly. To
achieve these goals in the clinic, a hCES2 assay that is simple,
accurate and can be used in high throughput formats is highly
warranted.
Carboxylesterases (CE) are a family of conserved a,b-fold
intracellular hydrolases that catalyze the hydrolysis and trans-
esterication of a variety of exogenous substrates containing
esters, amides, carbamates and thioester groups.1 Humans have
two major CE known as human carboxylesterase 1 (hCES1) and
human carboxylesterase 2 (hCES2), which both act as phase-I
drug metabolizing enzymes but differ in tissue distribution
and substrate specicity.1 Importantly, the activity of CE,
particularly hCES2 (expressed mainly in gastrointestinal tract,
minor expression in liver), was found to be responsible for the
activation of several anticancer agent prodrugs.2 For example,
the prodrug Gemcitabine requires hCES2-mediated hydrolysis
before it can be phosphorylated to the active triphosphate form
that ultimately inhibits DNA synthesis.3,4 The prodrug pentyl
PBABC-Doxaz is rst activated by hCES2 to the anthracycline
antitumour drug, Doxazolidine, a DNA cross-linking agent used
to kill cancer cells.5,6 Finally, the chemotherapeutic prodrug
aDepartment of Chemical and Physical Sciences, University of Toronto Mississauga,
Mississauga, ON L5L 1C6, Canada. E-mail: andrew.beharry@utoronto.ca
bDepartment of Clinical Cancer Prevention, The University of Texas MD Anderson
Cancer Center, Houston, TX, USA
The gold standard to measure hCES2 activity uses para-
nitrophenyl acetate (p-NPA), whereby active CEs will hydrolyze
the acetate group, producing para-nitrophenol and an absor-
bance increase at 405 nm. Although direct in activity, the assay
can only be used aer the cells are lysed, and is non-selective for
measuring hCES2 activity (e.g. active hCES1 will also produce
a signal). Other methods, such as immunoblotting, mass
cDepartment of Surgical Oncology, The University of Texas MD Anderson Cancer
Center, Houston, TX, USA
dDepartment of Gastrointestinal Oncology, H. Lee Moffitt Cancer Center, Tampa, FL,
USA
† Electronic supplementary information (ESI) available. See DOI:
10.1039/c9sc00283a
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spectrometry, liquid chromatography, and proteomic tech- conversion of the amine to an amide via ligation to a pyruval or
niques are oen used, although they are costlier, labor inten- benzyl moiety would cause a blue shi in the absorption spec-
sive, non-selective, and inaccurate as they oen correlate hCES2 trum, and a red-shi (i.e. recovering the properties of the parent
RNA or protein expression levels with enzymatic activity.12–15 dye) upon hCES2-mediated hydrolysis of the amide bond.
Fluorescent chemosensors are an attractive alternative to Compound
3 was synthesized according to established
conventional methods due to their simplicity, low detection methods27,28 (see ESI, Scheme 2†), and then coupled with pyr-
limit, non-invasive nature, and potential for real-time cellular/ uvyl chloride (see ESI, Scheme 3†), or benzoyl chloride (see ESI,
tissue applications. Perhaps the most common chemosensor Scheme 4†), to yield probes 1 and 2, respectively.
for hCES2 is uorescein diacetate, which is non-uorescent in
The photophysical properties and stability of probes 1 and 2,
its spirocyclized form, but becomes uorescent upon hCES2 and compound 3 in the absence of hCES2 were rst investigated
mediated hydrolysis of the acetate groups to produce uores- and are summarized in Table 1. As expected, probes 1 and 2
cein.16 Although this probe has been used to measure hCES2 exhibited blue shis in their absorption spectra compared to
activity in cells, it suffers from high false positives due to the parent dye, 3. Interestingly, conversion of the amine to an
background hydrolysis in the cellular media and false negatives amide did not diminish its uorescence, but rather resulted in
due to quenching of uorescein from certain cellular compo- a blue-shi towards the yellow spectrum region with overall
nents.16,17 Several other uorescent probes have been designed similar brightness to 3. To determine aqueous stability, each
that contain a uorophore conjugated via an ester linkage with probe was excited at their corresponding excitation maxima and
a benzoyl moiety.18–21 These probes were found to be selective the uorescence emission at 605 nm was monitored as a func-
for hCES2, but were sensitive to environmental factors (e.g. tion of time. We reasoned that any potential instability would be
temperature and pH), producing substantial background signal due to the presence of the aryl amide linkage, which could
when introduced in cells. To overcome these issues, several undergo background, uncatalyzed hydrolysis to generate the
ratiometric uorescent probes were constructed to more accu- parent dye, 3. We found that probe 1 exhibited a measurable
rately quantify hCES2 activity in living cells.20–24 However, these increase in uorescence at 605 nm when incubated under
ꢀ
probes either have aqueous solubility issues, absorb and emit at physiological conditions (PBS pH 7.4, 37 C), however probe 2
short wavelengths, suffer from photostability issues or are showed no change in uorescence under the same conditions
poorly selective for hCES2 over hCES1.21,25 Moreover, the use of (Fig. S1†). We hypothesized the difference in stability may be
these probes in patient-derived xenogras has yet to be due to the electron-withdrawing carbonyl functionality neigh-
demonstrated.
boring the carbonyl of the amide bond in probe 1, which is
To address this, we describe the design, synthesis and replaced by an electron donating phenyl moiety in probe 2
characterization of a uorescent chemosensor that responds to making the carbonyl carbon less electrophilic. Thus, given its
hCES2 activity ratiometrically in long wavelength emission superior aqueous stability, enzymatic and cellular studies were
channels (i.e. yellow and red), and with high selectivity over its conducted using probe 2.
related family member, hCES1. Our probe exhibits high water
To determine if probe 2 can undergo hCES2 catalyzed
solubility, photostability and cell permeability, and can be used hydrolysis, we rst monitored its UV-Vis spectrum as a function
to quantify hCES2 activity in cultured cancer cells as well as in of time. Upon addition of hCES2 (110 nM) we observed
pancreatic cancer patient-derived tissue samples.
