Laboratory of Pharmacology and Chemistry, National Institute of
Environmental Health Sciences, National Institutes of Health, Research
Triangle Park, North Carolina 27709
Quinacrine uptake
and distribution were studied in a primary culture of rat choroid
plexus epithelial cells using conventional and confocal fluorescence
microscopy and image analysis. Quinacrine rapidly accumulated in cells,
with steady-state levels being achieved after 10-20 min. Uptake
was reduced by other organic cations, e.g., tetraethylammonium (TEA),
and by KCN. Quinacrine fluorescence was distributed in two cytoplasmic
compartments, one diffuse and the other punctate. TEA efflux
experiments indicated that more than one-half of intracellular organic
cation was in a slowly emptying compartment. The protonophore monensin
both emptied that TEA compartment and abolished punctate quinacrine
fluorescence, suggesting that a large fraction of total intracellular
organic cation was sequestered in acidic vesicles, e.g., endosomes.
Finally, quinacrine-loaded vesicles were seen to move within the
cytoplasm and to abruptly release their contents at the blood side of
the cell; the rate of release was greatly reduced by the microtubule disrupter nocodazole.
compartmentation; confocal microscopy; endosomes; microtubules; monensin; vesicle fusion
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INTRODUCTION |
THE CHOROID PLEXUS ACTIVELY transports xenobiotics and
metabolic wastes from cerebrospinal fluid (CSF) to blood for subsequent elimination via urine or bile (reviewed in Refs. 5 and 8). Like kidney
and liver, choroid plexus possesses a specific transport system for
organic cations and weak organic bases. However, compared with kidney
and liver, little is known about the cellular mechanisms that drive
organic cations across choroid plexus. One reason for this is that
small tissue size, complex morphology, and inaccessibility have been
significant impediments to the study of transport mechanisms in choroid
plexus. To circumvent these difficulties, Villalobos et al. (10)
recently developed a procedure to isolate and culture choroid plexus
epithelial cells from neonatal rats. By morphological, biochemical, and
functional criteria, these cells were shown to grow with the apical
(CSF) side facing the medium and the blood side attached to the
support. Villalobos et al. (10) also demonstrated that these choroid
plexus cell monolayers grown on a solid support, like choroid plexus
slices from neonatal and adult rats, exhibit specific and concentrative
uptake of the model organic cation tetraethylammonium (TEA). Because of
the orientation of the cells in the monolayer, the uptake of TEA
corresponds to the first step in transport of organic cations from CSF
to blood.
In the present study, conventional and confocal fluorescence microscopy
and digital image analysis were used to investigate the second step in
organic cation transport across choroid plexus cells, movement through
the cytoplasm. To do this, we followed the uptake and intracellular
distribution of a fluorescent organic base, quinacrine. The data show
that quinacrine uptake was mediated by the same process responsible for
TEA uptake. Once in the cells, quinacrine partitioned between cytoplasm
and an acidic vesicular compartment, which accounted for a substantial
fraction of total organic cation accumulation. Quinacrine-loaded
intracellular vesicles were mobile and appeared to release their
contents at the blood side of the cell.
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MATERIALS AND METHODS |
Chemicals.
[14C]TEA bromide (53 mCi/mmol) was obtained from American Radiolabeled Chemicals (St. Louis,
MO). Quinacrine, TEA, and nocodazole were purchased from Sigma Chemical
(St. Louis, MO). All other chemicals were obtained from commercial
sources and were of the highest quality available.
Cell culture.
Three- to five-day-old Fischer rats reared in the animal facility at
the National Institute of Environmental Health Sciences (Research
Triangle Park, NC) were used in these studies. Animals were
anesthetized in the cold with CO2
before decapitation and removal of the brain. For each preparation,
lateral and fourth plexuses from a total of 30-36 neonatal rats
were removed and placed in ice-cold DMEM-Ham's F-12 medium (DMEM/F-12)
supplemented with penicillin (100 U/ml) and streptomycin (100 µg/ml). Cell isolation procedures are given in detail by
Villalobos et al. (10). Isolated epithelial cells were suspended in MEM
with D-valine substituted for L-valine and
with 10% Nu-Serum IV and growth promoters (triiodothyronine,
PGE1, forskolin, and epidermal
growth factor). Cells were plated at a density of 4.5 × 105
cells/cm2 on individual wells of
24-well tissue culture plates
([14C]TEA
uptake), glass dual-chamber microscope slides
(immunostaining), or 4 × 4-cm glass coverslips in culture dishes
(quinacrine accumulation). Cells were maintained at 37°C in
humidified 95% air-5% CO2. On day
3, unattached cells were removed as
the initial plating medium was replaced with medium containing 5%,
rather than 10%, Nu-Serum IV. From
day 5 on, cells were maintained with DMEM/F-12 medium containing 5% Nu-Serum
IV and growth promoters (10). Medium was changed every 2-3 days.
Cells were used for experiments on days
9-11.
Conventional fluorescence microscopy.
Glass coverslips (4 × 4 cm) with attached cells were mounted in
covered Bionique chambers under an atmosphere of 95% air-5% CO2. The medium in the chamber was
an artificial CSF (aCSF; in mM: 118 NaCl, 3 KCl, 0.7 Na2PO4,
18 NaHCO3, 2 urea, 0.8 MgCl2, 1.4 CaCl2, and 12 glucose, pH 7.4),
which also contained 5 µM quinacrine and, when indicated, transport
inhibitors. Experiments were carried out at room temperature. The
chamber containing cells was placed on the stage of a Nikon Diaphot
inverted microscope fitted with epifluorescence optics, fluorescence
objectives [Nikon ×40, numerical aperture (NA) 1.4; Olympus
×60 oil, NA 1.3], a 100-W mercury lamp, and a fluorescein
filter set (Nikon B-1A; 460- to 485-nm band-pass excitation filter,
510-nm dichroic filter, and 515-nm long-pass emission filter). To
minimize photobleaching, a neutral density filter that passed only 1 or
10% of the excitation light was kept in the light path, and
fluorescence measurements were made over periods of ~1-2 s.
