Analysis of the Transmembrane Topology and Membrane Assembly of the GAT-1 gamma -Aminobutyric Acid Transporter*

(Received for publication, January 23, 1997, and in revised form, March 24, 1997)

Janet A. Clark Dagger

From the Laboratory of Cell Biology, National Institute of Mental Health, Bethesda, Maryland 20892-4090

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
REFERENCES


ABSTRACT

The transmembrane topology of the Na+- and Cl--dependent gamma -aminobutyric acid transporter GAT-1 has been studied using protein chimeras in Xenopus oocytes. A series of COOH-terminal truncations was generated to which a prolactin epitope was fused. Following expression of transporter-prolactin chimeras in Xenopus oocytes, the transmembrane orientation of each chimera was determined by testing for protease sensitivity in an oocyte membrane preparation. Data from protease protection assays with GAT-1-prolactin chimeras has shown that residues in the loops connecting hydrophobic domain (HD)3 and HD4 and HD7 and HD8 are accessible to protease in the cytoplasm and suggest the presence of pore loop structures which extend into the membrane from the extracellular face. Such pore loop structures may be involved in the formation of the substrate-binding pocket. Studies presented herein confirm that the NH2 and COOH termini are cytosolic and hydrophobic domains span the membrane in a manner consistent with the predicted hydropathy model for Na+- and Cl--dependent transporters. These data also provide insight into GAT-1 transmembrane assembly and suggest that a complex series of topogenic sequences directs this process. A potential pause-transfer sequence has been identified and may be responsible for the translocational pausing observed in this study.


INTRODUCTION

The Na+- and Cl--dependent transporters are a family of complex integral membrane proteins responsible for the reuptake of many neurotransmitters, amino acids, osmolytes, and a variety of other substrates (1-3). The GAT-1 aminobutyric acid (GABA)1 transporter was the first member of this family for which a cDNA was isolated and characterized (4). Subsequent to identification of a cDNA for the norepinephrine transporter (NET) (5), comparison of the NET amino acid sequence with that of GAT-1 revealed a significant degree of homology and indicated that these transporters might be members of a larger transporter family. The extent and diversity of this transporter family was revealed by the identification of cDNAs for several members with the use of homology cloning strategies. Hydropathy analyses of NET and GAT-1 primary sequences predict that these proteins share many structural features. These include a motif of 12 hydrophobic regions connected by hydrophilic loops; a large extracellular loop connecting hydrophobic domains (HDs) three and four which contains potential sites for N-linked glycosylation; cytoplasmic localization of the amino terminus, based on the lack of a recognizable signal sequence; and cytoplasmic localization of the carboxyl terminus (1-3). In a small number of orphan members of this transporter family, the predicted structure differs in the presence of a larger hydrophilic loop connecting HDs seven and eight with potential sites for N-linked glycosylation (6-8). Some features of this predicted model have been confirmed. Immunocytochemical studies with sequence-directed peptide antibodies have shown that the amino and carboxyl termini of NET are located in the cytoplasm (9). In the same study, the large loop connecting HD3 and HD4 and the loop connecting HD7 and HD8 were shown to be extracellular (9). In addition, several groups have confirmed that the sites for N-linked glycosylation in the large loop connecting HD3 and HD4 are utilized in the norepinephrine, glycine, and serotonin transporters (10-13), consistent with the extracellular placement of this loop. While some features of the predicted model for the transmembrane topology of these transporters have been confirmed, detailed experimental testing of the model has only recently begun (14, 15).

Knowledge of the transmembrane topology of these important proteins is necessary for understanding how they function. Complex or polytopic integral membrane proteins, such as the Na+- and Cl--dependent transporters, are synthesized and assembled into their native structure at the endoplasmic reticulum (ER) (16). Assembly of polytopic proteins is thought to be directed by a series of topogenic sequences which interact with the ER translocation machinery (17, 18), resulting in translocation and integration of the nascent chain across and into the ER membrane. The sidedness of the hydrophilic loops connecting HDs can be utilized to infer the cellular location of the loops and the transmembrane orientation of the HDs. Protease protection assays of nascent chains in vesicles prepared from ER membranes are one approach that takes advantage of the sidedness of integral membrane proteins. This method has been used to study the transmembrane topology of several proteins including the human P-glycoprotein (19-21), the GluR3 glutamate receptor (22), and nicotinic acetylcholine receptor subunits (23). Study of the transmembrane topology of MDR1 (19-21) and GluR3 (22) has shown that models based on hydropathy analyses can be misleading and has resulted in the generation of novel topological profiles for these proteins.

