Rate of Lipid Peroxyl Radical Production during Cellular Homeostasis Unraveled via Fluorescence Imaging

Article abstract of DOI:10.1021/jacs.7b08036

Reactive oxygen species (ROS) and their associated byproducts have been traditionally associated with a range of pathologies. It is now believed, however, that at basal levels these molecules also have a beneficial cellular function in the form of cell signaling and redox regulation. Critical to elucidating their physiological role is the opportunity to visualize and quantify the production of ROS with spatiotemporal accuracy. Armed with a newly developed, extremely sensitive fluorogenic α-tocopherol analogue (H₄BPMHC), we report herein the observation of steady concentrations of lipid peroxyl radicals produced in live cell imaging conditions. Imaging studies with H₄BPMHC indicate that the rate of production of lipid peroxyl radicals in HeLa cells under basal conditions is 33 nM/h within the cell. Our work further provides indisputable evidence on the antioxidant role of Vitamin E, as lipid peroxidation was suppressed in HeLa cells both under basal conditions and in the presence of Haber-Weiss chemistry, generated by the presence of cumyl hydroperoxide and Cu²⁺ in solution, when supplemented with the α-tocopherol surrogate, PMHC (2,2,5,7,8-pentamethyl-6-hydroxy-chromanol, an α-tocopherol analogue lacking the phytyl tail). H₄BPMHC has the sensitivity needed to detect trace changes in oxidative status within the lipid membrane, underscoring the opportunity to illuminate the physiological relevance of lipid peroxyl radical production during cell homeostasis and disease.

Full text of DOI:10.1021/jacs.7b08036

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Article
Rate of Lipid Peroxyl Radical Production During Cellular
Homeostasis Unravelled Via Fluorescence Imaging
Lana E. Greene, Richard Lincoln, and Gonzalo Cosa
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/jacs.7b08036 • Publication Date (Web): 18 Oct 2017
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Rate of Lipid Peroxyl Radical Production During Cellular Homeosta-
sis Unravelled Via Fluorescence Imaging
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Lana E. Greene, Richard Lincoln, and Gonzalo Cosa
Department of Chemistry and Centre for Self Assembled Chemical Structures (CSACS/CRMAA), McGill University,
801 Sherbrooke Street West, Montreal, Quebec H3A 0B8, Canada.
ABSTRACT: Reactive oxygen species (ROS) and their associated byproducts have been traditionally associated with a range
of pathologies. It is now believed however, that at basal levels, these molecules also play a beneficial cellular function in the
form of cell signalling and redox regulation. Critical to elucidating their physiological role is the opportunity to visualize
and quantify the production of ROS with spatiotemporal accuracy. Armed with a newly developed, extremely sensitive
fluorogenic α-tocopherol analogue (H4BPMHC), we report herein the observation of steady concentrations of lipid peroxyl
radicals produced in live cell imaging conditions. Imaging studies with H4BPMHC indicate that the rate of production of
lipid peroxyl radicals in HeLa cells under basal conditions is 33 nM/h within the cell. Our work further provides indisputable
evidence on the antioxidant role of Vitamin E, as lipid peroxidation was supressed in HeLa cells both under basal conditions
and in the presence of Haber-Weiss chemistry -generated by the presence of cumyl hydroperoxide and Cu2+ in solution-
when supplemented with the α-tocopherol surrogate, PMHC (2,2,5,7,8-pentamethyl-6-hydroxy-chromanol, an α-tocoph-
erol analogue lacking the phytyl tail). H4BPMHC has the sensitivity needed to detect trace changes in oxidative status within
the lipid membrane, underscoring the opportunity to illuminate the physiological relevance of lipid peroxyl radical produc-
tion during cell homeostasis and disease.
The deleterious-beneficial duality of reactive oxygen
INTRODUCTION
species (ROS) is an equally intricate emerging hypothesis.
It is almost 100 years since Vitamin E was first discovered
by Evans and Bishop, who reported its role as a necessary
dietary factor.1 The antioxidant properties of Vitamin E
were recognized shortly after.2-3 Mechanistic studies in ho-
mogeneous solution and model lipid membranes, pio-
neered by Ingold and coworkers over 35 years ago, showed
that Vitamin E molecules, and in particular α-tocopherol,
act as scavengers of lipid peroxyl radicals effectively termi-
nating lipid chain autoxidation.4-6 Notwithstanding its
clear antioxidant role, the biological action of α-tocopherol
in vitro and in vivo has proven to be complex and at times,
disputed, interweaving the antioxidant role with a cell sig-
naling role.7
Indeed, overproduction of ROS has been linked to several
pathologies such as neurodegenerative diseases, inflamma-
tion, and cancer.10, 17-18 While it is evident that ROS can
damage lipids and compromise membrane integrity lead-
ing to disease, low levels of ROS have been shown to be
beneficial by enhancing the defense mechanism of cells,
promoting the expression of antioxidants, proteins, and
enzymes.19-21
We postulate that as part of cellular homeostasis, a
steady state concentration of lipid peroxyl radicals is pro-
duced. In this regard, the ability to observe lipid peroxyl
radical formation under standard physiological conditions
would enable exploring the spatiotemporal nature and
quantity of ROS production, and its physiological and po-
tentially beneficial role. It would further enable visualizing
the failure in keeping ROS homeostasis in disease and un-
der cellular pathological conditions. Monitoring the gener-
ation of lipid peroxyl radicals in both healthy and patho-
logical conditions would ultimately provide a way to un-
derpin the biological mechanisms in the generation and
elimination of lipid peroxyl radicals and the associated
physiological implications.
α-Tocopherol is a chain-breaking antioxidant with ben-
eficial roles in treating diseases associated with oxidative
stress such as non-alcoholic fatty liver diseases,8 athero-
sclerosis,9 ataxia and other neurodegenerative diseases.10-11
There is, however, a significant body of work supporting
non-antioxidant roles for α-tocopherol in vivo such as gene
regulation.12-14 Whether these roles are truly non-antioxi-
dant is up for debate, as oxidative stress and formation of
lipid peroxyl radicals might also account for the so called
“non antioxidant roles”.7, 15 Further complexities arise when
considering the potentially detrimental or pro-oxidant ef-
fects of α-tocopherol.16
The study of both ROS and α-tocopherol in vitro and in
vivo is a difficult pursuit. While metabolite and product
studies may be useful for studying many biological phe-
nomena, they may not provide information on the kinetics
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of lipid peroxidation and its inhibition by α-tocopherol Overall, a major milestone has been achieved, the sensi-
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(temporal resolution), nor do they provide information on
the location of ROS formation and build-up (spatial reso-
lution). In particular, the regeneration of α-tocopherol
from the tocopheroxyl radical by water soluble antioxi-
dants (a key characteristic of its potent antioxidant
power),22 further complicates the utility of product studies,
and has in part imperiled the acceptance that the α-to-
copherol mechanism of action in vivo rests on its lipid sol-
uble antioxidant properties.
tivity needed to detect very small changes in oxidative sta-
tus within the lipid membrane, underscoring the oppor-
tunity to illuminate the physiological relevance of lipid
peroxyl radical production during cell homeostasis and
disease.