Epifluorescence images were acquired through the microscope side port
by use of a Hamamatsu 2400 or a Paultek charge-coupled device (CCD)
video camera connected to an 8-bit video image capture card (Scion
Video Image LG-3 with 4 megabytes of onboard memory) in an Apple
Macintosh Centris 650 computer. The video card could capture and
average up to eight full frames (640 × 480 pixels) at the video
rate (30 frames/s). Incoming images were displayed on a high-resolution
computer monitor (Apple) using image capture and analysis software
[National Institutes of Health (NIH) Image 1.58], and
8-frame averages were computed and stored on an Olympus optical disk
recorder for later analysis.
To make a measurement, dye-loaded cells in the chamber were viewed
under reduced, transmitted light illumination. A field was selected,
and an epifluorescence image was acquired by averaging eight frames. We
have found using confocal and conventional video microscopy systems and
glass capillary tubes filled with solutions of known concentrations of
fluorescent solutes that the relationship between image fluorescence
and dye concentration is approximately linear (Ref. 3 and Miller,
unpublished data). Calibration of the present systems with quinacrine
showed a similar linear relationship over a 100-fold range of
concentrations, from 0.25 to 25 µM. However, with concentrations
>25 µM, the relationship became nonlinear, suggesting
self-quenching. For example, when the quinacrine concentration was
raised from 25 to 50 µM, measured fluorescence increased by 70%
rather than by 100%. Quinacrine fluorescence was also pH dependent, e..g., reducing buffer pH from 7.4 to 6.5 decreased fluorescence by
60%. Because of the many uncertainties in relating cellular fluorescence to actual compound concentration in cells with complex geometry, data are reported here as average measured pixel intensity rather than as estimated fluor concentration.
Confocal fluorescence microscopy.
Cells in a Bionique chamber were mounted on the stage of a Zeiss model
410 inverted laser scanning confocal microscope and viewed through a
Zeiss ×100 oil immersion objective (NA 1.4). To collect
fluorescent images, quinacrine-loaded cells were illuminated by an
Ar-Kr laser at 488 nm. A 510-nm dichroic filter was positioned in the
light path, and a 515-nm long-pass emission filter was placed in front
of the detector. In some experiments, cells were loaded with quinacrine
and the mitochondrial probe rhodamine 123. These cells were illuminated
by both 488- and 568-nm laser lines. A double dichroic filter was used;
quinacrine fluorescence was collected through a 515- to 565-nm
band-pass filter and rhodamine fluorescence through a 590-nm long-pass filter.
Each collected image was a single 8-s scan. To minimize photobleaching,
images were collected at 20% laser power with neutral density filters
passing only 3-30% of the light. Preliminary experiments showed
that, under these conditions, tissue autofluorescence was undetectable
and photobleaching was minimal, i.e., fluorescence intensities in cells
were reduced by <5% in consecutive 8-s scans. Confocal images (512 × 512 × 8 bits) were viewed on a high-resolution monitor,
saved to optical disk, and transferred to the Power Macintosh computer
for analysis.
Image analysis.
Average cellular quinacrine fluorescence was determined from stored
images. Each cell was outlined, and the average fluorescence intensity
was measured using NIH Image software as described previously (2). One
must keep an important caveat in mind when interpreting these
measurements of cellular fluorescence intensity. The fluorescence signal from a probe is sensitive to probe environment, e.g., pH and
solvent polarity. As a result, the relationship between probe fluorescence and concentration could and does (see
Quinacrine accumulation) vary with the region of
the cell.
Two procedures were used to analyze vesicle dynamics. First, the
movement of fluorescent vesicles within cells was followed over time
using confocal microscopy. Cells were incubated for 30 min in medium
with 5 µM quinacrine. The microscope was focused at roughly the
center of the cells (z-axis), and
confocal images were acquired at 10-s intervals over a total of 5 min.
The motion of individual fluorescent vesicles could be followed over
time by displaying the sequence as a movie. To display this
three-dimensional data set in two dimensions, the stack of images was
projected onto a single plane using a Silicon Graphics Octane
workstation and Vox Blast software (Vaytek). This software also allowed
us to treat the stack as a volume (with
x, y,
and time as the three dimensions) and to tilt the volume
around the x-axis and visualize the
displacement (tracks) of individual fluorescent vesicles through time.
Second, rates of vesicle "flashing" were measured using
conventional fluorescence video microscopy. Cells were loaded in medium with 5 µM quinacrine. The plane of focus was at the cell-coverslip interface, and images of a field of one or two cells were acquired at
one-half the video rate (15 frames/s) over a total of 5-10 s. To
determine flashing rate, each image in a sequence was subtracted pixel
by pixel from the next image (NIH Image). The resulting difference
image was scaled to an average pixel intensity of 128. Flashes were
evident in difference images as areas of black on a field of gray. The
resulting time series of difference images could then be viewed as a
movie, and the number of black areas that appeared could be counted.
TEA uptake and efflux.