a decrease in absorbance at 413 nm followed by an increase at
548 nm, with a nal spectrum resembling that of compound 3
(Fig. S2†). Given that probe 2 has weak absorbance at 548 nm,
we reasoned that excitation at 548 nm (where 3 would have
maximum absorbance) would lead to an increase in uores-
cence as 3 is produced. Indeed, we found that addition of
110 nM hCES2 led to a 40-fold increase in uorescence emission
at 605 nm, with a uorescence spectrum matching that of the
expected product, 3 (Fig. 1), which was further validated by
HPLC analysis of the reaction mixture (Fig. S3†). Similarly,
exciting 2 at its maximum absorbance at 413 nm resulted in
a decrease in emission at 562 nm upon addition of hCES2
(Fig. 1). Similar uorescence responses were also observed as
Results and discussion
Design, synthesis and evaluation of uorogenic hCES2 probes
To design probes that can report on hCES2 activity, we focused
on developing a strategy for coupling a uorescence change to
the CE hydrolysis reaction. With this in mind, we designed two
probes containing the pyruval (1) or benzoyl (2) group conju-
gated to the dye, 6-amino-1-phenalenone (3). Although the
photophysical properties of 3 have not been thoroughly char-
acterized,26 it was selected since it contains an exocyclic amine
in conjugation with the pi system of the dye. We reasoned that
Table 1 Photophysical properties of hCES2 chemosensors
lex
lem
Stability
in PBS
3 (Mꢁ1 cmꢁ1
)
(nm)
FF
(nm)
Brightness (Mꢁ1 cmꢁ1
)
3 (AP)
Probe 2 (Benz-AP)
Probe 1 (Pyr-AP)
12 736
8880
8872
548
413
413
0.20 ꢂ 0.01
0.39 ꢂ 0.1
0.16 ꢂ 0.03
605
562
550
2547 ꢂ 150
3500 ꢂ 900
1420 ꢂ 200
Yes
Yes
No
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Fig. 1 Ratiometric fluorescence response of probe 2 for hCES2
activity. Top, scheme of probe 2 upon hCES2 hydrolysis of the aryl
amide bond produces the dye 3 and benzoic acid as a by-product.
Bottom, time-dependent fluorescence emission spectra upon exci-
tation at 413 nm (left) and excitation at 548 nm (right) results in
a decrease and increase in yellow and red fluorescence, respectively,
upon reaction with hCES2 (insets, plot of emission maxima as a func-
tion of time). Scans are taken at 10 minute intervals at 37 ꢀC. Probe 2 ¼
11 mM, hCES2 ¼ 110 nM.
Fig. 2 Probe 2 selectivity for hCES2 and its response to CE inhibitors.
Incubation of probe 2 with 110 nM of either hCES1, AChE, BChE,
Cathepsin S or Cathepsin B resulted in no fluorescence increase. Pre-
incubation of hCES2 with the inhibitors LPA or BNPP resulted in
a substantial reduction in overall fluorescence compared to the positive
control. Fluorescence at 605 nm (Ex. 548 nm) was measured after 1
hour at 37 ꢀC. The positive control (“hCES2”) was normalized to 100%
activity. Experiments were performed in triplicates. Statistical signifi-
cance is calculated using unpaired t test compared with hCES2 (**p <
0.01, *p < 0.05). LPA ¼ loperamide, BNPP ¼ bis(4-nitrophenyl)phos-
phate, AChE ¼ acetylcholinesterase, BChE ¼ butyrylcholinesterase.
low as 5 nM hCES2 with an expected slower rate (Fig. S4†).
Lastly, probe 2 and compound 3 were found to be photostable
with constant irradiation for 1 hour and soluble in PBS up to 75
mM and 140 mM, respectively (Fig. S5†).
we rst incubated probe 2 (20 mM) with the pancreatic cell line,
SU.86.86, known to contain no detectable amounts of hCES2 by
Western blot (Fig. 3).11 Aer a 3 hour incubation, cells were
washed and imaged in the yellow channel for probe 2 and the
red channel for the expected hCES2 product, 3.
To validate the uorescence response is in fact due to hCES2
activity, we pre-incubated hCES2 with general CE covalent
inhibitor bis(4-nitrophenyl)phosphate (BNPP) or a hCES2-
specic non-covalent inhibitor loperamide (LPA) followed by
addition of probe 2. We observed an overall lower uorescence
increase at 605 nm (Fig. 2 and S6†) suggesting that our probe is
in fact responding to hCES2 activity and that the uorescence
response observed is due to hCES2 catalyzed hydrolysis of the
aryl amide bond to produce 3 (Fig. 1). Lastly, we determined if
probe 2 is selective for hCES2 over other major human car-
boxylesterase isoforms, mainly hCES1, which have been shown
to act on similar substrates.23 Although active on p-NPA
(Fig. S7†), addition of hCES1 (110 nM) to probe 2 led to no
signicant increase in uorescence emission at 605 nm (Fig. 2,
S8 and S9†). Further increasing the concentration of hCES1 up
to 440 nM also did not produce any product (Fig. S9†). These
results suggest probe 2 is highly selective for hCES2 over hCES1
As shown in Fig. 3 (Fig. S11†), yellow uorescence appeared
to be diffuse in the cytosol with no uorescence detected in the
nucleus. Switching to the red channel revealed low red uo-
rescence relative to the yellow channel (Fred/Fyellow ¼ 0.60 ꢂ
0.080), suggesting that most of probe 2 is present relative to the
product dye, 3. Next, we added probe 2 to SU.86.86 cells that
contain a vector overexpressing hCES2 (CES2 OE) (Fig. 3).11
Incubation of probe 2 under the same conditions, we now
observed high red uorescence and low yellow uorescence
(Fred/Fyellow ¼ 1.4 ꢂ 0.13), indicating a signicant portion of
probe 2 has been converted to 3 (Fig. 3 and S11†). To control for
any effects a vector could have on our uorescence signals, we
also incubated probe 2 with parental SU.86.86 containing an
empty vector (“control”). As seen in Fig. S12,† control and
parental cell lines yielded similar uorescence ratios. To ensure
the difference between CES2 OE and parental is due to the
action of hCES2, we pre-incubated CES2 OE cells with the
hCES2 specic LPA inhibitor, followed by incubation of probe 2
for 3 hours. We observed a signicant decrease in the Fred/Fyellow
ratio (Fred/Fyellow ¼ 0.37 ꢂ 0.060) compared to cells untreated
with LPA suggesting cellular hCES2 activity is responsible for
the conversion of probe 2 to compound 3 (Fig. 3 and S11†).
Treating the parental SU.86.86 with LPA did not result in a lower
ratio suggesting there is no signicant level of active hCES2,
consistent with previous Western blots.11 In addition to ratio-
metric imaging, we also performed single red channel
with a measured Km and kcat of 6.6 ꢂ 3.7 mM and 0.26 ꢂ 0.04 sꢁ1
,
respectively (Fig. S10†). Finally, probe 2 tested against other
hydrolases also did not produce a signicant uorescent
response (Fig. 2).
hCES2 activity in pancreatic cancer cells
Given the selective ratiometric uorescent response of probe 2
for hCES2, we asked whether it can be used to measure hCES2
activity in cultured cancer cells. Previous work by Capello et al.
have shown that hCES2 activity in pancreatic cancers is a crucial
determinant for the effectiveness of the anti-cancer drug, Iri-
notecan.11 To test whether the new probes might address this,
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Fig. 3 Measuring hCES2 activity in pancreatic cancer cells. Probe 2 (20 mM) was incubated for 3 hours with the pancreatic cancer cell line,
SU.86.86. Top row, probe 2 with SU.86.86 cells containing a vector overexpressing CES2 (CES2 OE) displayed strong red over yellow fluo-
rescence, indicating high conversion of probe 2 to product 3. Bottom row, probe 2 with parental SU.86.86 showed weak red fluorescence but
high yellow fluorescence indicative of poor conversion of probe 2 to 3. Far top right, Western blot showing hCES2 expression levels. Far bottom
right, bar graph quantifying the ratio of red over yellow fluorescence for CES2 OE and parental. Light gray bars indicate pretreatment with the
CES2 inhibitor, LPA, dark gray bars are no pretreatment (images in Fig. S9†). Ratios are an average of 3 regions in an 8-chambered well plate.
