Enzymatic activation of nitro-aryl fluorogens in live bacterial cells for enzymatic turnover-activated localization microscopy

Article abstract of DOI:10.1039/c2sc21074f

Many modern super-resolution imaging methods based on single-molecule fluorescence require the conversion of a dark fluorogen into a bright emitter to control emitter concentration. We have synthesized and characterized a nitro-aryl fluorogen which can be converted by a nitroreductase enzyme into a bright push-pull red-emitting fluorophore. Synthesis of model compounds and optical spectroscopy identify a hydroxyl-amino derivative as the product fluorophore, which is bright and detectable on the single-molecule level for fluorogens attached to a surface. Solution kinetic analysis shows Michaelis-Menten rate dependence upon both NADH and the fluorogen concentrations as expected. The generation of low concentrations of single-molecule emitters by enzymatic turnovers is used to extract subdiffraction information about localizations of both fluorophores and nitroreductase enzymes in cells. Enzymatic Turnover Activated Localization Microscopy (ETALM) is a complementary mechanism to photoactivation and blinking for controlling the emission of single molecules to image beyond the diffraction limit.

Full text of DOI:10.1039/c2sc21074f

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Enzymatic activation of nitro-aryl fluorogens in live
bacterial cells for enzymatic turnover-activated
localization microscopy†
Cite this: Chem. Sci., 2013, 4, 220
Marissa K. Lee,a Jarrod Williams,b Robert J. Twieg,b Jianghong Raoc
a
*
and W. E. Moerner
Many modern super-resolution imaging methods based on single-molecule fluorescence require the
conversion of a dark fluorogen into a bright emitter to control emitter concentration. We have
synthesized and characterized a nitro-aryl fluorogen which can be converted by a nitroreductase
enzyme into a bright push–pull red-emitting fluorophore. Synthesis of model compounds and optical
spectroscopy identify a hydroxyl-amino derivative as the product fluorophore, which is bright and
detectable on the single-molecule level for fluorogens attached to a surface. Solution kinetic analysis
shows Michaelis–Menten rate dependence upon both NADH and the fluorogen concentrations as
expected. The generation of low concentrations of single-molecule emitters by enzymatic turnovers is
used to extract subdiffraction information about localizations of both fluorophores and nitroreductase
enzymes in cells. Enzymatic Turnover Activated Localization Microscopy (ETALM) is a complementary
mechanism to photoactivation and blinking for controlling the emission of single molecules to image
beyond the diffraction limit.
Received 25th July 2012
Accepted 5th October 2012
DOI: 10.1039/c2sc21074f
www.rsc.org/chemicalscience
to specic cellular locations9,10 has been developed as an alter-
native to FPs, and although challenges remain for cell entry and
Introduction
Recently developed microscopies (e.g., PALM, FPALM, STORM)
bypass the optical diffraction limit (DL) by successive imaging
and localization of sparse subsets of single-molecule (SM) u-
orophores.1–3 Using these “superresolution” (SR) imaging
methods, structures can be imaged with spatial resolution 5–10
times smaller than the DL of z200 nm for visible wavelengths.
Bright emitters are essential for this SR strategy, because the
localization resolution improves as 1/ON, with N the number of
photons detected before photobleaching or per switching cycle.4
Using emitters whose concentration can be controlled ensures
that each frame during an acquisition has only a sparse set of
emitting molecules, even if the object of interest is densely
labelled.5 Though most previous SR experiments in cells have
used photocontrollable uorescent proteins (FPs)6–8 because of
their genetic targeting capabilities, these emit (on average) 10-
fold fewer photons before photobleaching than good small-
molecule uorophores.5 Targeting small organic uorophores
washout, the chemical properties, spectral range, and photo-
physics are more readily modied by organic synthesis.11
For SR imaging, the emission from organic uorophores can
be controlled optically by reversible photoswitching12 (oen in the
presenceofadditiveslikethiols,oroxidizing/reducingagents)3,13,14
or by irreversible photoactivation.15 Non-optical methods to
control uorescence emission include ligand binding,16 and
uorescence quenchers.17 Fluorogenic substrates have enabled
detailed single-molecule enzymology studies18 but without
extracting subdiffraction information. We present a small-mole-
cule uorophore synthetically modied to form an enzymatically
activated and cell-permeable uorogen, and demonstrate Enzy-
maticTurnoverActivatedLocalizationMicroscopy(ETALM), anew
SR imaging method, in bacterial cells. ETALM differs from other
uorescence modulation strategies using a chemical reduction19
because the enzymatic turnover continually generates new uo-
rophores. This activation is irreversible and the new uorophores
do not undergo multiple switching cycles. The emitting concen-
trationcanbesetattherequiredlowlevelforSRimagingsimplyby
limitingtheuorogenconcentrationorslowingtheenzymaticrate
using inhibitors.
aDepartment of Chemistry, Stanford University, Stanford, California 94305, USA.