On day
11, uptake of
[14C]TEA by cells
plated on 24-well tissue culture plates was measured as described
previously (10). Briefly, cells were rinsed and preincubated in aCSF
for 1 h at 37°C. Transport was initiated by replacement of
preincubation buffer with 1 ml aCSF containing the labeled substrate;
cells were incubated for 0-90 min. All incubations were conducted
at 37°C under 95% air-5%
CO2. To terminate uptake,
transport buffer was removed, and the cells were rinsed with 3 ml
isotope-free aCSF. Within the transport well, cells were solubilized in
1 N NaOH for 1 h and neutralized with 1 N HCl. An aliquot of the
solubilized cell suspension (800 µl) was retained for determination
of protein by a Bio-Rad microassay using BSA as standard. The remainder
of the cell suspension was transferred to a scintillation vial for counting. Cellular TEA was calculated in picomoles per milligram protein.
For efflux experiments, cells were loaded with 50 µM
[14C]TEA as described
above, rinsed twice with TEA-free aCSF, and incubated in TEA-free aCSF.
At each sampling time, an aliquot of the efflux medium was removed, and
the cells were rinsed and processed for counting as described above.
Immunostaining.
Cells on glass tissue culture chamber slides (Nunc, Naperville, IL)
were fixed for 10 min in 2% formaldehyde and 0.1% glutaraldehyde in
PHEM buffer (in mM: 60 PIPES, 24 HEPES, 5 EGTA, and 1 MgSO4, pH 6.9), permeabilized with
1% Triton X-100 in PHEM buffer for 10 min, and exposed to a primary
polyclonal, anti-tubulin antibody (rabbit; Sigma) and
fluorescein-labeled secondary antibody (Kirkegaard and Perry) as
described previously (10). Slides were sealed with Agua-PolyMount
(Polysciences, Warrington, PA) and allowed to dry overnight in the
dark. Immunostained cells were viewed using the Zeiss model 410 confocal scanning laser microscope.
Statistics.
Data are given as means ± SE. Means were considered to be
statistically different when P < 0.05 as determined by the appropriate paired or unpaired
t-test.
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RESULTS |
Quinacrine uptake and distribution.
A representative conventional epifluorescence micrograph of choroid
plexus cells that had been incubated for 10 min in medium containing 5 µM quinacrine is shown in Fig.
1A.
Notice that the distribution of fluorescence within the cells is not
uniform. There are three components to cellular fluorescence: one is
nuclear, a second is cytoplasmic and diffuse, and a third is
cytoplasmic and punctate. Although punctate fluorescence is seen
throughout the cytoplasm, it is most concentrated in the perinuclear
region, which is the thickest part of the cell. Figure
1B shows that adding the organic
cation TEA to the medium substantially reduced overall cellular
fluorescence. From these micrographs, it appears that TEA had the
greatest effect on the fluorescence intensity of the punctate
compartment.

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Fig. 1.
Conventional epifluorescence micrographs of choroid plexus cells after
10 min of incubation in medium with 5 µM quinacrine
(A) or 5 µM quinacrine + 1 mM
tetraethylammonium (TEA; B).
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Confocal microscopy confirmed these general features of quinacrine
distribution in control cells. Comparison of differential contrast
interference (DIC) and confocal images of quinacrine-loaded cells
showed that each punctate site of fluorescence could be localized to a
vesicular structure in the DIC image (Fig.
2). However, not all vesicular structures
seen in the DIC image were fluorescent. To further characterize the
vesicular compartment that accumulated quinacrine, cells were incubated
to steady state in medium with 5 µM quinacrine and then exposed to
0.5 µM rhodamine 123, a fluorescent dye that accumulates in
mitochondria. Figure 3 shows a
representative image of double-labeled cells. The quinacrine and
rhodamine 123 labeling patterns differed in two important respects:
1) punctate sites of localization
appeared to be compact for quinacrine (red image), whereas they were
elongated for rhodamine 123 (green image), and
2) little colocalization of the
labels [which would appear as yellow pixels in the red-green-blue
(RGB) image] could be seen. Clearly, mitochondria did not
accumulate quinacrine.

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Fig. 2.
Differential interference contrast
(left) and confocal
(right) images of quinacrine-loaded
choroid plexus cells.
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Fig. 3.
Double-labeled confocal image of choroid plexus cells. Cells were
incubated to steady state in medium with 5 µM quinacrine and then
briefly exposed to 0.5 µM rhodamine 123. This image shows quinacrine
labeling in red and rhodamine 123 labeling in green. Areas of
colocalization appear as yellow. Rhodamine 123 labeled elongated
structures, whereas quinacrine labeled roughly circular structures.
Little colocalization is evident, i.e., the 2 dyes appear to label
different structures.
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We acquired conventional epifluorescence images from cells incubated in
medium containing 5 µM quinacrine, and from these we measured average
cellular fluorescence intensity. Figure 4 shows the time course of quinacrine accumulation. Cellular fluorescence increased initially and reached a steady state within 10-20 min. At steady state, the model substrate for the renal organic cation transport system, TEA, reduced average cellular fluorescence by >50%
and the metabolic inhibitor, KCN, reduced fluorescence by ~75%.
Other experiments showed that average, steady-state cellular fluorescence was reduced in a concentration-dependent manner by TEA
(Fig.
5A).
