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Am J Physiol Renal Physiol 290: F1118-F1126, 2006. First published December 20, 2005; doi:10.1152/ajprenal.00462.2005
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Functional influence of N-glycosylation in OCT2-mediated tetraethylammonium transport

Ryan M. Pelis, Wendy M. Suhre, and Stephen H. Wright

Department of Physiology, College of Medicine, University of Arizona, Tucson, Arizona

Submitted 23 November 2005 ; accepted in final form 14 December 2005


   ABSTRACT
TOP
 ABSTRACT
METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
OCT2, an organic cation transporter critical for removal of many drugs and toxins from the body, contains consensus sites for N-glycosylation at amino acid position 71, 96, and 112. However, the extent to which these sites are glycosylated by the cell, and the influence glycosylation has on OCT2 function, remains unknown. To address these issues, the acquisition of N-glycosylation was disrupted by mutating the amino acid asparagine (N) to glutamine (Q) at these sites in the rabbit ortholog of OCT2, which was expressed in Chinese hamster ovary cells. Disruption of N-glycosylation followed by Western blotting indicated that each site is indeed glycosylated and that OCT2 contains no other sites of N-glycosylation. Plasma membrane expression (determined by surface biotinylation) of the N112Q mutant, but not N71Q or N96Q mutants, was fourfold lower than that of wild-type OCT2, and unglycosylated OCT2 (N71Q/N96Q/N112Q) was sequestered in an unidentified intracellular compartment. The N71Q, N96Q, and N112Q mutants had a higher affinity (~2-fold) for tetraethylammonium (TEA). Maximum transport rate was reduced in the N96Q (3-fold) and N112Q (5-fold) mutants, but not the N71Q mutant, and unglycosylated OCT2 failed to transport TEA (associated with its absence in the plasma membrane). Whereas the reduction in maximum transport rate of the N112Q mutant is consistent with its reduced plasma membrane expression, the lower rate of the N96Q mutant, which appeared to traffic properly, suggests that glycosylation at N96 increases the transporter turnover number.

SLC22A2; organic cation; rabbit

ORGANIC CATION TRANSPORTERS (OCTs) in the SLC22A gene family transport a variety of organic cations (OCs), including clinically important therapeutics (e.g., cimetidine) and environmental toxins (e.g., nicotine). Three OCTs (OCT1, OCT2, and OCT3) have been cloned, and each appears to be restricted to the basolateral membrane of barrier epithelia (32). In the renal proximal tubule, an important site for controlling plasma levels of OCs, all three OCTs are expressed and provide an essential pathway for OC uptake, the first step in tubular secretion (32). Depending on the prevailing set of electrical and chemical gradients, OCT-mediated transport can occur either via electrogenic facilitated diffusion (i.e., driven by the negative membrane potential) or OC/OC exchange (4, 5, 13). In humans, OCT2 (SLC22A2) has been postulated to be the greatest contributor to tubular OC secretion since its expression (as indicated by mRNA and protein levels) predominates over that of OCT1 and OCT3 (19). However, both OCT1 and OCT3 may play significant roles in the renal handling of select OCs (e.g., those transported inefficiently by OCT2) (32, 34).

OCT1, 2, and 3 belong to a larger family of solute carriers (OCT family), which include the OCTNs (OCTN1–3) and OATs (OAT1–5). Members of the OCT family are generally 550–560 amino acids in length and have common structural features, including 12 putative transmembrane-spanning domains, intracellular COOH and NH2 termini, that place these transporters into the larger group of related proteins, the major facilitator superfamily (MFS) (27).

The recent elucidation of high-resolution crystal structures of two MFS transporters, i.e., LacY (1) and GlpT (12), and the suggestion that all MFS transporters may share a common fold (23), has ushered the application of homology modeling methods to the development of hypothetical three-dimensional structures of a number of MFS transport proteins, including the rat ortholog of OCT1 (26) and the rabbit ortholog of OCT2 (38). These models have primarily focused on the organization and alignment of residues within the 12 transmembrane-spanning helices and, in conjunction with studies employing site-directed mutagenesis and computational methods, have been used to identify amino acid residues within a large hydrophilic cleft that play significant roles in substrate binding. Significantly, these homological models have offered no insight into either the structure or function of either the long cytoplasmic loop that is a common element of most MFS proteins or the long extracellular loop that is a unique property of all members of the OCT transporter family (33). Although the placement of this loop between TMHs 1 and 2, both of which appear to comprise a portion of the hydrophilic cleft region in which substrate binding has been proposed to occur (26, 38), suggests that the long extracellular loop may exert a significant influence on OCT structure, little is known of the influence of the long extracellular loop on OCT function. It is hypothesized that alterations to the structure of the long extracellular loop will influence the functional parameters of OCT2.