Magnification is 20ꢃ. Scale bars ¼ 50 mm. Statistical significance is calculated using unpaired t test compared with CES2 (***p < 0.001).
measurements (no washing), which yielded similar results shows PDX112 displaying a higher red/yellow emission ratio
(Fig. S13†). Lastly, we found that incubation of our probe for 3 (median Fred/Fyellow ¼ 0.98) compared to PDX121 (median Fred
/
hours up to 20 mM did not cause any cell toxicity (Fig. S14†).
Fyellow ¼ 0.34; p < 0.0001). H&E staining conrmed the presence
of cancerous tissue (Fig. S15†) while Western blots conrmed
the presence of CES2, with signicantly higher levels found in
PDX112 over PDX121 (Fig. 5). Although PDX121 shows no
detectable expression, uorescence reveals some degree of
hCES2 activity consistent with what we observed with parental
SU.86.86 also containing no detectable hCES2 expression and
with previous reports.11 Single channel measurements also
showed a higher median uorescence for PDX112 over PDX121,
however there is increased variability in PDX121 signals due to
a relatively larger interquartile range compared to ratiometric
measurements (Fig. S16†). Finally, it is worth noting that
addition of probe 2 to mostly stroma or necrotic tissues
produced low ratiometric signals (Fig. S17†) suggesting some
CES2 activity consistent with previous reports.6
We have developed a small-molecule ratiometric uorescent
chemosensor that can report on the activity of hCES2 in
pancreatic cancer cells and in patient-derived xenogras.
Although other hCES2 uorescent chemosensors have been
reported, probe 2 described here overcomes several limitations
that have hindered previous hCES2 chemosensors from
providing accurate and practical measurements of hCES2
activity.18–22,25,32–34 First, probe 2 can be readily synthesized
following the facile coupling of 3 to benzoyl chloride and
demonstrates enhanced selectivity for hCES2 over hCES1 to the
extent where Michaelis–Menten parameters for hCES1 could
not be measured.21,24,33,34 Second, our sensor demonstrates
improved solubility in aqueous media, with a maximal use of
0.5% organic solvent as opposed to 50% organic solvent used in
other reported probes.18–22,25 Thirdly, in addition to being
a ratiometric chemosensor, probe 2 also has the advantage of
hCES2 activity in pancreatic cancer patient-derived xenogras
Next, we determined whether probe 2 can be used to measure
hCES2 activity in pancreatic cancer tissue samples. We rst
obtained control vector and CES2 OE SU.86.86 orthotopic
xenogra tumors as frozen tissue slices (10 mm in thickness) on
microscope slides. We added 100 mL of probe 2 directly on the
area containing the tissue, then captured red channel uores-
cence readings every 30 min. In both tissue samples, we
observed a time dependent increase in red uorescence,
however, the CES2 OE orthotopic tissues showed an overall
larger uorescence increase aer 3 hours (p < 0.0001) (Fig. 4A).
Aer 3 hours, tissues were washed 3ꢃ with PBS to capture
uorescence from both red and yellow emission channels. The
average uorescence intensity was collected at 3 hours at 12
different regions of a given tissue sample, then their ve-
number summary distribution was plotted to construct a box
and whisker plot (Fig. 4B). Consistent with cell culture experi-
ments, the CES2 OE SU.86.86 tissues displayed higher red/
yellow ratio (median Fred/Fyellow ¼ 2.71) compared with control
vector tissues (median Fred/Fyellow ¼ 0.44; p < 0.0001). We further
performed H&E staining which indeed conrmed the presence
of cancerous tissue (Fig. S15†).
Lastly, we obtained patient-derived xenogras (PDX) which
have previously served as reliable models for predicting anti-
cancer efficacy of therapeutic agents.29–31 We applied probe 2 to
tissue samples from two pancreatic cancer patients (PDX112
and PDX121) and captured ratiometric images following the
same procedure we established for orthotopic xenogras. Fig. 5
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Fig. 4 Detecting hCES2 activity in orthotopic SU.86.86 xenografts. A few drops of probe 2 (20 mM) was added to a SU.86.86 frozen tissue slice
(10 mm in thickness) on a microscope slide and imaged. (A) Single channel red fluorescence as a function of time whereby probe 2 added to
SU.86.86 CES2 OE tissues (top row) led to larger time dependent increase in fluorescence, compared to control vector SU.86.86 tissues (bottom
row). (B) Ratiometric images obtained after 3 hour incubation. Tissues were washed 3 ꢃ with PBS to obtain images in the yellow channel. A higher
red/yellow ratio was observed for CES2 OE orthotopic xenografts compared to the respective control. Average fluorescence was collected at 12
different regions of the tissues at 3 hours to construct a box plot (far right). Magnification is 20ꢃ. Scale bars ¼ 50 mm. Statistical significance is
calculated using the Mann–Whitney–Wilcoxon test compared to CES2 OE (****p < 0.0001).
monitoring at longer wavelength emission channels (i.e. red nm).20,25,32 Overall, these properties of probe 2 render it by far
and yellow) compared to previous sensors, thereby minimizing superior to other reported hCES2 probes.
interference from auto-uorescence and improving tissue
The gold standard for measuring cellular hCES2 activity is
penetration depth.18,19,32 Furthermore, excitation of our sensor the colorimetric p-NPA assay, which requires cell lysis prior to
in the visible region reduces photodamage compared to several measurements. The act of lysis not only adds additional steps,
other hCES2 uorescent probes excited in the UV range (<400 but can also lead to deactivation of enzymes in the process.
Fig. 5 Detecting hCES2 activity in patient-derived xenografts. A few drops of probe 2 (20 mM) was added to a tissue slice (10 mm in thickness) on
a microscope slide and ratiometric images were acquired for after 3 hours. Far top right, Western blot showing hCES2 expression levels. Far
bottom right, average fluorescence collected at 12 different regions of the tissues at 3 hours to construct a box plot. Magnification is 20ꢃ. Scale
bars ¼ 50 mM. Statistical significance is calculated by the Mann–Whitney–Wilcoxon test (****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05).
