Introduction

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Many important endogenous metabolites (e.g., choline), therapeutic drugs (amiloride) and agricultural agents (paraquat) are organic cations. Accumulation of these compounds within the body may alter a wide variety of cellular functions. However, rapid elimination of organic cations from plasma by the liver and kidney protects the body from their potentially toxic effects. In addition, smaller subcompartments of extracellular fluid, such as the intraocular fluid and the CSF, are functionally and physically distinct from the plasma. These compartments, like the body as a whole, are protected by specialized systems that minimize accumulation of both endogenous and foreign cations through their active transport into the plasma for subsequent clearance from the body via the renal and hepatic systems (Pritchard and Miller, 1993).

Although organic cation transport mechanisms in the kidney and liver have been well characterized, experimental assessment of the mechanisms used in other tissues, such as the choroid plexus, is limited by the small size and anatomic inaccessibility of the transporting epithelia. Some basic information about choroid plexus organic cation transport has been obtained using in situ ventriculocisternal perfusion (Miller and Ross, 1976; Lanman and Schanker, 1980) and preparations of isolated choroid plexus (Tochino and Schanker, 1965a, 1965b) and apical membrane vesicles (Whittico et al., 1990). However, these techniques do not provide direct access to both faces of the intact choroid plexus epithelium and therefore limit the experimental ability to probe the mechanisms of organic cation transport across the CSF-blood barrier. Thus, we sought an alternative experimental model for the study of transepithelial transport of organic cations by the choroid plexus. Recently, researchers have begun to use primary cultures of choroid plexus epithelial cells to examine other physiological aspects of this tissue, such as synthesis and secretion of transferrin (Tsutsumi et al., 1989), transcytosis of thyroxine (Southwell et al., 1993) and serotonin receptor-linked phosphoinositide hydrolysis (Burris et al., 1991). The objective of this study was to evaluate the morphological and physiological properties of a primary culture system of choroid plexus epithelial cells and its capacity to serve as an in vitro model to access the cellular mechanisms of organic cation transport across the CSF-blood barrier.

	Materials and Methods

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Chemicals. ¹⁴C-TEA bromide (53 mCi/mmol) and ³H-L-proline (40 Ci/mmol) were obtained from American Radiolabeled Chemicals, (St. Louis, MO). [³H]Polyethylene glycol (1.1 mCi/g) was obtained from New England Nuclear NEN Research Products (Boston, MA). Triiodothryonine, prostaglandin E₁, forskolin and epidermal growth factor were obtained from Sigma Chemical Co. (St. Louis, MO). All other chemicals were obtained from commercial sources and were of the highest grade available.

Animals and tissue harvest. Three- to 5-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 under hypothermic conditions with CO₂ before decapitation and removal of the brain. Lateral and fourth plexi from a total of 30 to 36 neonatal rats were removed and placed in ice-cold DME/F12. All media were supplemented with penicillin (100 units/ml) and streptomycin (100 µg/ml).