Significant reductions in steady-state cellular fluorescence were also
seen with verapamil, darstine, paraquat, and tetrapentylammonium
(TPA), all organic cations (Fig. 5). The organic anion
p-aminohippurate, at 1 mM, was without
effect (not shown). From these data, we can construct a rough order of
inhibitory potency: verapamil = paraquat = TPA > TEA > darstine. We verified certain of these results using confocal
microscopy. Cells were incubated in medium with 5 µM quinacrine
without or with 100 µM TEA or 100 µM TPA, and confocal images were
acquired after 30 min. Average, steady-state cellular fluorescence
intensity was reduced 44 ± 3% by TEA and 55 ± 2% by TPA;
these values are similar to those seen in experiments with conventional
optics. In those cultures exposed to TEA or TPA, confocal microscopy
showed that fluorescence intensity in both the diffuse and punctate
compartments was reduced relative to controls. For example, in control
cells, fluorescence intensities in the cytoplasm and in the punctate compartment averaged 57 ± 6 and 176 ± 19 fluorescence units,
respectively; in cells incubated with 100 µM TEA, corresponding
values were 35 ± 4 and 95 ± 10 fluorescence units (both
significantly lower than controls, P < 0.01).

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Fig. 4.
Time course of quinacrine accumulation by choroid plexus cells. Cells
were incubated in medium with 5 µM quinacrine, without (control) or
with TEA or KCN. At times indicated, conventional fluorescence images
were acquired. Results (means ± SE of fluorescence intensity for
30-47 cells) are from 1 experiment representative of experiments
carried out with 2 cultures. TEA and KCN significantly reduced
fluorescence intensity at all times tested
(P < 0.01).
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Fig. 5.
Inhibition of quinacrine uptake by organic cations. Cells were
incubated for 30 min in medium with 5 µM quinacrine, without
(control) or with indicated additions (concentrations in µM). Each
set of bars shows results (means ± SE of fluorescence intensity for
35-50 cells) from 1 experiment representative of experiments
carried out with at least 2 cultures. All organic cations tested
significantly reduced fluorescence intensity
(P < 0.01). TPA,
tetrapentylammonium; VERAP, verapamil; DARST, darstine; PARAQ,
paraquat.
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It is important to note that, for these choroid plexus cells in
culture, the same general order of inhibitor effectiveness was found in
the present experiments in which quinacrine accumulation was followed
with fluorescence microscopy and in experiments in which
[14C]TEA uptake was
measured (10). This agreement and the finding that quinacrine
accumulation was reduced by TEA suggest that uptake of quinacrine and
TEA may be mediated by the same carrier protein. To investigate this
point further, we determined the effects of quinacrine on the 30-min
uptake of 10 µM
[14C]TEA. Quinacrine
at 50 and 100 µM reduced TEA uptake by 18 and 40%, respectively.
Taken together, the inhibition data for quinacrine accumulation
(present study) and for TEA uptake (Ref. 8 and present study) indicate
that choroid plexus cells grown on solid supports accumulate both
substrates through a common transporter specific for organic cations.
Organic cation efflux.
Both conventional and confocal fluorescence images of quinacrine
distribution in choroid plexus cells showed that a substantial fraction
of cellular fluorescence was associated with a punctate compartment.
Previous studies from this laboratory suggest an identity for that
compartment. Using endosomes isolated from rat renal cortex, Pritchard
et al. (6) demonstrated organic cation (TEA) uptake that was specific
and ATP dependent and was abolished by protonophores. The mechanism of
organic cation accumulation involved acidification of the
intraendosomal space by a proton ATPase and TEA/proton exchange. To
determine whether accumulation by an acidic vesicular compartment might
contribute to organic cation accumulation in choroid plexus cells, we
loaded cells to steady state in medium with 10 µM
[14C]TEA, removed the
cells to TEA-free medium, and followed cell TEA content for 30 min in
the absence (control) and presence of the protonophore monensin. Figure
6 shows that, in controls, TEA efflux was
initially rapid but then slowed. From 15 to 30 min, little additional
TEA was lost from the cells, and <50% of total cellular TEA had been
lost to the bath after 30 min. In contrast, when monensin was added 10 min into the experiment, TEA efflux subsequently increased, and at 30 min the total amount lost was twice that in cells not exposed to
monensin (Fig. 6). Even without detailed kinetic analysis, the control
data indicate that about two-thirds of cellular TEA was in a cellular
compartment from which efflux was very slow. Monensin rapidly released
TEA from that compartment.

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Fig. 6.
Time course of efflux of TEA from choroid plexus cells. Cells were
first loaded with 50 µM
[14C]TEA. After 30 min, they were washed twice in TEA-free medium and then incubated in
TEA-free medium; in 1 set of cultures, 5 µM monensin was added at 10 min (arrow). TEA content of medium samples and cell extracts was
determined by liquid scintillation counting, and data (means ± SE
of 3 experiments) are expressed as amount of TEA remaining in cells.
Monensin significantly reduced TEA content at both times tested
(P < 0.01).
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Fluorescence microscopy of quinacrine-loaded cells incubated in
quinacrine-free medium showed that monensin had two effects on
intracellular fluorescence distribution patterns (Fig.
7). First, monensin rapidly increased
overall cellular fluorescence. Preliminary measurements of average
cellular fluorescence showed that it had increased two to three times
within 2 min of monensin exposure. This increase in cellular
fluorescence occurred even though cells were in quinacrine-free
medium. Second, monensin greatly reduced vesicular fluorescence
(Fig. 7). Because quinacrine fluorescence is quenched by low pH and at
high concentrations (see Conventional fluorescence
microscopy), the increase in overall cellular
fluorescence caused by monensin most likely reflects alkalinization of
the vesicular compartment by the protonophore and movement of
quinacrine into the cytoplasm, where fluorescence was no longer
quenched.

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Fig. 7.
Effects of monensin on quinacrine fluorescence in choroid plexus cells.
Cells were loaded for 30 min in medium containing 5 µM quinacrine and
then transferred to quinacrine-free medium with no additions (control)
or with 20 µM monensin. A:
conventional epifluorescence image of control cells 5 min after
transfer. Substantial diffuse and punctate fluorescence is evident.