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Moreover, p-NPA is not selective for hCES2 and so the presence emissions (red and yellow). Probe 2 can be used to measure
of related esterases such as hCES1 would lead to an over- hCES2 activity in live pancreatic cancer cells with no cytotoxicity
estimation of hCES2 activity. In contrast to p-NPA, probe 2 at active concentrations. We also show for the rst time the use
undergoes a ratiometric uorescence response highly selective of a uorescent chemosensor for detecting hCES2 in pancreatic
for hCES2 over hCES1 activity. We show that probe 2 is cell cancer patient-derived xenogras. The developed assay raises
permeable and can readily respond to levels of hCES2 activity in the possibility of predicting patient response to Irinotecan-
live pancreatic cancer cells in just a few hours. Given the based therapy in a high throughput clinical setting.
simplicity and accuracy of our assay, we envision that probe 2
can be used in conjunction with ow cytometry to map hCES2
activity across various pancreatic cancer cell lines to make
Experimental section
General information
stronger correlations between activity and Irinotecan potency,
as was previously generated using p-NPA assay. Probe 2 may also
be used to discover new connections between hCES2 activity
and anticancer drug resistance in a variety of other cancer cell
lines in more of a throughput manner compared to other
methods. Finally, given the change in uorescence against
known existing CE inhibitors such as BNPP and LPA, probe 2
may also be used to discover new inhibitors or activators of
hCES2 directly in its native cellular environment.
Lastly, we demonstrate for the rst time the use of a uo-
rescent chemosensor for detecting hCES2 activity in patient-
derived xenogras. Previous work by Capello et al. found that
hCES2 expression is highly variable amongst pancreatic cancer
patients.11 They further demonstrated that the potency of the
anti-cancer drug Irinotecan is highly dependent on the active
hCES2. In our work, we showed that probe 2 can detect hCES2
activity within intact orthotopic SU.86.86 xenogras. Consistent
with cells in culture, probe 2 was capable of reporting the ex-
pected high level of hCES2 in CES2 OE tissues compared to
control vector tissues. Moreover, adding probe 2 to PDX
samples revealed hCES2 activity consistent with expression
levels via Western blots. Unfortunately, only one patient
(PDX112) out of the two had been treated with FOLFIRINOX,
and so we could not determine how the measured hCES2
activity correlates with survival rates. Given the simplicity of the
assay demonstrated here, it seems feasible that probe 2 can be
used to measure hCES2 activity on a large number of patient
tissues whereby their clinical response to Irinotecan had been
recorded. We then envision that one could determine
Chemicals and solvents were purchased from either Fisher
Scientic or Sigma-Aldrich unless otherwise stated and were
used without further purication. All reactions were conducted
under argon environment and in oven-dried glassware. Reac-
tions were monitored by silica gel TLC plates and were viewed
with a low intensity short/long UV wavelength lamp. 1H and 13
C
NMR spectra were taken on Bruker 400 MHz NMR spectrom-
eter. HRMS were acquired from the AIMS Mass Spectrometry
Laboratory at the Department of Chemistry, University of Tor-
onto. High performance liquid chromatography (HPLC) was
performed on a LC-20AT Shimadzu liquid chromatograph,
equipped with an SPD-M20A VD diode array detector, a CBM-
20A VP system controller, and a C18 semi-prep column.
Human CES2 (expressed in HEK293 cells, C-terminal His Tag)
was purchased from Sino Biological Incorporated (10380-
H08H). Human CES1 (expressed in E. coli, N-terminal His tag)
was purchased from Abbexa Limited (abx168930). All stock
solutions of probe 1, probe 2, and 3 were prepared by dissolving
each sample in 1 : 1 ACN/HyClone Cell Culture Grade Water.
Stock solutions of bis(4-nitrophenyl)phosphate (BNPP) and
loperamide (LPA) were prepared by dissolving each inhibitor in
DMSO (Molecular Probes™ DMSO, Anhydrous). Absorbance
spectra were obtained on a Shimadzu UV-1800 UV Spectro-
photometer. Fluorescence emission spectra were obtained on
a Shimadzu RF-6000 Spectro Fluorophotometer.
Synthesis of phenalenone (1H-phenalen-1-one)
a threshold for hCES2 activity that is required to elicit certain The synthesis of phenalenone was carried out as previously
survival rates. Such data may even rst be obtained in high described.27 Aluminum chloride (40 g, 0.3 mol) and naphtha-
throughput on PDXs that have been treated with anti-cancer lene (16.9 g, 0.1 mol) were adꢀded to anhydrous DCM (100 mL).
drugs, using recently developed live tissue sensitivity assays.35 The mixture was cooled to 0 C. Cinnamoyl chloride (13 g, 0.1
ꢀ
Finally, given that ratiometric signals were also observed in mol) was added at 0 C and the reaction was stirred at room
stroma and necrotic tissue, we note that uorescent chemo- temperature overnight. The mixture was then reuxed for 3 h,
sensing should be complemented by H&E staining to rule out cooled with iced hydrochloric acid, and ltered. The ltrate was
false positives/negatives. Future work will also aim to apply extracted with chloroform. The solids were repeatedly boiled
probe 2 directly to fresh biopsied tissue samples, where, by with hot chloroform (3 ꢃ 200 mL) and ltered until the ltrate
predicting hCES2 activity, could improve pancreatic cancer became colourless. All organic extracts were combined, dried
patient stratication for Irinotecan-based therapy.
over anhydrous sodium sulfate, and subjected to a rotary
evaporator to give a yellow solid (54%). The crude product was
further puried by column chromatography (petroleum ether/
ethyl acetate 10 : 1). H-NMR (400 MHz, CDCl3): d ¼ 8.63 (d, J
Conclusions
1
We have developed a small molecule ratiometric uorescent ¼ 7.34 Hz, 1H), 8.20 (d, J ¼ 8.07 Hz, 1H), 8.02 (d, J ¼ 8.44 Hz,
chemosensor that directly measures hCES2 activity. Probe 2 is 1H), 7.66–7.82 (m, 3H), 7.59 (t, J ¼ 7.70, 1H), 6.74 (d, J ¼ 9.90 Hz,
readily synthesized, soluble in aqueous media, photostable, and 1H). 13C-NMR (101 MHz, CDCl3): d ¼ 185.71, 141.83, 134.95,
can selectively monitor hCES2 activity at long wavelength 132.17, 131.97, 131.42, 130.40, 129.48, 129.25, 127.84, 127.55,
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127.15, 126.67. ESI-MS (m/z): [M + H]+ calculated for C13H8O puried by ash silica chromatography (100% DCM to DCM/
181.06 Da, found 181.02 Da.
MeOH 9 : 1) and further puried by HPLC (gradient of 5 to
100% water/ACN with 0.1% formic acid over 35 min), resulting
1
Synthesis of 6-amino-1-phenalenone (3; 6-amino-1H-
phenalen-1-one; AP)
in a yellow solid (52%). H NMR (400 MHz, CDCl3): d ¼ 8.72–
8.73 (d, 1H), 8.52 (s, 1H), 8.41–8.43 (d, J ¼ 7.8 Hz, 1H), 8.31–8.34
(d, 1H), 8.03–8.05 (d, 2H), 7.88–7.90 (t, 1H), 7.83–7.85 (d, J ¼
8.0 Hz, 1H), 7.77–7.79 (d, J ¼ 9.8 Hz, 1H), 7.66–7.68 (d, 1H),
7.62–7.64 (d, 2H). 13C-NMR (100 MHz, CDCl3): d ¼ 171.21,
162.39, 153.14, 142.20, 134.56, 132.49, 131.94, 130.97, 130.16,
127.36, 126.00, 125.90, 113.16, 112.51. HRMS (m/z): [M + H]+
calculated for C16H11NO3 300.09, found 300.1020.