Cell culture. Tissue was suspended in dissociation buffer that contained 137 mM NaCl, 2.7 mM KCl, 0.7 mM Na₂PO₄, 5.6 mM glucose, 10 mM HEPES (pH 7.4) and 1 mg/ml Pronase and 0.5 mg/ml DNAse I (Boehringer-Mannheim, Indianapolis, IN) (Crook et al., 1981). Cell dispersion was accomplished by incubating the tissue-enzyme mixture at 37°C and intermittently triturating and aspirating the mixture. Pooled aliquots of cells were filtered through 100 µM nylon mesh, and the filtrate was centrifuged and washed twice with DME/F12. Final resuspension was in DME/F12 with 10% Nu-Serum IV (Collaborative Biochemical, Bedford, MA). The cell suspension was plated for 2 to 3 hr (37°C, 95% air/5% CO₂) to allow for fibroblastic cell attachment. Unattached epithelial cells and residual fibroblastic cells were then aspirated, centrifuged and resuspended in minimum essential medium with D-valine substituted for L-valine (GIBCO BRL, Grand Island, NY) and with 10% Nu-Serum IV, 1.5 µM triiodothryonine, 100 ng/ml prostaglandin E₁, 10 µM forskolin and 50 ng/ml epidermal growth factor. Removal of L-valine from cell media inhibits growth of fibroblastic cells (Gilbert and Migeon, 1975). Cells were plated at a density of 4.5 × 10⁵ cells/cm² on solid support (i.e., individual wells of 24-well tissue culture plates, 2 cm²/well, or glass dual-chamber microscope slides, 2 cm²/chamber) or in Falcon Cyclopore membrane inserts (0.64 cm²) precoated with E-C-L cell attachment matrix (10 µg/cm²; Promega, Madison, WI) in 24-well culture plates with identical medium in the lower chamber. Cells were maintained at 37°C in humidified 95% air/5% CO₂. At ~72 hr (day 3), unattached cells were removed as the initial plating media was replaced with similar media that contained 5% instead of 10% Nu-Serum IV. Beginning on day 5, cells were maintained with DME/F12 media containing 5% Nu-Serum IV and the growth promoters listed above. Medium was changed every 2 to 3 days.

TEM. Samples were fixed in Karnovzky's fixative (2% p-formaldehyde and 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.3). Each sample was post fixed for 1 hr in 1.0% OsO₄ in 0.1 M sodium cacodylate buffer, dehydrated through a graded series of ethanol solutions and two changes of propylene oxide and embedded in Spurr's low-viscosity resin. Sections (0.06 µm) were mounted on Formvar-coated slot grids, stained with methanolic uranyl acetate and lead citrate and viewed by TEM on a Phillips 300 electron microscope.

Fixation and cell processing for immunofluorescent staining. Cells were isolated as described above and plated on glass tissue culture chamber slides (Nunc, Naperville, IL). On days 7 to 10, epithelial cells were incubated with 2% formaldehyde and 0.1% glutaraldehyde in PHEM buffer (60 mM PIPES, 24 mM HEPES, 5 mM EGTA, 1 mM MgSO₄, pH 6.9) and fixed for 10 min at room temperature (22°C). Fixed tissue was rinsed with PHEM buffer and then permeabilized with 1% Triton X-100 in PHEM buffer for 10 min. After the tissue was rinsed with PHEM buffer, it was incubated without or with primary antibody for 90 min at 37°C. Next, the tissue was washed four times with PHEM buffer to remove unbound antibody. It was then incubated in the dark with a fluorescein-conjugated secondary antibody for 60 min at 37°C. Finally, the labeled epithelium was sealed with Agua-PolyMount (Polysciences, Warrington, PA), and allowed to dry overnight in the dark. Fixed epithelial cells were viewed by a confocal scanning laser microscope (Zeiss Model 410) with an Ar-Kr laser (488-nm line) and FITC filter set (488 nm, 515 nm emission filter).

Cell uptake studies. On day 11, uptake of radiolabeled substrate by cells plated onto 24-well tissue culture plates was measured. Cells were rinsed and preincubated with aCSF containing 118 mM NaCl, 3 mM KCl, 0.7 mM Na₂PO₄, 18 mM NaHCO₃, 2 mM urea, 0.8 mM MgCl₂, 1.4 mM CaCl₂ and 12 mM glucose, pH 7.4, for 1 hr at 37°C. Transport was initiated by replacement of preincubation buffer with 1 ml of aCSF containing ³H-labeled or ¹⁴C-labeled substrate; cells were incubated for 0 to 90 min. Specific modifications of transport buffers are described in the figure legends. Unless otherwise stated, all incubations were conducted at 37°C in a 95% air/5% CO₂ environment. To terminate uptake, transport buffer was removed, and the cells were rinsed with 3 ml of isotope-free aCSF. Within the transport well, cells were solubilized in 1 ml of 1 N NaOH for 1 hr and neutralized with 1 N HCl. An aliquot of the solubilized cell suspension (800 µl) was retained for determination of protein by a BioRad (Hercules, CA) microassay using bovine serum albumin as a standard. The remainder of the cell suspension was transferred to a scintillation vial. Radioactivity was counted as disintegrations per minute by a liquid scintillation counter using an external quench correction. Uptake of radiolabeled isotope was calculated as picomoles of radiolabeled substrate per milligram of protein. For each sample, cell water volume was calculated from total protein, using the experimentally determined conversion factors: 3.85 × 10⁷ mg of protein/cell and 3.75 × 10⁶ µl of water/cell. [³H]Polyethylene glycol (molecular weight, 4000; 0.1 mg/ml transport buffer) was used to correct for extracellular fluid compartments (<10%). After correction for extracellular substrate, cellular accumulation was calculated from substrate radioactivity per microliter intracellular water and the transport buffer specific activity and expressed as the cell-to-medium concentration ratio.

