1 Departments of Medicine and Physiology, Cardiovascular Research Institute, University of California, San Francisco, California, 94143-0521; and 2 Faculty of Chemistry, A. Mickiewicz University, 60-780 Poznan, Poland
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ABSTRACT |
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Limitations of available indicators [such as
6-methoxy-N-(3-sulfopropyl)quinolinium
(SPQ)] for measurement of intracellular Cl are their relatively dim
fluorescence and need for ultraviolet excitation. A series of
long-wavelength polar fluorophores was screened to identify compounds
with Cl
and/or
I
sensitivity, bright
fluorescence, low toxicity, uniform loading of cytoplasm with minimal
leakage, and chemical stability in cells. The best compound found was
7-(
-D-ribofuranosylamino)-pyrido[2,1-h]-pteridin-11-ium-5-olate (LZQ). LZQ is brightly fluorescent with excitation and
emission maxima at 400-470 and 490-560 nm, molar extinction
11,100 M
1 · cm
1
(424 nm), and quantum yield 0.53. LZQ fluorescence is quenched by
I
by a collisional
mechanism (Stern-Volmer constant 60 M
1) and is not affected
by other halides, nitrate, cations, or pH changes (pH 5-8). After
LZQ loading into cytoplasm by hypotonic shock or overnight incubation,
LZQ remained trapped in cells (leakage <3%/h). LZQ stained cytoplasm
uniformly, remained chemically inert, did not bind to cytoplasmic
components, and was photobleached by <1% during 1 h of continuous
illumination. Cytoplasmic LZQ fluorescence was quenched selectively by
I
(50% quenching at 38 mM
I
). LZQ was used to
measure forskolin-stimulated
I
/Cl
and
I
/NO
3
exchange in cystic fibrosis transmembrane conductance regulator
(CFTR)-expressing cell lines by fluorescence microscopy and microplate
reader instrumentation using 96-well plates. The substantially improved
optical and cellular properties of LZQ over existing indicators should
permit the quantitative analysis of CFTR function in gene delivery
trials and high-throughput screening of compounds for correction of the
cystic fibrosis phenotype.
cystic fibrosis transmembrane conductance regulator; cystic fibrosis; 6-methoxy-N-(3-sulfopropyl)quinolinium; chloride transport; fluorescence; high-throughput screening
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INTRODUCTION |
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SEVERAL HALIDE-SENSITIVE fluorescent indicators have
been introduced to study the functional properties of
Cl transporters in
biomembrane vesicles, proteoliposomes, living cells, and epithelia (for
review, see Refs. 28 and 30). The archetype indicator
6-methoxy-N-(3-sulfopropyl)quinolinium
(SPQ) (11) is a polar quinolinium compound whose fluorescence is
quenched by Cl
and
I
by a collisional
mechanism. SPQ and related quinolinium indicators have been useful in
studying the cystic fibrosis transmembrane conductance regulator (CFTR)
Cl
channel expressed in
native and transfected cell models (4, 6, 7, 21, 22) and, recently, in
assaying functional CFTR delivery in human gene therapy trials (10,
20). Various SPQ derivatives have been synthesized for specific
applications including cell-permeable/trappable compounds (3),
dual-wavelength Cl
indicators for ratio imaging (12), dextran-linked conjugates (1), and
fiberoptic halide sensors (15).
Although used in numerous studies of CFTR function in cell culture
models, the existing quinolinium-based halide indicators have imperfect
optical and physical properties that limit their utility in more
challenging applications, including the analysis of CFTR function in
gene therapy trials and high-throughput drug screening. SPQ and related
indicators have relatively dim fluorescence in cells (molar extinction < 6,000 M1 · cm
1;
quantum yield < 0.1) and require ultraviolet excitation (excitation 320-365 nm; emission 420-460 nm). Significant technical
limitations include the need for sensitive detection instrumentation
with high numerical aperture optics, and photodynamic cell injury and background autofluorescence resulting from ultraviolet excitation. In
addition, the quinolinium halide indicators are quenched by intracellular proteins and organic anions, resulting in decreased indicator sensitivity to cytoplasmic
Cl
and undesired indicator
sensitivity to cell volume changes (5, 26). We previously synthesized
long-wavelength Cl
indicators containing the acridinium chromophore (2); although these
indicators are useful to measure
Cl
transport in liposomes
and biomembrane vesicles, they are chemically unstable in cytoplasm, a
problem that could not be overcome by derivatization.
