1 Department of Physiology, Faculty of Science, Prince of Songkla University, Songkla 90112, Thailand; and 2 Department of Physiology, College of Medicine, University of Arizona, Tucson, Arizona 85724-5051
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ABSTRACT |
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To examine directly in real
time the efflux of organic compounds [e.g., organic anions (OAs) such
as fluorescein (FL)] across the luminal membrane of isolated, perfused
renal tubules during net secretion, we devised an approach utilizing a
recently developed epifluorescence microscopy system for continuous
monitoring of fluorescence in the collected perfusate. To illustrate
this approach, we measured the luminal efflux rate of FL in mineral
oil-covered, isolated, perfused S2 segments of rabbit renal proximal
tubules. The washout profile of FL showed a deviation from linearity at time 0 when plotted on a semilog scale, indicating that the
luminal efflux of FL was a saturable process. We were able for the
first time to determine the kinetic parameters of luminal efflux [FL concentration at one-half maximal FL efflux
(Ktlumen) of ~560 µM and maximal
rate of FL efflux across the luminal membrane
(Jmaxlumen) of ~635
fmol · min1 · mm
1]. From
the present study, we conclude that the transport step for OAs across
the luminal membrane of OAs is a carrier-mediated process. This
approach will work to measure luminal transport in real time for any
secreted organic compound that is sufficiently fluorescent to be
measured with commonly available, highly sensitive optical equipment.
transport; kidney; p-aminohippurate; fluorescein
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INTRODUCTION |
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AN ORGANIC ANION
(OA) transport process in the proximal tubules of vertebrate kidneys is
responsible for the excretion of a wide variety of OAs (or weak organic
acids that exist as anions at physiological pH), among which are
potentially toxic substances, including endogenous metabolic waste
products, drugs, and xenobiotics (18, 19). The general
process, for which p-aminohippurate (PAH) and fluorescein
(FL) are prototypes, involves transport into the cells against an
electrochemical gradient at the basolateral membrane and movement from
the cells into the lumen down an electrochemical gradient
(18). Transport into the cells at the basolateral membrane is a tertiary active process, the final step of which is the transport of an OA into the cells against its electrochemical gradient in exchange for a dicarboxylate (DC) [physiologically, -ketoglutarate (
-KG)] moving down its electrochemical gradient by means of a recently cloned OA/DC exchanger (17, 25, 31). The
outwardly directed gradient for
-KG appears to be maintained through
a combination of intracellular metabolism and Na+-coupled
secondary active uptake of
-KG across the basolateral membrane. This
model, first based on studies with renal basolateral membrane vesicles
(17, 25), has now been shown to function in intact renal
proximal tubules from mammals and reptiles (2, 3, 24, 26, 27, 31,
36).
In contrast to the information now available on the mechanism of basolateral transport, very little is known about the process involved in OA efflux across the luminal membrane during net secretion. This movement is down an electrochemical gradient and must be mediated in some fashion to account for the relatively high apparent permeability of the membrane to these hydrophobic substances (7). Studies with rabbit and pig brush-border membrane vesicles are consistent with the idea that the luminal efflux of OAs involves electrogenic-facilitated diffusion (10, 37), but evidence for saturability of this process is still questionable. Although a diverse group of OA transporters located on the luminal membrane has recently been cloned, sequenced, and partially characterized (13, 21, 22, 35), they appear to interact preferentially with large-molecular-weight anionic conjugates and, therefore, appear unlikely to be responsible for the secretion of PAH, FL, and other low-molecular-weight OAs. Aspects of the specificity of luminal PAH efflux from rat proximal tubule cells were examined in an in vivo study by Ullrich and Rumrich (34). However, it has not been possible to measure the kinetics of the luminal transport process (for any organic electrolyte transporter) while it is occurring in the intact renal proximal tubule.
To approach the problem of studying a luminal transport process directly in intact renal tubules, we employed a recently developed epifluorescence microscopy system for continuous monitoring of fluorescence in perfusate collected from tubules isolated and perfused in vitro. We modified the use of this system to measure directly the time course of the washout of FL (or other fluorescent compounds) across the luminal membrane. We showed that the kinetics of a luminal efflux process could be determined from these washout curves. In the process of illustrating this approach, we also clearly demonstrated for the first time that the luminal efflux of OAs involves a saturable process.