RESULTS AND DISCUSSION
H4BPMHC Preparation
9
To tackle the challenge of rapidly trapping peroxyl radi-
cals with minimum amounts of a chemoselective fluoro-
genic probe under conditions that trace amounts of reac-
tion could be detected from the background; both the ki-
netics of peroxyl radical trapping and the sensitivity of the
probe had to be optimum.
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In order to monitor the production of lipid peroxyl radi-
cals within lipid membranes in real-time and in a non-dis-
ruptive manner, we have developed fluorogenic α-tocoph-
erol analogues.23-27 These sensitive probes preserve the
chromanol moiety of α-tocopherol, and bear a boron di-
pyrromethene (BODIPY) lipophilic tail, mimicking the re-
activity of α-tocopherol while undergoing emission en-
hancement upon reaction. The probes provide a sensitive
marker of lipid peroxyl radical production under chemical
stress, amenable to fluorescence high throughput and im-
aging studies.25-27
Kinetics were optimized by choosing the chromanol
moiety of α-tocopherol as a trap segment, as with our pre-
viously designed fluorogenic probes. α-Tocopherol is the
most potent naturally occurring lipid soluble antioxidant6
and the choice of its chromanol head ensured rapid peroxyl
radical trapping. Tethering a BODIPY dye to the chroma-
nol head both provided a fluorescent reporting element
but also ensured that the probes retained the membrane
partition characteristic of α-tocopherol.24, 26-27
Currently lacking is the necessary sensitivity to monitor
the production of lipid peroxyl radicals in live cells under
basal conditions, devoid of chemical stressors such as me-
thyl viologen, Fe3+ or other sources of ROS. Fundamentally,
the problem at stake is detecting a highly reactive species
produced in trace amounts. This brings a conundrum in
the experimental design: a large amount of ROS scavenger
is needed to efficiently trap the short-lived ROS, yet the
scavenger concentration must be small to reliably detect
relative changes in consumed scavenger over time, signifi-
cant enough that the ensuing signal may be decoupled
from background.
Sensing in our previous designs exploited photoinduced
electron transfer (PeT)29 from the chromanol head to the
BODIPY core; where PeT ensured effective deactivation of
the probe. Upon scavenging of two lipid peroxyl radicals,
oxidation of the trap to a chromanone or chromoquinone
deactivated PeT, restoring the BODIPY emission (fluoro-
genicity).24, 26-27 Despite a number of attempts however, we
were not able to surpass ~80-fold and ~40-fold emission
enhancements in homogeneous solution and model lipid
membrane studies, respectively.26
Armed with a newly developed, extremely sensitive, flu-
orogenic α-tocopherol analogue, we report herein the ob-
servation of steady concentrations of lipid peroxyl radical
produced during cellular homeostasis. Our work relies on
a fluorogenic α-tocopherol analogue that undergoes a
~1,000-fold fluorescence enhancement upon scavenging
peroxyl radicals. Our work also addresses the influence of
serum albumins. These proteins may mask subtle changes
in ROS production by transporting α-tocopherol and its
oxidized product (and the fluorogenic α-tocopherol ana-
logue) in and out of the cell, smearing the trapping of lipid
peroxyl radicals by α-tocopherol analogues.
Optimum sensitivity was achieved in this work upon in-
creasing the efficiency of PeT via tuning the redox proper-
ties of the BODIPY core. The tuning was achieved by re-
moving the methyl groups at the 1 and 7 positions of the
BODIPY. This modification increased the redox potential
of the dye from -1.73 V (for the previously reported
H2BPMHC) to -1.41 V for the newly designed chromophore
(H4BPMHC, see Figure 1), as determined from electro-
chemical experiments (Supporting Information Figure S1).
To ensure membrane partitioning was not compromised,
the methyl groups at the 3 and 5 position of the BODIPY
core in H2BPMHC were swapped with propyl-chlorides in
H4BPMHC. These moieties additionally provide a reactive
handle for future modifications.
Our results indicate that the rate of production of lipid
peroxyl radicals in live cell imaging studies is at an estimate
of 33 nM/h within the cell. Given that cell culture condi-
tions are oxidatively stressing,28 this rate provides an upper
bound to the concentration of lipid peroxyl radicals pro-
duced during cellular homeostasis. Our work further pro-
vides indisputable evidence on the antioxidant role of Vit-
amin E, as lipid peroxidation was suppressed in cells both
under basal conditions and in the presence of Haber-Weiss
chemistry -generated by the presence of cumyl hydroper-
oxide and Cu2+ in solution- when supplemented with the
α-tocopherol surrogate PMHC (2,2,5,7,8-pentamethyl-6-
hydroxy-chromanol, an α-tocopherol analogue lacking the
phytyl tail).
By providing a highly exergonic electron transfer reac-
tion from the chromanol moiety to the BODIPY, we en-
sured that the fluorescence quantum yield of the BODIPY
dye in the new probe was extremely low, ɸf ≤ 0.001, within
detection error, prior to lipid peroxyl radical scavenging.
This represents a ~1,000-fold reduction in ɸf when com-
pared to the parent BODIPY chromophore H4BCH3, (the
BODIPY core in H4BPMHC) with a ɸf = 0.94. It also sets
the ceiling for fluorescence enhancement (and dynamic
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lowing our established protocol.26 In addition, the fluores-
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cent control, H4BCH3, was synthesized from 3 and acetyl
chloride.30
H4BPMHC Spectroscopic Properties
The absorption and fluorescence maxima, the extinction
coefficient (ε), the fluorescence decay lifetime (τdec) and
the fluorescence quantum yield (ɸf) for H4BPMHC, its ox-
idized form (H4BPMHCox, see details below for its genera-
31
tion and Scheme S1)26,
and the fluorescent control
9
H4BCH3 are listed in Table 1. Recorded absorption and flu-
orescence maxima (Figure S2) are in accordance with typi-
cal values for a BODIPY dye.30, 32 The very rapid decay of
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H4BPMHC precluded the measurement of a reliable τdec
.
Based on its low quantum yield, and the τrad value for the
-9
⁄
control compound, H4BCH3 (τrad= τdec ɸf ~6 × 10 s), we
predict τdec ~6 × 10-12 s for H4BPMHC. Similar τdec values
(~5.6 × 10-9) recorded for H4BPMHCox and its control com-
pound H4BCH3 in turn indicated that oxidation fully re-
stored the emissive properties of the BODIPY core, placing
the emission enhancement at 1,000-fold, upon oxidation.
The 1,000-fold enhancement is actually estimated from the
experimentally determined fluorescence emission quan-
tum yield of the probe before oxidation “Φ” , and that after
oxidation “Φ(ox)”. The latter is in turn determined from the
ratio of the fluorescence decay rate constant of the probe
after oxidation “kdecay ”, and the radiative decay rate
(ox)
constant of the BODIPY control utilized “krad”. In other
words, Φox = krad/kdecay . The value of krad is obtained
(ox)
from the control compound H4BCH3, and should not be
affected by the presence of the chromanol ring either in its
reduced or oxidized form. While the emission quantum
yield recorded for H4BPMHCox at 0.700 is short of the ex-
pected ~0.940 (the value recorded for H4BCH3), the dis-
crepancy may reflect a lack of full conversion to the oxi-
dized product and/or partial degradation of the BODIPY
core under the oxidation conditions utilized.