The purpose of this project was to identify/synthesize bright,
long-wavelength fluorescent halide indicators for assay of CFTR-mediated anion conductance in living cells. A specific goal was to
develop a sensitive, robust anion transport assay suitable for
high-throughput drug screening. The desired specifications of the
indicator(s) included: high
Cl and/or
I
sensitivity, bright
fluorescence with excitation wavelength > 400 nm and emission
wavelength > 500 nm, minimal photobleaching, no cellular toxicity,
rapid loading into cytoplasm, uniform distribution and chemical
stability in cytoplasm, and slow leakage out of cells. A series of
candidate fluorescent compounds were screened for these strict
criteria. The compound luminarosine (LZQ), a
pyrido[2,1-h]-pteridin, satisfied all of the above
requirements. LZQ was characterized in vitro and in CFTR-expressing
cells, and a microplate reader assay of functional CFTR was established.
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METHODS |
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Materials. Forskolin, 8-(4-chlorophenylthio)adenosine 3',5'-cyclic monophosphate sodium salt (CPT-cAMP), IBMX, nigericin, valinomycin, and carbonyl cyanide m-chlorophenylhydazone were purchased from Sigma (St. Louis, MO). Rhodamine 110 and rhodamine green were purchased from Molecular Probes (Eugene, OR). Acridine yellow, acridine orange, 9-aminoacridine, 6-aminoquinoline, benzo[c]cinnoline, phenazine, resorufin, and phenosafranine were obtained from Aldrich (Milwaukee, WI). 2,7-Diphenylpyrido[3,2-g]quinoline was provided by Dr. Helmut Quast, University of Wuerzburg, Germany.
Organic synthesis.
Benzo[c]cinnoline and phenazine were quaternized with
methyl iodide to give compounds V and VI, respectively (see Fig.
1 and Table
2). Compound VII was synthesized by
reaction of 2,7-diphenylpyrido[3,2-g]quinoline with
trimethyloxonium tetrafluoroborate. Acridine yellow and acridine orange
were quarternized with dimethyl sulfate to give compounds IX and X,
respectively. 6-Aminoquinoline and 9-aminoacridine were quaternized
with propane sultone to give compounds XI and XII, respectively.
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Compounds I (LZQ) and II (LMQ) were synthesized by phototransformation
of
N-[9-(2',3',5'-tri-O-acetyl--D-ribofuranosyl)-purin-6-yl]pyridinium chloride in aqueous solutions (24, 25).
2',3',5'-Tri-O-acetyl-inosine (0.5 mmol) was dissolved in 2.5 ml of dry pyridine and reacted with
0.75 mmol 4-chlorophenyl dichlorophosphate in the dark for 24 h at room
temperature. Pyridine was then removed by evaporation, and the product
{N-[9-(2',3',5'-tri-O-acetyl-
-D-ribofuranosyl)-purin-6-yl]pyridinium chloride} was dissolved in water, neutralized, washed with
chloroform, and lyophilized. Purity was confirmed by TLC (yield 70%).
Two liters of a 0.3 mM aqueous solution of this compound (at pH
5.8-6.2) was deaerated and irradiated at >300 nm wavelength in a
Pyrex, air-cooled photoreactor. After conversion to the intermediate N-[5-formamido-6-(2',3',5'-tri-O-acetyl-
-D-ribofuranosyl)-purin-6-yl]pyridinium chloride was complete as determined by TLC, a concentrated aqueous solution of the sensitizer (9-methylpurin-6-yl) pyridinium perchloride (0.15 M) was added and irradiation was continued after adjusting the pH
to 7.5. Photoconversion to the bright yellow
2',3',5'-tri-O-acetyl-luminarosine was checked by absorption spectrophotometry. After completion of the
photoconversion, the reaction mixture was filtered and extracted with
chloroform, and the chloroform was evaporated (yield 90%). The product
was purified using silica gel column chromatography. Deacetylation to
LZQ was carried out using triethylamine in dry methanol for 72 h (yield
85%). LMQ was prepared by heating
2',3',5'-tri-O-acetyl-luminarosine in 0.1 M aqueous TCA to 90°C for 3 h (yield 70%).