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METHODS |
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Chemicals. Spectral grade FL and neutral tetramethylrhodamine-dextran (40,000 mol wt) were purchased from Molecular Probes (Eugene, OR). Light mineral oil and all other chemicals were purchased from commercial sources and were of the highest purity available.
Solutions. A modified rabbit Ringer solution, used throughout the studies as dissection buffer, superfusion bathing buffer, and perfusing solution, consisted of the following (in mM): 110 NaCl, 25 NaHCO3, 5 KCl, 2Na2HPO4, 1.8 CaCl2, 1 MgSO4, 10 sodium acetate, 8.3 D-glucose, 5 L-alanine, 4 lactate, and 0.9 glycine; it was adjusted to pH 7.4 with HCl or NaOH. This solution was gassed continuously with 95% O2-5% CO2 to maintain the pH. The bathing medium also contained 3 g/100 ml neutral dextran (40,000 ± 3,000 mol wt) to approximate the plasma protein concentration. The osmolality of the solution was ~290 mosmol/kgH2O.
Preparation of isolated tubules. New Zealand White rabbits, purchased from Myrtle's Rabbitry (Thompson Station, TN), were killed by intravenous injection of pentobarbital sodium. The kidneys were flushed via the renal artery with an ice-chilled solution containing 250 mM sucrose and 10 mM HEPES, adjusted to pH 7.4, with Tris base. They were then gently removed and sliced transversely by using a single-edge razor. A kidney slice was placed in a petri dish containing ice-chilled dissection buffer aerated with 95% O2-5% CO2. Dissection of tubules from a slice was performed manually from the cortical zone without the aid of enzymatic agents. All dissections were performed at 4°C, but all experiments were performed at 37°C. We used only proximal S2 segments in this study because the S2 segment of the rabbit proximal tubule is the primary site of OA secretion (38).
Perfusion of tubules. The basic in vitro perfusion technique used in these studies was the standard one described previously (5, 6), with the collecting pipette modified so that it had a length of uniform diameter that could be positioned parallel to the bottom of the bathing chamber to serve as a flow-through cuvette (27). Briefly, in this process, each isolated tubule was transferred into a custom-made, temperature-controlled chamber with a coverslip as the bottom. Both tubule ends were held in glass micropipettes, and the tubule was perfused at a rate of ~10-15 nl/min through a micropipette with its tip centered in the tubule lumen. The chamber was continuously superfused with bathing medium at ~3 ml/min, and the temperature of the incoming solution was controlled at 37°C (27).
Determination of FL in collected perfusate. This process has been described previously (27). Briefly, the perfusion chamber was mounted on the stage of an inverted microscope (Olympus IMT-2) fitted with epifluorescence optics. A ×60 oil-immersion objective (1.4 numerical aperture, Olympus) was used to focus excitation light from a 100-W mercury arc lamp and to collect fluorescence emitted from the solution in the collecting pipette. The intensity of excitation light was reduced by a 2.0 neutral-density filter (Oreil, Stratford, CT). FL was excited at 490 ± 10 nm by using a band-pass filter (Oriel) for this wavelength. The excitation light was reflected to the sample with a 490 DRLP dichroic filter (Omega Optical, Brattleboro, VT) which passed >90% of emitted light above 505 nm. The emission fluorescence was first limited to an area of 50 µm diameter by an iris diaphragm. The emission beam was filtered by using a 520 ± 10-nm band-pass filter (Oriel) and counted simultaneously by a photomultiplier tube (Hamamatsu model HC 120-03; Bridgewater, NJ) in photon-counting mode. The fluorescence intensity was integrated at 1-s intervals and saved for subsequent analysis with a MSC II data-acquisition microcomputer interface and software purchased from Oxford Instrument (Oak Ridge, TN). Concentrations of FL were determined from a standard curve constructed at the end of each experiment by retrograde infusion of known concentrations of FL into the collecting pipette, while the bathing solution contained either bathing solution only or bathing solution plus the appropriate concentration of FL (27). The background fluorescence of FL in the bathing medium during transport studies was determined by infusing perfusion solution alone into the collecting pipette, while the bathing solution contained the appropriate concentration of FL for that experiment. The autofluorescence and the appropriate FL background counts were subtracted from the counts obtained during net secretion to yield the absolute FL counts in the collecting pipette. The photon count was then converted into concentration from the standard curve.