E-mail: wmoerner@stanford.edu
bDepartment of Chemistry and Biochemistry, Kent State University, Kent, Ohio, 44240,
Nitroreductase (NTR) enzymes are a family of NAD(P)H-
dependent avoproteins ubiquitous in bacteria and capable of
reducing aryl-nitro compounds to their corresponding hydroxyl-
amino or amino derivatives.20,21 Previous work has used a NTR
USA
cDepartment of Radiology, Stanford University, Stanford, California 94305, USA
† Electronic supplementary information (ESI) available: Full experimental details,
product characterization, selected NMR spectra, and additional gures are
provided. See DOI: 10.1039/c2sc21074f
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enzyme to reduce a nitro-uorogen into a cancer therapeutic22–24 Solution measurements were performed in ethanol on puried
but the resulting uorophore was not sufficiently bright or compounds to allow comparison with our previously
emitted at too short a wavelength to allow SM imaging in cells. published work on DCDHF-type uorophores.15
A poly-
This study uses the commercially available E. coli nsfB NTR21 (methylmethacrylate) (PMMA) polymer lm in toluene was used
(>90 % pure, see ESI Fig. S1†) which has been well-studied in a bulk spectroscopic measurement to show the increase of
because of its potential use in cancer therapeutic delivery.25
uorescence quantum yield (FF) of 2 and 3 in more rigid envi-
We show below that NTR converts the nitro-aryl uorogen of ronments. PMMA polymers, though used previously to charac-
the present study 1 into a bright organic uorophore, suitable terize the azido-DCDHF uorogen,15 are less relevant to
for SR in live bacterial cells (Scheme 1). The emissive product behaviour inside a cell, but measurements were performed in
belongs to the dicyanomethylenedihydrofuran (DCDHF) class PMMA to compare 2 to prior work. As is the case for the other
of push–pull uorophores where an electron-poor DCDHF DCDHF uorophores,26 the uorescence quantum yield FF of 2
acceptor head is connected to an electron-rich donor, usually an increases in the more constrained polymer environment. In
amine, through a conjugated p network.26 The DCDHF uo- order to assess the potential of 2 for cell imaging, more bio-
rophores are detectable on the SM level because they emit logically relevant conditions were used for microscopy
millions of photons before photobleaching and have high measurements of the photobleaching quantum yield (FB) and
uorescence quantum yields in viscous environments, the average number of total photons emitted (Ntot). Here, we
including membranes and gelatin.27 Importantly, DCDHF u- used a primarily aqueous polymer lm (1% poly(vinyl alcohol)
orophores can enter living cells fairly easily because they are (PVA) in water) to immobilize the uorophores. Other in vitro
neutral, and they emit at long wavelengths to avoid environments like poly-lysine or gelatin either did not provide
autouorescence.27
sufficient immobilization, or had an unacceptably high uo-
rescence background at the wavelengths used.
Importantly for single-molecule based superresolution
imaging strategies, SMs of 2 were detectable when immobilized
in the aqueous PVA polymer lm (Fig. 2A), conrmed by the
observation of single-step photobleaching to the background
level. Movie S1† shows SMs of 2 in PVA, and representative time
traces of the SMs appear in Fig. S7.† The average number of
photons detected before photobleaching (Ntot) was measured
for 2 and 3 (Fig. 2A and B, Table 1) using the survival proba-
bility. The survival probability PN (Fig. 2B) is the ratio of the
number of SMs mn surviving aer a given number of emission
events N to the total number of molecules M in the measure-
ment set (see ESI†).15,30 This approach gives comparable results
to histogramming the detected photons31 but avoids binning
artifacts. To convert photons detected to total emitted photons,
the detected value is divided by the optical system collection
efficiency32 of 0.04.
Aer photophysical characterization of separately synthe-
sized 1–3, we used microscopy to measure the uorescence
generated by the NTR-reaction on a surface in vitro. Compound
4, a N-hydroxy-succinimide (NHS) derivative of 1 (Chart 1) was
covalently attached to poly-lysine coated coverslips through
reaction of NHS with the surface amines. Aer washing off the
unreacted 4, NADH and NTR enzyme were added in buffer and
allowed to react with the attached 4. Dramatic uorescence
turn-on was observed aer this addition (Fig. 2D and inset).