Figure 1. A) Structure of α-tocopherol and previously reported
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fluorogenic tocopherol analogue, H2BPMHC. B) Proposed
off/on sensing mechanism of the new fluorogenic probe,
H4BPMHC, relying on PeT.
range) of the new probe in going from reduced to oxidized
form at 1,000-fold (vide infra).
Synthesis of H4BPMHC required for new pyrroles to be
prepared, yet otherwise followed previously reported pro-
tocols26 involving the condensation of two equivalents of
the desired pyrroles with a chromanol acid chloride and
subsequent deprotection of the chromanol moiety, depro-
tonation of the resulting chromanol-dipyrrin, and chela-
tion with boron (see Scheme 1). The challenge in designing
pyrrole 3 rested on the necessity to preserve the alpha-po-
sition of the pyrrole free, for condensation, while also af-
fixing the reactive propyl chloride handle. Our synthesis of
3 comprised a set of 4 mild reactions that circumvented the
need for protections. Starting from pyrrole-2-carboxalde-
hyde, we first performed a Wittig reaction with methyl (tri-
phenylphosphoranylidene)acetate to add the two new car-
bons to form the propyl chain. The resulting alkenyl-ester,
1, was then reduced over two steps (dehydrogenation and
borane reduction) to give the desired alkyl-alcohol, fol-
lowed by the Appel reaction to give the propyl-halogen, 3.
While all steps are reasonably high yielding, caution had to
be followed in removing the solvent, as the pyrrole prod-
ucts are very volatile. With the new pyrrole in hand, our
desired probe, H4BPMHC was synthesized over 4 steps fol-
Table 1. Photophysical properties of the novel
BODIPY dyes presented herein.
Abs
Em
ε
τdec
ɸf
(ns)
λmax
(nm) (nm)
λmax
(cm-1M-
)
1
H4BPMHC
509
509
520
520
8.2 × 104 NA
0.001
a
H4BPMHCox
5.68 0.700 -
1.0b
H4BCH3
502
516
7.9 × 104 5.60 0.940
a The extinction coefficient is expected to be the same before
and after oxidation. bThe range reflects the experimental
measurement where BODIPY degradation post-activation
may have occurred, and an estimation based on krad and
kdecay
values.
(ox)
Reactivity of H4BPMHC with azo initiators
In order to test the reactivity and validate the sensitivity
of our new probe towards ROS, we subjected H4BPMHC
to peroxyl radicals generated at a steady state while moni-
toring the fluorescence intensity enhancement.
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Scheme 1. Synthesis of H4BPMHC
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The intensity-time trajectories were recorded in the ab-
sence and, when stated, presence of competing antioxi-
dants: either α-tocopherol or 2,2,5,7,8-pentamethyl-6-
chromanol (PMHC) to record the fluorescence enhance-
ment and the relative rate constants of H4BPMHC vs α-
tocopherol and PMHC. In our experiments, peroxyl radi-
cals were generated at a constant rate of Rg = 9 × 10−9 Ms−1
via thermolysis at 37 °C of a 200 µM solution of the radical
initiator, 2,2-azobis(4-methoxy-2,4-dimethyl valeronitrile)
(MeO-AMVN, see Scheme S1), in air-equilibrated acetoni-
trile.26, 31
The larger enhancements observed in samples supple-
mented a priori with additional antioxidant (PMHC or α-
tocopherol) are two-fold: 1) sacrificial scavenging of any
radicals initially present in the solvent by the added anti-
oxidant which would otherwise pre-activate the probe giv-
ing an artificially high initial fluorescence intensity and ac-
cordingly a reduced value for the enhancement24 and 2) the
inner filter effect due to the large absorbance of 10 µM
H4BPMHC.
The relative reactivity towards peroxyl radicals of
H4BPMHC
and
the
competing
antioxidants
A fluorescence enhancement of 400 to 700-fold was rec-
orded for H4BPMHC in the presence of peroxyl radicals
(Figure 2). This sensitivity is consistent with the deactiva-
tion of PeT following oxidation of the chromanol moiety.
(kinhH4BPMHC/kinhTOH, where TOH stands for α-tocopherol or
PMHC) was determined from a kinetic analysis based on
Equation 1,26 where I∞, It, and I0 are the intensity of
H4BPMHC at the maximum, time t, and at t = 0 and τ is
the time required to fully consume the antioxidant load.
The relative antioxidant activity determined for
H4BPMHC is on par with that of both α-tocopherol and
PMHC (Supporting Information, Figure S3). At about 90%
of the reactivity of the competing antioxidants, coupling
the chromanol segment to the BODIPY moiety thus did
not diminish the peroxyl radical trapping reactivity of the
former.
700
600
500
400
300
1M H4BPMHC + 9M -Toc
200
1M H4BPMHC + 9M PMHC
100
퐻4퐵푃푀퐻퐶
10M H4BPMHC
−퐼
−푙푛 (퐼∞ 푡) = − 푘
푙푛 (1 − 푡)
Equation 1
ꢀꢁℎ
푇푂퐻
퐼
∞
−퐼
0
푘
휏
0
ꢀꢁℎ
0
2000
4000
6000
8000
Influence of serum albumin in monitoring H4BPMHC re-
sponse
Time (s)
Prior to testing the sensitivity of H4BPMHC in live cells
under various levels of oxidative stress, and stimulated by
a recent report illustrating the solubility and increased flu-
orescence of BODIPY dyes in the presence of bovine serum
albumin (BSA)33 we investigated the effect of fetal bovine
serum (FBS) on the partitioning of our BODIPY fluoro-
phores into adherent cells. Here, partitioning is reflected
by the fluorescence of our control dye, H4BCH3, observed
in cells using fluorescence microscopy. This was a critical
Figure 2. Fluorescence-time trajectories of H4BPMHC (1 µM
or 10 µM as indicated) in the presence of the radical initiator
MeO-AMVN (200 µM) and supplemented with and without
antioxidants (PMHC and α-tocopherol, 9 µM). Solutions were
excited at 485 nm and emission was monitored at 520 nm. Flu-
orescence enhancement times, τ, were 2730 s, 2870 s, and 2460
s for H4BPMHC and tocopherol, H4BPMHC and PMHC, and
10 µM H4BPMHC respectively.
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Figure 3. Images of HeLa cells acquired over 30 minutes upon the addition of 100 nM H4BCH3 in LCS, DMEM or DMEM supple-
mented with 10% FBS. Images were acquired using widefield fluorescence microscopy with 20x magnification, λex = 488 nm (0.09
mW), emission was spectrally filtered with a ZET480/640m emission filter.
exercise to optimize staining conditions toward maximiz-
ing sensitivity.
fit to an exponential growth curve (Supporting Infor-
mation, Figure S5A). The dimmer intensity with FBS is thus
not the result of kinetic control in partitioning. We may
conclude that FBS, while facilitating the solubility of the
control dye, effectively prevented its partitioning into the
cells by providing a competing hydrophobic environment
for H4BCH3 to reside into. This is consistent with the re-
ports of BODIPY complexation by BSA.33
HeLa cervical cancer cells (ATCC CCL-2) on fibronectin
treated coverslips were used to compare the membrane
partitioning of our fluorescent control BODIPY, H4BCH3
(100 nM) when utilizing different imaging buffers (all con-
taining 0.33% DMSO vehicle) including Live Cell Imaging
Solution (LCS), and Dulbecco’s Modified Eagles Medium
(DMEM) supplemented with and without 10% FBS.