Measurement of luminal efflux rate of FL. To examine the rate of efflux of FL across the luminal membrane (JFLlumen) as an isolated process, we employed a new method. We first exposed a perfused tubule to 50 µM FL in the bath. FL at this concentration is ~10 times higher than the Kt, where Kt is the FL concentration that produces one-half of the maximal transepithelial secretion rate (Jmax), for its net transepithelial secretion, as reported previously (27). Therefore, we expected that exposure of a perfused proximal tubule to this concentration of FL in the bath would raise the intracellular concentration of FL enough so that, at steady-state net secretion, the luminal transporters for FL would be saturated or partially saturated. When net secretion had reached steady state, the bathing solution was switched to water-saturated light mineral oil that had been prebubbled with 95% O2-5% CO2 and prewarmed to 37°C. Because FL has negligible lipid solubility, covering a tubule with mineral oil effectively limits FL efflux from the tubule cells to that which occurs across the luminal membrane. When we did not wish to limit efflux to that occurring across the luminal membrane only, we replaced the FL-containing bathing medium with FL-free standard bathing medium.
The rate of efflux of FL across the luminal membrane alone, JFLlumen (in fmol · min
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(1) |
Data presentation. When numerical results are presented to illustrate the method, they are summarized as means ± SE. The n value is the number of experiments. Each experiment involved a single tubule from a different animal.
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RESULTS |
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Washout profile of FL from isolated perfused S2 segment of rabbit
renal proximal tubules.
Using our system for continuously monitoring FL fluorescence in
collected perfusate (27), we examined the washout of FL from perfused tubules. Figure 1 shows the
consecutive washout profiles obtained from a single isolated, perfused
S2 segment of rabbit renal proximal tubule. The tubule was exposed to
50 µM FL in the bath until steady-state secretion of FL was achieved (5-8 min). The bathing medium was then switched to either a
FL-free standard solution or mineral oil. The washout using standard
buffer solution always yielded a similar profile, confirming that the tubule was still intact after an exposure at high concentration of FL
and that the efflux of FL during the subsequent exposure to mineral oil
was not an artifact. The semilog plots of the time course of FL
concentration in the collected perfusate during exposure to standard
FL-free medium in the bath were nonlinear and, significantly, displayed
a concave-upward curvature (Fig. 1). This suggested that FL washout
under these conditions (which permitted both luminal and peritubular FL
efflux from the preloaded tubule cells) was not a simple first-order
process but, rather, a multiexponential process. The efflux profile
obtained with mineral oil covering the tubule (thereby limiting FL
efflux to the luminal membrane) was different (Fig. 1). While also
being nonlinear, it revealed a convex-upward curvature consistent with
a saturable process (Fig. 1). These data demonstrate directly, for the
first time, that the luminal transport step associated with secretion
of a low-molecular-weight OA is a saturable process.
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Determination of kinetic parameters,
Ktlumen and Jmaxlumen, for the
luminal efflux of FL.