Our microscopy-based experiments could detect SMs of 2
and see uorescence from the NTR-reaction, but do not quan-
tify the NTR reaction rate. We used bulk spectroscopic
measurements in solution to nd the NTR reaction kinetic rates
(Fig. 3) and used the more water-soluble nitro derivative 5
(Chart 1) instead of 1 because 1 aggregates in higher concen-
tration regimes. (Kinetic measurements using 1 at lower
concentrations are shown in the ESI, Fig S8†). NTR follows a
ping-pong bi–bi mechanism33 (eqn (1)) where the rate per
enzyme (ni/E) depends on both NADH and nitro substrate 5
concentrations with two Michaelis constants:34,35
Results and discussion
We previously reported a photoactivatable azido-DCDHF15,28
suitable for SR in bacterial and mammalian cells.29 To create
another dark DCDHF uorogen, the electron-rich amine of the
emissive form can be replaced with an electron-poor nitro group
which eliminates uorescence and shis the absorbance to the
blue. Then, reduction of 1 in buffer by the NTR enzyme in the
presence of NADH creates the red-shied emitter (Fig. 1).
The product uorophore in these bulk spectroscopic
measurements could be either the hydroxyl-amino-DCDHF 2 or
the amino-DCDHF 3 (Scheme 1). The product of the NTR reac-
tion was determined to be 2 instead of 3 by comparing the bulk
emission and excitation spectra (in buffer) of separately
synthesized 2 and 3 with the reaction mixture (Fig. 1B). There is
close spectral overlap between the NTR reaction mixture
product and 2.
The bulk and SM photophysical characteristics of molecules
1–3 were measured in solution and in a polymer lm (Table 1).
Scheme 1 Reaction of 1 with NTR and NADH.
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Fig. 1 (A) Spectral change during the NTR reaction with 1 and NADH over z10 minutes showing red-shifted absorption and fluorescence increase. Solid curves show
absorbance of the reaction mixture at different time points, broken red curve is absorbance of 1 alone. Dashed lines are fluorescence emission (500 nm excitation)
before and after the NTR reaction. No fluorescence turn-on or absorbance change is observed without all three components, see ESI Fig. S2 and S3.† (B) Normalized
excitation (solid curves and empty symbols, collected at emission max) and emission (dashed curves and solid symbols, 500 nm excitation) spectra of NTR reaction
mixture (triangles) compared to separately synthesized 2 (circles) and 3 (squares). All spectra measured in 10 mM Tris–HCl (pH 7.5), for more details, see ESI.†
When B. subtilis cells are incubated with 5 mM concentrations
vi
E
kcat½5ꢀ½NADHꢀ
¼
(1)
½5ꢀ
of 1, the emission spots from the large number of enzymatically
generated diffusing uorophores overlap, and the cells appear
bright so that distinct single molecules cannot be observed. By
lowering the incubation concentration of 1 to <nM levels, sepa-
rated uorescent spots can be observed turning on and
Km ½NADHꢀ þ KmNADH½5ꢀ þ ½5ꢀ½NADHꢀ
The initial rate of product formation, monitored by absorp-
tion at 500 nm, was measured for a range of initial concentra-
tions of 5 and NADH (Fig. 3). These rates were t to eqn (1),
yielding t parameters kcat of 224 ꢁ 40 sꢂ1 and KMNADH of 13 ꢁ
3 mM and a K[M5] of 4.9 ꢁ 1 mM, consistent with prior work.34
Aer conrming the NTR-reaction uorescence turn-on on a
surface and measuring the bulk NTR rate, we explored the NTR-
reaction scheme for turnover of the uorogen in bacterial cells.
NTR enzymes are endogenous in many bacteria. When several
bacterial species (E. coli, Caulobacter crescentus, Bacillus subtilis)
are incubated with mM concentrations of 1, entire cells become
uorescent (Fig. 4A and ESI Fig S9†). This effect was most
dramatic in wild type B. subtilis cells, which are known to have
enhanced response to nitro compounds.36,37 To support the
hypothesis that this uorescence arises from a NTR reduction
analogous to the reaction studied in vitro, 1 and an excess of
menadione, a competitive NTR inhibitor, were added simulta-
neously to the bacteria. Under this incubation condition, the
uorescence of individual cells was reduced by 14ꢃ to just
above background levels (Fig. 4B and E). No additives were
needed to make 1 cell permeable.
Table 1 Photophysical characterization of molecules 1–3 in ethanol (unless
otherwise stated)a
Fig. 2 (A) SM fluorescent spots of 2 (verified by one-step photobleaching) in
PVA film (532 nm excitation, 91 W cmꢂ2). (B) Survival probability histogram of 2
doped into PVA films at nM concentrations with an average 71 000 detected
photons, corresponding to an average of 1.7 ꢃ 106 photons emitted by each SM.