Cell culture. For fluorescence microscopy experiments, 3T3 fibroblasts (control and CFTR expressing), T84 cells, and Calu-3 cells were cultured on 18-mm round coverslips in DMEM H21 medium supplemented with 5% FCS (3T3 fibroblasts) or 10% FCS (Calu-3 cells), penicillin (100 U/ml), and streptomycin (100 µg/ml). T84 cells were grown in DMEM/ F-12, Ham's nutrient mix (1:1) containing 5% FCS, penicillin (100 U/ml), and streptomycin (100 µg/ml). Cells were grown at 37°C in 95% air-5% CO2 and used when 90% confluent. For microplate reader assays cells were cultured on Costar 96-well black plates with a clear flat bottom and used when nearly confluent.
Fluorescence microscopy. Fluorescence microscopy measurements were performed as described previously (5). Briefly, coverglasses were mounted in a 0.5-ml perfusion chamber in which the cell-free glass surface made contact with an oil immersion objective [Nikon ×40 magnification, numerical aperture (NA) 1.3]. Cell fluorescence was excited at 365 ± 10 nm (SPQ) or 420 ± 10 nm (LZQ and LMQ). Emitted fluorescence was detected by a photomultiplier in a Nikon inverted epifluorescence microscope using a 410-nm dichroic mirror and 420-nm barrier filter (SPQ) or a 455-nm dichroic mirror and 500-nm barrier filter (LZQ). For cells colabeled with SPQ and LZQ, SPQ fluorescence was excited at 365 ± 10 nm and was detected using a 410-nm dichroic mirror and 455 ± 30 nm interference filter. Confocal fluorescence micrographs were obtained using a Nipkow wheel confocal microscope and cooled charge-coupled device camera detector using a ×60 oil immersion objective (Nikon, NA 1.4). LZQ-labeled cells were viewed using a fluorescein filter set (excitation 480 ± 15 nm, emission 520 ± 20 nm).
Fluorescence microplate reader measurements. Microplate reader measurements were performed in a BMG Fluostar microplate reader (BMG LabTechnologies, Durham, NC) equipped with two syringe pumps for automated solution additions. After cell loading with LZQ, extracellular LZQ was washed using a Labsystems Cellwash-4 (Franklin, MD). LZQ fluorescence was excited using a liquid fiberoptic and 425 ± 15 nm interference filter (HQ, Chroma Optical), and emitted fluorescence was collected using a liquid fiberoptic and 530 ± 15 nm filter. The fiberoptic was positioned just below the plate. Fluorescence was recorded continuously in 2-s intervals, each representing the average of 100 20-µs pulses of the xenon illumination lamp source.
Transport measurements. Table 1 lists
the solution compositions and protocols for the transport measurements.
Cells were labeled with LZQ or SPQ by hypotonic shock (loading
buffer/water 1:1 containing 2 mM LZQ or 7.5 mM SPQ) for 5 min
at room temperature or by overnight incubation with indicators in the
cell culture medium. Extracellular indicator was removed by washing
just before measurements. For fluorescence microscopy, glass coverslips
were then mounted in a perfusion chamber (flow ~2 ml/min) in which solutions were exchanged using a four-way valve. For microplate reader
measurements, cells in 96-well plates were bathed in 20 µl of the
first buffer (see Table 1), and 160 µl of the
Cl-containing,
I
-free solution were
injected into the well to drive
Cl
/I
exchange (protocol 1). For
stimulation of CFTR using protocol 2,
60 µl of a 1:8 mixture of the first buffer and the second buffer containing 80 µM forskolin, 2 mM CPT-cAMP, and 400 mM IBMX were added.