The high degree of sequence homology that marks the long extracellular loop of OCT family members suggests that it plays an important role in structure/function of these transporters. In addition to a highly conserved set of cysteine residues (6 in all OCTs and 4 in all OATs), the long extracellular loop is also highly glycosylated. The influence of N-glycosylation of the long extracellular loop on activity of OAT1 and OAT4 has received some attention (29, 39). Both of the human orthologs of OAT1 and OAT4 were found to be glycosylated at multiple sites in the long extracellular loop. In hOAT1, disruption of N-glycosylation by replacing asparagine with glutamine at each individual site had no effect on p-aminohippurate (PAH) uptake into HeLa cells expressing the transporter. Elimination of glycosylation at all four sites, however, caused retention of the transporter in an intracellular compartment, thus preventing PAH uptake. Similarly, disruption of N-glycosylation at all four sites in hOAT4, either by mutation or tunicamycin treatment, resulted in impaired trafficking of the protein. With the use of mutant Chinese hamster ovary (CHO)-Lec cells lacking various enzymes required for glycosylation processing, it was shown that processing of glycosylation from a mannose-rich type to a complex type is associated with an increased affinity of hOAT4 for its substrate, estrone sulfate.

The influence of N-glycosylation has not been established for any OC transporter in the OCT family. Of the three putative N-glycosylation sites found in all OC transporters, only one is homologous to a site found in the OATs. Thus the present study examined the functional influence of N-glycosylation on OCT2-mediated transport. The findings reported here demonstrate that N-glycosylation of OCT2 has a profound effect on plasma membrane expression, substrate affinity, and the maximum rate of substrate transport.


   METHODS
TOP
 ABSTRACT
 METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Site-directed mutagenesis. The rabbit ortholog of OCT2 was subcloned into the pcDNA3.1 expression vector as described previously (37). To facilitate immunolocalization of wild-type (WT) transport protein and mutant proteins expressed in transfected cells, the V5 epitope tag (amino acid sequence, GKPIPNPLLGLDST; nucleotide sequence GGT AAG CCT ATC CCT AAC CCT CTC CTC GGT CTC GAT TCT ACG) was added to the COOH terminus of OCT2 using PCR amplification. Mutations of the V5-tagged OCT2 sequence were introduced by site-directed mutagenesis using the Quick Change system (Stratagene, La Jolla, CA) following the manufacturer's instructions. Plasmid DNA was prepared using standard methods (Genesee Scientific, San Diego, CA), and sequencing was used to confirm the presence of mutations and absence of PCR artifacts.

Cell culture and transfection. CHO cells were grown in plastic culture flasks at 37°C in humidified atmosphere (5% CO2). Kaighn's modification (F12K) medium supplemented with 10% fetal calf serum served as the growth medium. Cultures were split every 3 days; 5 x 106 cells were transfected (by electroporation; BTX ECM 630) at 260 V (time constant of ~25 ms) with 10 µg of salmon sperm (Invitrogen, Carlsbad, CA) and 10 µg of plasmid containing the cDNA sequence for the desired construct. Cells were seeded in a T-75 flask following transfection and maintained under G418 (1 mg/ml; Research Products International, Mt. Prospect, IL) selection pressure. Cells were plated at either 275,000 cells/well (12-well plate) for transport and immunocytochemistry, 550,000 cells/well (6-well plate) for surface biotinylation, or 1 x 106 cells per a 10-cm plate for preparation of crude membranes. In initial transport experiments, the amount of tetraethylammonium (TEA) uptake was especially low in some of the mutants, making kinetic analysis difficult. To coax the transporter to the plasma membrane, cells expressing WT and mutant transporters were cultured in DMSO (2% for ~36 h), a known chemical chaperone (3, 21, 28). Cells were confluent ~48 h after plating, at which time experiments were conducted. Immediately before the experiments, cells were rinsed three times (15 min each) with culture media to remove the DMSO. In a single experiment, treatment of CHO cells expressing WT OCT2 with DMSO had no effect on the affinity of the transporter for TEA (53 vs. 57 µM) but increased the maximum rate of transport (45 to 90 fmol·mg protein–1·min–1), an effect consistent with a greater abundance of transporter at the plasma membrane. Cells containing WT and mutants of OCT2 were at the same passage number during all experiments, and several successive passages were used during the course of the study.

Measurement of transport. CHO cells grown to confluence in 12-well plates were rinsed twice (15 min each) with Waymouth's buffer (WB; in mM: 135 NaCl, 13 HEPES-NaOH, pH 7.4, 28 D-glucose, 5 KCl, 1.2 MgCl2, 2.5 CaCl2, and 0.8 MgSO4) at room temperature, followed by incubation in WB containing 1 µCi/ml [3H]TEA (0.05 µM; synthesized by the Southwest Environmental Health Sciences Center, University of Arizona, Tucson, AZ), and in some cases, increasing concentrations of unlabeled TEA. After a predetermined amount of time, the "transport buffer" was removed, and the wells were rinsed three times with 2 ml of ice-cold WB to stop transport. The cells were then solubilized in 400 µl of 0.5 N NaOH with 1% SDS (vol/vol), and the resulting lysate was neutralized with 200 µl of 1 N HCl. Accumulated radioactivity was determined by liquid scintillation spectrometry (Beckman model LS3801).