That FBS in solution competes with cellular membranes
for the lipophilic dye H4BCH3 (and presumably also for the
lipophilic fluorogenic probe H4BPMHC) was additionally
illustrated by a series of experiments where the staining so-
lution of H4BCH3 containing no FBS was removed, and ad-
dition of either fresh LCS, DMEM, or DMEM supple-
mented with 10% FBS took place. In the latter case, the
control dye was rapidly depleted from the cell membranes,
while only minor intensity reductions took place in the for-
mer two cases (no FBS added), see Figure 4. Final intensi-
ties of the solution containing 10% FBS were 50-fold lower
than when FBS was not added (Figure S4B). Intensity-time
trajectories of the partitioning out of H4BCH3 are shown in
Figure S5B and clearly indicate a rapid and significant drop
in intensity in the presence of FBS.
The partitioning was recorded using widefield fluores-
cence (WF) microscopy. Frames 100 ms in duration were
captured every minute over 30 minutes to minimize photo-
bleaching from competing with the fluorescence increase
due to partitioning of the dye into the cell.
Qualitatively, the presence of FBS resulted in a signifi-
cant drop in intensity as may be observed in Figure 3 where
images acquired at the same time intervals in each of the
three buffers are displayed. Consistent with this observa-
tion, the corrected total cell fluorescence (CTCF) of the im-
ages acquired after 30 minutes for samples without FBS are
similar (Suplementary Information, Figure S4A), while the
CTCF of the sample containing FBS is 5 to 7-fold lower, in-
dicating less dye partitioned into the cell in the presence of
FBS. In all three buffers, staining of various organelle lipid
membranes of the cell was clearly observed as is typical of
the lipophilic BODIPY scaffold.34-36 Importantly, partition-
ing rate constants in all buffers were very similar (0.15-0.18
min-1), taking about 20 minutes for the control dye to par-
tition fully, as shown from the intensity-time trajectories
Previous imaging studies in DMEM supplemented with
FBS precluded us from using dye concentrations below 1
µM. Lengthy incubation times (e.g > 30 minutes) were ad-
ditionally required. By utilizing buffers free from FBS, we
reduce the incubation time thus preventing masking reac-
tions occurring during the incubation period. Additionally,
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Figure 4. Images of HeLa cells acquired over 30 minutes upon the replacement of 100 nM H4BCH3 in LCS, with fresh media (LCS,
DMEM or DMEM supplemented with 10% FBS). Images were acquired using widefield fluorescence microscopy with 20x magni-
fication, λex = 488 nm (0.09 mW), emission was spectrally filtered with a ZET480/640m emission filter.
utilizing buffers free from FBS enables us to reduce the
probe concentration, by at least one order of magnitude to
100 nM. This is key when small concentration of analytes,
such as lipid peroxyl radicals in this work (and concomi-
tantly small amounts of probe) are expected to be pro-
duced (to react). Working with minimal amounts of the
fluorogenic probe H4BPMHC ensures maximizing signal
to background, enabling us to observe fluorogenic antiox-
idant consumption in real-time upon production of trace
amounts of peroxyl radicals (vide infra).
antioxidant status both under Haber-Weiss chemistry and
in the absence of chemical stressors (see Figure 5).
HeLa cells were stained with a 100 nM solution of
H4BPMHC for 10 min, washed and the buffer replenished.
A combination of cumene hydroperoxide (100 µM) and
copper(II) sulfate (10 µM) was used as a source of ROS un-
der Haber-Weiss conditions. Cumene hydroperoxide re-
acts with copper via the Haber-Weiss reaction to produce
cumoxyl and cumoperoxyl radicals which may in turn lead
to lipid peroxidation and the production of lipid peroxyl
radicals.37 Additives, consisting of 1 µM and 10 µM solutions
of PMHC pre-incubated for 30 min followed by cell stain-
ing for 10 min, and washing, were also studied with and
without Haber-Weiss reagents. While one may argue that
α-tocopherol should be the preferred choice, both PMHC
and the latter have the same radical trapping reactivity, yet
α-tocopherol has poor solubility and cell penetration.
Thus, we use PMHC in our work, as a source of lipid solu-
ble antioxidant. Unstained cells were imaged to determine
the cell autofluorescence at the laser power and gain set-
tings used.
Ultimately, incubation of H4BPMHC and H4BCH3 (100
nM) for 10 min in LCS in the absence of FBS provided opti-
mal conditions in terms of signal to background. LCS was
chosen as the incubation buffer for all subsequent experi-
ments due to its low background fluorescence, and the rel-
atively good cell viability when compared to unsupple-
mented DMEM.
Rate of lipid peroxyl radicals generated during cellular
homeostasis and under Haber-Weiss conditions
Armed with newly developed H4BPMHC and working
with optimized imaging conditions, we next sought to ex-
plore the antioxidant status of live cells both when exposed
to high levels of oxidative stress in the form of lipid peroxyl
radicals generated via Haber-Weiss chemistry, but also un-
der conditions of cellular homeostasis (no chemical stress-
ors). We also explored how the chromanol compound
PMHC, related to α-tocpherol in Vitamin E, affected the
H4BPMHC underwent a 30-fold enhancement in the
CTCF value in the presence of the Haber-Weiss reagents,
computed by comparing the sample before and after 50-
minute incubation with the reagents (see Supporting In-
formation section, Figure S6). A prominent yet smaller flu-
orescence enhancement of 24-fold and 10-fold was in turn
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Figure 5. A) Images of HeLa cells in the presence of 100 nM H4BPMHC (Probe) with/without PMHC acquired after 50 minutes
of treatment with (stressed) or without (healthy) cumene hydroperoxide (100 µM) and copper(II) sulfate (10 µM). Images were
acquired using widefield microscopy with 20x magnification, λex = 488 nm (0.09 mW), emission was spectrally filtered with a
ZET480/640m emission filter. Inset shows compressed DIC images for the same region. B) Corrected total cell fluorescence vs
time determined from images acquired over a 45 minute period with 100 nM H4BPMHC in healthy cells and cells exposed to
Haber-Weiss reagents.
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recorded after 50 minutes for samples also stressed with
the Haber-Weiss reagents, yet supplemented with 1 µM
and 10 µM PMHC respectively when compared to time zero
(see Figure S6). Importantly, enhancements in the absence
of Haber-Weiss reagents were smaller than 3-fold (vide in-
fra). Further, no change in fluorescence was observed for
our control dye, H4BCH3 in the presence of either PMHC
or Haber-Weiss reagents (Figure 6B). Close inspection of
images acquired with 100× magnification following treat-
ment with Haber-Weiss reagents for 50 minutes (see Sup-
porting Information section, Figure S7) reveals that
H4BPMHC is localized in the various cell lipid membranes
consistent with its lipophilic nature. A similar distribution
is also apparent from the dim images prior to addition of
Haber-Weiss reagents
(and correspondingly, conversions) of 13%, 12% and 5.5%,
respectively, relative to the intensities recorded with
H4BCH3. PMHC thus exerted a concentration dependent
protective effect.
