Interaction of cations, anions, and weak base quinine with rat
renal cation transporter rOCT2 compared with rOCT1
Petra
Arndt1,*,
Christopher
Volk1,*,
Valentin
Gorboulev1,
Thomas
Budiman2,
Christian
Popp1,
Isabel
Ulzheimer-Teuber1,
Aida
Akhoundova1,
Stefan
Koppatz1,
Ernst
Bamberg2,
Georg
Nagel2, and
Hermann
Koepsell1
1 Institute of Anatomy of the Bayerische
Julius-Maximilians-Universität, 97070 Würzburg; and
2 Max-Planck-Institute of Biophysics, 60596 Frankfurt,
Germany
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ABSTRACT |
The rat
organic cation transporter (rOCT)-2 was characterized by electrical and
tracer flux measurements compared with rOCT1. By applying choline
gradients to voltage-clamped Xenopus oocytes expressing
rOCT2, potential-dependent currents could be induced in both
directions. Tracer flux measurements with seven organic cations
revealed similar Michaelis-Menten constant values for both
transporters, with the exception of guanidine. In parallel experiments
with rOCT2 and rOCT1, inhibition of tetraethylammonium transport by 12 cations, 2 weak bases, corticosterone, and the anions
para-amminohippurate,
-ketoglutarate, and probenecid was characterized. The IC50 values of many inhibitors were
similar for both transporters, whereas others were significantly
different. Mepiperphenidol and O-methylisoprenaline showed
an ~70-fold lower and corticosterone a 38-fold higher affinity for
rOCT2. With the use of these inhibitors together with previous
information on cation transporters, experimental protocols are proposed
to dissect out the individual contributions of rOCT2 and rOCT1 in
intact proximal tubule preparations. Inhibition experiments at
different pH levels strongly suggest that the weak base quinine
passively permeates the plasma membrane at physiological pH and
inhibits rOCT2 from the intracellular side.
polyspecific cation transporter; rat organic cation
transporter; organic cation transporter subtypes; substrate
specificity
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INTRODUCTION |
THE TISSUE
CONCENTRATION of endogenous cations, cationic drugs, or
xenobiotics is influenced by polyspecific organic cation transporters
(OCTs) in kidney, liver, small intestine, and brain (5, 17, 18,
31). In 1994, we cloned the rat OCT1 (rOCT1), which proved to be
the first member of a rapidly growing family of polyspecific
transporters (9, 18). This family contains a subfamily of
electrogenic cation transporters, a subfamily containing OCTs and the
sodium-dependent carnitine transporter, and a subfamily of organic
anion transporters (OATs) (2, 18). The subfamily of
electrogenic cation transporters is composed of the transporter subtypes OCT1, OCT2, and OCT3. OCT1 was first cloned from
rat (9) and subsequently isolated from mouse
(27), human (6, 41), and rabbit
(30). By employing immunohistochemistry, rOCT1 was
localized to the sinusoidal membrane of hepatocytes (20) and to basolateral membranes of renal proximal tubules (15, 28). Tracer flux and electrical measurements in Xenopus
oocytes showed that OCT1 from rat and human is electrogenic,
translocates a variety of small cations, and is inhibited by some large
cations that are not transported (3, 4, 9, 22).
OCT2 is a closely related subtype that shares ~70% amino acid
identity with OCT1. OCT2 has been cloned from rat (24),
human (6), pig (8), and mouse
(21). It is mainly expressed in the kidney but has also
been detected in neurons (3). After OCT2 from pig, human,
and rat was characterized in several papers (6, 10, 11, 15,
24-26, 29, 34), controversial issues concerning
localization and function have been solved. Preliminary immunohistochemical experiments suggested that human OCT2 in the kidney
is localized at the luminal membrane of distal tubules (6)
and supported the speculation that OCT2 in kidney might be an apical
electroneutral proton/cation antiporter that mediates cation efflux
into the tubular lumen (8, 10, 11). However, electroneutral proton/cation antiport function was ruled out by functional measurements that showed that OCT2 from human and rat can be
driven by the membrane potential (3, 6, 18, 29) and that
substrate uptake and efflux via OCT2 from rat (rOCT2) is not influenced
by pH gradients (29). Because OCT2 in rat kidney has been
localized unequivocally to the basolateral membrane of proximal
tubule cells (15, 28), it is reasonable to
assume that OCT2 is also localized at the basolateral membrane in
humans (for further discussion, see Ref. 15). The
overlapping localization of rOCT1 and rOCT2 in the proximal tubule
demanded their detailed functional comparison to determine their
specific physiological functions. The previous studies suggested
similar affinities and transport rates for some cations and showed a
different affinity for procainamide (24-26, 34). At
variance, it was concluded in one report that choline is not
transported by rOCT2, and that histamine and guanidine are transported
by rOCT2 but not by rOCT1 (11).
Preliminary experiments showed that choline is actually transported by
rOCT2. Furthermore, we observed that the affinity of rOCT1 for cations
was influenced by the membrane potential and varied by a factor of up
to 10 in different expression systems or batches of oocytes (Ref.
4 and unpublished data). Therefore, we decided to express
rOCT1 and rOCT2 in Xenopus laevis oocytes from the same
batch and carefully compare their cation selectivity by transport
measurements that are performed within 4 h (side-by-side experiments; see METHODS). Our results showed that both
rOCT1 and rOCT2 interacted with all tested cations and that both
transporters exhibited similar affinities for most of them. However,
several cations with dramatically different affinities were identified. Together with previous data on affinities of some substrates and inhibitors of other cation transporters that are also expressed in the
proximal tubule (16, 37-40), these findings provide a rationale to distinguish rOCT1 and rOCT2 in functional studies on
intact proximal tubule preparations.
Because OCTs and OATs belong to the same transporter family (2,
18), we further investigated whether organic anions may interact
with OCTs and thereby influence their function. In these experiments, a
low-affinity interaction of para-aminohippurate (PAH),
-ketoglutarate (AKG), and probenecid with rOCT1 and rOCT2 was
demonstrated. Moreover, we investigated the interaction of larger, more
hydrophobic cations like cyanine863 and the weak base quinine, which
are both high-affinity inhibitors of the OCTs, with rOCT2 and rOCT1 at
different substrate concentrations. With these compounds, a
noncompetitive inhibition was observed. However, this does not
necessarily mean that the OCTs contain allosteric cation binding sites.
In the case of quinine, we were able to demonstrate that quinine
diffuses through the lipid bilayer and interacts with the intracellular
conformation of the cation binding site of rOCT2, which explains the
observed noncompetitive type of inhibition from the outside. These data
may be of biomedical importance, inasmuch as drugs that are weak bases
or hydrophobic membrane-permeant cations may cause prolonged inhibition
of OCTs.
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METHODS |
Cloning of rOCT2.
For cloning of rOCT2, the cDNA of rOCT1 (9) was used as a
probe to screen a rat kidney
gt10 cDNA library under high-stringency conditions (6). A positive cDNA with a restriction pattern different from rOCT1 was cloned into pBluescript II SK(
) and sequenced on both strands using the dideoxynucleotide chain-termination method. The sequence was submitted to GenBank/EBI Data Bank with the
accession number X98334 (4). The derived amino acid sequence is identical to the sequence reported by Gründemann et
al. (8) but differs in two amino acids from the original sequence reported by Okuda et al. (24). In the latter
sequence, Asn332 is replaced by Lys, and Phe335
by Ile. For expression in X. laevis oocytes, rOCT2 was
subcloned into the pOG2 vector containing untranslated regions of the
Xenopus
-globin gene (4). For the expression
in human embryonic kidney (HEK)-293 cells, rOCT2 was subcloned into the
pRcCMV vector (Invitrogen, Groningen, The Netherlands).
Expression of rOCT2 and rOCT1 in Xenopus oocytes and HEK-293
cells.