Immunocytochemistry. CHO cells grown to confluence on coverslips in 12-well plates were washed with PBS (137 mM NaCl, 2.7 mM KCl, 8.0 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.3). All subsequent washes were performed in triplicate at room temperature in PBS. Cells were fixed in ice-cold 100% methanol for 20 min, washed, and incubated for 1 h with mouse anti-V5 antibody (Invitrogen) diluted (2 µg/ml) in PBS. The cells were washed and incubated for 1 h in the dark with FITC-conjugated goat anti-mouse IgG (Invitrogen) diluted to 2 µg/ml in PBS. The cells were washed before staining of the nuclei with propidium iodide (5 µg/ml in PBS; Sigma, St. Louis, MO) for 10 min. Cells were washed again and the coverslips were mounted onto microscope slides using Dako fluorescent mounting media (Dako, Carpenteria, CA). A confocal microscope (Nikon PCM 2000 scan head fitted to a Nikon E800 microscope) was used for detection of immunoreactivity in CHO cells.

Preparation of crude membranes from rabbit renal proximal tubules. Rabbit renal proximal tubules were isolated from adult New Zealand White rabbits (15–20 wk of age) as described previously (10). The use of rabbits followed the Guide for the Care and Use of Laboratory Animals as adopted and promulgated by National Institutes of Health. Crude membranes were prepared using a modification of the method of Ogihara et al. (20). Briefly, isolated tubule pellets (frozen and stored at –20°C) were homogenized in a homogenization buffer (in mM: 230 sucrose, 5 Tris·HCl, pH 7.5, 2 EDTA) containing protease inhibitors (in µM: 200 AEBSF, 0.16 aprotinin, 4 leupeptin, 8 bestatin, 3 pepstatin A, 2.8 E-64; Sigma). The homogenate was centrifuged for 15 min at 3,000 g (4°C), and the supernatant was retained and further centrifuged at 100,000 g for 30 min. The resulting pellet was resuspended in a buffer containing (in mM) 150 KCl, 300 mannitol, and 10 HEPES, pH 7.5. Protein concentration was determined by the Bradford method (Bio-Rad, Hercules, CA). Crude membrane proteins were diluted to 1–2 µg/µl in Laemmli sample buffer (31.2 mM Tris·HCl, pH 6.8, 2.5% beta-mercaptoethanol, 1% SDS, 12.5% glycerol, 0.005% bromophenol blue; Bio-Rad).

Preparation of crude membranes from CHO cells. CHO cells were rinsed twice with PBS and the confluent monolayer was scraped from the 10-cm dish using a cell scraper. The cells were resuspended in 20 ml of PBS and pelleted by centrifugation (230 g) for 10 min at 4°C. The cell pellet was resuspended in 1 ml of lysis buffer (50 mM mannitol, 1 mM Tris-base, pH 7.4 with HEPES) containing the aforementioned protease inhibitors by passing it 20 times through a 27-gauge needle. Insoluble cellular material was removed by centrifugation at 100 g for 5 min at 4°C. The supernatant was centrifuged for 30 min at 15,800 g (4°C), and the resulting pellet was resuspended (by vortexing) in lysis buffer. Protein concentration was determined by the Bradford method. Crude membrane proteins were diluted to 1–2 µg/µl in Laemmli sample buffer.

Isolation of plasma membrane proteins by cell surface biotinylation. The method described here is a minor modification of that used by Tanaka et al. (29). The membrane-impermeant biotinylating reagent sulfosuccinimidyl 2-(biotinamido)-ethyl-1,3-dithiopropionate (sulfo-NHS-SS-biotin; Pierce Biotechnology, Rockford, IL) was used to examine plasma membrane expression of OCT2. All solutions were kept ice-cold throughout the procedure, and long incubations were conducted on ice with gentle shaking. Cells were initially washed three times with 2 ml of PBS/CM solution (containing in mM; 137 NaCl, 2.7 KCl, 8 Na2HPO4, 1.5 KH2PO4, 0.1 CaCl2, and 1 MgCl2, pH 8.0 with NaOH) followed by a 20-min incubation in Sulfo-NHS-SS-biotin diluted to 0.5 mg/ml in PBS/CM. The sulfo-NHS-SS-biotin was removed and the cells were incubated a second time for 20 min with freshly prepared sulfo-NHS-SS-biotin. After biotinylation, the cells were rinsed once briefly with 3 ml of PBS/CM containing 100 mM glycine followed by another 20-min incubation in the same solution. The cells were then lysed in 400 µl of lysis buffer (150 mM NaCl, 10 mM Tris·HCl, 1% Triton X-100, 1% sodium deoxycholate, and 0.1% SDS, pH 7.4) containing protease inhibitors for 1 h, and centrifuged at 15,800 g (4°C) for 30 min. The pellet was resuspended and protein concentration of the resulting cell lysate was determined by the bicinchoninic acid method (BCA; Pierce). Differences in protein concentration of lysates from CHO cells expressing WT and mutants of OCT2 were adjusted until equal. This step was included to ensure that equivalent amounts of biotinylated plasma membrane protein from WT and mutants of OCT2 were loaded onto the streptavidin-agarose beads. Fifty microliters of streptavidin-agarose beads (Pierce) were added to the lysates and incubated overnight at 4°C with constant mixing. After extensive washing with the above lysis buffer, 50 µl of Laemmli sample buffer were added, and the proteins were eluted from the beads by boiling (100°C) for 10 min.