Time-resolved fluorescence measurements. Fluorescence lifetime and anisotropy decay measurements were carried out in the frequency domain by cuvette fluorometry using a Fourier transform fluorometer (48000 MHF; SLM Instruments, Urbana, IL) or by fluorescence microscopy using epifluorescence microscope optics in place of the cuvette compartment (29). For microscopy measurements, impulse-modulated vertically polarized light (442 nm, He-Cd laser 35 mW) was reflected onto the sample by a 510-nm dichroic mirror and objective lens; emitted fluorescence was filtered by a 515-nm barrier filter and passed through a rotatable analyzing calcite polarizer. In some experiments, a biconcave lens was introduced just in front of the dichroic mirror to diverge beam diameter in the focal plane to ~40 µm.
Analysis of lifetime and time-resolved anisotropy was performed by a comparative approach. Generally six pairs of measurements (each acquisition 8 s) were obtained, comparing sample and reference (fluorescein in 0.1 N NaOH, lifetime 4.0 ns) for measurement of lifetime, and parallel and perpendicular orientations of the emission polarizer for measurement of anisotropy decay. The phase-modulation data consisted of phase angles and modulation ratios at 40 discrete, equally spaced modulation frequencies (5-200 MHz). The analyzing polarizer was positioned at the magic angle for lifetime measurements. Additional details of the data acquisition and analysis routines were described previously (29). Median phase and modulation values for paired data were analyzed by nonlinear least squares for determination of lifetimes and rotational correlation times.
Computations. In vitro fluorescence quenching studies were carried out at peak excitation and emission wavelengths. Microliter aliquots of the sodium salt of the quenching anion (1 M) were added to indicator solution in 5 mM sodium phosphate (pH 7.2, unless indicated otherwise). Fluorescence intensities in the absence (Fo) and presence (F) of quencher anion ([Q]) were measured to give the Stern-Volmer constant (Kq) as defined by: Fo/F = 1 + Kq[Q].
For determination of the absolute
I transport rates, the time
course of intracellular I
concentration
([I
]) was
computed from the time course of SPQ or LZQ fluorescence using the
modified Stern-Volmer equation
[I
] = {([Fo
Fb]/[F
Fb])
1}/KI,
where Fo is fluorescence intensity in the absence of I
,
Fb is background fluorescence of
unlabeled cells, and
KI the intracellular quenching constant for
I
(see
RESULTS).
I
transport rates were
calculated from the slope of a linear regression of the first 10 time
points after changing perfusate composition.
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RESULTS |
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Long-wavelength fluorophores were screened for sensitivity to
Cl and
I
and for intracellular
properties (loading, intracellular distribution, leakage, stability).
Table 2 summarizes the optical and cellular properties of some of the fluorophores tested. The available
Cl
indicators are based on
fluorescence quenching of the quinolinium chromophore and similar
positively charged heterocycles. Various classes of compounds were
tested with extended conjugation, one or more heteroatoms, and linear
or angular geometry. Several rhodamines were tested based on reports
indicating rhodamine sensitivity to
I
(31). Although several of
the compounds tested showed good sensitivity to
Cl
(compounds V, VI) and
I
in vitro, they were not
useful as intracellular indicators because of instability, nonuniform
intracellular distribution, and/or rapid leakage out of cells as given
in Table 2. Of the compounds tested, the
pyrido[2,1-h]-pteridins LZQ and LMQ had potentially useful
optical and cellular properties and were characterized further.
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Figure 2A
shows the fluorescence spectra of LZQ. LZQ (and LMQ, not shown) have
broad excitation and emission peaks with maxima at 424 and 528 nm,
respectively. The molar extinction coefficient of LZQ was 11,100 M1 · cm
1
at 424 nm, and the quantum yield in the absence of
I
was 0.53. Figure
2B is a Stern-Volmer plot for
quenching of LZQ and LMQ by
I
and
Cl
, with data for SPQ shown
for comparison. In water (or saline), LZQ fluorescence was quenched by
I
with a Stern-Volmer
constant of 60 M
1. LZQ
fluorescence was not quenched (Stern-Volmer constant < 1 M
1) by
Cl
,
NO
3, phosphate, acetate,
Na+, and
K+. For comparison, Stern-Volmer
constants for SPQ quenching are 118 M
1
(Cl
) and 282 M
1
(I
, not shown). LZQ
fluorescence and I
sensitivity were not affected by pH changes in the range 4.5-8. Fluorescence lifetime analysis indicated a 7.1-ns lifetime for LZQ in
the absence of I
, which
decreased in proportion to fluorescence intensity with increasing
[I
], indicating
a collisional mechanism. Stopped-flow measurements showed that LZQ
fluorescence responds in <1 ms to rapid addition and removal of
I
(not shown), as expected
for a collisional quenching mechanism.