B: conventional epifluorescence image
of monensin-treated cells 5 min after transfer. Note loss of punctate
fluorescence and increase in overall cellular fluorescence intensity.
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Vesicle dynamics.
To document the movements of quinacrine-containing intracellular
vesicles, choroid plexus cells were incubated to steady state in medium
with 5 µM quinacrine and then observed using fluorescence microscopy.
Two time frames were employed: in the slow time frame, events occurring
over tens of seconds to minutes were followed; in the rapid time frame,
events occurring over tens of milliseconds were followed.
Confocal microscopy was used to record the slow movement of fluorescent
vesicles. In these experiments, a single optical section was taken
every 3-10 s at a level of the cell that was 2-4 µm above
the coverslip, roughly at the middle of the cell. These time sequences
were displayed sequentially on the computer monitor as a movie showing
vesicle dynamics within a narrow volume of focus (depth <1 µm).
About 10-30% of the vesicles moved little in recordings lasting
2-5 min. The remaining vesicles showed several types of movement.
About 50% of the vesicles exhibited saltatory movements, which
resulted in no obvious net change in position within an optical slice.
About 20-40% of the vesicles clearly changed position. Some were
lost from view, suggesting that they had moved out of the optical
section. Some appeared suddenly, suggesting that they had moved into
the section from a volume of cytoplasm above or below. Others moved
within the plane. For some of these, the path of movement through the
cytoplasm appeared to be nearly linear, suggesting vesicle movement
along a cytoskeletal element.
Figure 8 shows some elements of vesicle
motion from one time sequence of confocal images. The first image gives
the position of vesicles at the start of the sequence (Fig.
8A). The second image is a
projection of all 30 images onto a single plane; it shows the
integrated movement of these vesicles over 90 s as lines or tracks
(Fig. 8B). Also shown are plots of
the movement of four of the vesicles (Fig.
8C) and of their linear velocities
for each sampling period (Fig. 8D).
Vesicles
1 and
2 show directed movement with a total
displacement in the 10- to 20-µm range.
Vesicles 3 and
4 show little net displacement but
rather appear to move within a restricted area. In spite of the
differences in net displacement, linear velocities calculated for each
sampling period are comparable for the four vesicles. Clearly, this
representative sequence of confocal images shows that, on the time
scale of tens of seconds, a substantial fraction of quinacrine-loaded
vesicles move within the cytoplasm.

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Fig. 8.
Vesicle motion within quinacrine-loaded choroid plexus cells. Cells
were incubated for 30 min in medium with 5 µM quinacrine. Confocal
images of a single field were acquired at 3-s intervals for a total of
1.5 min (plane of focus at nucleus; 1.4-s scans of a 174-line region of
interest). A: first image of time
sequence. Numerous fluorescent vesicles are evident in cytoplasm.
B: a projection of all 30 images onto
a single plane. Vesicle motion within optical section is seen as a
series of white tracks, some of which extend for several µm.
C: analysis of movement of individual
vesicles. Four vesicles were selected
(vesicles
1-4),
and their positions were followed and plotted.
D: linear velocities of
vesicles
1-4
for each consecutive sampling period.
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In initial experiments concerned with more rapid elements of vesicle
dynamics, we used conventional epifluorescence optics to follow cells
for periods of 5-10 s under continuous illumination from a Hg lamp
(images collected at 15 frames/s). With appropriate neutral density
filters to attenuate the light and a sensitive CCD camera to monitor
cell fluorescence, little photobleaching of quinacrine was observed. In
the course of these experiments, we were surprised to see repeated,
local, transient spreading of fluorescence that appeared to arise from
some of the vesicles. These were most evident when a
high-numerical-aperture objective was focused at the level of the
cell-glass support interface. In these polarized cells, the plane of
focus was at the plasma membrane that would face the blood side of the
choroid plexus epithelium. Because of the transient nature of these
changes in fluorescence pattern and because of an initial increase in
local fluorescence, we called the phenomenon flashing. Figure
9 shows a representative sequence of video
frames acquired at 15 frames/s. The arrows indicate areas in which
flashes have occurred. The flashes appeared to arise from single
vesicles or clusters of vesicles and remained visible for two or three
frames (100-200 ms). Close examination of electronically magnified
image sequences showed that the phenomenon involved two phases, an
initial increase in fluorescence followed by a rapid decline (Fig.
10A).
Other vesicles in close proximity do not appear to be altered (Fig. 10,
B-F).

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Fig. 9.
Sequence of video images demonstrating phenomenon of "flashing."
Cells were incubated for 30 min in medium with 5 µM quinacrine and
viewed continuously with conventional epifluorescence optics. Plane of
focus was at cell-coverslip interface. Images were acquired at rate of
15 frames/s for 7 s and numbered sequentially. Arrows point to regions
within cell where fluorescence intensity has transiently increased.
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Fig. 10.
Fate of single quinacrine-loaded vesicles.
Top: sequence of images shows a
magnified and enhanced region from a time series collected as in Fig.
7. Arrow points to a vesicle that exhibits an increase in fluorescence
intensity followed by an abrupt decrease. Notice that fluorescence
intensities of neighboring vesicles appear unchanged.
Bottom: vesicle fluorescence (average
measured pixel intensity) vs. time for vesicle indicated by arrow at
top
(A) and other vesicles nearby
(B-F).
Increase in fluorescence intensity in
A is limited by signal saturation at a
value of 255. Broken line in A
indicates background level of diffuse cytoplasmic fluorescence.