Tissue uptake studies. Lateral and fourth plexi were harvested as described above. Tissue from a single animal (~1 mg wet wt.) was held in 2 ml of ice-cold, oxygenated aCSF in an individual vial. Transport was initiated by replacement of the dissection buffer with 2 ml of fresh aCSF containing ¹⁴C-TEA, and the vial was immediately gassed with O₂ for ~15 sec. Tissue was incubated for 0 to 90 min at 37°C in a shaking water bath. Uptake was terminated by removing the tissue from the transport buffer and immediately blotting it to remove excess buffer. Tissue was then placed on a preweighed foil disk for determination of tissue wet weight (±0.001 mg). The disk and tissue were placed in a desiccator for 24 hr, and dry weight was determined (±0.001 mg). Finally, the disk was transferred to a scintillation vial containing 2 ml of water to disrupt the cells and extract the radioactivity. After 24 hr, radioactivity was determined as previously described. Tissue accumulation of ¹⁴C-TEA was expressed as pmol/mg of dry wt. [³H]Polyethylene glycol (molecular weight, 4000; 0.1 mg/ml transport buffer) was used to correct for the fraction of ¹⁴C-TEA within extracellular fluid compartments (~15%). Accumulation of substrate was calculated from radioactivity per milligram dry weight tissue and the medium specific activity and expressed as the T/M (i.e., disintegrations per minute per gram of wet weight/disintegrations per minute per milliliter transport buffer).

Statistical analysis. Uptake was measured in triplicate in cells from at least three separate culture preparations (n = 3). For tissue uptake studies, measurements were made separately in plexus tissue isolated from at least four animals. Data are presented as mean ± S.E. Control and experimental means values were compared by Student's t test for paired observations and were deemed to be significantly different when the value was P < .05.

	Results

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Cell morphology. The electron micrograph in figure 1 shows the polarity of intact choroid plexus tissue. The apical pole of the intact epithelium, which faces the CSF within the ventricular compartment, is characterized by abundant microvilli and occasional cilia. Tight junctions between epithelial cells physically separate the ventricular membrane from the basolateral membrane, which faces the interstitial fluid. Cells dispersed from choroid plexus and plated on either solid or permeable support grew as differentiated monolayers. Within 1 week, cells appeared confluent with a "cobblestone" organization characteristic of epithelial cells (fig. 2A). Electron microscopic examination of cultured plexus cells revealed features very similar to the intact tissue. As shown in figure 2B, the cells were polarized, and the surface membrane opposite the plating surface (i.e., the apical membrane) possessed numerous microvilli and occasional cilia. Tight junctions were present between cells, and the cells had basal nuclei, numerous mitochondria and a perinuclear golgi apparatus.

View larger version (155K): [in this window] [in a new window]   Fig. 1.   TEM of a cross section of intact rat choroid plexus epithelium, showing the polarized ultrastructure of the intact epithelium. Note the intercellular tight junctions (TJ), the microvillous ventricular membrane that interfaces the CSF in situ and the proximity of the nuclei to the basolateral membrane.