For expression in Xenopus oocytes, pOG2 vector containing
rOCT2 and pRSSP vector containing rOCT1 (4) were
linearized with NotI and MluI, respectively, and
sense cRNAs were transcribed with the use of T7 or SP6 RNA polymerase
as described earlier (36). Xenopus oocytes were
defolliculated with collagenase A and stored for several hours in Ori
buffer [5 mM 3-(N-morpholino)propanesulfonic acid-NaOH, pH
7.4, 100 mM NaCl, 3 mM KCl, 2 mM CaCl2, and 1 mM MgCl2] containing 50 mg/l gentamycin. Before cRNA
injection, the oocytes were incubated for 5-15 min in hyperosmolar
Ori buffer (130 mM NaCl). The oocytes were then injected with 50 nl
H2O/oocyte with or without 10 ng of rOCT2 or rOCT1 cRNA.
For a comparison between rOCT2 and rOCT1 in side-by-side experiments,
cRNAs of rOCT2 or rOCT1 were injected within 3 h into oocytes from
the same batch. For transporter expression, the oocytes were incubated at 16°C in Ori buffer containing 50 mg/l gentamycin (2-3 days for tracer flux measurements, or 3-5 days for electrical
measurements). For transient expression of rOCT2 in mammalian cells,
HEK-293 cells grown in Dulbecco's modified Eagle's medium
supplemented with 10% fetal calf serum were transfected with the empty
vector pRcCMV or with pRcCMV containing rOCT2, using the FuGENE 6 reagent from Roche Molecular Biochemicals (Mannheim, Germany).
Tracer uptake measurements.
Uptake measurements of radioactive cations into oocytes of X. laevis and into HEK-293 cells were performed as described
(3). To compare transport by rOCT2 and rOCT1 expressed in
the same batch of oocytes in side-by-side experiments, uptake by both
transporters was measured within 4 h. For uptake measurements, the
oocytes were incubated at 19°C for 10 min or 1 h with the
radioactive substrates in the absence or presence of inhibitors, and
the uptake was stopped with ice-cold Ori buffer. It was verified that
the uptake of 400 µM tetraethylammonium (TEA) into oocytes expressing rOCT2 or rOCT1 was linear for 90 min (data not shown). Measurements of
choline uptake in transiently transfected HEK-293 cells were performed
2 days after transfection when the cells had reached confluence. The
cells were washed with phosphate-buffered saline (PBS), suspended by
shaking, collected by 10-min centrifugation at 1,000 g, and
suspended at 37°C in PBS. For uptake, the cells were incubated
between 1 and 60 s in PBS (37°C) with or without 100 µM
tetrapentylammonium (TPeA). Uptake was stopped by addition of ice-cold
PBS containing 100 µM quinine (stop solution), and the cells were
washed three times with ice-cold stop solution.
Electrophysiology.
Electrical measurements were performed on Xenopus oocytes
that had been kept at 16°C in Ori buffer, or on oocytes that had been
incubated for 12 h in Ori buffer containing 10 mM choline. The
two-electrode voltage-clamp measurements were performed as described
earlier (3). The oocytes were continuously superfused at
room temperature (~3 ml/min). For the determination of
current-voltage relationships, steady-state currents were averaged
during the last 100 ms of 500 ms of voltage pulses that were applied at
a frequency of 0.4 Hz from a holding potential of
50 mV.
Calculation and statistics.
In Xenopus oocytes, substrate dependence of uptake and
maximal velocity (maximal transport rate;
Vmax) values were calculated from
8-10 oocytes incubated without inhibitor minus 8-10 oocytes incubated with inhibitor per data point. The uptake in the presence of
different inhibitor concentrations was measured from 8-10 oocytes incubated with a specific inhibitor concentration per data point. In
HEK-293 cells, uptake was calculated from four determinations in the
absence and four determinations in the presence of inhibitor. For each
data point, the mean ± SE was calculated. From individual uptake
experiments with different substrate concentrations, apparent Michaelis-Menten constant (Km) ± SD values
were calculated by fitting the Michaelis-Menten equation to the data.
For some substrates, the Michaelis-Menten equation was fitted to each
of three or four individual experiments, and means ± SD of the
Km values were calculated. IC50
values were calculated from individual dose-response curves by fitting
the Hill equation for multisite inhibition to the data. For inhibitors
with largely different affinities, three to five individual inhibition
experiments were performed side by side, and means ± SD of the
IC50 values were calculated. The unpaired Student's
t-test was employed to evaluate the significance of differences between means of Km,
IC50, or Vmax values. The straight lines shown (see Figs. 4, 6, 7, and 9) were obtained by linear regression analysis.
Materials.
[3H]choline (2.6 TBq/mmol) and
[3H]histamine (1.9 TBq/mmol) were obtained from Amersham
Buchler (Braunschweig, Germany). [14C]TEA (1.9 TBq/mmol),
[14C]- guanidine (2.0 GBq/mmol),
[3H]1-methyl-4-phenylpyridinium (3.1 TBq/mmol),
[3H]quinine (0.56 TBq/mmol), and
[3H]quinidine (0.54 TBq/mmol) were purchased from
Biotrend (Köln, Germany).
[3H]N'-methylnicotinamide (0.11 TBq/mmol) was
purchased from ICN Biochemicals (Meckenheim, Germany). Unlabeled
cyanine-863, decynium-22, quinine, quinidine, desipramine,
procainamide, N'-methylnicotinamide (NMN), PAH, probenecid,
and AKG were obtained from Sigma (Deisenhofen, Germany);
tetramethylammonium, tetrapropylammonium, tetrabutylammonium (TBuA),
TPeA, and corticosterone from Fluka (Neu-Ulm, Germany); 3-O-methylisoprenaline from Boehringer (Ingelheim,
Germany); and mepiperphenidol from Merck Sharp and Dohme (Rahway,
NJ). The other chemicals were obtained as described earlier
(4).
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RESULTS |
Electrical properties of choline uptake by rOCT2.
Superfusion of X. laevis oocytes expressing rOCT2 that were
clamped to
50 mV with choline, TEA, NMN, and procainamide induced inward currents as has been described for rOCT1
(4). These currents were not observed in
water-injected control oocytes (e.g., Fig.
1A). The inward currents
induced by 10 mM choline or other transported cations could be
completely inhibited by 200 µM quinine or 100 µM cyanine863 (data
not shown). In oocytes with high transport activity that were
superfused for 60 s with 10 mM choline, a transient outward
current was observed after the removal of choline from the bath
solution (Fig. 1C). This outward current was inhibited by
200 µM quinine (see right trace in Fig. 1C). It
may represent electrogenic efflux of choline that had been taken up by
the oocyte (see Fig. 2). Current-voltage
(I-V) curves from the water-injected oocyte in
Fig. 1A or from the rOCT2-injected oocyte in the
left part of Fig. 1C are presented in Fig. 1,
B and D, respectively. The choline-induced inward
currents observed in rOCT2-expressing oocytes (see difference curves)
are in perfect keeping with the idea that the choline uptake by rOCT2
is electrogenic. These currents increased with more negative membrane
potential, indicating that the rate-limiting step is voltage dependent.

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Fig. 1.
Choline-induced inward currents in Xenopus
oocytes expressing rat organic cation transporter (rOCT)-2.
Xenopus oocytes were injected with water (control) or 10 ng
of rOCT2 cRNA and incubated for 4 days. The oocytes were
voltage-clamped and superfused with Ori buffer ( ) or
with Ori buffer containing 10 mM choline ( ).
A: steady-state currents in water-injected oocyte clamped at
50 mV. B: current-voltage (I-V)
curves from the control oocyte in A. C:
steady-state currents in 2 rOCT2-expressing oocytes clamped at 50 mV.
The trace on the left is from an oocyte that was superfused
with Ori buffer without and with 10 mM choline. The superimposed traces
on the right are from another oocyte. In a first superfusion
period (solid line), this oocyte was superfused with Ori buffer or Ori
buffer containing 10 mM choline. After 15 min of superfusion with Ori
buffer (not shown), the same oocyte was superfused first with Ori
buffer containing 10 mM choline and then with Ori buffer containing 200 µM quinine (dotted line). D: I-V
curves that were obtained during superfusion of the left
oocyte in C.
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Fig. 2.
Choline-induced outward currents in Xenopus
oocytes expressing rOCT2. Xenopus oocytes from the same
batch as in Fig. 1 were injected with water (control) or 10 ng of rOCT2
cRNA and incubated for 3.5 days in Ori buffer and for 12 h in Ori
buffer containing 10 mM choline. A: water-injected oocyte
that was clamped to 50 mV and superfused with Ori buffer containing
10 mM choline ( ) or with Ori buffer ( ).