In this study the transmembrane topology of GAT-1 has been studied as a representative member of the Na+- and Cl--dependent transporter family. While the data suggest that the transmembrane topology of GAT-1 HDs is not significantly different from that predicted by hydropathy analysis, the data show that the positioning of some extracellular loops may differ from the predicted model. Protease protection assay data has shown that residues in the loops connecting HD3 and HD4 and HD7 and HD8 of GAT-1-prolactin fusion proteins are accessible to protease in the cytoplasm. These residues may extend into a central pore from the extracellular face forming pore loop structures. These data also suggest that coordinate actions of several topogenic sequences are necessary for translocation and membrane integration of GAT-1.


EXPERIMENTAL PROCEDURES

Materials

[3H]GABA was purchased from DuPont NEN. Restriction enzymes, Vent polymerase, and PNGase F were purchased from New England Biolabs. Taq polymerase was purchased from Promega. SDS-polyacrylamide gels, MultiMark protein standards, and Mark12 protein standards were from Novex.

Plasmid Constructions

GAT-1 GABA transporter cDNA in pBluescript plasmid (pBSSKII(-)), generated as described previously (24), was digested with XhoI and XbaI and subcloned into plasmid JG3.6 (25) for the wild-type construct (pGATA). Polymerase chain reaction was used to generate a construct, pFLAGA-N in which the 5'-noncoding region of pGATA was replaced with a KpnI site followed by a consensus Kozak sequence, an ATG, sequence for the FLAG epitope tag, and nucleotides 4-20 of GAT-1. Amplification of GAT-1 with the FLAG oligonucleotide paired with an oligonucleotide directed to nucleotides 559-582 of the coding sequence yielded a fragment which was digested with KpnI and EcoRI and ligated into digested pGATA to create pFLAGA-N. pFLAGA-C, encoding GAT-1 with the FLAG epitope fused at the carboxyl terminus, was generated by amplification of GAT-1 with an oligo directed against nucleotides 1641-1671 paired with an oligo directed against nucleotides 1780-1797, sequence for the FLAG epitope, a TGA, and an XbaI site. The fragment was digested with XmaI and XbaI and ligated into digested pGATA.

Truncated GAT-1 constructs were generated using polymerase chain reaction with an oligo containing a KpnI site and directed against the 5'-noncoding region (nucleotides -146 to -126) paired with oligos directed against defined regions in the GAT-1 coding sequence and containing a BstEII site (putative extracellular loop 1, EL1, 211-234; putative cytoplasmic loop 1, CL1, 319-339; EL2, 610-627; CL2, 694-711; EL3, 838-855; CL3, 937-954; EL4, 1094-1110; CL4, 1225-1242; EL5, 1348-1365; CL5, 1450-1467; EL6, 1579-1596; see Fig. 1, A and B). Polymerase chain reaction fragments were digested with KpnI and BstEII and ligated into digested JGPRO. JGPRO was created by ligating an EcoRI/SphI fragment from pSPSp+1L.ST.gG.pT, obtained from W. R. Skach and V. R. Lingappa, into digested JG3.6 creating a vector bearing the prolactin epitope tag with a BstEII site engineered just prior to the sequence encoding the tag. PRO, a construct with full-length prolactin, was generated by ligation of a PstI and HindIII fragment from a prolactin construct (BPI) received from W. R. Skach and V. R. Lingappa into pBSSKII(-). Sequence analysis of constructs confirmed the absence of mutations introduced by polymerase chain reaction in all but two constructs, EL5 and CL5. In CL5 a single mutation of C to T (478 in the GAT-1 coding sequence) located in the large loop connecting HD3 and HD4 results in a Pro to Ser change. A nonmutant CL5, Delta CL5, was generated by digesting pGATA and CL5 with EcoRI and AvrII and replacing the mutant fragment in CL5 with that from pGATA. Data obtained from proteinase K experiments with Delta CL5 were identical to those obtained with CL5. In EL5 a mutation of T to A (985 in the GAT-1 coding sequence) is located in HD7 and results in a Cys to Ser change, and a C to G (1131 in the GAT-1 coding sequence) is located in HD8 and results in an Ile to Met change. The T to A mutation in EL5 was corrected by digesting EL5 and CL4 with EcoRI and AvaI and replacing the mutant fragment in EL5 with that from CL4 generating Delta EL5. Data obtained from proteinase K experiments with Delta EL5 were identical to those obtained with EL5. Correction of the mutations in Delta CL5 and Delta EL5 was confirmed by sequencing of the constructs. Sequencing was performed by NAPCORE (Nucleic Acid/Protein Research Core Facility, The Children's Hospital of Philadelphia).