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LZQ and LMQ were loaded into cells efficiently by hypotonic shock or
passive incubation. The confocal fluorescence micrograph in Fig.
3B shows
LZQ fluorescence labeling of the aqueous compartments in cytoplasm and
nucleus. The green/yellow LZQ fluorescence was quite uniform and
remarkably more intense than the blue fluorescence of SPQ (Fig.
3A), even though a substantially
higher SPQ concentration was used. All cell types tested, including
epithelial cells (T84, Calu-3, JME,
LLC-PK1, Madin-Darby canine
kidney) and nonepithelial cells (3T3 fibroblasts, Chinese hamster ovary
cells, HeLa cells), could be labeled with LZQ and showed uniform
labeling of the cytoplasm and nucleoplasm. As was found for SPQ, LZQ
and LMQ were nontoxic to cells in assays of cell growth (2 mM in
culture medium for 72 h). TLC of lysates from LZQ-loaded cells showed
that LZQ remained chemically stable in cells.
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It was found that LZQ had better cytoplasmic retention properties than
LMQ, probably because of its sugar moiety. Figure
4A, top, shows the time course of LZQ
fluorescence during perfusion with physiological saline at 23°C. A
small group of 20-30 cells was illuminated continuously, and
fluorescence was detected by a photomultiplier using a ×40, 1.3 NA objective. Cellular fluorescence decreased slowly at a rate of
2.5%/h. To determine the relative contributions of indicator leakage
vs. photobleaching to the slow decline in fluorescence, the perfused
cells were illuminated intermittently (Fig.
4A,
bottom). The decline in fluorescence
was not different from that during continuous illumination, indicating
the absence of photobleaching under the low-light-level conditions
employed here. Similar LZQ loading and leakage properties were found
for multiple different epithelial and nonepithelial cells, as well as
for cells grown on plastic and on porous Transwell filters. Figure
4B shows the time course of cellular
LZQ fluorescence in response to repeated addition and removal of 50 mM
I. There were large,
reversible changes in fluorescence with signal-to-noise ratios
generally >500:1. Background fluorescence (in nonlabeled cells) was
generally <5% of the fluorescence of LZQ-labeled cells.
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A calibration study was done to determine the intracellular sensitivity
of LZQ to I. 3T3
fibroblasts expressing CFTR were perfused with solutions containing
high K+ concentration and
ionophores to equalize solution and intracellular [I
]. LZQ
fluorescence changed reversibly in response to changes in solution
I
(Fig.
4C). Figure
4D shows the Stern-Volmer analysis
indicating a Stern-Volmer constant of 26 M
1 for quenching of
intracellular LZQ by I
,
which is better than that of 12-18
M
1 for quenching of
intracellular SPQ by Cl
(5). Fluorescence lifetime analysis was done to determine whether LZQ
fluorescence is quenched in cells by substances other than
I
. Nanosecond lifetimes
were measured by frequency-domain microfluorometry. Figure
5A shows a
phase-modulation plot indicating a single-component LZQ lifetime in
cells of 6.5 ± 0.3 ns (n = 4) in the absence of I
, close to that of 7.1 ns
for LZQ in water. This finding contrasts with results for SPQ, where
the SPQ fluorescence lifetime was eightfold decreased in cells (in the
absence of Cl
) vs. water
(5). It is concluded that LZQ fluorescence is quenched little by
intracellular components, a significant advantage over SPQ.