B: I-V curves from the oocyte in
A. C: an rOCT2-expressing oocyte that was
superfused first with Ori buffer containing 10 mM choline
( ), then with Ori buffer without choline
( ), again with Ori buffer containing 10 mM choline, and
finally with Ori buffer containing 100 µM cyanine863
( ). D: I-V curves from
the oocyte presented in C. Similar data as in C
and D were obtained when the superfusion protocol with and
without choline was repeated with the same oocyte.
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By employing inside-out patches from Xenopus oocytes, we
recently showed that rOCT2 is able to mediate cation-induced outward currents (1). These measurements are technically difficult and have serious limitations. For example, the transport activity of
rOCT2 in excised membrane patches varied largely between different patches and decreased dramatically with time. For these reasons it was
impossible to compare the magnitude of rOCT2-mediated inward and
outward currents in the presence of cation gradients. To circumvent these problems, we tried to demonstrate cation gradient-induced outward
currents by rOCT2 in intact oocytes. We preloaded water-injected control oocytes and rOCT2-expressing oocytes with choline by incubating them for 12 h in 10 mM choline. When control oocytes and
rOCT2-expressing oocytes were clamped to
50 mV and superfused with 10 mM choline, no significant quinine-inhibitable currents were detected
(data not shown). When choline-preloaded oocytes were superfused with choline-free buffer, however, a large outward current was observed in
rOCT2-expressing oocytes but not in water-injected control oocytes
(Fig. 2, A and C). This rOCT2-mediated outward
current was abolished when 100 µM cyanine863 or 200 µM quinine were
added to the bath (data not shown). I-V curves
from water-injected control oocyte and rOCT2-expressing oocyte of Fig.
2, A and C, are shown in Fig. 2, B and
D. The rOCT2-mediated outward current that was induced by
the outwardly directed choline gradient and could be inhibited by
cyanine863 (see difference curve in Fig. 2D) was decreased
at more negative membrane potentials. Taken together, these findings
show that rOCT2 is able to mediate electrogenic choline transport in
both directions, as has been shown for rOCT1 (22). In
rOCT2-expressing oocytes clamped to
50 mV, the outward currents
obtained by preloading the oocytes in 10 mM choline and superfusing
with choline-free buffer were higher than the inward currents that were
obtained by superfusion of nonpreloaded oocytes with 10 mM choline
(compare, for example, Figs. 1 and 2).
In two batches of Xenopus oocytes clamped to
50 mV, we
compared the induced inward and outward currents by rOCT2. In these oocytes, the choline-induced inward current, measured as in Fig. 1, was
29 ± 3 nA (means ± SD, n = 10), whereas the
current induced by removal of choline from the bath solution, measured
as in Fig. 2, was 127 ± 22 nA (means ± SD,
n = 10). By performing an analogous experiment with two
batches of oocytes expressing rOCT1, we obtained choline-induced inward
currents of 9 ± 2 nA (means ± SD, n = 4) and currents induced by choline removal of 26 ± 1 nA (means ± SD, n = 5). The data show that rOCT1 and rOCT2 may
mediate significant choline efflux at normal membrane potential if the
transmembrane concentration gradient is high enough.
Next, we measured the concentration dependence of choline-induced
currents at three different membrane potentials. Figure 3 shows an experiment where
rOCT2-expressing oocytes that were not preloaded were clamped to 0,
50, or
90 mV and superfused with various concentrations of choline.
By fitting the Michaelis-Menten equation to the data, apparent
Km values of 0.70 ± 0.09 mM (0 mV),
0.38 ± 0.05 mM (
50 mV), and 0.27 ± 0.04 mM (
90 mV) and apparent maximal current (Imax) values
of 136 ± 15 nA (0 mV), 232 ± 8 nA (
50 mV), and 282 ± 13 nA (
90 mV) were determined. The data indicate that the
Km and Imax for
electrogenic choline uptake are potential dependent, as has been
described for cation transport by rOCT1 and rOCT3 (4, 16).
Inward currents could also be induced when rOCT2-expressing oocytes
were superfused with other substrates. When rOCT2-expressing oocytes
were clamped to
50 mV and superfused with saturating concentrations
of TEA (1 mM), guanidine (10 mM ), choline (10 mM), procainamide (10 mM), NMN (10 mM), histamine (10 mM), and 1-methyl-4-phenylpyridinium (MPP) (100 µM), the following relative inward currents were obtained (mean ± SD from 3-8 oocytes from 2 oocyte batches, relative
to the TEA-induced current): TEA 1.0 ± 0.10, guanidine 2.14 ± 0.24, choline 1.96 ± 0.45, NMN 1.16 ± 0.24, histamine
1.12 ± 0.18, MPP 0.71 ± 0.14, and procainamide 0.12 ± 0.01.

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Fig. 3.
Concentration dependence of choline-induced inward
currents in Xenopus oocytes expressing rOCT2 measured at
different membrane potentials. The oocytes were first superfused with
Ori buffer; clamped to 90, 50, or 0 mV; and then superfused with
Ori buffer containing various concentrations of choline. Mean
values ± SE of 5 oocytes are presented. The Michaelis-Menten
equation was fitted to the data.
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Concentration dependence and Vmax of cation uptake by
rOCT2 compared with rOCT1.
In Xenopus oocytes injected with rOCT2 mRNA, the uptake of
[3H]NMN, [14C]TEA, [3H]MPP,
[14C]histamine, and [3H]guanidine was at
least 10 times higher than in water-injected control oocytes, showed
substrate saturation, and was inhibitable by >95% with specific
inhibitors (100 µM cyanine-863 or 100 µM decynium-22). In
water-injected control oocytes, the uptake of these cations showed no
saturation and was insensitive to cyanine-863 and decynium-22 (data not
shown). In control oocytes, a higher uptake was observed for choline
compared with the other cations (Fig.
4A). However, this uptake
increased linearly with the substrate concentration and was insensitive
to cyanine-863. By fitting the Michaelis-Menten equation to
cyanine-863-inhibitable uptake of TEA, choline (Fig. 4B),
NMN, and MPP in single experiments performed on different oocyte
batches of rOCT2-expressing oocytes, apparent Km
values of 91 µM (TEA), 0.6 mM (choline), 0.25 mM (NMN), and 17 µM
(MPP) were obtained. These values are similar to those that have been
determined for rOCT1 by tracer flux in Xenopus oocytes (7, 9, 17) and are summarized in Table
1. For the uptake of histamine and
guanidine by rOCT2 and rOCT1, the substrate dependence was measured in
four side-by-side experiments that were performed as described in
METHODS (Table 1, values with index "c"). For rOCT2 and
rOCT1, apparent Km values for histamine uptake
were similar: 0.28 ± 0.05 mM (rOCT2) vs. 0.30 ± 0.12 mM
(rOCT1). The Km values for guanidine uptake,
however, differed by a factor of 10: 0.17 ± 0.06 mM (rOCT2) vs.
1.7 ± 0.7 mM (rOCT1) (P < 0.05 for difference).

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Fig. 4.
Concentration dependence of [3H]choline
uptake measured in oocytes of Xenopus laevis injected with
water (A; control) or rOCT2 cRNA (B). One hour of
[3H]choline uptake at different choline concentrations
was measured in the absence (open symbols) and presence (solid symbols)
of 100 µM cyanine-863. Mean values from 8-10 oocytes ± SE
are presented. The straight lines were calculated by linear regression
analysis (r2 > 0.98). The top
curve in B was obtained by fitting the Michaelis-Menten
equation plus a linear uptake component to the data. The broken line in
B was obtained by fitting the Michaelis-Menten equation to
the cyanine-863-inhibited fractions of the choline uptake rates. For
cyanine-863-inhibitable uptake mediated by rOCT2, a Michaelis-Menten
constant (Km) value of 0.6 ± 0.1 mM and a
maximal transport rate (Vmax) value of 0.70 ± 0.04 nmol · oocyte 1 · h 1
were estimated.