The synthesis of 6-amino-1-phenalenone was carried out as
previously described, with some modications.28 To a solution
of phenalenone (5 g, 0.03 mol) in sulphuric acid was added
a dropwise mixture of nitric acid and sulphuric acid. The
reaction mixture was stirred for 3 h at room temperature. Water
was added and the mixture was extracted with dichloro-
methane. The combined extracts were dried with Na2SO4,
ltered and concentrated. The crude mixture was dissolved in
dioxene at room temperature. Tin chloride (25 g, 0.11 mol) and
acetic acid were slowly added and stirred for 2 days. The reac-
tion mixture was neutralized by sodium hydroxide, and extrac-
ted with ethyl acetate. The extract was puried by column
chromatography (ethyl acetate/hexanes to 100% ethyl acetate)
to yield a bright red solid (15%). 1H NMR (400 MHz, MeOD): d ¼
8.65 (d, J ¼ 6.24 Hz, 1H), 8.60 (d, J ¼ 6.97 Hz, 1H), 7.79–7.61 (m,
4H), 6.72 (d, J ¼ 8.07 Hz, 1H), 6.46 (d, J ¼ 9.17 Hz, 1H). 13C NMR
(101 MHz, MeOD): d ¼ 186.69, 160.43, 155.47, 145.25, 139.46,
132.96, 131.25, 131.15, 129.59, 125.94, 122.03, 117.03, 109.84.
DART-MS (m/z): [M + H]+ calculated for C13H9NO 195.0684 Da,
found 196.07598 Da.
hCES2 UV-Vis assays
Enzyme assays were performed in a 1 cm path length, 60 mL
quartz cuvette. Stock solutions of probes were diluted in DPBS
containing magnesium chloride and calcium chloride to a nal
concentration of 11 mM. Enzyme was added to yield a nal
concentration of 110 nM. To conrm hCES1 was active, 110 nM
hCES1 was added to 2 mM p-NPA in DPBS at 37 ꢀC and the UV-
Vis spectra was measured in 30 s intervals.
HPLC analysis of enzyme reaction
Probe 2, 3, and the hCES2 reaction with probe 2 were analyzed
by HPLC. Samples of puried probe 2 and 3 were prepared to
110 mM in ACN/0.1% formic acid. The hCES2 reaction was
prepared byꢀincubating 11 mM probe 2 with 110 nM hCES2 in
DPBS at 37 C for 3 h. The reaction mixture was concentrated
10ꢃ to achieve 110 mM product and dissolved in ACN/0.1%
formic acid for HPLC analysis. HPLC was performed on a LC-
20AT Shimadzu liquid chromatograph, equipped with an SPD-
Synthesis of probe 1 (2-oxo-N-(1-oxo-1H-phenalen-6-yl)
propanamide)
To a solution of pyruvic acid (50 mL, 0.705 mmol) in anhydrous
DCM (250 mL), oxalyl chloride (75 mL, 0.886 mmol) was added at
room temperature followed by a few drops of anhydrous DMF.
Aer stirring at room temperature for 30 min, the mixture was
concentrated by evaporation, resulting in pyruvyl chloride as an
oil. Sodium hydride (5.1 mg, 0.127 mmol) was added to
a mixture of AP (8 mg, 0.041 mmol) in DMF (750 mL) and was
stirred for 20 min. The mixture was then added dropwise to the
pyruvyl chloride oil and was stirred overnight. The solvent was
removed in vacuo and the residual solid was puried by ash
silica chromatography (100% DCM to DCM/MeOH 9 : 1). The
compound was further puried by HPLC (gradient of 5 to 100%
water/ACN with 0.1% formic acid over 35 min) to afford probe 1
M20A VD diode array detector,
a CBM-20A VP system
controller, and an ACE Equivalence 5 C18 column (250 ꢃ 3 mm
ID). The mobile phase consisted of gradient of 5–100% water/
ACN with 0.1% formic acid over 35 min using a ow rate of
0.2 mL minꢁ1
.
Water solubility
Stock solutions of probe 2 and compound 3 were prepared in
50% ACN/50% water. Dilutions were performed to yield solu-
tions ranging in concentration from 10 nM to 142 mM, with the
concentration of ACN maintained at less than 1%. UV-Vis
spectra were taken in DPBS in a 60 mL quartz cuvette and
absorbance at lmax (413 nm for probe 2, 548 nm for 3) was
plotted against concentration. The maximum linear response
was used to quantify the solubility limit.
1
as a yellow solid (46%). H-NMR (400 MHz, CDCl3): d ¼ 8.69–
8.71 (d, J ¼ 7.3 Hz, 1H), 8.45–8.47 (d, J ¼ 7.9 Hz, 1H), 8.28–8.30
(d, J ¼ 8.9 Hz, 1H), 7.87–7.91 (t, 1H), 7.78–7.80 (d, J ¼ 7.9 Hz,
1H), 7.72–7.75 (d, J ¼ 9.7 Hz, 1H), 6.70–6.73 (d, J ¼ 9.7 Hz, 1H),
2.67 (s, 3H). 13C-NMR (100 MHz, CDCl3): d ¼ 192.43, 171.33,
157.66, 143.07, 133.31, 130.06, 128.39, 124.72, 112.04, 111.93,
22.91, 1.03. HRMS (m/z): [M + H]+ calculated for C16H11NO3 hCES2 uorescence assays
266.07 Da, found 266.08 Da.
Enzyme assays were performed using a 1 cm path length, 60 mL
quartz cuvette. Stock solutions of probe were diluted in DPBS
containing magnesium chloride and calcium chloride to a nal
Synthesis of probe 2 (N-(1-oxo-1H-phenalen-6-yl)benzamide)
Sodium hydride (5.1 mg, 0.127 mmol) was added to a mixture of concentration of 11 mM. Enzyme was added to a nal concen-
AP (8 mg, 0.041 mmol) in DMF (750 mL) and stirred for 20 min. tration of 110 nM and the uorescence intensity was measured
Aer dropwise addition of benzoyl chloride (82 mL, 0.699 at 605 nm with 1 s intervals at a scan speed of 600 nm minꢁ1
ꢀ
mmol), the mixture was stirred overnight and the solvent was and 3 nm excitation and emission bandwidths at 37 C. Emis-
removed under reduced pressure. The crude product was sion spectra were collected before and aer the time course at
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excitation wavelengths of each probe and 3. Ratiometric uo- was calculated. Assuming that this concentration is equal to the
rescent properties were measured by collecting emission concentration of probe before the addition of probe, the molar
spectra from 0 to 160 min in 10 min intervals at excitation extinction coefficient was determined by the Beer–Lambert law.
wavelengths of 413 nm and 548 nm.