View larger version (143K): [in this window] [in a new window]   Fig. 2.   Micrographs of primary cultures of rat choroid plexus epithelial cells. A, Light micrographs of cells plated on solid support, showing the cobblestone appearance of confluent cultured epithelia. B, TEM of cells plated on permeable membrane coated with extracellular matrix, showing that ultrastructure of cultured epithelium resembles that of intact choroid plexus epithelium [i.e., tight junctions (TJ), apical microvilli, golgi (G) and basally located nuclei].

Biochemical polarization. Choroid plexus epithelial cells grown on glass microscope slides were processed for immunofluorescent staining to examine the localization of plasma membrane transport proteins, Na⁺,K⁺-ATPase and the facilitative glucose transporter (GLUT-1) (fig. 3). In the intact murine choroid plexus epithelium, Na⁺,K⁺-ATPase is restricted to the ventricular, or apical, membrane (Ernst et al., 1986), and GLUT-1 is confined to the basolateral membrane (Harik et al., 1990; Farrell et al., 1992). In cultured plexus epithelial cells incubated with primary antibody against Na⁺,K⁺-ATPase and a secondary fluorescent antibody, marked immunoreactivity was present within the apical membrane, but none was observed at the basolateral membrane (fig. 3A). In contrast, faint and diffuse fluorescence was observed in cells incubated only with the secondary antibody. In those cells incubated with primary antibody against rat brain GLUT-1 glucose transporter and a fluorescein-labeled secondary antibody, immunoreactivity was abundant at the basolateral membrane but not at the apical membrane (fig. 3B). Fluorescent staining in cells treated only with secondary antibody was minimal and diffuse. Immunostaining for Na⁺,K⁺-ATPase and GLUT-1 was conducted under similar fixation and detergent-solubilization regimens; thus, the distinct localization of Na⁺,K⁺-ATPase to the apical membrane was not the result of limited access to the basolateral membrane.

View larger version (21K): [in this window] [in a new window]   Fig. 3.   Immunofluorescence staining of (A) Na+,K+-ATPase and (B) GLUT-1 brain glucose transporter in primary cultures of rat choroid plexus epithelial cells plated on glass slides. Fixed and permeabilized cells were incubated with polyclonal antibody for Na+,K+-ATPase (A) or monoclonal antibody for GLUT-1 (B) and subsequently incubated with a fluorescein-conjugated secondary antibody. A, Immunoreactivity for Na+,K+-ATPase was restricted to the apical membrane. Arrow, piece of debris adjacent to the apical surface that is useful as a marker for the apical boundary between cells and bathing medium. B, Immunoreactivity for GLUT-1 was restricted to the basolateral membrane (cellular pole adjacent to the plating surface). Arrows, apical (free) surface of the cells.

Transport of organic substrates. Based on the morphological polarization and localization of marker transport proteins, the exposed membrane of plexus epithelial cells grown on solid support was the ventricular membrane. The functional integrity of other transport proteins known to be present in the ventricular membrane of plexus epithelium was examined by measuring uptake of organic substrates by cells grown on solid support (i.e., in plastic 24-well tissue culture plates). It had been demonstrated previously that neutral amino acids, including L-proline, are actively accumulated across the ventricular membrane of both intact (Coben et al., 1971; Lorenzo and Culter, 1969; Wright, 1972a) and cultured (Ramanathan et al., 1996) choroid plexus by a ouabain-sensitive, sodium-coupled transport mechanism. Thus, 30-min uptake of 20 nM ³H-L-proline by cultured plexus epithelial cells was measured (fig. 4). Tissue content of L-proline was 24.2 ± 6.0 nmol/mg of protein/30 min; the mean cell-to-medium ratio was 161.6 ± 73.7 under control conditions. Reduction of external sodium concentration from 138 to 20 mM (i.e., isosmotic replacement of external NaCl with N-methyl-D-glucamine chloride) decreased cellular uptake of proline by ~70% (7.1 ± 0.3 nmol/mg of protein/30 min). In the presence of sodium, ouabain (1 mM) reduced uptake by 50% (11.3 ± 1.0 nmol/mg of protein/30 min). Sodium/L-proline symport at the apical, or luminal, membrane of the renal proximal tubule is electrogenic (Chesney et al., 1991). In cultured plexus epithelial cells, an increase in external potassium concentration (from 3 to 30 mM KCl), which should depolarize membrane potential (Zeuthen and Wright, 1981), reduced L-proline uptake by ~45% (13.0 ± 0.7 nmol/mg of protein/30 min).