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Next, we compared the maximum transport rates of different cations
mediated by rOCT2 compared with rOCT1. In each of five side-by-side
experiments (see METHODS), oocytes from a single batch were
injected within 3 h with 10 ng/oocyte of either rOCT1 or rOCT2
cRNA, and after 3 days, the Vmax values for
cyanine-863-inhibitable uptake of TEA, choline, NMN, histamine,
guanidine, and MPP were measured within 4 h. During this time
period, electrical measurements were performed on five oocytes from
each individual experiment. rOCT2-expressing oocytes clamped at
50 mV
were superfused with 0.65 mM TEA, and the TEA-induced currents were
determined in the absence and presence of 200 µM quinine. In the
tracer flux experiments, the uptake rate of 0.65 mM TEA expressed by
rOCT2 was three times higher than the uptake of 0.65 mM TEA expressed
by rOCT1 [0.47 ± 0.06 (rOCT2) vs. 0.15 ± 0.06 (rOCT1), in
nmol · oocyte
1 · h
1;
P < 0.01 for difference]. By superfusion of oocytes
expressing rOCT2 or rOCT1, with 0.65 mM TEA clamped at
50 mV,
sevenfold higher currents were obtained with rOCT2 [28.6 ± 5.2 nA (rOCT2) vs. 4.0 ± 1.2 nA (rOCT1)]. The greater ratio of the
TEA-induced currents expressing rOCT2 and rOCT1 compared with the
ratios of the respective [14C]TEA uptake rates
may be explained by a more pronounced membrane depolarization in the
course of TEA uptake in rOCT2-expressing oocytes. Because the tracer
flux measurements were performed with nonclamped oocytes, the degree of
oocyte depolarization depends on the magnitude of the expressed
electrogenic cation uptake. Membrane depolarization would lead to a
reduction of the apparent Vmax.
To compare the uptake rates of different transported cations in a given
isoform, we normalized the maximal uptake rates for various individual
cations to the maximum TEA uptake rate measured in the respective
experiment, which was set to 1.0 (Table 1). For rOCT2, the
Vmax values of guanidine, choline, and histamine were significantly higher than the Vmax of TEA,
whereas the Vmax of MPP was significantly lower.
In contrast, for rOCT1, only the Vmax of
guanidine was significantly higher than that of TEA. Making the
assumption that substrate binding was not rate limiting in our
Vmax measurements, the comparison between the
normalized Vmax values by rOCT2 and rOCT1 shows
that rOCT2 has a higher turnover number, relative to TEA, for choline
and histamine than rOCT1. At variance, the turnover number of rOCT2 for
MPP, relative to TEA, is significantly smaller compared with rOCT1. The
data suggest that rOCT2 and rOCT1 have different rank orders in
turnover numbers for different cations in the following way:
guanidine = choline > histamine > NMN = TEA > MPP for rOCT2, and guanidine > histamine = NMN = TEA = choline > MPP for rOCT1.
Recently, Gründemann et al. (11) reported that the
uptake of TEA, MPP, histamine, creatinine, and guanidine in HEK-293 cells was increased after transfection with rOCT2 compared with control
cells, whereas the uptake of choline was not. To determine whether the
expression of choline uptake was dependent on the expression system, we
transiently transfected HEK-293 cells with rOCT2 and measured the time
course of [3H]choline uptake in the absence and presence
of 100 µM TPeA. Figure 5 shows that the
rOCT2-transfected HEK-293 cells exhibited a significantly higher
TPeA-inhibitable uptake of 50 µM [3H]choline than
HEK-293 control cells that were transfected with the empty vector. The
observation that a fraction of the choline uptake into control cells
could also be inhibited by TPeA may be explained by the fact that
HEK-293 cells express small amounts of endogenous human OCT1
and show low activity of organic cation transport (Ref. 19
and unpublished data).

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Fig. 5.
Expression of rOCT2-mediated choline uptake in human
embryonic kidney (HEK)-293 cells. HEK-293 cells were transiently
transfected with the pRcCMV vector containing rOCT2 or with the empty
vector. At 2 days after transfection, the time course of uptake of 50 µM [3H]choline was measured in the absence and presence
of 100 µM tetrapentylammonium (TPeA). The TPeA-inhibitable uptake
after various incubation times is indicated. Mean values ± SE
(n = 4) are shown. The 60-s uptake in the presence of
100 µM TPeA was 100 ± 5 and 92 ± 4 pmol/mg protein in
control cells and rOCT2-expressing cells, respectively.
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Inhibition of rOCT2 and rOCT1 by organic cations and weak bases.
To compare specificity and affinity of various inhibitors in rOCT2 and
rOCT1, we measured concentration-inhibition curves for uptake of 10 µM TEA at various concentrations of several cations and weak bases at
pH 7.4. The low concentration of TEA was used to minimize the substrate
effect on the IC50 values of competitive inhibitors. For
the comparison between rOCT2 and rOCT1 in side-by-side experiments,
oocytes from single batches were injected with the respective two
cRNAs, and after 3 days' incubation transport was measured within
4 h. These precautions are important because the affinity of rOCT2
and rOCT1 for cations is influenced by the membrane potential (Fig. 3
and Ref. 4), and the membrane potential may differ between
oocyte batches. Inhibition experiments with choline and NMN revealed
IC50 values for rOCT2 and rOCT1 that were in the same range
as the apparent Km values shown in Table 1
(choline: rOCT2 0.34 ± 0.04 mM, rOCT1 0.35 ± 0.5 mM; NMN:
rOCT2 0.27 ± 0.01 mM, rOCT1 0.27 ± 0.02 mM). Comparable
IC50 values have been reported by Okuda et al.
(25) for choline, but these authors obtained IC50 values for NMN that were more than five times higher.
For the inhibition of TEA uptake by MPP, we estimated IC50
values of 46 ± 5 µM (rOCT2) and 24 ± 10 µM (rOCT1).
Similar values have been reported earlier (25). In Table
2, IC50 values for the inhibition of TEA uptake by nine additional cations and by the weak
bases quinine and quinidine are summarized. The mean IC50 values ± SD presented for TPeA, tetrapropylammonium, quinidine, and tetramethylammonium were calculated from individual side-by-side experiments with rOCT2 and rOCT1. Because no large difference between rOCT2 and rOCT1 became apparent, these experiments were not
repeated. For cyanine-863, quinine, guanidine, procainamide, mepiperphenidol, and O-methylisoprenaline, we performed
three to five side-by-side experiments and calculated the mean
IC50 values. For cyanine-863, quinine, procainamide,
mepiperphenidol, and O-methylisoprenaline, 5 to 70 times
higher IC50 values were obtained for rOCT2 compared with
rOCT1. In contrast, the IC50 value for guanidine was more
than 20 times smaller for rOCT2 than for rOCT1. It is noteworthy that
quinine exhibited a significantly lower affinity for rOCT2 than for
rOCT1, whereas the quinine enantiomer quinidine had a similar affinity
to both transporters.
Investigation of quinine for transport by rOCT2 and rOCT1.
To compare rOCT2 and rOCT1 with respect to quinine uptake, the uptake
of [3H]quinine was measured side by side in
water-injected control oocytes, rOCT2-expressing oocytes, and
rOCT1-expressing oocytes. In addition to pH 7.4, these measurements
were also performed at pH 6 to reduce the nonspecific uptake of quinine
that correlates with the fraction of uncharged quinine. Quinine has an
acid dissociation constant (pKa) value
of 8.4 (23), and the fraction of uncharged quinine is 10%
at pH 7.4 and <1% at pH 6.0. For the uptake of 0.1 µM
[3H]quinine in the absence or presence of 100 µM
cyanine863, the following values (mean values ± SE, in
pmol · oocyte
1 · h
1) were
obtained: control pH 7.4, 0.34 ± 0.01 vs. 0.33 ± 0.01; control pH 6.0, 0.06 ± 0.01 vs. 0.02 ± 0.01; rOCT2 pH 7.4, 0.32 ± 0.03 vs. 0.32 ± 0.01; rOCT2 pH 6.0, 0.05 ± 0.01 vs. 0.01 ± 0.01; rOCT1 pH 7.4, 0.83 ± 0.05 vs.