Michaelis–Menten enzyme kinetics assays
hCES2 uorescence inhibition assays and selectivity
55 nM CES2 was added to various solutions of probe 2 (0–66 mM)
prepared in DPBS buffer. Fluorescence intensity was collected at
605 nm (548 nm excitation) with 1 s intervals for 10 min at
37 ꢀC. Fluorescence intensity rates were converted to rates of
product formation (nM minꢁ1) by a uorescence calibration
curve (uorescence intensity versus concentration of AP
produced (mM)) generated from the reaction of 66 mM probe 2
with 55 nM CES2, assuming linearity. Michaelis–Menten kinetic
parameters (Km) were calculated by nonlinear regression anal-
ysis of the Michaelis–Menten equation (GraphPad Prism) by
Vmax½Sꢄ
For inhibition assays, 1 mM BNPP or LPA was incubated with
110 nM CES2 at 37 ꢀC in a DPBS/1% DMSO solution for 30 min.
11 mM probe 2 was then added to the solution and uorescence
emission at 605 nm was measured. For selectivity assays,
110 nM enzyme was added to 11 mM probe 2 in DPBS in a 60 mL
quartz cuvette and uorescence emission at 605 nm was
ꢀ
measured for 1 h at 37 C.
Fluorescence quantum yield
tting to the model V ¼
, where V represents enzyme
Km þ ½Sꢄ
The uorescence quantum yields of probe 1 and probe 2 were
determined using the uorophore perylene (FF ¼ 0.94 in
cyclohexane) as a standard due to overlap between the absorp-
tion spectra at the 413 nm excitation band of the probes.36
Separate solutions of perylene in cyclohexane and probe 1 or
probe 2 in DPBS were prepared to yield an absorbance of 0.05 at
366 nm. Emission spectra of each sample were taken with
excitation at 366 nm (1.5 nm bandwidth) and emission was
measured over the 376–800 nm range (1 nm emission band-
width) at a 60 nm minꢁ1 scan speed. The emission spectra were
integrated by Riemann sums at 1 nm intervals of the midpoint
of the uorescence intensity values from 376 to 700 nm. To
determine the uorescence quantum yield of 3, the uorophore
rhodamine B (FF ¼ 0.77 in ethanol) was used as a standard due
to its similar absorption prole.37 Emission spectra solutions of
rhodamine B in ethanol and 3 in DPBS with uorescence values
between 0.04 and 0.05 at 542 nm were measured with excitation
at 542 nm (1.5 nm bandwidth) from 552 nm to 800 nm (1 nm
bandwidth). The Riemann sum of the uorescence values at the
midpoint of 1 nm intervals from 552 nm to 710 nm was taken as
the integration of the emission spectra. Fluorescence quantum
yields were calculated according to the following equation:
velocity, Vmax represents maximal enzyme velocity, [S] repre-
sents substrate concentration, and Km represents substrate
concentration required to achieve half of the maximal enzyme
Vmax
velocity. kcat was determined by the equation kcat
¼
, where
½Eꢄ
kcat represents the turnover number and [E] represents the
concentration of enzyme.
Cell culture
Parental SU.86.86 cells were purchased from ATCC. Manipu-
lated SU.86.86 cell lines were provided by Dr Samir Hanash
from MD Anderson Cancer Center and were cultured as previ-
ously described.11 For overexpression, CES2 was cloned into
pLenti-C-Myc-DDK-IRESPuro (OriGene, Rockville, MD) vector,
an empty vector was used as a control. Lentiviral infections were
performed using 293LTV cells (Cell Biolabs, Inc., San Diego, CA)
as producers of viral supernatants. 293LTV cells were grown on
10 cm plates to 70% conuence and cotransfected with lenti-
viral DNA and the helper vectors pCMVD8.74 and pVSV-G
(Clontech, Mountain View, CA) using Lipofectamine 3000 (Life
Technologies), according to the manufacturer's instructions.
The medium was harvested 48 h post-transfection, ltered
through a 0.45 mm lter and used to cross-transduce cancer
cells in the presence of 8 mg mLꢁ1 Polybrene (Sigma-Aldrich).
Subsequently clones were selected by puromycin. All cells
were maintained in a 5% CO2 atmosphere at 37 ꢀC. SU.86.86
cells were grown in Roswell Park Memorial Institute (RPMI)
1640 Medium supplemented with 10% fetal bovine serum and
1% penicillin/streptomycin/antimycotic (complete growth
medium).
Fsample
¼
(Fstandard)(Isample/Istandard)(Astandard/Asample)(nsample2/nstandard
)
2
where F represents the uorescence quantum yield, I repre-
sents integrated uorescence intensity, A represents the absor-
bance at the specied wavelength, and n is the refractive index
of the solvent. All measurements were repeated in triplicate.
Molar extinction coefficient
3 (1.0 mg) was dissolved in 1 : 1 ACN/water (1.00 mL) to make
a stock solution. The slope of absorbance measurements of
Microscopy imaging
a series of dilutions of the stock solution was taken to determine All uorescence microscopy images and corresponding bright-
the molar extinction coefficient according to the Beer–Lambert eld images were obtained on an Olympus IX73 microscope. All
law. To determine the molar extinction coefficient of the probes, probe 2 cell images were obtained using a lter cube consisting
0.6 mL of probe was diluted in DPBS to a total volume of 60 mL of 380–420 nm excitation bandpass lter, a 484 nm dichroic
and an absorbance spectrum was obtained. 110 nM CES2 was mirror, and a 513–604 nm emission bandpass lter. Cell images
added and absorbance spectra were recorded in 10 min inter- of 3 were acquired using a lter cube comprising 503–557 nm
vals. The concentration of 3 upon the completion of the reaction excitation bandpass lter, a 580 nm dichroic mirror, and a 600–
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700 nm emission bandpass lter. All cells were plated in 8- tumors dissected from mouse subcutaneous tissue under sterile
chambered wells (Lab-Tek® Chambered #1.0 Borosilicate Cov- conditions. Approximately one-third of the tumor was prepared
erglass System) at a density of 5 ꢃ 104 cells per well one day for paraffin embedment, one-third was cut into quadrants and
before imaging. The cells were washed once with DPBS, and placed in liquid nitrogen for future study (F1), and other one-
were incubated in Opti-MEM with or without 100 mM LPA for third was injected into NOD/SCID mice again to generate
1 h. 20 mM probe 2 was then diluted into the appropriate wells additional generations (F2, F3) of direct xenogra tumors
and the cells were incubated for 3 h. The cells were washed ve applied in this study. For orthotopic xenogra models, a total of
times in DPBS and were imaged in DPBS. All images were 106 viable cells in 50 mL of complete growth medium with 50%
analysed using Olympus CellSens Dimension V1.18. H&E growth factor-reduced Matrigel were injected into the pancreas
images were acquired from a Nikon Eclipse 50i Microscope with of 4 to 6 week-old female nude mice; a total of ve mice were
a Nikon DS-Ri1 camera and NIS-Elements BR3.0 soware.