Enzymatic deglycosylation with peptide N-glycosidase F. For deglycosylation, proteins isolated by biotinylation or proteins from crude membranes were denatured at 100°C in Laemmli buffer for 10 min. The denatured proteins were then incubated for 1 h at 37°C in 1x G7 reaction buffer, 1x NP-40, and 500 U of peptide N-glycosidase F (PNGase F; New England BioLabs, Ipswich, MA).

SDS-PAGE, Western blotting, and densitometry. Proteins were separated on 10% SDS-PAGE gels and electrophoretically transferred to a polyvinylidene difluoride membrane. The membrane was blocked for 1 h in blocking buffer [5% nonfat dry milk in PBS-T (PBS containing 0.05% Tween 20)] at room temperature, followed by overnight incubation (4°C) with either mouse anti-V5 antibody (0.1 µg/ml; Invitrogen) or chicken anti-rabbit OCT2 antibody (0.2 µg/ml; Aves, Tigard, OR) diluted in blocking buffer. After extensive washing with PBS-T, the membrane was incubated with either horseradish peroxidase-conjugated goat anti-mouse (0.01 µg/ml) or goat anti-chicken IgG (0.001 µg/ml) diluted in blocking buffer. Following extensive washing in PBS-T, the membrane was incubated in SuperSignal West Femto Maximum Sensitivity Substrate (Pierce), and the secondary antibody was detected on high-performance chemiluminescence film (Amersham Biosciences, Buckinghamshire, UK).

Densitometry was used to examine differences in the level of expression of WT and mutants of OCT2 at the plasma membrane of CHO cells. Immunoreactive intensity of individual bands was determined from scanned images using ImageJ 1.34s (National Institutes of Health). To obtain the level of immunoreactivity, a band on a Western blot was selected, and ImageJ scanned the selected band, averaging the 8-bit gray scale values that were located on each horizontal line. The dimensions of the selection area were set to encompass the largest band on an individual blot, and the selection area was kept the same for each subsequent band analyzed. The average eight-bit gray scale values on each horizontal line were summed to obtain cumulative eight-bit gray scale values for each band. The average background gray scale value was subtracted from each horizontal line average to standardize for differences in background between Western blots.

Statistics. All data are expressed as means ± SE, with calculations of standard errors based on the number of separate experiments conducted on cells at a different passage number. Statistical comparisons were performed using an unpaired t-test (ProStat 3.81; Poly Software Intl., Pearl River, NY).


   RESULTS
TOP
 ABSTRACT
 METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Molecular weight and glycosylation profile of OCT2. Based on the deduced amino acid sequence and secondary structure of OCT2 (human, mouse, pig, and rabbit orthologs), three consensus sites [NX(S/T)] for N-glycosylation (at amino acid 71, 96, and 112) reside in the long extracellular loop between transmembrane domains 1 and 2 of the protein (Fig. 1). However, whether these sites are glycosylated by the cell, and the influence N-glycosylation may have on OCT2 function, remains unknown. To address these issues, N-glycosylation was disrupted by changing the amino acid asparagine (N) to glutamine (Q) at these sites in the rabbit ortholog of OCT2 by site-directed mutagenesis, a strategy used by many other studies that have examined the role of N-glycosylation on membrane transporter function (2, 7, 29). In constructing OCT2 DNA, a V5-epitope tag was placed on the COOH-terminal end of the WT protein and the mutant constructs to facilitate their immunolocalization. The Western blots in Fig. 2, A and B, were probed with an anti-V5 antibody and show OCT2 from crude membranes (Fig. 2A) and plasma membranes (proteins isolated by surface biotinylation; Fig. 2B) from CHO cells expressing the WT transporter. In the crude membranes, which presumably consist of both plasma membranes and membranes from intracellular sources (e.g., endoplasmic reticulum and Golgi), two bands with apparent molecular masses of 85 and 57 kDa were evident. Plasma membrane proteins pulled down with the membrane-impermeant biotinylating reagent sulfo-NHS-SS-biotin and streptavidin-coupled agarose beads produced an 85-kDa band on Western blots. No immunoreactivity was present when sulfo-NHS-SS-biotin was omitted (not shown), showing that OCT2 does not interact nonspecifically with streptavidin, as was shown to occur for the Na+-sulfate cotransporter, NaSi-1 (16). Treating the membrane fractions with PNGaseF produced a decrease in apparent molecular mass of both the 85-kDa band and the 57-kDa band to a common apparent size of ~47 kDa, suggesting that the deglycosylated protein found in both the plasma membrane and in intracellular compartments was the same size. When crude membranes isolated from CHO cells expressing WT OCT2 were probed with an anti-rabbit OCT2 antibody (Fig. 2C), a staining pattern identical to that obtained with the anti-V5 antibody was achieved (Fig. 2A), demonstrating the specificity of both antibodies. The molecular weight profile of OCT2 from crude membranes prepared from rabbit renal proximal tubules was nearly identical to that of rabbit OCT2 expressed in CHO cells (Fig. 2D). Noteworthy is the observation that the 57-kDa band from OCT2 expressed in CHO cells was absent from crude membranes of rabbit renal proximal tubule.