0.32 ± 0.02; and rOCT1 pH 6.0, 0.16 ± 0.02 vs. 0.02 ± 0.01. In water-injected control oocytes, a very small but significant
(P < 0.01) cyanine-863-inhibitable uptake of quinine
became visible at pH 6.0 that was not detected at pH 7.4. After
expression of rOCT2, no additional cyanine-863-inhibitable uptake could
be observed. This result was confirmed by electrical measurements. By
superfusion with 100 µM quinine at pH 7.4 or pH 6.0, no inward
currents could be detected in rOCT2-expressing oocytes that were
clamped at
40 mV. In the same oocytes, regular-sized inward currents
>80 nA were observed on superfusion with 10 mM choline. In
rOCT1-expressing oocytes at pH 7.4, significant (P < 0.001) cyanine-863-inhibitable quinine uptake was detected as reported earlier (22). In rOCT1-expressing oocytes at pH
6.0, the cyanine-863-inhibitable quinine uptake was smaller compared with that at pH 7.4, but it was still significantly larger
(P < 0.001) than in control oocytes. The data show
that rOCT2 does not mediate detectable transport of quinine. In
contrast, a small amount of quinine uptake was induced on expression of
rOCT1 (see DISCUSSION).
Inhibition of rOCT2 and rOCT1 by quinine, decynium22, and TPeA in
the presence of different substrate concentrations.
The concentration dependence of [14C]TEA uptake by
rOCT2 and rOCT1 was measured in the presence of various concentrations
of NMN, quinine, decynium22, and TPeA (Figs.
6 and 7).
Similar results were obtained for rOCT2 and rOCT1. For the inhibition
of TEA uptake by the transported cation NMN, a competitive type of
inhibition was obtained (see Figs. 6A and 7A). In
contrast, a noncompetitive type of inhibition was observed for the weak
base quinine (see Figs. 6B and 7B).
Noncompetitive inhibition of TEA uptake by rOCT2 and rOCT1 was also
shown for the permanently charged cations decynium-22 (shown for rOCT2
in Fig. 6C) and for cyanine-863 (data not shown). For rOCT1,
we demonstrated noncompetitive inhibition of TEA uptake also for the
permanently charged cation TPeA (see Fig. 7C). The kinetics
for TPeA inhibition of TEA uptake by rOCT2 were not determined. Electrical measurements suggested that neither cyanine-863 nor decynium-22 nor TPeA is transported to a significant extent by rOCT2 or
rOCT1, since no inward current could be induced when nonpreloaded
oocytes expressing rOCT2 or rOCT1 were clamped at
50 mV and
superfused with 100 µM of cyanine-863, decynium-22, or TPeA. In
contrast, currents >80 or 7 nA were induced when the oocytes
expressing rOCT2 or rOCT1 were superfused with 10 mM choline, respectively.

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Fig. 6.
Kinetics for the inhibition of rOCT2-expressed
tetraethylammonium (TEA) uptake by N'-methylnicotinamide
(NMN), quinine, and decynium22. Xenopus oocytes were
injected with rOCT2 cRNA and incubated for 2 days in Ori buffer. The
uptake of [14C]TEA at various bath TEA concentrations was
measured after 1 h of incubation in the absence of additional
organic cations or in the presence of various concentrations of NMN
(A), quinine (B), and decynium-22 (C).
The data in A-C were obtained in 3 different
oocyte batches. They are plotted according to Hofstee
(12). Means ± SE from 8-10 oocytes are
presented. The lines were calculated by linear regression analysis
(r2 > 0.94). The data show competitive
inhibition of rOCT2-expressed TEA uptake by NMN and noncompetitive
inhibition of TEA uptake by quinine and decynium22.
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Fig. 7.
Kinetics for the inhibition of rOCT1-mediated TEA uptake
by NMN, quinine, and TPeA. Xenopus oocytes were injected
with rOCT1 cRNA and incubated for 2 days in Ori buffer.
[14C]TEA uptake was determined after 1 h of
incubation in the presence of various TEA concentrations plus the
indicated concentrations of NMN (A), quinine (B),
and TPeA (C). The data are presented as in Fig. 6. They show
competitive inhibition of rOCT1-expressed TEA uptake by NMN and
noncompetitive inhibition of TEA uptake by quinine and TPeA.
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Interaction of the weak base quinine with rOCT2.
To elucidate whether quinine may interact with an allosteric binding
site, we tried to exclude other reasons for the observed noncompetitive
inhibition of rOCT2. We investigated whether the noncompetitive type of
inhibition with quinine may result from an irreversible inactivation of
rOCT2 by quinine or whether it may result from an intracellular
interaction of quinine with rOCT2. This latter possibility was
considered, since quinine has a pKa value of 8.4 (23), so that ~10% of quinine is uncharged at pH 7.4 and may readily cross the lipid bilayer via nonionic diffusion. A
quinine-induced inactivation of rOCT2 during the transport measurements in Fig. 6B could be excluded as a reason for the observed
noncompetitive type of inhibition. First, electrical measurements in
rOCT2-expressing oocytes revealed that the inhibition of TEA-induced
inward currents was reversible when the oocytes had been incubated for
1 h with 100 µM quinine (data not shown). Second, when
electrical measurements were performed within several minutes, a
noncompetitive inhibition of TEA uptake by quinine was also observed.
In Fig. 8, rOCT2-expressing oocytes
clamped at
40 mV were superfused with various concentrations of TEA in the absence and presence of 30 µM quinine. The apparent Km value for TEA was not changed by quinine, and
quinine inhibited the inward currents induced by 50, 200, or 2,000 µM
TEA by 65, 65, and 67%, respectively. When this experiment was
repeated with four other oocytes, similar results were obtained. During
superfusion with 50, 200, or 2,000 µM TEA, the degree of inhibition
by 30 µM quinine was nearly identical at the three TEA
concentrations, showing a maximal difference of 2%.

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Fig. 8.
Inhibition of TEA-induced inward currents by quinine at
pH 7.4. Xenopus oocytes were injected with 10 ng rOCT2 cRNA
and incubated for 4 days. Then, the oocytes were clamped at 40 mV and
superfused with Ori buffer (pH 7.4) containing various concentrations
of TEA, 30 µM quinine, or 30 µM quinine plus various concentrations
of TEA. It can be seen that the relative concentration dependence of
TEA-induced inward currents was not changed in the presence of quinine.
Whereas quinine did not induce an inward current in nonpreloaded
oocytes (see addition of quinine at right), it inhibited
TEA-induced outward currents that were observed after TEA removal (see
addition of quinine at left). Vhold,
holding potential.
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To evaluate whether quinine may inhibit rOCT2 from the intracellular
side, we determined whether the TEA uptake by rOCT2 was correlated to
nonionic diffusion of quinine across the lipid bilayer. We measured the
passive uptake of quinine at different pH values and tried to correlate
passive quinine uptake with the IC50 values for the
inhibition of expressed TEA uptake by quinine in the bath. At pH 8.4, 7.4, and 6.0, ~50, 10, and <1%, respectively, of quinine is
uncharged and may permeate the lipid bilayer by nonionic diffusion. Figure 9A shows the uptake of
0.1 µM [3H]quinine into Xenopus oocytes at
different pH values. The uptake rates at different pH values observed
in water-injected oocytes and in rOCT2-cRNA injected oocytes were not
significantly different. Because the uptake increased strongly with
increasing pH in the bath, it probably includes diffusion of uncharged
[3H]quinine through the plasma membrane. The prolonged
increase of pH-dependent [3H]quinine uptake is not
understood. It may be explained by a slow equilibration of cytosolic
quinine with yolk lipids that represent a large lipophilic compartment
(see DISCUSSION). In Fig. 9B, we determined the
apparent IC50 values for quinine inhibition of TEA uptake
at bath pH 6.0, 7.4, and 8.4. The IC50 values increased with decreasing pH from 1.5 ± 0.3 µM at pH 8.4, over 8 ± 2 µM at pH 7.4, to 38 ± 8 µM at pH 6.0. Because the membrane
potential was only slightly affected by these pH changes (60.3 ± 4.6 mV at pH 8.4, 61.3 ± 4.8 mV at pH 7.4, 54.1 ± 2.0 mV at
pH 6.0; means ± SD, n = 6), the affinity changes
may not be explained by a potential sensitivity of quinine binding to
rOCT2. Electrical measurements under voltage-clamped conditions
excluded this possibility more unequivocally. Figure
10 shows experiments in which oocytes
expressing rOCT2 were clamped at
40 mV and superfused with Ori buffer
adjusted to pH 6.4 or 8.4 that contained 1 mM TEA or 1 mM TEA plus
quinine at various concentrations. For the inhibition by quinine,
apparent IC50 values of 3.9 ± 0.3 and 74 ± 20 µM were estimated at pH 8.4 and 6.4, respectively. This indicates
that the pH effects on the quinine inhibition are already observed
within 1 min and that they are not induced by changes of the membrane
potential. The observation that higher IC50 values were
obtained in the electrical than in the tracer flux measurements may
reflect less complete intracellular equilibration of quinine in the
electrical measurements. Interestingly, the time for reactivation of
transport from quinine inhibition was significantly longer at pH 8.4 than at pH 6.4 (see Fig. 10). The data are consistent with the
interpretation that quinine diffuses in its uncharged form through the
lipid bilayer and inhibits rOCT2 from the intracellular side. We
wondered whether the high IC50 value at pH 6.0 reflects the
low intracellular concentration of quinine at this pH or represents
low-affinity binding of quinine to rOCT2 from the extracellular side.