used for each experimental condition. The condition of mice
was monitored two times weekly over a period of 5 weeks. As the
mice tumor burden increased, mice were euthanized immedi-
ately upon becoming moribund, as required by our institutional
animal care guidelines. All tissues were harvested and ash
frozen with liquid nitrogen, then stored at ꢁ80 ꢀC until needed.
Using a cryotome, the harvested tissues were cut into 10 mm
thick slices and placed in glass microscope slides. These were
kept frozen until ready for the assessment of CES2 activity. All
slides were imaged before addition of probe 2 to determine
normal tissue uorescence. Slides were incubated with 100 mL
Cell viability assays
SU.86.86 Parental, IRES control vector and CES2 overexpressing
cells were each seeded at a density of 5000 cells per well in 48-
well plates (Thermo Scientic Nunclon™ Delta Surface) and
were incubated in supplemented RPMI 1640 Medium overnight
at 37 ꢀC in 5% CO2 atmosphere. The media was discarded; cells
were washed once with DPBS, replaced with solutions of probe 2
or 3 in complete growth media at different concentrations (0–20
mM), and were incubated for 3 h. The media was discarded; the
cells were washed once with DPBS, and were replaced with 250
mL of a 0.5 mg mLꢁ1 solution of 3-(4,5-dimethylthiazol-2-yl)-2,5-
diphenyltetrazolium bromide (MTT) in complete RPMI.
Following 2 h of incubation, the solution was removed and
replaced with 250 mL DMSO to dissolve the produced formazan.
Absorbance values were then read by a Tecan Innite M1000
ꢀ
of 20 mM probe 2 at 37 C for 3 h. Slides were then imaged for
red uorescence measurements. Ratiometric measurements
were taken aer washing three times with DPBS. H&E staining
was performed by established protocols.40
Statistical analysis
plate reader. Cell viability (CV) was determined by CV ¼ Asample
/
All data in bar graphs are expressed as mean ꢂ standard devi-
ation of triplicate measurements. Statistical differences were
determined using a Student's t-test via GraphPad Prism 7.00 for
Mac OS X (GraphPad Soware, La Jolla California, USA), with p <
0.05 considered statistically signicant. All box-and-whisker
plots of mean uorescence intensity were designed using R
Studio (version 3.4.4), with boxes indicating the rst and third
quartiles, and bolded horizontal lines in the boxes representing
the median value. Upper whiskers represent the maximum and
lower whiskers represent the minimum values. All cell and
tissue uorescence data have normal cell or tissue uorescence
for each channel (uorescence with no probe) subtracted
respectively. To determine statistical differences in median
values, Mann–Whitney–Wilcoxon rank sum test was performed
using R studio, with p < 0.05 considered statistically signicant.
Acontrol ꢃ 100%, where Asample and Acontrol are the absorbance
values of the experimental group (treated with probe or AP) and
control group (not treated with probe or AP) at 560 nm respec-
tively, with cell viability of the control group adjusted to 100%.
Western blot analysis
Cell pellets were lysed in RIPA buffer. Tumor tissues were cut
into small pieces and disrupted with a Dounce homogenizer in
ꢀ
RIPA buffer at 4 C for use in all procedures. The lysates were
then separated by SDS–PAGE and subjected to Western blotting.
The following antibodies were used: CES2 (Sigma-Aldrich,
HPA018897), and b-actin (Sigma-Aldrich).
Animal studies, tissue preparation and imaging
Orthotopic and patient-derived xenogras were provided by Dr
Samir Hanash and Dr Jason Fleming from MD Anderson Cancer
Center. Animal experiment protocols were reviewed and
approved by The University of Texas MD Anderson Cancer
Center IRB and in accordance with the Guidelines for the Care
and Use of Laboratory Animals published by the NIH (Bethesda,
MD). Heterotopic engrament of patient tumors into immu-
nodecient mice and expansion of direct xenogra tumors were
performed as previously described.38,39 Briey, excised patient
tumor tissues were obtained at surgery at The University of
Texas MD Anderson Cancer Center and mechanically minced
into fragments (<1 mm). Five tumor fragments were individu-
ally placed into formed tissue pockets. Once tumors reached
1.2 cm in greatest diameter, the mice were sacriced and the
Conflicts of interest
The authors declare no conicts of interest.
Acknowledgements
A. A. B. acknowledges support from Natural Sciences and
Engineering Research Council of Canada (NSERC Discovery).
Notes and references
1 S. C. Laizure, V. Herring, Z. Hu, K. Witbrodt and R. B. Parker,
Pharmacotherapy, 2013, 33, 210–222.
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Chem. Sci.

View Article Online
Chemical Science
Edge Article
2 D. Wang, L. Zou, Q. Jin, J. Hou, G. Ge and L. Yang, Human 22 Q. Jin, L. Feng, D. D. Wang, Z. R. Dai, P. Wang, L. W. Zou,
carboxylesterases: a comprehensive review, Acta Pharm. Sin.
B, 2018, 8, 699–712.
Z. H. Liu, J. Y. Wang, Y. Yu, G. B. Ge, J.-N. Cui and
L. Yang, ACS Appl. Mater. Interfaces, 2015, 7, 28474–28481.
3 S. E. Pratt, S. Durland-Busbice, R. L. Shepard, K. Heinz- 23 Z. M. Liu, L. Feng, G. B. Ge, X. Lv, J. Hou, Y. F. Cao, J. N. Cui
Taheny, P. W. Iversen and A. H. Dantzig, Clin. Cancer Res.,
2013, 19, 1159–1168.
4 L. de Sousa Cavalcante and G. Monteiro, Eur. J. Pharmacol.,
2014, 741, 8–16.
and L. Yang, Biosens. Bioelectron., 2014, 57, 30–35.
24 S. J. Park, H. W. Lee, H. R. Kim, C. Kang and H. M. Kim,
Chem. Sci., 2016, 7, 3703–3709.
25 L. Feng, Z.-M. Liu, J. Hou, X. Lv, J. Ning, G. B. Ge, J. N. Cui
and L. Yang, Biosens. Bioelectron., 2015, 65, 9–15.