Figure 1
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  		Fig. 1. Secondary structure model of OCT2, based on a homology model of the 3-dimensional structure of OCT2 (38). Putative N-glycosylation sites (at asparagine 71, 96, and 112) present in the long extracellular loop between transmembrane domains 1 and 2 are shown as treelike structures.

 

Figure 2
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  		Fig. 2. Western blot of V5-epitope-tagged wild-type OCT2 from Chinese hamster ovary (CHO) cell crude membranes (A and C) and plasma membranes (B), and OCT2 from crude membranes prepared from rabbit renal proximal tubules (D). Plasma membrane proteins were isolated by surface biotinylation. The proteins were treated with (+) or without (–) peptide N-glycosidase F (PNGase F). Western blots were either probed with a mouse anti-V5 antibody (A and B) or a chicken anti-rabbit OCT2 antibody (C and D). The position of the molecular mass markers (in kDa) is indicated (arrows).

 
Location of N-glycosylation in OCT2. Although the above results indicate that OCT2 is a glycosylated protein, it is not known whether any of the putative N-glycosylation sites contribute to this glycosylation. Figure 3A shows a Western blot of the WT and mutant transporters (except the N71Q/N96Q/N112Q mutant) that were expressed at the plasma membrane. It should be noted that the amount of protein loaded onto the gel differed for the WT and each of the mutants. Thus the level of immunoreactivity is not a direct reflection of the abundance of transporter at the plasma membrane. Evident from Fig. 3 are the differences in the apparent molecular weights of the WT and mutant transporters, with the following general trend: WT > individual mutants (N71Q, N96Q, and N112Q) > double mutants (N71Q/N96Q and N71Q/N112Q). Interestingly, N-glycosylation at each individual site does not contribute equally to the apparent molecular weight of the mature peptide. Enzymatic deglycosylation caused the apparent molecular masses of each mutant protein to shift to the same point, ~47 kDa. Because the triple mutant could not be detected by surface biotinylation, crude membranes were used instead. The size of the triple mutant appeared to be the same as that of unglycosylated OCT2 (i.e., PNGase F treated), and there was no shift of the triple mutant on enzymatic deglycosylation (Fig. 3B). From these data it can be concluded that N71, N96, and N112 are all glycosylated and that OCT2 contains no other sites of N-glycosylation.


Figure 3
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  		Fig. 3. Western blot of wild-type (WT) and mutants of OCT2 either expressed at the plasma membrane of CHO cells (A; WT and the N71Q, N96Q, N112Q, N71Q/N96Q, N71Q/N112Q mutants) or from crude membranes prepared from CHO cells (B; WT and the N71Q/N96Q/N112Q mutant). Putative N-glycosylation sites were disrupted by mutating asparagine (N) to glutamine (Q). Plasma membrane proteins were isolated by surface biotinylation. Proteins were treated with (+) or without (–) peptide N-glycosidase F (PNGase F). Western blots were probed with the mouse anti-V5 antibody. The position of the molecular mass markers (in kDa) is indicated (arrows).

 
Immunocytochemical localization of WT OCT2 and mutants lacking varying degrees of N-glycosylation. The presence in crude membranes of two unique immunoreactive OCT2 bands, and the loss of the lower molecular weight band following surface biotinylation, suggested that a fraction of transporter protein was retained in one or more intracellular compartments, in addition to being successfully trafficked to the plasma membrane. It is clear from Fig. 4 that the WT transporter (OCT2; green) is prominent in both the plasma membrane and in a compartment that is perinuclear (nuclei; red). The N71Q and N96Q mutants exhibited a staining pattern that was similar to the WT transporter. In contrast, the N112Q mutant, both of the double mutants (N71Q/N96Q, N71Q/N112Q), and the triple mutant (N71Q/N96Q/N112Q), were mainly localized to an intracellular compartment, with either no staining or only limited staining visible at the plasma membrane.


Figure 4
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  		Fig. 4. Immunocytochemical localization of OCT2 expressed in CHO cells following elimination of putative N-glycosylation sites. Putative N-glycosylation sites were disrupted by mutating asparagine (N) to glutamine (Q). Cells were incubated with mouse anti-V5 antibody followed by FITC-conjugated goat anti-mouse antibody for detection of OCT2 (green). Nuclei were stained with propidium iodide (red). Insets: highlight the representative distribution of immunoreactive protein within cells expressing each of the OCT2 constructs.