(C) Schematic of in vitro experiment where 4 (structure shown in Chart 1) was
covalently attached to poly-lysine coverslips immersed in PBS buffer (pH 7.4)
containing free NADH and NTR enzyme. Enzymatically generated fluorophores
remained attached to surface lysines. (D) Bulk fluorescence at the surface after
NADH and NTR addition. Inset: bulk fluorescence of 4 before NTR reaction (same
contrast). Bulk fluorescence of all control incubation conditions is shown in
Fig. S6.† sample and control details in ESI.†
c
labs (nm),
l
SM Ntot
(ꢃ106)
3 (dm3 molꢂ1 cmꢂ1
)
(nm) FF
FB (ꢃ10ꢂ6
)
b
1
2
3
390, 17 020
497, 15 300
570, 54 100
n/a
609
613
n/a
n/a
n/a
1.7
2.3
0.021, 0.27d 5.6
0.025, 0.39d 4.1
a
b
See ESI† and ref. 28 for details on measurements and calculations.
Bulk quantum yield of permanent photobleaching, measured in
aqueous PVA polymer lm. Average total number of photons emitted
for SMs in PVA polymer lm. Fluorescence quantum yield in PMMA.
c
d
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Fig. 4 (A) B. subtilis incubated with 5 mM of 1. Left image: average fluorescence;
right image: corresponding white light image of cells. All cells become fluores-
cent; cells in the center are brighter because the laser excitation spot is more
intense in the center. Images in the controls (B)–(D) are structured similarly: left
image is average fluorescence at same contrast scale as (A), and right image is
corresponding white light image. (B) B. subtilis incubated with both 1 (5 mM) and
menadione (0.2 mM), showing a dramatic fluorescence reduction. The images in
(C) show cells incubated with only menadione, and part (D) shows untreated B.
subtilis. (E) Quantification of number of photons detected per cell (ꢄ300 cells per
Chart 1 Structures of nitro-DCDHF derivatives 4 and 5 used for surface immo-
bilization and kinetic experiments, respectively. Bulk absorbance and fluorescence
changes from the NTR reaction are shown in ESI Fig. S4 and Fig. S5, respectively.†
Table S1 in ESI† summarizes photophysical characteristics for these molecules.
condition) for each case. All scale bars are 2 mm, imaging intensity is 2.5 kW cmꢂ2
.
All conditions were incubated for 30 minutes. For more details on incubation and
imaging conditions, see ESI.†
subsequently photobleaching (Fig. 5A and insets). Even though
the true size of the emitting molecule is z1 nm, the diameter of
the SM uorescent spot is z200 nm, on the order of the bacterial
cell width, due to the DL. These uorescent spots correspond to
SMs, veried by one step photobleaching to background levels.
At the laser intensity used here (15 kW cmꢂ2), photobleaching is
fast relative to the 100 ms exposure time, as shown by repre-
sentative time-traces of the SM emission (Fig. 5A).
We now seek to demonstrate that SR information can be
extracted from the enzyme turnovers using ETALM.‡ As is well-
known, with separated SM uorophores, the tted center of the
DL spot is an estimate of the true position of the emitting
molecule. By lowering the incubation concentration, the total
number of enzymatically generated uorophores is easily
decreased. The incubation concentration of 1 used to create
Fig. 5A was slightly too low for SR imaging, as only one SM spot
would appear in a given DL region every 8–30 seconds. The
incubation concentration of 1 was increased very slightly to
1.5 nM, so new SMs were generated at a reasonable rate during
continuous imaging. A DL image composed of the average of
many imaging frames is shown in Fig. 5B. It should be noted
that our interpretation of SR reconstructions could still be
complicated by blurred spots arising from diffusion of the free
uorophores aer catalysis. To minimize this effect, the
number of long-lived uorophores is reduced by using high
intensity reading light, and the camera exposure time is
reduced to 8 ms per frame. In the limit of very high laser
intensity, each newly generated SM uorophore should be
photobleached before it has time to diffuse away from the NTR
enzyme. In this limit, the extracted SM positions would be
samplings of the NTR enzyme position over time as it moves
through the cell, generating uorescent molecules. In this
initial demonstration, we only approached this limit in inten-
sity, thus in our SR reconstructions (Fig. 5C–F and Movie S2†),
two populations of localizations are observed: (i) punctate spots
(dashed boxes in Fig. 5C and E) and (ii) membrane localizations
(e.g., seen easily in Fig. 5C). For case (i), aggregation could cause
the punctate spots, but 1 is added at the nM level and was only
found to aggregate at the mM regime in separate experiments.