Fig. 1. Predicted transmembrane topology of GAT-1 and GAT-1-prolactin chimeras. A, schematic of the transmembrane topology of GAT-1 GABA transporter as determined by Kyte-Doolittle hydropathy analysis of the amino acid sequence (4). Barrels represent proposed hydrophobic domains and potential sites for asparagine-linked glycosylation are denoted by tree-like structures. Points of truncation for each of the GAT-1-prolactin chimeras are noted. EL, extracellular loop; CL, cytoplasmic loop. B, GAT-1-prolactin chimeras used to map the transmembrane topology of GAT-1. Darkened ovals represent proposed hydrophobic domains. Shaded rectangles designate the prolactin epitope which has been fused in-frame to the COOH terminus of defined locations in GAT-1. Sites predicted to be N-glycosylated are denoted by a line with a circle. The cross-hatched region of the PRO construct represents the endogenous signal peptide sequence and the white box is the intervening prolactin coding sequence.
[View Larger Version of this Image (20K GIF file)]

RNA Transcription

mRNA was transcribed from the GAT-1 fusion protein constructs with 1 µg of linearized DNA using T7 RNA Polymerase according to the mMessage mMachineTM protocol (Ambion, Inc.).

Xenopus laevis Oocyte Expression

X. laevis were purchased from Xenopus (Ann Arbor, MI) and oocytes were dissected and prepared as described previously (26). 25 µCi of Trans35S-label (ICN Biomedicals Inc.) (0.25 µl of 10 × concentrated solution) was added to 1 µl of 250 ng/µl transcript and 50 nl of this solution injected per oocyte. Following incubation at 18 °C for 4-6 h, oocytes were pooled and homogenized on ice in homogenization buffer (0.25 M sucrose, 50 mM potassium acetate, 5 mM magnesium acetate, 1.0 mM dithiothreitol, 50 mM Tris, pH 7.5). Following homogenization CaCl2 was added to 10 mM final concentration.

Protease Protection Assay

Proteinase K (Boehringer Mannheim) was added to aliquots of oocyte homogenates (0.2 mg/ml final) in the presence or absence of 1% Triton X-100 and incubated on ice for 1 h. Protease was inactivated by addition of 10 mM phenylmethylsulfonyl fluoride (Sigma) in Me2SO, dilution with 10 volumes of 1% SDS, 0.1 M Tris, pH 8.0, and boiling for 5-10 min. Samples were diluted with 4 volumes of 1.25 × RIPA buffer (187.5 mM NaCl, 1.25% Triton X-100, 1.25% deoxycholate, 62.5 mM Tris, pH 8.0, 2.5 mM EDTA, 0.125% SDS) and set at 4 °C with rocking ~12-16 h. Samples were centrifuged at 14,000 × g for 15 min and proteins immunoprecipitated from the supernatant as described below.

Carbonate Extraction

Oocyte homogenates were extracted with sodium carbonate as described by Fujiki et al. (27) and Skach et al. (21). Briefly, homogenates were diluted in 400 volumes in either 0.1 M sodium carbonate, pH 11.5, or 0.1 M Tris, pH 7.5, and set on ice for 30 min. Membranes were pelleted by centrifugation at 50,000 rpm (230,000 × g) for 30 min using a 70.1 Ti rotor (Beckman). Proteins in the supernatants were precipitated with 15% trichloroacetic acid and pelleted. Trichloroacetic acid pellets and membrane pellets were dissolved in 1% SDS, 0.1 M Tris, pH 8.0, 100 µM 4-(2-aminoethyl)benzenesulfonyl fluoride (ICN Biomedicals, Inc.) and samples were boiled for 5-10 min. Following addition of 4 volumes of 1.25 × RIPA buffer, transporter-prolactin chimeras were immunoprecipitated as described below.

PNGase and Endo H Treatment

Oocyte homogenates were denatured and peptide:N-glycosidase F (PNGase, 1000 units) or endoglycosidase H (Endo H, 2500 units) treatment performed at 37 °C for 1 h, according to the New England Biolabs protocol provided with the enzymes. Samples were solubilized in 1 × RIPA buffer, as described above, and subsequently immunoprecipitated.