 
Western blot analysis of the plasma membrane expression of WT OCT2 and mutants lacking varying degrees of N-glycosylation. The qualitative data presented in Fig. 4 led to the supposition that N-glycosylation influences trafficking of OCT2 to the plasma membrane. To test this hypothesis, the level of expression of WT OCT2 and each of the mutants at the plasma membrane was examined using surface biotinylation followed by Western blot analysis (densitometry) (Fig. 5). Whereas WT OCT2 and the N71Q and N96Q mutants displayed a similar level of expression, plasma membrane expression of the N112Q mutant was fourfold lower than that of the WT transporter. Although present at the plasma membrane, the abundance of each double mutant was also greatly reduced compared with that of WT OCT2 (5- to 6-fold), and the unglycosylated transporter could only be detected on Western blots when the X-ray film was exposed for very long time periods (not shown). When expression of OCT2 was examined in crude membranes, which presumably represents the majority of the transporter in the cell at the time of membrane isolation, the total amount of the WT and mutant transporter protein (except the N71Q/N96Q/N112Q mutant) was comparable (Fig. 6). Immunoreactivity corresponding to the triple mutant was greatly reduced.


Figure 5
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  		Fig. 5. Effect of disrupting putative N-glycosylation sites on the level of plasma membrane expression of OCT2. Putative N-glycosylation sites were disrupted by mutating asparagine (N) to glutamine (Q). Proteins expressed at the plasma membrane were isolated by surface biotinylation. Western blots were probed with the mouse anti-V5 antibody. Densitometry analysis (top) was performed on 4 different Western blots. Data are presented as a percentage of WT and are means ± SE. *Significantly different from WT (P < 0.05, t-test). Bottom: representative Western blot.

 

Figure 6
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  		Fig. 6. Western blot of total OCT2 expression in crude membranes prepared from CHO cells expressing WT and glycosylation-deficient mutants of OCT2. Putative N-glycosylation sites were disrupted by mutating asparagine (N) to glutamine (Q). Western blots were probed with the mouse anti-V5 antibody. The position of the molecular mass markers (in kDa) is indicated (arrows).

 
Kinetic analysis of TEA transport by WT OCT2 and mutants lacking varying degrees of N-glycosylation. To determine whether removal of N-glycosylation alters OCT2 function, the kinetics of TEA transport into CHO cells expressing WT OCT2 and deglycosylated mutants were measured. Figure 7, left, shows the time course of [3H]TEA uptake into CHO cells expressing WT OCT2. Uptake was approximately linear for 30 min and was almost completely blocked by 2.5 mM unlabeled TEA.


Figure 7
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  		Fig. 7. Left: time course of [3H]TEA uptake into CHO cells expressing WT OCT2. Shown is a representative experiment of 0.05 µM [3H]TEA uptake done in duplicate at each time point. Uptake was measured in the absence and presence of 2.5 mM unlabeled TEA. Right: kinetics of TEA transport in CHO cells expressing WT OCT2. Shown is a representative experiment of duplicate measures of 0.05 µM [3H]TEA uptake (at 2 min) measured in the presence of unlabeled TEA (0–2,500 µM). Lines were fit to the data using a nonlinear regression algorithm (Sigma Plot 2001).

 
The kinetics of TEA transport into CHO cells expressing WT OCT2 are shown in Fig. 7, right. Increasing concentrations of unlabeled TEA in the transport medium reduced the rate of transport by a process adequately described by the Michaelis-Menten equation for competitive interaction of labeled and unlabeled substrate (18):

Formula
where J is the rate of [3H]TEA transport from a concentration of labeled substrate equal to [*T]; Jmax is the maximum rate of transport; Kt is the TEA concentration that results in half-maximal transport (Michaelis constant); [T] is the concentration of unlabeled TEA; and C is a constant representing the component of total TEA uptake that is not saturable over the concentration range tested. This nonsaturable component likely reflects the combined influence of diffusive flux, nonspecific binding, and/or incomplete rinsing of the cell-layer. In four separate experiments, the Jmax and Kt values for TEA transport mediated by WT OCT2 were 7.4 ± 1.4 pmol·cm–2·min–1 and 109 ± 13.3 µM, respectively (Fig. 7B and Table 1). Removal of glycosylation at N71 had no effect on Jmax, but reduced the affinity for TEA by 45% (Table 1). Individual removal of glycosylation at N96 or N112, and simultanous removal at N71 and N96 (N71Q/N96Q), caused reductions in both Jmax (3- to 5-fold) and Kt (~50%). The nonsaturable component of TEA uptake for cells expressing the WT and mutant transporters was not different (data not shown). In two separate experiments, mediated TEA uptake was undetectable at 2 min in cells expressing either the N71Q/N112Q or N71Q/N96Q/N112Q mutants (data not shown). To determine whether the reduction in Jmax resulted from changes in expression of OCT2 at the plasma membrane, Jmax values (data in Table 1) were normalized to the amount of OCT2 protein expressed at the plasma membrane (data in Fig. 5). Whereas the reductions in Jmax of the N112Q mutant were adequately described from its diminished abundance at the plasma membrane, the level of expression of the N96Q and N71Q/N96Q mutants could not explain their low maximum rates of transport (Fig. 8). After correcting for membrane expression, the maximum rate of transport exhibited by the N96Q and N71Q/N96Q mutants was nearly the same and was ~2.6-fold less than that measured for WT OCT2.