To approach this question, we measured the concentration dependence of
TEA uptake at pH 6.0 without an inhibitor and in the presence of 30 or
100 µM quinine (Fig. 11). When the
Michaelis-Menten equation was fitted to the data, a competitive type of
inhibition was observed. The apparent Vmax
values were not significantly changed by quinine; the apparent Km values, however, increased with increasing
quinine concentration. The following Vmax values
(in pmol · oocyte
1 · h
1)
and Km values (in mM) were estimated: absence of
quinine, Vmax = 725 ± 72, Km = 0.21 ± 0.04; 30 µM quinine,
Vmax = 827 ± 64, Km = 0.31 ± 0.05; and 100 µM
quinine, Vmax = 844 ± 80, Km = 0.66 ± 0.09. With the use of the
Km of TEA in the absence of quinine, and
assuming a competitive type of inhibition, inhibitor constant (Ki) values for quinine of 55 and 45 µM were calculated from the Km values
determined in the presence of 30 and 100 µM quinine, respectively.
The data suggest that quinine interacts at pH 6.0 with the
outward-facing cation binding site of rOCT2.

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Fig. 9.
Passive quinine uptake into Xenopus oocytes
and inhibition of rOCT2-mediated TEA uptake by quinine at different pH
values. Xenopus oocytes from 1 batch were injected with
water ( ) or rOCT2 cRNA ( ,
, , and ) and incubated
for 3 days. A: oocytes were incubated at room temperature at
pH 6.0, pH 7.4, or pH 8.4 in Ori buffer containing 0.1 µM
[3H]quinine. After the indicated time intervals, the
oocytes were washed with ice-cold Ori buffer and analyzed for
radioactivity. B: rOCT2-expressing oocytes were incubated at
room temperature at pH 6.0, 7.4, or 8.4 in Ori buffer containing 10 µM [14C]TEA plus the indicated concentrations of
quinine. After 10 min of incubation, the oocytes were washed and
analyzed for radioactivity. The measurements at different pH were
normalized for TEA uptake in the absence of quinine. The passive
quinine uptake (A) increased with increasing pH, whereas the
IC50 values for quinine inhibition of rOCT2-expressed
uptake (B) decreased with increasing pH.
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Fig. 10.
Quinine inhibition of TEA-induced inward currents
mediated by rOCT2 at pH 8.4 (A) and 6.4 (B).
Xenopus oocytes were injected with 10 ng of rOCT2 cRNA and
incubated for 4 days. The oocytes were clamped at 40 mV and
superfused with Ori buffer at pH 8.4 or 6.4. Inward currents were
induced by addition of 1 mM TEA and inhibited by various concentrations
of quinine as indicated. At pH 8.4, the IC50 value for the
inhibition of TEA-induced inward current by rOCT2 is smaller compared
with that at pH 6.4, and the reactivation of TEA-induced current is
more rapid.
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Fig. 11.
Effect of quinine on the concentration dependence of
rOCT2-mediated [14C]TEA uptake measured at pH 6.0. Xenopus oocytes were injected with rOCT2 cRNA and incubated
for 2 days in Ori buffer. The uptake of [14C]TEA was
measured after 10 min of incubation of the oocytes at pH 6.0 in Ori
buffer containing the indicated concentrations of TEA and quinine. The
Michaelis-Menten equation was fitted to the data. They suggest
competitive inhibition of rOCT2-mediated TEA uptake by quinine at pH
6.0.
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Interaction of the permanently charged, nontransported cation TBuA
with rOCT2.
In an attempt to compare the above-described effects of quinine with
effects of a permanently charged inhibitor with a comparable affinity
that may not permeate the lipid phase of the plasma membrane, we
investigated the inhibition of rOCT2-mediated cation uptake by TBuA. To
test whether TBuA is transported by rOCT2, we compared the inward
currents in rOCT2-expressing oocytes clamped to
50 mV that were
superfused with saturating concentrations of choline (10 mM) or TBuA
(0.5 mM). Although the choline-induced currents in these experiments
were very high (295 ± 5 nA, mean ± SD, n = 3), no currents could be induced by 0.5 mM TBuA (<2 nA,
n = 3). The data indicate that TBuA is not translocated
by rOCT2 or that it is translocated with a Vmax
that is <1/100 of the Vmax for
choline. The inhibition of TEA-induced inward currents by TBuA in the
bath is shown in Fig. 12. Figure
12A shows that 1 mM TBuA partially inhibited inward currents
that were induced by 5 or 20 mM TEA, and that the inhibition by TBuA
was decreased when the TEA concentration was increased. After removal
of TBuA from the bath, the inhibition disappeared within the latency of the buffer exchange (see arrow in Fig. 12A). Figure
12B shows current measurements in Xenopus oocytes
clamped to
40 mV that were superfused with different concentrations
of TEA either without TBuA or in the presence of 100 or 350 µM TBuA.
The curves suggest competitive inhibition of rOCT2-mediated TEA uptake
by TBuA. Assuming a competitive type of inhibition, and using the
Km value for TEA obtained in the absence of TBuA
(150 µM), Ki values of 12 and 18 µM were
calculated from the measurements performed in the presence of 100 and
350 µM TBuA, respectively. Finally, we determined whether the
IC50 values for TBuA inhibition of rOCT2-mediated uptake of
10 µM [14C]TEA were dependent on the pH in the bath, as
had been observed for the inhibition of TEA uptake by quinine. With the
use of the same experimental conditions and fitting procedure as for
the experiments in Table 2, IC50 values of 17.2 ± 3.8 µM at pH 6.0, 24.9 ± 5.7 µM at pH 7.4, and 18.2 ± 6.8 µM at pH 8.4 were estimated. This indicates that the inhibition of
rOCT2-mediated TEA uptake by TBuA is not dependent on the pH in the
bath. The data suggest that TBuA is a nontransported inhibitor of rOCT2
that interacts with the outward-facing substrate binding site.

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Fig. 12.
Inhibition of rOCT2-mediated inward currents by
tetrabutylammonium (TBuA) induced by various concentrations of TEA.
Xenopus oocytes were injected with rOCT2 cRNA and incubated
as in Fig. 11. A: steady-state currents in an
rOCT2-expressing oocyte clamped at 40 mV. The oocyte was superfused
with Ori buffer, with Ori buffer containing TEA, or with Ori buffer
containing TEA plus TBuA. B: TEA-induced inward currents
from oocytes expressing rOCT2 that were clamped at 40 mV. Each oocyte
was first superfused with various TEA concentrations in the absence of
TBuA. Then the oocytes were superfused with different TEA
concentrations in the presence of 100 µM TBuA (n = 3, mean values ± SE) or 350 µM TBuA (1 oocyte). The data were
normalized to the currents induced by 20 mM TEA, and the
Michaelis-Menten equation was fitted to the data. They show rapid
inhibition of TEA-induced currents by TBuA (A) and suggest
that the inhibition of TEA-induced current is competitive
(B).