CTCF-time trajectories were next retrieved from the data
set by plotting the CTCF evolution with time for all condi-
tions explored (Figure 5B). For cells bearing the Haber-
Weiss reagents and no PMHC, the initial intensity in-
creased in a linear fashion (reaching a 30-fold increase
within 50 minutes). In contrast, for cells bearing the Ha-
ber-Weiss reagents and 1 µM PMHC, an induction period
in fluorescence intensity was observed followed by a linear
increase with a similar rate to that observed for the sample
with no antioxidant additive. The observed induction pe-
riod is a hallmark of inhibition of oxidation of H4BPMHC
by PMHC, under competitive kinetic conditions. The in-
duction period was prominent (~10 min) when samples
were supplemented with 10 µM PMHC, yet the intensity af-
ter the induction increased at a smaller rate than when no
antioxidant was present indicating that inhibition was fol-
lowed by retardation in the oxidation of H4BPMHC.
To provide a benchmark value for how much peroxyl
radical was trapped by H4BPMHC after 50 min incubation
with Haber-Weiss reagents (both with and without addi-
tive PMHC), we compared CTCF values for cells stained
with H4BPMHC vs H4BCH3. Under the assumption that
both probes partition equally well within cells (i.e. their
concentrations are similar), and that the fluorescence life-
time of H4BPMHCox in membranes is on par with that of
H4BCH3, the CTCF recorded with H4BCH3 would corre-
spond to 100% conversion of H4BPMHC to its oxidized,
fully emissive, form upon trapping of two peroxyl radicals.
Figure 6 shows CTCF values normalized by frame exposure
time and obtained under otherwise identical conditions.
Samples containing 100 nM H4BPMHC and supplemented
with 0, 1 µM and 10 µM PMHC achieved final intensities
Results of CTCF evolution with time underscore that
PMHC, within the cellular milieu, is outcompeting
H4BPMHC in scavenging ROS produced by cumene hy-
droperoxide and copper. Qualitatively, similar results (in-
duction periods) were previously observed in liposomes ex-
posed to a source of peroxyl radicals and stained with a less
sensitive, yet equally reactive and lipophilic analogue of
H4BPMHC.26 Our current work shows that PMHC, lacking
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Figure 6. CTCF of H BPMHC (A) and H BCH (B) in HeLa cells. Intensities were determined after 50 minutes of treatment
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3
with/without stressor. Dye = 100 nM in either H BPMHC or H BCH , stressor = 100 μM cumene hydroperoxide and 10 μM cop-
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per(II) sulfate. Images were acquired using widefield microscopy with 20x magnification, λex = 488 nm (0.09 mW), emission was
spectrally filtered with a ZET480/640m emission filter. CTCF of B was corrected by the exposure time used to match that of A.
Error bars shown are standard deviations from the CTCF of 3 different regions of cells.
a lipophilic tail, and thus provided with higher mobility
within the cell membrane lipid milieu, is more effective in
scavenging radicals at the cellular level than H4BPMHC -
and presumably α-tocopherol as H4BPMHC and the latter
are expected to have similar reactivities within the mem-
Alternatively, partitioning of PMHC within the cell is re-
duced compared to the probe, where upon incubation of
both PMHC and H4BPMHC, and following washing of the
solution and refilling with fresh LCS buffer, PMHC would
more readily escape cellular confinement given the lack of
a lipophilic tail.
brane.26 While such an outcome had been described with
model lipid membranes22,
it is truly remarkable that
26
CTCF intensity-time trajectories acquired with 100 nM
H4BPMHC and no Haber-Weiss reagents in turn shed
light on basal levels of lipid peroxyl radicals produced dur-
ing cellular homeostasis. In healthy cells, there was a nota-
ble gradual fluorescence increase with changes in CTCF
values of 1.00 x 105 CTCF and 0.75 x 105 CTCF over 50
minutes for the samples supplemented with either no or 1
µM PMHC, and for the sample with 10 µM PMHC respec-
tively. In the context of the benchmark, emission intensity
recorded under identical conditions for 100 nM H4BCH3, at
a value of 1.00 × 107 CTCF, we may estimate that the
amount of peroxyl radicals produced under basal condi-
tions over 50 min resulted in at most 1% of the probe being
oxidized. If we approximate the concentration of dye in-
side a cell at 1.7 µM (see Supporting Information for discus-
sion), we may place the rate of production of lipid peroxyl
radicals under basal conditions at 33 nM/h within a cell (2
equivalents of ROS required to oxidize the probe). We note
that different cell lines, imaging conditions etc., would
yield different rates of lipid peroxyl radical generation, in
that sense, it was the aim of this work to report what are
typically considered as standard imaging conditions.
within the complex cell environment it still holds. It is in-
deed the case that should PMHC outcompete the probe in
scavenging peroxyl radicals, or in turn, regenerate the
probe giving in turn PMHC phenoxyl (chromanoxyl) radi-
cal, one would observe the same induction period. Here
PMHC would be the final sink of radical character, regard-
less whether it donates its H-atom directly to a peroxyl rad-
ical or indirectly through the probe.38-39
An intriguing aspect, defying explanation, is the con-
trasting times required to oxidize H4BPMHC, and compar-
atively, to consume the antioxidant additive PMHC. The
time evolution of CTCF with Haber-Weiss reagents reveals
that while 50 min were required to oxidize ~10% of the
probe, ~5 min elapsed in the induction period with PMHC,
for experiments where probe and antioxidant were incu-
bated at 100 nM and 1 µM, respectively. During the induc-
tion period, we postulate, all PMHC within the cell was de-
pleted leading next to a temporal evolution characteristic
of no PMHC. Should H4BPMHC and PMHC have similar
partitions within the cell lipid milieu, these numbers
would indicate that 100-fold more PMHC reacts roughly in
one tenth of the time than H4BPMHC. This large differ-
ence conflicts with the 10-fold larger reactivity toward per-
oxyl radicals recorded for PMHC vs both an analogue of
H4BPMHC and α-tocopherol, when in lipid membranes.26
Regarding the smaller slope recorded for cells whose me-
dia was supplemented with 10 µM PMHC, we may postu-
late that excess PMHC suppressed the oxidation of
H4BPMHC in healthy cells. In line with PMHC protection,
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analysis of the fluorescence intensity also shows a noticea- treated with an onslaught of ROS, but more importantly
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ble difference in fluorescence between healthy samples
pre-supplemented with PMHC vs those supplemented
with no PMHC, about 2-fold larger for the latter, Figure 6A.
Presumably some initial oxidation of H4BPMHC may oc-
cur while incubation, washing and exploration of the
chambers prior to image acquisition, where oxidation
would be enhanced when no PMHC is present.
allowed us to detect lipid peroxidation at basal levels and
in the presence of additional antioxidant. Importantly, we
showed indisputable evidence of the antioxidant role of
Vitamin E in vitro as lipid peroxidation was suppressed in
the presence of additional PMHC in HeLa cells under du-
ress, and under normal cell culture conditions. The latter
indicates potential complications that can arise from using
cultured cells in experiments, especially when looking at
the production of ROS. These experiments are typically
conducted in the absence of antioxidants and the outcome
of lipid peroxyl radical formation, we believe, should be
considered in interpreting results.