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  		Table 1. Kinetics of TEA transport by OCT2 following disruption of putative N-glycosylation sites

 

Figure 8
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  		Fig. 8. The maximum OCT2-mediated transport rate (Jmax) of TEA in WT and glycosylation-deficient mutants of OCT2 before and after correcting for their respective levels of expression at the plasma membrane (data presented in Fig. 5). Data are presented as a percentage of WT. Jmax values before correcting for protein levels (filled bars) were calculated from the data in Table 1 as follows: (Jmax of mutant/Jmax of WT) x 100. Jmax values after correcting for protein expression (open bars) were calculated as follows: [(Jmax of mutant/Jmax of WT) x (gray scale of WT/gray scale of mutant) x 100)].

 

   DISCUSSION
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 ABSTRACT
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Several mammalian orthologs (human, rat, mouse, rabbit, and pig) of OCT2 have been cloned, and each (except the rat ortholog) has consensus sites for N-glycosylation at amino acid position 71, 96, and 112. The rat ortholog of OCT2 is an exception, lacking a consensus site at position 112. Despite the potential for OCT2 to be glycosylated, the extent and location of glycosylation, and the role glycosylation may have on OCT2 transport activity, have not been addressed. In the present study, the rabbit ortholog of OCT2 and several mutants deficient in the ability to acquire glycosylation at the putative sites were studied. Immunocytochemical localization in CHO cells showed expression of WT OCT2 both in the plasma membrane and an unidentified intracellular compartment. Western blotting of crude membranes from CHO cells expressing the WT transporter revealed two bands with apparent molecular masses of 85 and 57 kDa, and enzymatic deglycosylation resulted in a shift to 47 kDa, which is slightly lower than expected (~55 kDa) based on the amino acid sequence. This is not surprising, however, given that membrane proteins frequently migrate faster in SDS-PAGE than molecular weight markers, which are soluble proteins (11). Only the 85-kDa band was observed when plasma membrane proteins labeled with the membrane-impermeant biotinylating reagent Sulfo-NHS-SS-biotin were precipitated with streptavidin-coupled agarose beads. These data suggest that the mature peptide expressed at the plasma membrane is heavily glycosylated. The 57-kDa band observed in crude membranes, but not in the plasma membrane, likely represents partially glycosylated OCT2 that is misfolded and is retained in an intracellular compartment. OCT2 from crude membranes prepared from rabbit renal proximal tubules showed a single band at ~85 kDa, nearly identical to the plasma membrane fraction of rabbit OCT2 expressed in CHO cells. Importantly, the low-molecular-mass band observed in CHO cell crude membranes was not present in rabbit renal proximal tubule, implying that the native tissue is efficient in processing the protein.

Typically, N-glycosylation at a single site contributes ~10–15 kDa to the molecular weight of a protein in SDS-PAGE (11, 22). Thus the approximate 30-kDa contribution of glycosylation to the molecular mass of OCT2 was a priori evidence for glycosylation at multiple sites. Indeed, blocking the acquisition of glycosylation at each individual consensus site (N71, N96, and N112) caused a reduction in molecular mass. The protein migrated to 47 kDa after removing all three glycosylation sites, and there was no further shift following enzymatic deglycosylation. This demonstrates that N71, N96, and N112 are all glycosylated and that OCT2 contains no other N-glycosylation sites. This is in contrast to the human ortholog of OAT1, which contains five consensus sites for N-glycosylation, although typically only four are used (29). Of these four sites in OAT1, only one (N112 in OCT2 and N97 in OAT1) is found in a homologous location within both OCTs and OATs. The presence in OCT2 of glycosylation at N71, N96, and N112 also confirms the extracellular placement of these amino acids in the postulated secondary structure model of OCT2 (38).

N-glycosylation is generally important for the folding, sorting, stability (resistance to proteolysis), and intracellular trafficking of proteins (24). Elimination of glycosylation at position N112, but not N71 or N96, caused reduced expression of OCT2 at the plasma membrane (Figs. 4 and 5). This effect differs from both the mouse and human orthologs of OAT1, where disruption of N-glycosylation at individual sites has no effect on plasma membrane expression (29). Removal of glycosylation at two sites of OCT2 (N71/N96 and N71/N112) greatly diminished plasma membrane expression, and compared with the other mutants (single and double), plasma membrane expression of unglycosylated OCT2 was negligible. This observation is consistent with numerous other studies, in which elimination of glycosylation at multiple sites in a protein was observed to disrupt targeting to the plasma membrane (e.g., 15, 29, 39). In crude membranes, which consist of plasma membranes and membranes from intracellular sources, expression of the WT transporter and each of the mutants, except the triple mutant, was comparable. The lower level of expression of unglycosylated OCT2 suggests either a lower rate of production or a higher rate of turnover for the protein. This latter possibility is exemplified by the observation that the maximum velocity of D-aspartate uptake into C6 astrocytoma cells is reduced by tunicamycin treatment, and this effect is reversed by protease inhibition, suggesting an increased rate of proteolysis of unglycosylated transporters responsible for D-aspartate uptake (9).