5 B. L. Barthel, R. C. Torres, J. L. Hyatt, C. C. Edwards,
M. J. Hateld, P. M. Potter and T. H. Koch, J. Med. Chem., 26 T. Bezrodna, A. Negrviko, Y. Bezrodnyi and L. Kosyanchuk, J.
2008, 51, 298–304. Mol. Liq., 2018, 267, 89–95.
6 B. L. Barthel, Z. Zhang, D. L. Rudnicki, C. D. Coldren, 27 Y. N. Zhang, Y. A. Feng, Z. Li and X. S. Shao, Chin. Chem.
M. Polinkovsky, H. Sun, G. G. Koch, D. C. F. Chan and
T. H. Koch, J. Med. Chem., 2009, 52, 7678–7688.
7 V. Charasson, R. Bellott, D. Meynard, M. Longy, P. Gorry and
J. Robert, Clin. Pharmacol. Ther., 2004, 76, 528–535.
Lett., 2017, 28, 1228–1231.
´
˜
28 M. B. Freijo, A. Lopez-Arencibia, J. E. Pinero,
G. McNaughton-Smith and T. Abad-Grillo, Eur. J. Med.
Chem., 2018, 143, 1312–1324.
8 K. Uchida, K. Otake, K. Tanaka, K. Hashimoto, S. Saigusa, 29 G. Bousquet, J. P. Feugeas, L. Ferreira, L. Vercellino,
K. Matsushita, Y. Koike, M. Inoue, M. Ueeda, Y. Okugawa,
Y. Inoue, Y. Mohri and M. Kusunoki, J. Pediatr. Surg., 2013,
48, 502–509.
9 C. Shaojun, H. Li, H. Haixin and L. Guisheng, Cancer Biol.
Ther., 2018, 19, 153–159.
N. Jourdan, P. Bertheau, C. de Bazelaire, E. Barranger and
A. Janin, Breast Cancer Res., 2014, 16, 401.
30 M. Hidalgo, E. Bruckheimer, N. V. Rajeshkumar, I. Garrido-
Laguna, E. De Oliviera, B. Rubio-Viqueira, S. Strawn,
M. J. Wick, J. Martell and D. Sidransky, Mol. Cancer Ther.,
2011, 10, 1311–1316.
10 K. Ohtsuka, S. Inoue, M. Kameyama, A. Kanetoshi,
˜
T. Fujimoto, K. Takaoka, Y. Araya and A. Shida, Lung 31 M. P. Morelli, E. Calvo, E. Ordonez, M. J. Wick,
Cancer, 2003, 41, 187–198.
B. R. Viqueira, P. P. Lopez-Casas, E. Bruckheimer,
A. Calles-Blanco, D. Sidransky and M. Hidalgo, J. Clin.
Oncol., 2012, 30, e45–e48.
11 M. Capello, M. Lee, H. Wang, L. Babel, M. H. Katz,
J. B. Fleming, A. Maitra, H. Wang, W. Tian, A. Taguchi and
S. M. Hanash, J. Natl. Cancer Inst., 2015, 107, djv132.
12 M. K. Ross, A. Boraziani, R. Wang, J. A. Crow and S. Xie, Arch.
Biochem. Biophys., 2012, 522, 44–56.
32 K. Yoshioka, T. Komatsu, A. Nakada, J. Onagi, Y. Kuriki,
M. Kawaguchi, T. Terai, T. Ueno, K. Hanaoka, T. Nagano
and Y. Urano, J. Am. Chem. Soc., 2015, 137, 12187–12190.
13 Y. Sato, A. Miyashita, T. Iwatsubo and T. Usui, Drug Metab. 33 Y. Wu, S. Huang, F. Zeng, J. Wang, C. Yu, J. Huang, H. Xie
Dispos., 2012, 40, 1389–1396. and S. Wu, Chem. Commun., 2015, 51, 12791–12794.
14 S. K. Quinney, S. P. Sanghani, W. I. Davis, T. D. Hurley, 34 Y. Zhang, W. Chen, D. Feng, W. Shi, X. Li and H. Ma, Analyst,
Z. Sun, D. J. Murry and W. F. Bosron, J. Pharmacol. Exp.
Ther., 2005, 313, 1011–1016.
15 H. J. Zhu, D. I. Appel, Y. Jiang and J. S. Markowitz, Drug
Metab. Dispos., 2009, 37, 1819–1825.
2012, 137, 716–721.
35 D. Roife, B. Dai, Y. Kang, M. V. Rios Perez, M. Pratt, X. Li and
J. B. Fleming, Clin. Cancer Res., 2016, 22, 6021–6030.
36 A. M. Brouwer, Pure Appl. Chem., 2011, 83, 2213–2228.
16 J. Wang, E. T. Williams, J. Bourgea, Y. N. Wong and 37 K. Rurack, in Standardization and Quality Assurance in
C. J. Patten, Drug Metab. Dispos., 2011, 39, 1329–1333.
17 J. M. Clarke, M. R. Gillings, N. Altavilla and A. J. Beattie, J.
Microbiol. Methods, 2001, 46, 261–267.
18 L. Feng, Z.-M. Liu, L. Xu, J. Ning, J. Hou, G.-B. Ge, J.-N. Cui
and L. Yang, Chem. Commun., 2014, 50, 14519–14522.
Fluorescence Measurements, ed. U. Resch-Genger, Springer-
Verlag, Berlin Heidelburg, Germany, 2008, pp. 101–146.
38 M. P. Kim, D. B. Evans, H. Wang, J. L. Abbruzzese,
J. B. Fleming and G. E. Gallick, Nat. Protoc., 2009, 4, 1670–
1680.
19 Q. Jin, L. Feng, D.-D. Wang, J.-J. Wu, J. Hou, Z.-R. Dai, 39 M. P. Kim, M. J. Truty, W. Choi, Y. Kang, X. Chopin-Lally,
S.-G. Sun, J.-Y. Wang, G.-B. Ge, J.-N. Cui and L. Yang,
Biosens. Bioelectron., 2016, 83, 193–199.
20 Z. M. Liu, L. Feng, J. Hou, X. Lv, J. Ning, G. B. Ge,
G. E. Gallick, H. Wang, D. J. McConkey, R. Hwang,
C. Logsdon, J. Abbruzzesse and J. B. Fleming, Ann. Surg.
Oncol., 2012, 19, S395–S403.
K. W. Wang, J. N. Cui and L. Yang, Sens. Actuators, B, 2014, 40 A. H. Fischer, K. A. Jacobson, J. Rose and R. Zeller, Cold
205, 151–157.
Spring Harb. Protoc., 2008, prot4986.
21 S. J. Park, Y. J. Kim, J. S. Kang, L. Y. Kim, K. S. Choi and
H. M. Kim, Anal. Chem., 2018, 90, 9465–9471.
Chem. Sci.
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