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Confocal microscopy was used to characterize the flashing phenomenon
further. Although confocal optics increase spatial resolution in three
dimensions and allow optical sectioning of cells, for most confocal
systems the price one pays for enhanced spatial resolution is limited
temporal resolution. For our microscope, useful full-frame images (512 scan lines) of quinacrine-loaded choroid plexus cells could only be
obtained at a rate of one every 2-4 s, too slow to be able to
investigate the flashing seen with conventional optics. However,
fluorescence from a single laser scan line could be imaged on a
millisecond time scale. Figure 11 shows
the results of scanning, at high magnification, a single line across a
quinacrine-loaded choroid plexus cell every 5 ms for a total of 2.5 s.
In this experiment, the microscope was focused at the interface between
the cell and the coverslip, and the confocal pinhole was closed down so
that the axial resolution of the system was maximal (0.5-1 µM).
The position of the scan line was chosen to include several fluorescent
vesicles, and each vertical line shows the history of one or more
vesicles over 2.5 s (512 scans). Many of the lines appear
uninterrupted, with little change in fluorescence intensity or position
with time. However, several show an abrupt broadening followed by a
sharp decline in fluorescence. The broadening represents the spread of
fluorescence into the cytoplasm surrounding the vesicle. In some cases,
punctate fluorescence is no longer detectable after this broadening.
Fluorescence intensity scans of single-vesicle tracks are shown in Fig.
12. Each track shows a roughly constant
level of fluorescence interrupted by a sharp decline. A small,
short-lived increase in local fluorescence is evident just before the
decline. Note that no flashing was evident when the plane of focus was
raised several micrometers into the cells (not shown). These confocal
images show the same vesicle-derived flashing seen with conventional
optics; they again demonstrate that the phenomenon was localized to the
region near or on the membrane at the blood side of the cells.

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Fig. 11.
Confocal analysis of flashing. Repeated scan of a single line across a
quinacrine-loaded cell. Plane of focus was at cell-coverslip interface,
and confocal pinhole was narrowed to give a thin optical section (<1
µm in depth with 1.4-numerical-aperture, ×100 objective used).
Scans are 2.5 ms apart. Vesicle histories appear as vertical lines.
Each flash is evident as a lateral spreading of fluorescence followed
by an abrupt decrease in fluorescence intensity.
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Using conventional fluorescence optics, the video system, and digital
image analysis, we collected time series of fluorescent images at 15 frames/s and, from these, measured the rate of flashing in fields of
choroid plexus cells. In all of these experiments, the microscope was
focused at the interface between the cells and the coverslip support.
To determine the rate of flashing within a time series, each image was
subtracted pixel by pixel from the next image in the series. Flashes
showed up in the difference images as areas of black on a field of
gray. The resulting time series of difference images could then be
viewed as a movie. The number of black spots arising during this movie
was taken as the number of flashes. Figure
13 shows flashing rates from a typical experiment in which choroid plexus cells were incubated in medium with
5 µM quinacrine and 2- to 5-s time series were acquired at various
times. The rate of flashing increased over the first 15 min of exposure
to quinacrine and then reached a plateau. On the plateau, the rate of
flashing was ~6 s
1. In 53 experiments using 32 coverslips from 8 cultures, the average control
rate after 15-45 min of exposure to quinacrine ranged from 3.5 to
8.2 s
1, with a grand mean
of 5.3 s
1.

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Fig. 13.
Analysis of flashing rate using conventional fluorescence microscopy.
Cells were incubated for time indicated in medium with 5 µM
quinacrine and viewed with conventional epifluorescence optics. Plane
of focus was at cell-coverslip interface. At each time, images were
acquired at rate of 15 frames/s for 5-7 s. To determine flashing
rate, each image in a sequence was subtracted pixel by pixel from next
image. Resulting difference image was scaled to an average pixel
intensity of 128. Flashes were evident in difference images as areas of
black on a field of gray. Resulting time series of difference images
could then be viewed as a movie, and black areas were counted.
A: representative pair of sequential
images (left and
middle) and resulting difference
image (right). Arrow, flash region.
B: rate of flashing as a function of
time in medium with quinacrine. Data are from 1 experiment that is
representative of experiments carried out with 3 cultures. Each point
gives flashing rate for a single 5- to 7-s sequence of 75-105
images.
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We previously speculated that endosome recycling, guided by
cytoskeletal elements, may contribute to organic cation secretion in
renal proximal tubule (6). To determine whether flashing in organic
cation-loaded vesicles involved microtubules, we loaded choroid plexus
cells to steady state in medium with 5 µM quinacrine, exposed the
cells to nocodazole, a microtubule disrupter, and determined rates of
flashing. Figure
14A
shows that 20 µM nocodazole caused a time-dependent decrease in
flashing rate. A significant reduction in the rate was seen after 20 min, and by 30 min the rate was reduced by 70%. Although 20 µM
nocodazole reduced flashing by >50%, it had only a small effect on
average cellular fluorescence (10-20% reduction; data not shown).
Figure 14B shows that nocodazole effects on flashing were concentration dependent, with a significant reduction occurring at 5 µM and a further reduction at 20 µM.

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Fig. 14.
Effects of nocodazole on flashing rate.
A: cells were preincubated in medium
containing 5 µM quinacrine. After 20 min, a short time sequence of
images was acquired as described in Fig. 11, and then nocodazole was
added to a concentration of 20 µM. Additional sequences were acquired
at times indicated. B: cells were
preincubated in medium containing 5 µM quinacrine for 20 min. Then
0-20 µM nocodazole was added, and image sequences were acquired
after 30 min. All image sequences were analyzed as described in Fig.