The influence of N-glycosylation on OCT2 function was determined by measuring the kinetics of TEA (an organic cation and OCT2 substrate) transport by WT OCT2 and the various glycosylation-deficient mutants. Given their low level of expression at the plasma membrane, it is not surprising that the N71Q/N112Q mutant and unglycosylated OCT2 failed to transport TEA. The maximum rate of TEA transport was diminished following removal of glycosylation at N96 and N112, but not N71. Seeing that the maximum rate of transport was reduced in both the N96Q and N112Q mutants, and that the N71Q/N112Q mutant did not transport TEA, cells expressing an N96Q/N112Q mutant were not generated. The fivefold reduction in the maximum rate of TEA transport exhibited by the N112Q mutant was well correlated with its level of expression at the plasma membrane (4-fold lower than WT OCT2). In contrast, removal of glycosylation at N96 had no effect on plasma membrane expression, suggesting that deglycosylation at this position reduces the turnover number of the transporter. Similarly, deletion of glycosylation in the {gamma}-aminobutyric acid (GABA) transporter has been shown to reduce its turnover number (6, 17). Removal of glycosylation at N71, N96, and N112 increased the affinity of OCT2 for TEA. This contrasts sharply with the results of other studies examining the loss of N-glycosylation on membrane transporter function, where substrate affinity was either found to decrease or not change following disruption of N-glycosylation (2, 16, 30, 35). Of course, OCT2 interacts with a wide variety of OCs, and the consequence of N-glycosylation on TEA transport observed in the present study may not apply to all substrates.

From predictions derived from homological modeling, transmembrane helices 1, 2, 4, 5, 7, 8, 10, and 11 of OCT form a hydrophilic cleft, the proposed region of substrate-protein interaction (26, 38). Numerous amino acids residing in the cleft have already been shown to play an essential role in substrate binding (e.g., 26, 38). However, the influence of the intracellular and extracellular loops on OCT substrate interaction is unknown. Here, we found that N-glycosylation of the long extracellular loop exerted a profound effect on TEA transport. This suggests that the long extracellular loop between transmembrane helices 1 and 2 influences the structural arrangement of amino acids in the substrate binding region. Structural studies show that N-glycosylation can affect the beta-turn content of proteins, and perhaps the formation of covalent and hydrogen bonds in backbone and side-chain atoms (25). Ciarimboli et al. (8) proposed that the intracellular loop between transmembrane domains 6 and 7 of OCT1 may also modulate the structure of the binding region. More in depth studies are required to determine how the intracellular and extracellular loops influence the organization of amino acids associated with interaction of substrate with OCTs.

Sulfo-NHS-SS-biotin is a charged molecule that does not permeate the plasma membrane readily (36). The reagent labels proteins by forming a covalent bond through the N-hydroxysulfosuccinimide ester group with the {epsilon}-amine of lysine residues. However, examination of the secondary structure model of OCT2 (Fig. 1; see Ref. 33 for a more explicit representation of this structure) revealed that there are no lysine residues in any of the extracellular loops. According to the three-dimensional model proposed by Zhang et al. (38), three lysines are present within the hydrophilic cleft. The ability of sulfo-NHS-SS-biotin to pull-down OCT2 in the present study suggests that the cleft containing the putative binding region is an aqueous compartment with access to the extracellular milieu. This hypothesis is also supported by experimental data showing that the OCT2 binding surface is accessible to substrates and inhibitors from either the intracellular or extracellular side of the plasma membrane (31).

Various heterologous expression systems have been used to study the function of membrane transport proteins. N-glycosylation of OCT2 was found to have a profound effect on plasma membrane expression, substrate affinity, and maximum velocity of substrate transport in the present study. Depending on both the species and cell type, the extent of glycosylation a protein receives may be different (14). For example, Kee et al. (15) found that when expressed in yeast (N17 strain), the organic solute transporter, Oatp-1, is functionally inactive, an effect attributed to insufficient glycosylation. Here, we examined N-glycosylation of the rabbit ortholog of OCT2 using CHO cells. Yet, with SDS-PAGE, the molecular weight profile of OCT2 when expressed in CHO cells was indistinguishable from OCT2 derived from native tissue (rabbit renal proximal tubule), consistent with the use of CHO cells as an expression system for studying N-glycosylation of renal transport proteins.

In summary, this investigation demonstrated that N-glycosylation is important for the transport function of OCT2, a process that exerts significant influence on body disposition of OCs, including many of clinical relevance. N-glycosylation at amino acid position 112 was found to be essential for trafficking of OCT2 to the plasma membrane, whereas N-glycosylation at amino acid position 96 exerted a marked influence on the kinetics of the transport cycle. The elevated affinity of OCT2 for TEA following removal of N-glycosylation suggests that the long extracellular loop influences the structural arrangement of amino acids in the region of the protein associated with substrate interaction.


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 ABSTRACT
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This work was supported in part by National Institutes of Health Grants DK-58251, ES-06694, and HL-07249.


   ACKNOWLEDGMENTS
 
The authors thank Dr. C. E. Groves for preparing crude membranes of rabbit renal proximal tubule.


   FOOTNOTES
 

Address for reprint requests and other correspondence: R. M. Pelis, Dept. of Physiology, College of Medicine, Univ. of Arizona, Tucson, AZ 85724 (e-mail: rpelis{at}email.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.


   REFERENCES
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