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Interaction of rOCT2 and rOCT1 with corticosterone and anions.
Next, we compared the concentration dependence of inhibition of TEA (10 µM) uptake by the uncharged compound corticosterone. In each of three
independent side-by-side experiments, rOCT2 and rOCT1 were expressed,
and the inhibition of expressed TEA uptake was measured within 4 h. The IC50 values were determined for each inhibition
curve. Significantly different mean values of 4.0 µM (rOCT2) and 151 µM (rOCT1) were calculated (Table 3).
The data indicate that corticosterone interacts with both rOCT2 and
rOCT1. They confirm an earlier reported IC50 value for
rOCT2 (40) and show that corticosterone has a 38-fold
higher affinity to rOCT2 compared with rOCT1. Previous measurements in
intact renal proximal tubules and isolated plasma membranes showed
that organic cation transport may be inhibited by organic anions
(14, 32, 33). Because indirect effects on the transport
activity could not be excluded in these experiments, we now
investigated whether anions may inhibit organic cation transport
expressed by rOCT2 or rOCT1 that is structurally similar to the OATs
(2, 18). After preliminary experiments had shown that
rOCT2- and rOCT1-mediated uptake of [14C]TEA was
partially inhibited by 1 mM probenecid, 1 mM
-ketoglutarate (AKG) or 1 mM para-aminohippurate (PAH), we measured the
concentration-dependent inhibition of 10 µM [14C]TEA
uptake by these anions. Similar data were obtained with rOCT2 and
rOCT1. When rOCT2 and rOCT1 were expressed in identical oocyte batches
(Table 3), the IC50 values calculated for rOCT2 vs.
rOCT1 were 0.7 vs. 0.7 mM (AKG), 4.5 vs. 1.3 mM (PAH), and 1.7 vs. 1.6 mM (probenecid). To determine whether the inhibition of cation uptake
by anions is due to a competitive type of interaction rather than to an
allosteric or indirect effect of the anions, we compared the
concentration dependence of TEA-induced inward currents in the absence
and presence of 2 mM PAH. This comparison was performed with 16 oocytes
from 5 different batches expressing rOCT2 that were clamped at
50 mV.
Although the TEA concentration required to induce half-maximal currents
in the absence of PAH showed a large variation between different oocyte
batches, in all experiments a mixed, mainly competitive type of
inhibition by PAH was obtained. At 2 mM PAH, the apparent
Km values increased more than twofold (231 ± 34%, P < 0.001), whereas the
Vmax values decreased slightly (21 ± 3%,
P < 0.01).
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DISCUSSION |
Functional comparison between rOCT2 and rOCT1.
In this study, two OCTs are compared that exhibit similar sites of
expression. In rat kidney, rOCT2 and rOCT1 proteins have been localized
to the basolateral membranes of renal proximal tubules (15,
28). rOCT1 protein is mainly expressed in S1 and S2 segments
with a low expression in the S3 segment, whereas rOCT2 is expressed in
S2 and S3 segments. Because we show in the present study that rOCT2
mediates electrogenic uptake of several structurally different cations,
as has been demonstrated for rOCT1, both transporters may participate
in basolateral uptake of cations in the proximal tubule. This
represents the first step in cation excretion. The efflux measurements
from the present study show, however, that rOCT2 can also mediate
significant electrogenic efflux of choline at physiological membrane
potentials if the outwardly directed choline gradient is high enough.
Because electrogenic efflux has been also described for rOCT1
(22), both transporters may also participate in
basolateral cation export during cation reabsorption. Our observation
that the amount of choline that was taken up during 1-min superfusion
of oocytes expressing rOCT2 with 10 mM choline was sufficient to
generate a transient outward current on switching to choline-free
superfusate at
50 mV (Fig. 1C) suggests a physiological
significance of this transport mode.
Our study shows that rOCT2 and rOCT1 have similar basic functional
properties but exhibit distinct differences that may entail specific
physiological functions. rOCT2 and rOCT1 exhibit similar Km values for many transported cations,
including choline, which has been claimed not to be transported by
rOCT2 (11). At variance for guanidine, rOCT2 has a 10-fold
lower Km value compared with rOCT1. Whereas for
seven organic cations, transport by both rOCT2 and rOCT1 was
demonstrated, we observed that some uptake of quinine was expressed by
rOCT1 but not by rOCT2. Earlier we explained the small amount of
quinine uptake by rOCT1 as binding to the transporter that may be
followed by endocytosis (22). Because we observed
virtually no uptake of quinine with rOCT2, and recent experiments
showed that the permanently charged quinine derivative [3H]N-methylquinine is transported by rOCT1
with a Vmax of nearly 20 pmol · oocyte
1 · h
1
but not by rOCT2 (Ref. 35 and unpublished data),
we no longer exclude the possibility that quinine belongs to the
transported substrates of rOCT1. Taking together the uptake and
electrical measurements with quinine performed so far, we would like to
suggest that rOCT1 mediates inwardly directed transport of quinine
under trans-zero conditions, but is
trans-inhibited by intracellular quinine. This would explain
why quinine is a potent inhibitor of the rOCT1-expressed transport of
other cations, why no rOCT1-expressed uptake of
[3H]quinine could be detected with quinine concentrations
>3 µM (22), and why we were not able to detect inward
currents when we superfused voltage-clamped oocytes expressing rOCT1
with quinine at pH 7.4 (22).
Without knowing the number of functionally expressed transporter
proteins, the turnover numbers of rOCT2 and rOCT1 cannot be compared
directly. For this reason, we compared the relative Vmax values of different cations in side-by-side
experiments, injecting the cRNAs of rOCT2 and rOCT1 into the same batch
of oocytes and performing the transport measurements within 4 h. Thereby, we tried to ascertain comparable assay conditions, e.g., with
respect to membrane potential of the oocytes and concentrations of
endogenous compounds that may interact with transport. In normalizing the Vmax values to the
Vmax of the prototypic cation TEA, relative turnover numbers of the two transporters were compared. Our data show
that the relative turnover numbers of rOCT2 and rOCT1 depend on the
structure of transported cations and that both transporters have
different relative turnover numbers for structurally different substrates. For example, the turnover number of rOCT2 for choline is
two times higher than for TEA, whereas the turnover number of rOCT1 for
TEA and choline is about the same.
Inhibitors and transported cations with 20- to 70-fold different
affinities for rOCT2 and rOCT1 have been identified that help to
determine the individual contribution of rOCT2 and rOCT1 to cation
secretion or reabsorption in vivo. For cation transport measurements in
rat kidney, the contribution of three additional cation transporters
that are also transcribed in the proximal tubule must be considered.
These transporters are the electrogenic cation transporter rOCT3
(16, 38, 40), the cation transporter rOCTN1
(37), and the carnitine/cation transporter rOCTN2
(39). The transcription of these transporters in proximal
tubule has been shown by in situ hybridizations, but the subcellular
localization of the respective proteins has not been determined. The
inhibitor profiles of rOCT3, rOCTN1, and rOCTN2 have been characterized in less detail. Interestingly, our identification of inhibitors with
largely different affinities for rOCT2 and rOCT1 and the previously
reported functional data on rOCT3, rOCTN1, and rOCTN2 suggest a
rationale for experimentally dissecting out the individual contributions of rOCT2 or rOCT1 to cation transport in the intact proximal tubule. It has been reported 1) that rOCTN2 has a
high affinity for carnitine (IC50 value 15.5 µM; Ref.
39), 2) that rOCTN1 and rOCTN2 do not interact
with 5 and 2.5 mM guanidine, respectively (37, 39),
3) that rOCTN1 has a very low affinity for MPP
(IC50 > 5 mM; Ref. 37), 4)
that corticosterone inhibits rOCT3 with about the same affinity
(IC50 = 4.9 µM) as rOCT2 (40), and
5) that
-estradiol inhibits rOCT3 with a IC50
value of 1.8 µM, whereas it inhibits rOCT2 with a IC50
value of 85 µM and does not inhibit rOCT1 at all (40).