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The similar LZQ lifetime in cells and water predicts that LZQ
fluorescence should be relatively insensitive to cell volume in the
absence of I. Figure
5B compares the time course of
intracellular SPQ and LZQ fluorescence in response to osmotically
induced changes in cell volume in the absence of
Cl
. Although SPQ
fluorescence decreased by 30 ± 2% (SE;
n = 3) on exposure to 600 mosmol/kgH2O, LZQ
fluorescence changed by only 5 ± 1% (in the absence of
I
). When cells were
exposed to hyposmotic medium (225 mosmol/kgH2O), SPQ fluorescence
increased by 15 ± 2%, whereas LZQ fluorescence increased by only 4 ± 1%.
The uniform cellular distribution of LZQ in Fig. 3A suggests that LZQ is in the aqueous phase of cytoplasm and nucleoplasm and binds little to intracellular proteins and lipids. Time-resolved anisotropy was measured to quantify intracellular LZQ binding. In water, LZQ rotated freely with a single-component rotational correlation time of 121 ± 3 ps (n = 3; not shown). For LZQ in cells, a plot of differential phase and modulation amplitude ratio (Fig. 5C) indicated a two-component anisotropy decay model with a major component (fractional amplitude 0.90-0.94) of 144 ± 10 ps, similar to that in water. Together these results indicate little binding of LZQ to intracellular components.
The utility of LZQ for functional measurement of CFTR expression was
tested. Figure
6A
compares the time course of LZQ and SPQ fluorescence in
forskolin-stimulated CFTR-expressing cells using identical
I/NO
3
exchange protocols. Although the amplitudes and curve shapes of the
data differed for SPQ vs. LZQ because of unequal indicator Stern-Volmer
constants, the computed I
influx rates [SPQ: 0.26 ± 0.03 mM/s; LZQ: 0.25 ± 0.01 mM/s (SE, n = 3)] and efflux
rates (SPQ: 0.08 ± 0.01 mM/s; LZQ: 0.09 ± 0.03 mM/s) were not
different. Figure 6B shows a
comparison of data obtained for cells loaded with LZQ and SPQ initially
loaded with I
, followed by
replacement of I
with
NO
3 in the absence of forskolin,
followed by addition of forskolin. The initial rates of
forskolin-stimulated I
efflux (SPQ: 0.09 ± 0.02 mM/s; LZQ: 0.08 ± 0.01 mM/s) were
similar. Measurements of
Cl
/NO
3
exchange were also carried out by measuring SPQ fluorescence using
cells labeled with SPQ alone vs. cells colabeled with SPQ and LZQ (Fig.
6C). The presence of LZQ (which is
quenched by I
but not by
Cl
) did not affect the
time course of SPQ fluorescence, indicating that LZQ did not itself
affect CFTR-mediated Cl
transport.
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Because LZQ fluorescence is sensitive to
I but not to
Cl
, it was possible to
measure
Cl
/I
exchange without the need to introduce
NO
3. Figure
7A shows
the kinetics of forskolin-stimulated
Cl
efflux on replacement of
solution Cl
by
I
measured by LZQ
fluorescence. A rapid decrease in LZQ fluorescence was seen in the
CFTR-expressing fibroblasts and T84 cells but not in control
fibroblasts. Figure 7B shows
cAMP-stimulated I
efflux in
response to replacement of solution
I
by
Cl
and addition of cAMP
agonists. There was a slow increase in fluorescence on solution
exchange, representing basal anion transport, followed by a prompt
increase in LZQ fluorescence on addition of cAMP agonists to the
CFTR-expressing cell types.
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The low rate of LZQ leakage out of cells and its bright long-wavelength
fluorescence permitted measurements of CFTR-mediated Cl/I
or
NO
3/I
exchange in a fluorescence microplate reader with cells cultured directly on 96-well plastic dishes. To establish a protocol for the
microplate reader assay, the background fluorescence of clear flat-bottom, 96-well plates obtained from different vendors was measured at LZQ excitation and emission wavelengths. Fluorescence signals [in instrument units ± SD (to assess interwell
variability) at gain typically used in transport assays] were
12,258 ± 386 (Falcon clear); 9,749 ± 568 (Nunc clear); 23,167 ± 755 (Greiner white); 39,782 ± 555 (Falcon white); 8,955 ± 116 (Packard Polyfibronics black); 9,582 ± 211 (Falcon
black); 6,989 ± 192 (Nunc black); 6,420 ± 562 (Greiner black);
and 5,068 ± 268 (Costar black). Costar black plates were used for
subsequent studies because of their relatively low background signal
and interwell variability. Background fluorescence was generally 5- to
10-fold lower than cellular LZQ fluorescence, with very little (<2%)
instrument background (with 96-well plate removed) or cellular
autofluorescence (measured in unlabeled cells). Another technical
consideration was selecting the flow rate and fluid volume added by the
syringe pumps to adapt the microscopy protocol in Fig.