In closing, we may compile the above results to establish
the sensitivity of H4BPMHC within cells. Overall, a 40-fold
fluorescence enhancement was observed between healthy
cells pre-supplemented with PMHC, and cells exposed to
Haber-Weiss reagents for 50 minutes without additional
antioxidant. Considering that in the latter case there was
only 13% conversion, a maximum enhancement would be
placed at ~300-fold. The discrepancy with the expected
1,000-fold enhancement may be rationalized in terms of
pre-oxidation in the dark periods of the experiment. We
note that should a 0.3% fraction of our pristine solution of
H4BPMHC undergo oxidation, and considering the en-
hancement is ~1,000-fold in going from reduced to
H4BPMHCox , such a small conversion would account for
an initial intensity 3-fold larger than that of pristine solu-
tion, and in turn a 1,000/3 or ~ 300 value for the observed
intensity enhancement. Care should be taken in avoiding
pre-oxidation, toward a quantitative analysis.
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In closing, we note that while the chemistry of α-tocoph-
erol and lipid peroxyl radicals has been studied extensively
in the last 35 years, their complex relationship in cellular
mechanisms remains very much unclear. We strongly be-
lieve that imaging studies with sensitive fluorogenic probes
capable of detecting subtle levels of lipid peroxidation,
while also mimicking the reactivity of tocopherol, like
H4BPMHC reported herein, will ultimately help unravel
the physiological role of both lipid peroxyl radicals and α-
tocopherol in vivo.
EXPERIMENTAL
Materials
HPLC grade solvents for spectroscopy and column chro-
matography were purchased through Fisher Scientific.
MeO-AMVN was purchased from WAKO Pure Chemicals
Ltd. Buffers, and growth media and other reagents for cell
culture and staining were purchased through Ther-
moFisher Scientific. All other chemicals were supplied by
Sigma-Aldrich, Co. and used without further purification.
CONCLUSION
Albeit lipid peroxyl radicals have long been known to be
formed at the cellular level,20, 40 their elusive nature has
prevented quantifying this highly reactive intermediate.
Their physiological role has thus remained obscured, and
that of α-tocopherol, its par excellence scavenger, contro-
versial as an antioxidant.
Instrumentation
By developing the highly sensitive fluorogenic α-tocoph-
erol analogue, H4BPMHC, we managed to overcome a ma-
jor limitation in detecting trace amounts of lipid peroxyl
radicals, that is, that the initial signal is small enough that
trace amounts of reaction can be monitored. The high im-
aging sensitivity we achieved is both the result of optimiz-
ing the construct, ensuring very rapid PeT takes place, am-
ply outcompeting emission by about 1,000-fold, and re-
moving serum albumins, proteins that will otherwise
scramble the distribution of lipophilic molecules including
BODIPY dyes in general, and H4BPMHC in particular.
Thus, serum albumins may replenish fresh probe and re-
move oxidized emissive probe, ultimately masking lipid
peroxidation. The latter results provide a note of caution
when using lipophilic dyes in live cell imaging in the pres-
ence of FBS/BSA. Our work ultimately calls for the avoid-
ance of FBS for cellular imaging of BODIPY fluorophores
(and likely other lipophilic small molecule fluorophores) to
gain better signal to background at low (nanomolar or sub-
micromolar) concentrations.
Absorption spectra were recorded using a Hitachi U-
2800 UV−vis−NIR spectrophotometer. Luminescence
spectra were recorded using a PTI QuantaMaster spectro-
fluorimeter using 1 cm × 1 cm quartz cuvettes. Spectra were
1
13
corrected for detector sensitivity. H NMR and C NMR
spectra were recorded on a Varian VNMRS 500, Varian
VNMRS 300 or a Bruker AVIIIHD 500. Electrospray ioniza-
tion (ESI) and atmospheric chemical ionization (APCI)
mass spectra were measured on a Bruker maXis impact.
Time-based fluorescence enhancements with radical in-
itiator
Time based fluorescent measurements were recorded on
a PTI QuantaMaster spectrofluorimeter equipped with a
four-position Peltier with motorized turret. 3 mL glass flu-
orimetry cells with 1 cm path lengths were used as sample
cells. The excitation and emission slits were set to 1.2 nm.
The temperature control was set to 37oC. Three samples
containing 10 µM antioxidant load were prepared in ace-
tonitrile (MeCN): 1) 10 µM H4BPMHC 2) 1 µM H4BPMHC
and 9 µM α-tocopherol, and 3) 1 µM H4BPMHC and 9 µM
PMHC. Samples were excited at 485 nm and emission was
collected at 520 nm. Initial fluorescence of the samples was
Our results in live cells underscore the potential of
H4BPMHC to investigate the physiological relevance of li-
pid peroxyl radical production during cell homeostasis and
disease. The resulting large dynamic range of H4BPMHC
allowed us to not only discern healthy cells from cells
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measured, and samples were then spiked with 200 µM rad- of the objectives where the beam was coupled into the mi-
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ical initiator, MeO-AMVN (2,2-azobis(4-methoxy-2,4-di-
methyl valeronitrile)). Fluorescence was then recorded
over time until the fluorescence intensity of all samples
reached a maximum plateau.
croscope objective using a multiband beam splitter
(ZT488/640rpc, Chroma Technology). The emission was
spectrally filtered with an emission filter (ZET488/640m).
For multi-channel imaging between the 488 channel and
DIC, a motorized filter block turret was used. The fluores-
cence emission was collected through the same objective
and captured on a back illuminated electron multiplying
charge coupled device (EM-CCD) camera (Andor iXon Ul-
tra DU-897).
Fluorescence quantum yield
Acetonitrile solutions of meso-acetoxymethyl-BODIPY
dye30 (H2BOAc, ɸf=0.87), or 8-phenyl pyrromethene
fluoroborate (H6BPh)41 (ɸf = 0.013) (structures in Figure S9)
were used as standards to calculate the quantum yields of
solutions of oxidized H4BPMHC and H4BCH3 or reduced
H4BPMHC respectively in MeCN. Solutions of H4BPMHC
were also supplemented with 5 µM PMHC to help to pre-
vent any unwanted oxidation. Absorption and emission
spectra of the dye standards and dyes of interest were
measured at five different concentrations. 200 µM MeO-
AMVN, a radical initiator, was added to H4BPMHC to ob-
tain H4BPMHCox where the absorption and emission spec-
tra were then measured once the maximum fluorescence
intensity was reached. The integrated intensity versus ab-
sorbance were then plotted and fitted linearly. Quantum
yields for the dyes with respect to the standard were ob-
tained from Equation 2, where Φx, Δ, and η refer, respec-
tively, to the fluorescence quantum yield, the slope ob-
tained from the above-mentioned plot, and the solvent re-
fractive index for the unknown (x) or standard (st) sample.
Emission spectra were recorded for solutions using excita-
tion and emission slits of 1.2 nm for solutions of H2BOAc,
H4BPMHCox and H4BCH3, and 4 nm for solutions of
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Cell culture
HeLa cells (ATCC CCL-2) were cultured in Dulbecco’s
Modified Eagle Medium (DMEM) containing high glucose,
L-glutamine, phenol red, and sodium pyruvate (Gibco),
supplemented with 10% fetal bovine serum (FBS) and 1%
penicillin-streptomycin. Cells were maintained at 37 °C
(5% CO2) in a humidified atmosphere and were trypsinized
and split 1/15 about twice a week when confluence of cells
was reached. Cells were passaged a maximum of 30 times
and tested for mycoplasma infection upon first passaging.