We hypothesize that the punctate spots arise from a “cloud” of
enzymatically generated uorophores photobleached before
they could travel very far from the enzyme. Notably, time-
coloring of the successive SM localizations from the uores-
cence movie sometimes shows apparent motion of the enzyme
(Fig. 5F). Considering the second population (ii), some emitters
can apparently survive for a longer time and are able to diffuse
to reach the membrane which represents a sink for the relatively
nonpolar 2. It is reasonable that a small fraction of uorophores
Fig. 3 3D Michaelis–Menten plot of initial rate of product formation (monitored
by change in absorption at 500 nm) in PBS (pH 7.4) versus various starting
concentrations of 5 and NADH. The structure of 5 is shown in Chart 1. The grid is
the fit of eqn (1) to the data points.
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Fig. 5 (A) Representative SM emission traces of enzymatically generated fluorophores in B. subtilis showing photons detected per 100 ms bin vs. time. For each graph,
the inset is a single imaging frame from the specific B. subtilis cell (cell outline in dashed green, SM plotted is in blue box). (B) DL average image of 3000 frames (24 s) of
B. subtilis incubated with 1 and corresponding white light image (inset). (C) SR reconstruction (average 17.9 ꢁ 0.3 nm statistical precision) from the cell boxed in blue
with both membrane localizations and punctate spots (blue dashed) consisting of many overlapping localizations. (D) Temporal progression of the localizations in the
punctate spots, with early localizations colored purple, and later localizations colored white. The lack of temporal progression in the localizations suggests that these
punctate spots correspond to the fluorescent cloud generated by two relatively stationary enzymes. (E) SR reconstruction from the green-boxed cell. (F) Temporal
progression of localizations in the green dashed box of (E) showing a defined temporal progression with later localizations toward the center of the cell (shown by
arrow), suggesting that this punctate spot is a single enzyme diffusing on the time scale of our experiment. Scale bars: 1 mm except 0.5 mm in (D and F). For more details
on incubation, imaging, fitting, and display conditions, see the ESI.†
could reach the protective environment of the membrane nascent uorophores, but the use of an inhibitor could also
without photobleaching given that the expected diffusion allow temporary suppression and tuning of the uorescence
coefficient for a small molecule in a bacterial cell is on the order generation rate.
of 5.0 ꢃ 10ꢂ7 cm2 sꢂ1
,
which could easily result in an rms
38
displacement of 1.5 microns during the 8 ms imaging time.
Membrane-bound uorophores are more easily localized Acknowledgements
because diffusion is slower39 and the DCDHF-type uorophores
are harder to bleach (since FF is larger in the membrane).27
This work was supported in part by grant no. R01-GM086196
from the National Institute of General Medical Sciences.
Samples of wild-type Bacillus subtilis, E. coli, and Caulobacter
crescentus were obtained from Jerod Ptacin in the Shapiro lab at
Stanford University. The protein gel was run with the help of
Steffi Duttler in the Frydman lab at Stanford University.
Conclusions
We have synthesized and characterized a nitro-uorogen which
can be enzymatically activated to produce a bright SM emitter.
The uorescent product was identied and characterized on a
bulk and SM-level in vitro and in cells. Unlike previous work
using enzymatic turnovers to generate uorescent SM prod-
ucts,40–42 we used the nascent uorescent molecules from the
NTR enzyme on the single-molecule level to explore the sub-
diffraction-scale organization of a cytosolic bacterial enzyme in
B. subtilis as well as the membrane locations where some of the
product molecules are found.
Considering extensions beyond bacteria, mammalian cells
oen have much lower levels of NTR-type enzymes20 or activity
and would be interesting targets for future work. In cells with
suitably low levels of endogenous NTR activity, the ETALM
scheme could be used as an orthogonal genetically encoded
strategy for SR imaging of protein structures of interest fused to
NTR. One particular advantage of this approach arises from the
fact that the precursor nitro-uorogen 1 is not uorescent, so no
washing steps are required. The activation of 1 is not light-
dependent, and samples do not require special handling under
red lights or in the dark. It may be useful in some contexts to use
the NTR scheme as a generator of uorescent molecules from a
source inside the bacterial cell to obtain the position of the
inner membrane.43 At the same time, one of the challenges of
this system is controlling the total amount of activated uo-
rophores. We used high intensity laser light to rapidly bleach
Notes and references
‡ Since many nitro compounds are known to be toxic to bacterial cells,20 the total
time of SR experiments from addition of 1 to imaging was kept as short as possible
(<10 minutes).
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