View larger version (61K): [in this window] [in a new window]   Fig. 4.   Effects of sodium, potassium and ouabain on 30-min uptake of L-proline by primary cultures of rat choroid plexus epithelial cells grown on solid support. Cells were incubated with 20 nM 3H-L-proline in aCSF containing 118 mM NaCl, 3 mM KCl, 0.7 mM Na2PO4, 18 mM NaHCO3 2 mM urea, 0.8 mM MgCl2, 1.4 mM CaCl2 and 12 mM glucose, pH 7.4 (Control). Experimental conditions were low Na+, NaCl isosmotically replaced with N-methyl-D-glucamine chloride; high K+, KC1 was increased to 30 mM; ouabain, 1 mM ouabain was added to control buffer. * P < .05 vs. control uptake.

Organic cation transport. The general properties of organic cation transport in cultured plexus epithelial cells were examined using ¹⁴C-TEA, the prototypic organic cation used in experimental characterization of organic cation transport in renal epithelium (Pritchard and Miller, 1993). The time course of 10 µM ¹⁴C-TEA uptake by isolated choroid plexus was monitored (fig. 5A); the organic cation was accumulated in a time-dependent manner. At 15 min, tissue TEA content was 53.2 ± 8.4 pmol/mg of dry wt., and the T/M was 1.7 ± 0.1. Tissue content progressively increased, reaching steady state at 60 min of 124.9 ± 0.5 pmol/mg of dry wt.; steady-state T/M values ranged from 4 to 6. TePA (100 µM), a potent inhibitor of TEA uptake in isolated renal proximal tubules (Groves et al., 1994; Groves and Wright, 1995), reduced 15-min tissue content of TEA by ~30% and steady state uptake by 60%. In the presence of TePA, T/M values were slightly greater than unity (15-min T/M = 1.2 ± 0.3; 60-min T/M = 1.6 ± 0.1). TePA-sensitive apical uptake of 10 µM ¹⁴C-TEA by cultured plexus cells grown on solid support was examined in a similar manner (fig. 5B). As in intact tissue, cellular TEA uptake was time dependent, progressively increasing from ~200 pmol/mg of protein at 5 min to 1000 pmol/mg of protein at 90 min in the absence of inhibitor. The corresponding cell-to-medium concentration ratios at these times were 3.2 ± 1.9 and 15.4 ± 7.6. Unlike intact tissue, an apparent steady state in TEA uptake was not reached by 90 min; however, TePA reduced uptake by 50% at 5 through 90 min.

View larger version (23K): [in this window] [in a new window]   Fig. 5.   Effects of TePA on time-dependent uptake of TEA by isolated rat choroid plexus and primary cultures of rat choroid plexus epithelial cells grown on solid support. A, Isolated tissue was incubated with 10 µM 14C-TEA and 0 or 100 µM TePA in aCSF. B, Similarly, cultured cells were incubated with 10 µM 14C-TEA and 0 or 1 mM TePA in aCSF. For isolated tissue and cultured cells, * P < .05 vs. uptake in the absence of TePA.

View larger version (46K): [in this window] [in a new window]   Fig. 6.   Effects of hypothermia and metabolic poisoning on 30-min TEA uptake by primary cultures of rat choroid plexus epithelial cells grown on solid support. Cells were incubated with 10 µM 14C-TEA in aCSF at 37°C (Control), at 4°C or at 37°C with 2 mM NaCN in aCSF. Transport buffer also contained 0 or 1 mM TePA. * P < .05 vs. uptake at 37°C in the absence of inhibitor. TePA did not significantly reduce TEA uptake by cells incubated at 4°C (P > .75) or by cells incubated with NaCN at 37°C (P > .96).