Thus transport by rOCT2 in proximal tubule may be analyzed by measuring
uptake of guanidine at a concentration <0.3 mM (excluding transport by
rOCTN1 and rOCTN2) in the presence of 10 µM
-estradiol (inhibition
of rOCT3) plus 200 µM O-methylisoprenaline (inhibition of
rOCT1). Transport by rOCT1 may be analyzed by measuring uptake of MPP
at a concentration <50 µM (excluding uptake by rOCTN1) in the
presence of 0.5 mM carnitine (inhibition of rOCTN2) plus 15 µM
corticosterone (inhibition of rOCT2 and rOCT3). These experimental conditions may be used to determine initial uptake rates of cation transport by rOCT1 or rOCT2 into proximal tubular cells. To determine the role of the individual cation transporters in transcellular cation
movement under steady-state conditions, nonmetabolized substrates
should be employed that are common for all transporters that may be
involved, and only compounds that are not transported themselves should
be used for the inhibition of individual transporters.
Interaction of rOCT2 and rOCT1 with anions and hydrophobic cations.
In the present paper, we show that the organic anions PAH, AKG, and
probenecid are low-affinity inhibitors of rOCT2 and rOCT1. It has been
shown previously through measurements on intact proximal tubules and
membrane vesicles that certain anions interact with organic cation
transport (14, 32, 33). A direct interaction of anions
with cation transporters has not been demonstrated, however. Because
the OATs and OCTs belong to the same protein family, share ~30%
identical amino acids, and have the same predicted membrane topology
(2, 18), the interaction of organic anions with the OCTs
is not surprising. The observation that the inhibition by PAH of
rOCT2-mediated cation transport was partially competitive with TEA
uptake suggests that the substrate binding sites of the OCTs and OATs
share structural features. Thus, for future in vivo measurements and
for drug therapy, the possibility of cross inhibition between
substrates or inhibitors of the organic anion and cation transporters
must be considered.
To investigate the possibility that the OCTs contain allosteric cation
binding sites that may add to the diversity of their interactions with
cations, we performed kinetic studies with hydrophobic cationic
inhibitors on rOCT2 and rOCT1. The investigated inhibitors were either
permanently charged cations (TPeA, decynium-22, or cyanine-863) or the
weak base quinine. With both transporters, similar results were
obtained. Competitive inhibition of TEA uptake was observed with the
transported low-affinity cation NMN, whereas a noncompetitive type of
inhibition was obtained with the high-affinity inhibitors quinine,
cyanine863, decynium22, and TPeA.
Interaction of rOCT2 and rOCT1 with quinine.
Mutagenesis studies with rOCT1 suggested that the noncompetitive
inhibitor TPeA binds close to the substrate binding site of rOCT1
(discussed in Ref. 7), and recent data with inside-out macropatches from Xenopus oocytes showed to our surprise
that quinine inhibits rOCT2 from the intracellular side in a
competitive manner (1). Thus we performed a more detailed
investigation on the interaction of quinine with rOCT2 to determine the
reason for the noncompetitive inhibition by quinine observed with
intact oocytes. Quinine represents a weak base that may permeate the lipid bilayer in its uncharged form. This allowed us to change the
passive membrane permeation experimentally by varying bath pH. Our
findings strongly suggest that quinine permeates the plasma membrane
via nonionic diffusion at pH 7.4 or more alkaline pH and inhibits rOCT2
from the intracellular side. We showed that passive quinine uptake into
oocytes increased with increasing pH, and that the IC50
value for quinine inhibition of rOCT2-mediated cation transport was
decreased from 8 to 1.5 µM when the bath pH was increased from 7.4 to
8.4. At pH 6, where >99% of quinine is positively charged, quinine
probably inhibits rOCT2 by interacting with the outwardly facing
substrate binding site, which has a relatively low affinity for
quinine. At this pH, a competitive inhibition of quinine was observed
that could be washed out more quickly than the inhibition at pH 8.4. The intracellular interaction of quinine with rOCT2 probably occurs at
an inwardly facing substrate binding site, which may have a higher
affinity for quinine than the outward conformation. This interpretation
is strongly supported by electrical measurements on inside-out
macropatches showing that quinine inhibited electrogenic cation efflux
through rOCT2 from the intracellular side with an IC50
value of ~1 µM (1). Because this inhibition was
competitive with TEA and choline at pH 7.4, quinine may interact from
the aqueous phase with the inward-facing substrate site of rOCT2,
rather than uncharged quinine from the lipid phase of the plasma
membrane with a hydrophobic domain of rOCT2. Further experiments are
necessary to determine whether the inhibition of rOCT2 reflects an
interaction of the uncharged or the positively charged form of quinine.
Another unsolved question is why the pH-dependent quinine uptake
increases linearly for 1 h, whereas the pH-dependent change of the
IC50 values for quinine inhibition of TEA uptake was almost complete after 1 min of incubation (see Fig. 10). Also, the
IC50 value for quinine inhibition of TEA uptake at pH 7.4 was similar when the incubation with quinine was performed for 10 min
or 1 h (compare Fig. 9B with Table 2). A possible
explanation for this apparent discrepancy is that uncharged quinine in
the bath equilibrates rapidly with the plasma membrane and the
intracellular aqueous phase where quinine may reach a
quasi-steady-state concentration within minutes. However, the total
amount of quinine in the oocyte increases over a much longer time
period, possibly because the uncharged form of quinine from the
intracellular aqueous phase, representing ~5% of total quinine at an
estimated intracellular pH of 7.1, is slowly trapped by diffusing into
the large pool of yolk lipids. Quinine could be also trapped in acidic
vesicular compartments. Trapping of quinine in an intracellular
compartment may also explain why the concentration of quinine in the
oocytes, after 1 h of incubation at pH 7.4, exceeds the estimated
equilibrium concentration, assuming a homogeneous aqueous compartment
within the oocytes, an oocyte volume of 0.5-1 µl, and an
intracellular pH of 7.1. It would be helpful to investigate the
intracellular distribution of quinine and to determine whether the
permanently charged nontransported inhibitors decynium22, TPeA, and
cyanine863 may also diffuse through the plasma membrane and interact
from the intracellular side of rOCT2, or whether the noncompetitive type of inhibition observed with these compounds has a different reason.
Although several questions have remained open, our data with quinine
show that nonionic diffusion of hydrophobic compounds must be taken
into account in drug therapy, since intracellular drugs may exhibit
high-affinity interactions with inward-facing substrate sites of plasma
membrane transporters and may lead to long-lasting inhibition that may
alter transporter stability.
In conclusion, the excretion and reabsorption of cations in the kidney
are mediated by the successive and parallel action of a set of
polyspecific cation transporters with overlapping substrate
specificities and membrane distributions. The functional role of the
individual transporters for the secretion and reabsorption of specific
endogenous and exogenous cations is not understood because the membrane
localization of most transporters is not known, their substrate
specificities have not been clarified, and the employed transport
mechanisms have not been elucidated in sufficient detail. The situation
is even more complex, since the transporters may be regulated
differentially, nontransported inhibitors may interact from both sides
of the plasma membrane, and the transporters may exhibit significant
interspecies differences in substrate specificity and localization. In
the present study, we showed that the two electrogenic cation
transporters rOCT2 and rOCT1 that exhibit an overlapping substrate
specificity and membrane localization can be functionally distinguished
with the help of inhibitors. In addition, data are presented that
suggest that rOCT2 may operate in both directions at physiological
membrane potentials and that the weak base quinine inhibits rOCT2 from the intracellular side.
 |
ACKNOWLEDGEMENTS |
We thank Irina Schatz for expert technical assistance, Michael
Christof for preparing the figures, and Bernhard M. Schmitt for
critically reading the manuscript.
 |
FOOTNOTES |
*
P. Arndt and C. Volk contributed equally to this work.
This study was supported by the Deutsche Forschungsgemeinschaft Grant
SFB 487/A4.
Address for reprint requests and other correspondence: H. Koepsell,
Anatomisches Institut der Universität, Koellikerstr. 6, 97070 Würzburg, Germany (E-mail: Hermann{at}Koepsell.de).
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. Section 1734 solely to indicate this fact.
Received 21 November 2000; accepted in final form 14 May 2001.
 |
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