Cell imaging of H4BCH3
24 h prior to day of imaging, cells were plated on 35 mm
glass imaging dishes (World Precision Instruments, Inc.)
pre-coated with fibronectin (1 μg/cm2), in DMEM contain-
ing growth factors. After 24 h, cells were then washed twice
with incubation buffer (LCS, DMEM or DMEM/10%FBS),
followed by the addition of 1 mL of fresh incubation buffer.
Cells were stained by the addition of 500 µL of the dye (300
nM) to give a final concentration of 100 nM in dye. The so-
lution of dye was prepared from the addition of 5 µL of a
30 µM stock solution in DMSO. This overall 300-fold dilu-
tion was to ensure the percentage of DMSO in media was
0.33% (v/v). Images of the samples were then collected
every minute (100 ms exposure time) over the course of 30
minutes.
H6BPh and H4BPMHC.
2
∆
푥
휙푥 = 휙푠푡
(
푥 ) × (휂
)
휂
푠푡
Equation 2
2
∆
푠푡
Fluorescence lifetime studies
The fluorescence lifetime measurements were carried
out using a Picoquant Fluotime 200 Time Correlated Single
Photon Counting setup employing a supercontiuum laser
(WhiteLase SC-400-4, Fianium, Beverly, MA). Excitation
wavelengths were spectrally separated from broadband
emission by a computer controlled acousto-optical tunable
filter (AOTF, Fianium). H4BPMHCox, and H4BCH3 were
excited at 485 nm, and emission was collected at 520 nm
and 516 nm respectively. The excitation rate was 10 MHz,
and the detection frequency was less than 100 kHz. Pho-
tons were collected at the magic angle.
Cell imaging of H4BPMHC or H4BCH3 in healthy and
stressed cells
24 h prior to imaging, 300 µL of 50 000 cells/mL were
plated into each well of an 8-well ibidi-Treat µ-slide in
DMEM containing growth factors. After 24 h, all wells were
washed twice with LCS, and either loaded with 200 µL LCS,
or 200 µL LCS containing either 1 µM PMHC or 10 µM
PMHC. After 30 minutes, wells to be stained were loaded
with 100 µL of a 300 nM stock of H4BPMHC or H4BCH3
bringing the final concentration to 100 nM in dye, and in-
cubated for 10 minutes. Wells were then washed with LCS
and loaded with either 300 µL of LCS (wells to be main-
tained as healthy) or 200 µL of LCS (wells to be stressed
later once slide is loaded onto the microscope). The slide
was then loaded onto the microscope, where once fixed in
place, 100 µL of LCS containing 300 µM cumene hydroper-
oxide and 30 µM copper(II) sulfate was added to the wells
to be stressed (100 µM and 10 µM final concentration of cu-
mene hydroperoxide and copper(II) sulfate respectively).
All wells were then imaged every 2 minutes (300 ms and 70
ms exposure for H4BPMHC and H4BCH3 respectively) over
the course of 50 minutes, where 3 single frames of each well
were then taken.
Microscopy
All fluorescence and differential interference contrast
(DIC) imaging of HeLa cells were performed using either a
wide-field objective-based total internal reflection fluores-
cence (TIRF) microscopy setup equipped with an oil-im-
mersion objective (Nikon CFI SR Apochromat TIRF 100x,
NA = 1.49) or a lower magnification wide-field microscopy
setup equipped with an air objective (Nikon CFI Plan Apo
VC 20x objective, NA = 0.75) consisting of an inverted mi-
croscope (Nikon Eclipse Ti) equipped with a Perfect Focus
System (PFS). A stage-top incubator (Tokai Hit) was used
to maintain the cells at 37 °C (5% CO2) in a humidified at-
mosphere. The excitation laser was 488 nm with powers of
0.09 mW (for 20x) and 0.04 mW (for 100x) measured out
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onto a silica flash column and eluted with 20% ethyl ace-
Image analysis
1
tate in hexanes affording the clean white solid product
All images were processed using the FIJI imaging pro-
cessing package.42 All quantifications were performed
adapting a previously published protocol.43 Equation 3 was
used to calculate the corrected total cell fluorescence
(CTCF):
1
2
3
4
5
(4.82 g, 97%). H-NMR (500 MHz; CDCl3): δ 9.56 (s, 1H),
7.64 (d, J = 15.9 Hz, 1H), 6.96 (q, J = 2.0 Hz, 1H), 6.60 (dd, J
= 4.7, 2.6 Hz, 1H), 6.31 (q, J = 3.0 Hz, 1H), 6.16 (d, J = 15.9 Hz,
1H), 3.81 (s, 3H). 13C-NMR (126 MHz; CDCl3): δ 168.9, 135.1,
128.5, 122.9, 114.7, 110.9, 110.5, 51.7 HRMS (ESI+) for
C8H9NO2Na (M+Na) calcd 174.0525, found 174.0522.
6
7
IntDensitytotal cell area − (Total Cell Area × BkgFluorescence)
8
퐶푇퐶퐹 =
Methyl 3-(1-H-pyrrol-2-yl)propanoate (2). 1 (4.82 g,
0.031 mol) was dissolved in methanol/THF (5:1, 170 mL) in
the presence of palladium on activated carbon (0.482 g,
10% wt/wt) under 1 atm of hydrogen. The reaction mixture
was stirred for 20 h at room temperature after which it was
filtered through celite. Solvent was then removed from the
filtered solution very carefully since the product was vola-
tile. Due to volatility, the exact mass of the product was not
quantified since residual solvent was present, and was used
without purification. 1H-NMR (500 MHz; CDCl3): δ 8.72 (s,
1H), 6.72 (td, J = 2.6, 1.6 Hz, 1H), 6.18 (q, J = 2.9 Hz, 1H),
6.01-5.99 (m, 1H), 3.77 (s, 3H), 2.99 (t, J = 7.1 Hz, 2H), 2.72
(t, J = 7.1 Hz, 2H). 13C NMR (126 MHz; CDCl3): δ 174.4, 130.9,
116.9, 108.1, 105.4, 51.9, 34.4, 22.7 HRMS (APCI+ for
C8H12NO2) (M+H) calcd 154.08626, found 154.08618.
푁
푐푒푙푙푠
9
Equation 3
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where IntDensitytotal cell area is the integrated intensity of
the pixels for all cells in the image, Total Cell Area is the
number of pixels of all of the cells, BkgFluorescence is the
mean fluorescence intensity per pixel for a region contain-
ing no cells and Ncells is the number of cells in the image.
Measurements and errors reported are from the average
and standard deviation CTCF obtained from at least 3 im-
ages per sample.
An ImageJ macro was utilized to identify the cells from
the DIC image. First, a variance filter of radius five pixels
was applied to the DIC image. The resulting image was
converted to a binary image and the Fill Holes function was
applied. The Analyze Particles function was next used to
identify and fill any remaining holes smaller than 20 pixels2
in the binary image, by setting their pixel values to 255. A
dilate function was then applied. The binary image was
then used to create a selection area for the region contain-
ing cells and the region without.
2-(3-Chloropropyl)-1H-pyrrole (3). 2 (0.031 mol, as-
sumed quantitative from previous step) was dissolved in
THF under argon at -78 °C. Borane in 1M THF (6.2 mL, 2
eq) was added dropwise to the solution. The reaction mix-
ture was allowed to warm to room temperature and stirred
for 20 h. The reaction was then quenched with methanol,
diluted with dichloromethane and washed twice with
brine. Organic fractions were dried over anhydrous sodium
sulfate and solvent was removed carefully under reduced
pressure. The resulting volatile alcohol (3-(1H-pyrrol-2-
yl)propan-1-ol) was then dissolved in dichloro-
methane/carbon tetrachloride (95:5, 150 mL) under argon.