11. Each point represents mean ± SE of flashing rate for 4-6
cultures.
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To establish that the concentrations of nocodazole used here actually
disrupted microtubules in choroid plexus cells, we incubated cells for
30 min in medium without (control) and with 5-10 µM nocodazole,
fixed and permeabilized the cells, and immunostained them using an
anti-tubulin primary antibody and a fluorescein-labeled secondary
antibody. Antibody distribution was visualized using confocal
microscopy. Control cells exhibited a well-developed filamentous
network of microtubules (Fig.
15A).
Nocodazole at 5 and 10 µM caused nearly total disruption of this
network (Fig. 15, B and
C). Other experiments with cells
labeled with anti-actin antibodies showed that nocodazole did not
disrupt microfilaments (not shown). Thus the action of this drug
appeared to be specific for microtubules.

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Fig. 15.
Immunostaining of tubulin in control and nocodazole-treated choroid
plexus cells. Cells were exposed to 0 (A), 5 (B), or 10 (C) µM nocodazole for 20 min,
fixed, and then immunostained for tubulin (see
MATERIALS AND METHODS).
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DISCUSSION |
Quinacrine accumulation.
It is well established that choroid plexus epithelial cells possess one
or more specific transporters for organic cations on the apical
(CSF-facing) plasma membrane and that these contribute to the transport
of positively charged xenobiotics and metabolites from CSF to blood (5,
8). However, little is known about subsequent steps in transport. In
the present study, we used a recently developed primary culture of rat
neonatal choroid plexus epithelial cells, a fluorescent organic cation,
fluorescence microscopy, and image analysis to investigate the
mechanisms by which organic cations move across the cellular
compartment. Conventional and confocal microscopy showed that the
distribution of quinacrine fluorescence within choroid plexus cells was
not uniform. Quinacrine was detected in both the nucleus and cytoplasm.
Within the cytoplasm, fluorescence was distributed between diffusive
and punctate compartments. The latter appeared to be vesicular, and the
fluorescence intensity in the vesicular compartment was substantially
higher than in bulk cytoplasm, suggesting accumulation of quinacrine by
these vesicles. A similar pattern of vesicular quinacrine accumulation was also found in intact choroid plexus from adult and neonatal rats
(Miller, unpublished observations). It is not surprising to find a
complex intracellular distribution pattern for quinacrine, since this
weak organic base has been shown to bind to a variety of polyanions,
including RNA, DNA, and ATP, and to accumulate within acidic
intracellular compartments (see, e.g., Refs. 4, 11, 13).
Apical uptake of quinacrine (average cellular fluorescence) by choroid
plexus cells was specific and energy dependent. Uptake was reduced by
several organic cations, including TEA, but not by the organic anion
p-aminohippurate. The order of
inhibitory effectiveness of the tested organic cations on total
quinacrine uptake paralleled that found for uptake of
[14C]TEA by Villalobos
et al. (10). In addition, we found that TEA reduced quinacrine uptake
and that quinacrine reduced
[14C]TEA uptake
(present study). Finally, analysis of confocal micrographs showed that
TEA and TPA reduced quinacrine fluorescence in both the cytoplasmic and
vesicular compartments. Taken together, these results are consistent
with TEA and quinacrine sharing a common, organic cation-specific entry
step at the apical (CSF) side of the choroid plexus cells.
In addition to providing direct evidence for quinacrine accumulation in
a vesicular compartment, the present results also suggest intracellular
compartmentation for the model organic cation TEA.
[14C]TEA efflux
experiments indicated that substantially more than one-half of organic
cation within choroid plexus cells was in a slowly exchanging
compartment that could be the same vesicular compartment that
accumulated quinacrine. By themselves, these findings have important
implications with regard to the energetics of organic cation transport
in choroid plexus. They argue that the cell-to-medium concentration
ratio calculated from tracer uptake experiments greatly overestimates
the actual cytoplasm-to-medium concentration ratio. As a result, cells
would be directing less metabolic energy to organic cation uptake at
the apical plasma membrane and more to uptake by vesicles. In addition,
the actual organic cation concentration gradient available to drive
transport from cytoplasm to blood should be less than that predicted
based on average cellular concentration, possibly requiring a higher than expected direct or indirect input of metabolic energy to overcome
the electrical potential energy barrier to efflux at the blood side
plasma membrane.
What is the nature of the vesicular compartment for organic cations?
Previous experiments with isolated endosomes from rat renal cortex and
liver showed ATP-dependent accumulation of organic cations (6, 9). Data
for endosomes from both tissues were consistent with organic cation
accumulation being due to proton/organic cation exchange that was
energetically coupled to a proton ATPase that acidified the vesicle
interior. Membrane-permeable protonophores, e.g., monensin, both
collapsed the pH gradient across the endosomal membrane and greatly
reduced organic cation uptake (6, 9). In the present experiments with
choroid plexus cells, we used monensin as a tool to implicate an acidic
intracellular compartment in TEA and quinacrine sequestration. TEA
efflux from preloaded cells was resolved into a fast component and a
very slow component, with the latter being the larger of the two. More
than one-half of the TEA in this slow cellular compartment was rapidly
released by monensin, suggesting that TEA was sequestered in an acidic intracellular compartment. It is important to note that, unlike quinacrine, which is a weak base, TEA is an organic cation.
Accumulation of TEA in an acidic compartment cannot be driven by
nonionic diffusion followed by pH trapping of the conjugate acid.
Rather, the present data for choroid plexus cells suggest that, as in
kidney (6) and liver (10), TEA accumulation in the acidic compartment
is specific, possibly mediated by a proton/organic cation exchanger.