7B to the microplate reader. Exchange of I
for
Cl
was effected in the
microplate reader by diluting the
I
-containing buffer with a
Cl
buffer. To minimize
rapid signal changes due to a rise in meniscus level and the associated
light scattering, cells were initially bathed in a minimum 20-µl vol
of the first buffer (Table 1) and diluted with 160 µl of the second
buffer. The rates of fluid addition by the automated syringe pump was
maintained at 150 µl/s to avoid bubbles (causing light scattering)
and displacement of the cells from the flat-bottomed plastic well.
Figure 8 shows microplate reader data.
Cells were loaded with 2 mM LZQ by a 5-min hypotonic shock, and
extracellular LZQ was removed by rinsing the wells.
Protocols 1 and
2 (Table 1) were compared for
measurement of CFTR-stimulated
I transport. Figure
8A shows rapid
I
influx in the
CFTR-expressing fibroblasts and T84 cells but not in control
fibroblasts. Figure 8B shows
cAMP-stimulated I
efflux in
T84 cells but not in a cell line (JME) expressing the
F508 CFTR.
Although the fluorescence signal-to-noise ratio of the microplate
studies was inferior to the microscopy data in Fig. 7, cAMP-stimulated
CFTR-mediated transport was clearly detected. As described in the
DISCUSSION, technical improvements
should further increase signal-to-noise ratio and thus assay
sensitivity.
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DISCUSSION |
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The goal of this study was to identify bright, long-wavelength halide
indicators suitable for analysis of CFTR function in gene delivery
trials and high-throughput drug discovery. With quinolinium-type
Cl indicators as a starting
point, the strict optical and cellular requirements of the indicator as
outlined in the Introduction limited the types of potentially useful
compounds. The best compound identified, LZQ, was
I
sensitive, brightly green
fluorescent, stable and uniformly distributed in cells, and well
retained in cells. The substantially improved optical properties and
cell retention of LZQ compared with existing halide indicators (like
SPQ, 6-methoxy-N-ethylquinolinium,
MQAE) permitted measurements of CFTR-mediated
Cl
/I
exchange with excellent signal-to-noise ratio without the need for
signal detection by microscopy.
LZQ fluorescence was quenched by
I by a collisional
mechanism in which its nanosecond fluorescence lifetime decreased in
proportion to quantum yield. A collisional quenching mechanism implies
a rapid, reversible response of LZQ fluorescence to changes in
[I
] without
static LZQ-I
binding.
Recently, analysis of the quenching mechanism of quinolinium-type indicators by anions indicated a charge-transfer mechanism in which
electron transfer occurs during the formation of a transient indicator-anion complex (13). Because the efficiency of a
charge-transfer quenching mechanism depends on anion redox potential
(19, 23), quinolinium indicators are quenched significantly by a wide
variety of anions, including intracellular organic anions and proteins. In contrast, the selective quenching of LZQ fluorescence by
I
suggests that
fluorescence quenching is dominated by a heavy-atom quenching mechanism
rather than a charge-transfer mechanism (17). The specificity of LZQ
fluorescence quenching by I
was supported by the similar nanosecond fluorescence lifetimes of LZQ
in cells vs. water and by the relative insensitivity of intracellular
LZQ fluorescence to changes in cell volume. The specificity of LZQ
fluorescence quenching constitutes a distinct advantage over SPQ.