Triphenylphosphine (8.1 g, 0.031 mol, 1 eq.) was added in
one portion. The reaction mixture was stirred for 3 h at 40
° C. Solvent was then carefully removed under reduced
pressure, and the resulting oil was loaded onto a silica flash
column and eluted with ethyl acetate/hexanes (20%) af-
fording an oil (2.15 g, 50% yield from compound 1, over 3
steps). 1H-NMR (300 MHz; CDCl3): δ 8.01 (s, 1H), 6.71 (d, J
= 1.4 Hz, 1H), 6.19 (d, J = 2.8 Hz, 1H), 6.00 (s, 1H), 3.60 (t, J
= 6.3 Hz, 2H), 2.82 (t, J = 7.3 Hz, 2H), 2.10 (t, J = 6.9 Hz, 2H).
13C NMR (75 MHz; CDCl3): δ 130.6, 116.6, 108.5, 105.5, 44.5,
32.5, 24.6 HRMS (APCI for C7H11ClN) (M+H) calcd
144.0580, found 144.0573.
Quantifying concentration of probe in cells
HeLa cells were seeded at 5 × 104 cells/mL in a 10 cm petri
dish (23.5 mL) and were allowed to grow for 24 h, approxi-
mately doubling in population (These concentrations and
volumes were used to reflect the ratio of concentration:vol-
ume:surface area of the ibidi dishes used for imaging). Cells
were then washed with LCS, and 23.5 mL of LCS containing
100 nM H4BCH3 was added for 10 minutes. The fluores-
cence of the supernatant was then measured. Simultane-
ously, a 10 cm petri dish containing no cells (control) was
treated the same way: 23.5 mL of LCS with 100 nM H4BCH3
was added for 10 minutes. The fluorescence of this control
was then measured and compared to the supernatant of
the sample containing cells. Once the LCS solution was re-
moved from the control dish, 2 mL of DMSO was then
added and the fluorescence of this solution was measured
to quantify the amount of dye which adhered to the dish.
With these values in hand, the amount of dye which parti-
tioned into cells was then quantified (see Section 4.7.3).
8-((6-Hydroxy-2,5,7,8-tetramethylchroman-2-yl)-
methyl)-1,5-di-(3-chloropropyl)-Pyrromethene
Fluoroborate (H4BPMHC). 2-(6-Methoxy-2,5,7,8-tetra-
methylchroman-2-yl)acetic acid26 (0.19 g, 0.7 mol) was dis-
solved in thionyl chloride (1 mL) under argon and stirred
at 50 °C for 1 h. The thionyl chloride was then removed by
co-evaporation with toluene (15 mL, 5 times) and the flask
was left on a vacuum pump overnight. The resulting acid
chloride was then dissolved in 7 mL of dichloromethane
under argon. Phosphorus oxychloride (0.33 mL, 3.4 mol)
was added dropwise followed by 3 (0.2 g, 1.4 mol) pre-dis-
solved in 2 mL of dichloromethane. The reaction mixture
Synthesis
Methyl (E)-3-(1H-pyrrol-2-yl)acrylate (1). Pyrrole-2-
carboxaldehyde (3.0 g, 0.031 mol) was dissolved in tetrahy-
drofuran (THF) (60 mL) under argon. Methyl (tri-
phenylphosphoranylidene)acetate (12.6 g, 0.037 mol, 1.2
eq.) was added to the stirring solution and the temperature
was increased to 60 °C. After 20 h the solvent was removed
under reduced pressure. The resulting solid was loaded
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was stirred for 22 h at 35 °C. The mixture was then diluted
26.1, 15.5 HRMS (ESI-) for C16H18N2BCl2F2 (M-H) calculated
357.09136, found 357.09181.
1
2
3
4
5
6
7
8
with dichloromethane, washed with brine, and dried over
sodium sulfate. Solvent was removed under reduced pres-
sure. The resulting red oil was dissolved in dichloro-
methane under argon and cooled to -10 °C. Borontribro-
mide (3.5 mL, 1M in dichloromethane, 3.5 mol) was added
dropwise. The reaction mixture was allowed to slowly
warm to room temperature over the course of 1.5 h when
the reaction was quenched carefully with water. The solu-
tion was extracted with ethyl acetate and the organic frac-
tions were washed with brine and dried over sodium sul-
fate. Solvent was removed under reduced pressure. The re-
sulting red oil was dissolved in 5 mL of dichloromethane
and 10 mL of toluene under argon at 0 °C. Diisopropylethyl-
amine (0.49 mL, 2.8 mol) was added dropwise. After 10
minutes, Boron trifluoride diethyl etherate (0.69 mL, 6
mmol) was added dropwise and stirred for 1 h. The mixture
was then quenched with water and extracted with ethyl ac-
etate. The organic fractions were washed with brine and
dried over sodium sulfate. Solvent was removed under re-
duced pressure. The resulting red solid was purified by
flash column chromatography (30% ethyl acetate in hex-
anes) followed by further purification by a preparatory sil-
ica plate eluted in dichloromethane giving an orange film
(30 mg, 8% yield). 1H-NMR (500 MHz; CDCl3): δ 7.12 (d, J
= 4.2 Hz, 2H), 6.32 (d, J = 4.2 Hz, 2H), 4.25 (s, 1H), 3.62 (t, J
= 6.6 Hz, 4H), 3.22-3.11 (m, 6H), 2.68 (qd, J = 13.3, 7.0 Hz,
2H), 2.22 (quintet, J = 7.3 Hz, 4H), 2.14 (s, 3H), 2.11 (s, 3H),
1.94 (s, 3H), 1.87 (t, J = 6.9 Hz, 2H), 1.28 (s, 3H). 13C NMR
(126 MHz; CDCl3): δ 159.5, 145.1, 144.8, 140.5, 136.1, 129.5,
122.9, 121.5, 118.6, 117.5, 116.6, 74.3, 44.3, 40.7, 33.0, 31.6, 26.1,
24.5, 20.7, 12.3, 12.2, 11.3 HRMS (ESI-) for C29H34BCl2F2N2O2
(M-H) calcd 561.2064 found 561.2076.
ASSOCIATED CONTENT
Supporting Information
The Supporting Information is available free of charge on the
ACS Publications website.
Figures S1-S9, Scheme S1, Equations S1 and S2, Approximation
1
of dye concentrations in cells, and H NMR and 13C NMR spec-
9
tra for compounds 1-3, 3-(1H-pyrrol-2-yl)propan-1-ol),
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H4BPMHC, and H4BCH
3 (PDF).
AUTHOR INFORMATION
Corresponding Author
* Email: gonzalo.cosa@mcgill.ca.
ACKNOWLEDGMENT
G.C. is grateful to the Natural Sciences and Engineering Re-
search Council of Canada (NSERC) and the Canadian Founda-
tion for Innovation (CFI) for funding; L.E.G. is thankful to Va-
nier Canada for a postgraduate scholarship; R.L. is thankful to
NSERC for a postgraduate scholarship.
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