As seen in similarly designed quinacrine efflux experiments, monensin
effects on quinacrine fluorescence appeared to be more complicated.
Monensin caused a rapid increase in cellular fluorescence followed by a
decline. In monensin-treated cells, punctate sites of intense
fluorescence were greatly reduced. During the efflux phase of the
experiment, the cells were in quinacrine-free medium, so the increase
in cellular fluorescence could not be due to increased uptake. The data
are consistent with the following explanation. The immediate effect of
monensin was to collapse the pH gradient across the vesicle membrane.
This had two consequences. First, because quinacrine fluorescence is
quenched by low pH (see Conventional fluorescence
microscopy), the intrinsic fluorescence of the dye increased and total cellular fluorescence rose. Second, with the increase in pH of the vesicle interior, the driving force for quinacrine accumulation was dissipated, and quinacrine diffused into
the cytoplasm and then rapidly out of the cells; an additional increase
in intrinsic fluorescence could have occurred as dye was released
(relief of self-quenching). Thus, in TEA and quinacrine efflux
experiments, monensin effects can be interpreted within the context of
the ionophore releasing sequestered organic cation from an acidic,
vesicular compartment, e.g., endosomes.
Note that the results of the quinacrine efflux experiments indicate
that for this fluorescent organic cation measurements of average
vesicular fluorescence clearly underestimate total cell quinacrine
content. As a result, imaging should not be used to quantitate the
actual total uptake of quinacrine by choroid plexus cells or the
distribution of quinacrine among the various cellular compartments. For
quinacrine, imaging should only be considered to be a semiquantitative
tool with which to characterize uptake and intracellular distribution.
Vesicle dynamics.
Fluorescence microscopy and image analysis have also allowed us to
investigate vesicle dynamics within choroid plexus cells. In these
experiments, only those vesicles that concentrated the fluorescent
organic cation, quinacrine, were visible. At present, we do not know
whether other vesicle populations within these cells behave similarly.
On a slow time scale (tens to hundreds of seconds), many
quinacrine-loaded vesicles were seen to move in three dimensions. Two
types of vesicle motion were observed: saltatory motion within a
limited volume and directed movements over several micrometers. We have
observed similar movement of vesicles in experiments with choroid
plexus slices from adult rats. Preliminary experiments with intact
tissue and cells in culture showed that vesicle motion was reduced
following exposure to the microtubule disrupter nocodazole (Miller,
unpublished observations). Further experiments are needed to determine
the mechanistic significance of this result. However, it could signify
that some of the vesicles that accumulate organic cations are
associated with microtubules and that the microtubules and associated
molecular motors provide the respective tracks and engines that direct
vesicle motion (1, 7, 12).
An unexpected finding of the present study was the phenomenon we have
called flashing. This was revealed in time series of images acquired
using both wide field and confocal fluorescence microscopy. Flashing
appeared as an abrupt but transient increase in fluorescence around a
quinacrine-loaded vesicle, followed by a spread of fluorescence through
the immediate area. Each event (local increase in fluorescence followed
by the spread of fluorescence) occurred on a time scale of 100-200
ms. After the flash, the fluorescence intensity of the original vesicle
was abolished or greatly diminished, although fluorescence intensities
in adjacent vesicles remained unchanged. Confocal microscopy confirmed
that flashing occurred in the immediate vicinity of the basal (blood
side) plasma membrane but not several micrometers into the cell
interior. Thus it is unlikely that these flashes represent release of
vesicle contents due to photodynamic damage. Preliminary experiments
with choroid plexus cells loaded with another fluorescent organic
cation, daunomycin, also revealed occasional vesicle-derived, abrupt
spreads of fluorescence; these were, however, more difficult to detect,
since daunomycin fluorescence is relatively insensitive to pH and there
was no initial increase in fluorescence (Miller, unpublished findings). No such phenomenon was seen in cells that were not exposed to fluorescent organic cations. Finally, nocodazole both disrupted the
microtubular network and reduced the rate of flashing. We interpret
these observations to mean that quinacrine was rapidly lost from
vesicles at the basal pole of choroid plexus cells and that this loss
was microtubule dependent.
We recently proposed a sequence of events whereby endosomal
sequestration of organic cations could contribute to transepithelial transport in renal proximal tubule (6):
1) endosomes, which are known to
cycle between the luminal membrane and intracellular organelles,
concentrate organic cations from the cytoplasm by an ATP-dependent
process, and 2) endosomal contents
are delivered to the urinary space when the endosomes fuse with the
luminal membrane. On the basis of the present results, we suggest that such a vesicle-based transport mechanism could contribute to the flux
of organic cations from CSF to blood in choroid plexus. In choroid
plexus cells, a substantial fraction of organic cation (TEA and
quinacrine) was sequestered within an acidic vesicular compartment. The
organic cation-containing vesicles were seen to move within the cells.
Vesicles near the plasma membrane at the blood side of the cells
appeared to release their contents; release was not seen in vesicles
removed from the plasma membrane. Finally, the effects of nocodazole on
microtubule integrity, vesicle motion, and quinacrine release suggest
that microtubules are involved in both phenomena, perhaps serving as
tracks along which vesicles coupled to molecular motors might move.
Although these results are indeed consistent with membrane fusion and
exocytosis, additional experiments are required to
1) definitively establish
vesicle-membrane fusion as the source of flashing and
2) directly demonstrate that the
vesicular phenomena reported here contribute to the flux of organic
cations from CSF to blood.
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: D. S. Miller, LPC, NIH/NIEHS, PO Box 12233, Research Triangle Park, NC 27709 (E-mail: miller{at}niehs.nih.gov).