LZQ was readily loaded into the aqueous compartment of cytoplasm by a
brief hypotonic shock or by overnight incubation. Loading by either
method produced brightly stained cells with excellent spatial
uniformity, as judged by confocal microscopy for epithelial cells. LZQ
has the most uniform intracellular distribution of the many dyes we
have studied. Time-resolved anisotropy measurements indicated that the
majority of LZQ molecules rotated freely and thus were not bound to
cytoplasmic components. Therefore, apart from its use as an
intracellular halide indicator, LZQ appears to be an excellent
aqueous-phase probe for analysis of cytoplasmic rheology (9) and
comparative dye distribution. The calibration of intracellular LZQ
fluorescence vs. I
concentration gave a Stern-Volmer constant of 26 M
1, indicating that LZQ
fluorescence is 50% quenched by 38 mM
I
. The lower apparent
Stern-Volmer constant for LZQ quenching by I
in cells vs. water may be
related to the crowded/mildly viscous intracellular environment and/or
to imperfect equalization of cytoplasmic and external
I
in the calibration
procedures. The similar fluorescence lifetimes of LZQ in cells vs.
water indicates that LZQ fluorescence quenching by components other
than I
is not responsible
for the decreased apparent Stern-Volmer constant. In any case, the
strong sensitivity of LZQ fluorescence to intracellular I
permits measurements of
Cl
/I
exchange using <10 mM I
.
The high intracellular fluorescence of LZQ in combination with its
red-shifted fluorescence spectra and
I
selectivity make it
superior to quinolinium-based
Cl
-indicators for
measurements of CFTR function.
Although we believe that LZQ is the best available halide indicator for
measurement of CFTR function, a few potential concerns are noted. This
report established the application of LZQ in a limited number of cell
types. LZQ loading, leakage, and
I sensitivity should be
established in each cell type and tissue studied. Although the
measurement of CFTR-mediated
I
transport (rather than
Cl
transport) is an
advantage because the non-CFTR cotransporters are generally
Cl
selective, possible
inhibitory effects of I
on
CFTR function have been noted (27). Despite this potential concern, the
majority of reported CFTR functional measurements utilizing SPQ
fluorescence or tracer efflux use
I
transport protocols
(reviewed in Ref. 18). In addition, because LZQ is a single-wavelength
indicator, it is not suitable for ratio imaging as would be required
for measurements of I
transport by cell cytometry. As accomplished recently for the quinolinium-based Cl
indicators (12), chemical modification and/or chromophore conjugation will be needed to develop dual-wavelength
I
indicators based on the
LZQ chromophore.
Notwithstanding these concerns, LZQ should find applications in studies
of CFTR gene delivery and drug screening. The use of SPQ has been
problematic for CFTR transport in freshly isolated nasal or tracheal
epithelial cells in gene therapy trials (reviewed in Ref. 18). The
efficient incorporation of LZQ and its bright fluorescence in cells
should make attractive the use of a fluorescent indicator assay as a
surrogate marker for functional CFTR gene delivery. The application of
LZQ for measurement of CFTR-mediated halide transport using automated
microplate reader technology establishes the basis for high-throughput
drug screening. The correction of F508 CFTR mistrafficking by low
temperature (8), chemical chaperones (3), and other agents (14) raises
the exciting possibility that high-potency modifiers of the cystic fibrosis phenotype might be identified. The data here were obtained on
a standard commercial fluorescence plate reader without modification. Substantial improvements in signal-to-noise ratio and assay sensitivity are anticipated on optimization of light source and detector stability, light collection optics, and fluid addition hardware.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Joachim Biwersi for advice and assistance in the fluorescence studies and manuscript preparation and Dr. Robert Beall of the National Cystic Fibrosis Foundation for direction, encouragement, and support.
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FOOTNOTES |
---|
This work was supported by grants from the National Cystic Fibrosis Foundation, by National Institutes of Health (NIH) Grants DK-43840, HL-60288, HL-59198, and DK-35124, and by NIH Gene Therapy Core Center Grant DK-47766.
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: A. S. Verkman, 1246 Health Sciences East Tower, Cardiovascular Research Institute, Univ. of California, San Francisco, San Francisco, CA 94143- 0521 (E-mail: verkman{at}itsa.ucsf.edu; http://www.ucsf.edu/verklab).
Received 28 April 1999; accepted in final form 25 June 1999.
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