Figure 2 shows the
JFLlumen determined from Eq. 1 (see METHODS) as a function of FL concentration
inside the tubule cells, [FL]cell. [FL]cell
was determined as follows. Knowing the perfusate collection rate
(~10-15 nl/min) and the time-dependent concentration profile of
FL in the collected perfusate after the tubule was covered with mineral
oil, we determined the amount of FL lost from the tubule cells over the
time course of the experiment. By integrating the amount of FL thus
determined over time, and deriving the volume of tubule cell water from
the tubule length as described previously (8), we were
able to calculate the concentration of FL inside the tubule cells at
time 0 of the washout period. In a similar fashion, we
determined the decreasing [FL]cell during the washout period. Then we determined the
JFLlumen for each incremental
decrease in cellular FL concentration. These data for each tubule were
fitted to the following Michaelis-Menten model by using a nonlinear
regression model (Enzfitter, Biosoft)
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(2) |
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DISCUSSION |
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In the present study, we demonstrated a novel technical and
analytical approach to real-time measurements of the efflux of fluorescent solutes from cell-to-lumen in intact, isolated perfused renal tubules. To illustrate the utility of this approach, we examined
the efflux of FL (as an example of an OA) across the luminal membrane
of isolated, perfused S2 segments of rabbit renal proximal tubules. We
clearly demonstrated that such efflux involves a saturable and,
therefore, apparently a carrier-mediated process. Moreover, using this
technique, we were able, for the first time, to determine the kinetic
parameters of a luminal OA transporter operating in the physiological
context of the intact, perfused tubule. Comparing the luminal transport
kinetic parameters (Ktlumen of ~560
µM and Jmaxlumen of ~635
fmol · min1 · mm
1) with
those of the basolateral OA/DC exchanger (Kt of
~10 µM and a Jmax of ~498
fmol · min
1 · mm
1 )
(29), we see that the luminal OA transporter has a much
lower affinity but slightly higher capacity than the basolateral
transporter. In conjunction with our previous report of the kinetic
parameters of net transepithelial OA secretion
(Kt of ~4 µM and a
Jmax of ~280
fmol · min
1 · mm
1)
(27), these data support the concept that the OA transport step at the basolateral membrane is rate limiting during net
transepithelial secretion.
In summary, we have introduced a novel approach for the measurement of the kinetics of luminal organic solute transport as it occurs in intact, perfused renal tubules. The method is applicable to the determination of the kinetics of the luminal exit of any solute that is sufficiently fluorescent to be measurable by using commonly available, highly sensitive optical equipment. As an example, we showed the efflux kinetics of FL, which is considered a prototype for general OA secretion, although its only proven interaction with a known transporter is with the basolateral OA/DC exchanger (ROAT1) (30). The transporter with which it interacts during efflux across the luminal membrane has not yet been identified. However, the activities of several other transporters that are suspected of contributing to the renal secretion of different structural classes of OA have been studied by using different fluorescent probes. These include Lucifer yellow [a substrate of ROAT1 and of at least two different secretory processes in the luminal membrane (11)], fluorescein-methotrexate [a suspected substrate of the luminal transporter, OAT-K1 (12)], ochratoxin A [a substrate of ROAT1 (36) and a suspected substrate of OAT-K1 (4)], and fluo 3 [a substrate of the luminal transporter, Mrp2 (16)]. In addition, several fluorescent cationic substrates, including daunomycin (14, 28), quinacrine (15), 4-(4-dimethylaminostyryl)-N-methylpyridinium (20, 32), and the novel compound [2-(4-nitro-2,1,3-benzoxadiazol-7-yl)aminoethyl]trimethylammonium (1), as well as fluorescent derivatives of verapamil (9) and cyclosporin (23), have been used to study organic cation transporters of the kidney and other tissues. All of these fluorescent probes would lend themselves to the study of the kinetics of the luminal efflux step in intact tubules with the approach we have described.
Of course, the use of this technique and the determination of the kinetic parameters are predicated on the assumption that the fluorescent substrate is free (i.e., neither bound nor sequestered) within the cells. We cannot be certain of this for all possible fluorescent substrates. However, in the case of FL, preliminary confocal microscopy data on rabbit S2 segments of proximal tubules under the buffer conditions used in these experiments have provided no evidence of binding or sequestration (S. Shpun and W. H. Dantzler, unpublished observations). In general, it appears that the increasing availability of optically active substrates that can access different renal transport pathways, when used in conjunction with the method described here, offers the promise of substantially increasing our understanding of the transport activity of a heretofore poorly accessible membrane for transport studies: the luminal membrane of renal tubules.
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ACKNOWLEDGEMENTS |
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We thank the Royal Thai Government for its support of A. Shuprisha during the course of this work.
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FOOTNOTES |
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This study was supported in part by National Institutes of Health Grants ES-06757; Training Grants HL-07249, NS-07309, and GM-08400; and Southwest Environmental Health Science Center Grant ES-06694.
Address for reprint requests and other correspondence: W. H. Dantzler, Dept. of Physiology, College of Medicine, Univ. of Arizona, Tucson, AZ 85724-5051 (E-mail: dantzler{at}u.arizona.edu).
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 7 January 2000; accepted in final form 13 July 2000.
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