View larger version (15K): [in this window] [in a new window]   Fig. 7.   Effects of external substrate concentration on 5-min TEA uptake by primary cultures of rat plexus epithelial cells grown on solid support. Cells were incubated for 5 min with 0 to 1000 µM 14C-TEA in aCSF at 37°C. The rate of TEA uptake (V) was determined in three primary culture preparations (n = 3). Data shown are from a representative experiment in which the calculated Km and Vmax values for TEA were 350 µM and 15.6 nmol/mg of protein/5 min. Inset, Lineweaver-Burke analysis [i.e., the inverse of the rate of uptake (1/V) vs. the inverse of TEA concentration (1/[TEA]) for this experiment].

View larger version (33K): [in this window] [in a new window]   Fig. 8.   Trans effects of unlabeled TEA on 30-min 14C-TEA uptake by primary cultures of rat choroid plexus epithelial cells grown on solid support. Cells were preloaded for 30 min with aCSF containing 0 or 5 mM unlabeled TEA-Br at 37°C. Cells were rinsed quickly three times with ice-cold aCSF and incubated for 30 min with 10 µM 14C-TEA in aCSF containing 0 or 1 mM TePA. TePA significantly reduced 14C-TEA uptake by cells preincubated without or with unlabeled TEA (P > .05). * P < .05 vs. control uptake (i.e., after preincubation with TEA-free aCSF).

View larger version (68K): [in this window] [in a new window]   Fig. 9.   Effects of organic ions on 30-min TEA uptake by primary cultures of rat choroid plexus epithelial cells grown on solid support. Cells were incubated with 10 µM 14C-TEA in aCSF containing no test compound (control) or 1 mM of each inhibitor. * P < .05 vs. control uptake.

	Discussion

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The extracellular fluid of the central nervous system is physically isolated from the plasma by the blood-brain barrier (i.e., the cerebral capillary endothelial layer) and by the CSF-blood barrier (i.e., the choroid plexus epithelial cells). CSF, which is secreted by the choroid plexus, is functionally continuous with the interstitial fluid bathing the central neurons and serves as a sink for metabolic precursors and waste products, neurotransmitters and drugs. Many important endogenous metabolites (e.g., choline), neurohormones (e.g., serotonin) and therapeutic agents (e.g., vinblastine) are organic cations. Elimination of these and other organic compounds from CSF may occur by wash out as fluid drains through the arachnoid villi into the dura sinuses or by diffusion across CSF-blood barrier. However, low membrane permeability for charged compounds limits exit via the latter route. Thus, an additional means of elimination is needed for clearance of ionized endogenous metabolites and xenobiotics from CSF. Active transport from CSF to plasma by the choroid plexus epithelium provides a critical route for CSF clearance of many organic ions (e.g., Mayer et al., 1960; Schanker et al., 1962). The small size, structural complexity and anatomic location of the choroid plexus have limited the experimental characterization of cellular mechanisms that mediate transport of organic cations across the CSF-blood barrier. This study evaluated the potential of primary culture of choroid plexus epithelial cells as an alternative in vitro model for characterizing cellular mechanisms of organic cation transport across the CSF-blood barrier based on morphological, biochemical and functional properties.

Cell morphology and biochemistry. As suggested by the work of other investigators who examined primary culture systems for plexus epithelium (Crook et al., 1981; Southwell et al., 1993; Tsutsumi et al., 1989), epithelial cells dispersed from rat neonate choroid plexus grew as differentiated monolayers on both solid and permeable support and showed morphological asymmetry characteristic of intact choroid plexus. For example, as shown in figure 2, cultured plexus cells possessed lateral tight junctions, an apical microvillous membrane, apically located mitochondria and basally located nuclei. Furthermore, the biochemical polarity of this primary culture system matched its structural polarity. In intact choroid plexus, the Na⁺,K⁺-ATPase is localized at the ventricular, microvillous membrane (Ernst et al., 1986; Quinton et al., 1973; Siegel et al., 1984; Wright, 1972b), whereas GLUT-1, the sodium-independent brain glucose transporter, is localized at the basolateral membrane (Farrell et al., 1992; Harik et al., 1990; Kalaria et al., 1988). Fluorescence microscopic images of cultured cells incubated with specific antibodies for these membrane marker proteins showed that immunoreactivity for GLUT-1 was confined to the basolateral membrane, whereas that for the sodium pump was restricted to the apical membrane (fig. 3). Thus, this primary culture system maintained structural and biochemical polarity similar to intact choroid plexus epithelium, such that the apical membrane, which would face the CSF-filled ventricular compartment in situ, was exposed in cells grown on solid support.

Functional polarity. The concentration of neutral amino acids in CSF is much lower than that in plasma (Dickinson and Hamilton, 1966; Bito et al., 1966). This transepithelial gradient is generated in part by the active uptake of neutral amino acids across the ventricular membrane of the choroid plexus by a sodium-driven cotransport mechanism (Coben et al., 1971; Lorenzo and Culter, 1969; Ross and Wright, 1984; Wright, 1972a). Cultured plexus cells grown on solid support accumulated the neutral amino acid L-proline in a sodium-dependent manner (fig. 4). Apical Na⁺/neutral amino acid cotransport in other epithelia (e.g., renal proximal tubule) is electrogenic (Chesney et al., 1991). Similarly, an increase in extracellular potassium concentration, which was previously shown to depolarize the ventricular membrane of choroid plexus cells (Zeuthen and Wright, 1981), markedly reduced L-proline uptake in cultured plexus cells (fig. 4). Ouabain, an inhibitor of Na⁺,K⁺-ATPase, which binds only to the ventricular membrane of intact choroid plexus (Quinton et al., 1973; Wright, 1972b) and reduces accumulation of neutral amino acids by isolated choroid plexus tissue (Coben et al., 1971), depressed cellular uptake of L-proline in the cultured cells (fig. 4). Ouabain inhibition of amino acid uptake by plexus cells grown on solid support was consistent with the observed localization of ventricular membrane marker enzyme Na⁺,K⁺-ATPase to the apical pole of the cultured epithelium (fig. 3). Thus, both ouabain sensitivity and potential sensitivity of sodium-dependent L-proline transport in cultured plexus epithelial cells indicated that this primary culture system has not only morphological and biochemical polarity but also functional polarity, qualitatively similar to that of intact choroid plexus.

Organic cation transport. Carrier-mediated transepithelial absorption of both endogenous and xenobiotic organic cations and bases, including choline, NMN, hexamethonium and cimetidine, from CSF has been demonstrated experimentally (Lanman and Schanker, 1980; Miller and Ross, 1976; Schanker et al., 1962; Suzuki et al., 1985). Similarly, these and several other organic cations, including TEA, serotonin and norepinephrine, are accumulated by isolated choroid plexus in vitro (Bárány, 1976; Hug, 1967; Miller and Ross, 1976; Suzuki et al., 1986; Tochino and Schanker, 1965b). However, the membrane transport mechanisms involved in accumulation and transepithelial absorption of organic cations by the plexus epithelium are still poorly understood. General aspects of organic cation transport across the apical membrane were examined in plexus epithelial cells grown on solid support. Cultured cells accumulated the model organic cation TEA in a time-dependent manner (fig. 5B). TePA, a high-affinity substrate for both basolateral and luminal organic cation carriers in the renal proximal tubule (David et al., 1995; Groves et al., 1994), inhibited apical TEA uptake (e.g., figs. 5B and 9), and TePA-sensitive ¹⁴C-TEA uptake was stimulated after preloading of cells with unlabeled TEA (fig. 8). Uptake was reduced when cells were incubated at 4°C (fig. 6). Thus, cultured plexus epithelium expressed energy-dependent, carrier-mediated organic cation transport.