Immunoprecipitation

Fusion proteins were immunoprecipitated with rabbit anti-ovine prolactin polyclonal serum (ICN Biomedicals, Inc.) at 1:500 for 4 h at 4 °C. Protein A Affi-Gel (50 µl) (Bio-Rad) was added and samples were set at 4 °C for 3 h followed by 3 washes with 1 × RIPA buffer and 2 washes with 0.1 M NaCl, 0.1 M Tris, pH 8.0. Proteins were eluted from Protein A Affi-Gel in 2 × Laemmli buffer (28) at 55 °C for 15 min. Samples were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (PAGE) and imaged on the Fujix PhosphorImaging system (Fuji).

Transfection of CV-1 Cells

CV-1 cells were plated at a density of ~2-3 × 105 cells per well in 24-well plates directly in the well or on 1.2-cm diameter coverslips. Cells were transfected with 1 µg of DNA/well using LipofectACE (Life Technologies, Inc.) at a ratio of 1:5 DNA/lipid (w/w). Cells were incubated with the DNA/lipid solution at 37 °C for 16-24 h at which time the medium was removed and replaced with complete medium. At 72 h post-transfection, cells were assayed for transport and immunocytochemistry was performed.

Transport Assay

Transfected cells were incubated with [3H]GABA (50 nM, 1 µCi/µl) in a modified Krebs-Ringer-HEPES buffer (24) in the presence of 100 µM aminooxyacetic acid, a GABA transaminase inhibitor, for 30 min at 37 °C. Uptake was stopped by placing cells on ice and washing with ice-cold assay buffer. Cells were solubilized in 1% SDS, and the amount of accumulated [3H]GABA was determined by liquid scintillation counting.

Immunocytochemistry

CV-1 cells transfected with pFLAGA-N or pFLAGA-C were rinsed with phosphate-buffered saline and fixed by incubation in 4% formaldehyde in phosphate-buffered saline for 30 min at room temperature. Fixed cells were incubated in blocking solution (1% bovine serum albumin, 3.3% normal goat serum in phosphate-buffered saline) in the presence or absence of 0.1% Triton X-100 for 60 min at room temperature. Cells were incubated in blocking solution for 16 h prior to addition of anti-FLAG M2 monoclonal antibody (Eastman Kodak Co.) at 1:300 in blocking solution. A 1-h incubation with primary antibody was followed by three 5-min washes with blocking solution. Incubation with Cy3-conjugated AffiniPure goat anti-mouse IgG (Jackson Immunoresearch Labs., Inc.), at 1:300-1:500 in blocking solution, was done for 1 h followed by three 5-min washes with blocking solution. Cells were then covered with coverslips after adding SlowfadeTM reagent (Molecular Probes). Observation of the fluorescence staining was performed with a laser scanning microscope (Zeiss, LSM 410, at the Light Imaging Facility, NINDS, Bethesda, MD).


RESULTS

Assembly of HD1 through HD6 of GAT-1 Requires Cooperative Actions of Topogenic Sequences

To examine the transmembrane topology of the GAT-1 GABA transporter a series of constructs were generated in which the transporter sequence was truncated at the COOH-terminal region of each of the putative hydrophilic loops connecting the putative HDs (see Fig. 1A). Each of the truncated transporter fragments were ligated into a vector (JGPRO) in-frame with the sequence for the carboxyl-terminal fragment of the secretory protein prolactin (codons 56-199). This prolactin fragment lacks intrinsic translocation activity and has previously been shown to serve as a faithful reporter for translocation when following topogenic sequences in chimeric proteins (17, 20, 22, 23). GAT-1-prolactin chimeras are diagrammed in Fig. 1B. Transcripts were made from the constructs and injected into X. laevis oocytes. Following incubation for 4-6 h, oocytes were homogenized. Endo H sensitivity, a reliable marker of protein residence in the ER membrane, was used to confirm chimera residence in the ER, and absence from the plasma membrane, at 4-6 h post-injection (data not shown). Because the chimeras have not exited the ER, it is evident that the membrane vesicles containing the chimeras display a single orientation. Furthermore, subcellular localization of GAT-1 expressed in oocytes as late as 2-5 days following transcript injection revealed that the vast majority of the transporter remained in intracellular compartments (29). Protease protection assays were used to determine the transmembrane spanning abilities of hydrophobic domains and the cellular location of regions connecting these domains. Nascent chains in the ER are oriented such that extracellular domains are located in the ER lumen and cytoplasmic domains are on the cytosolic face of the ER membrane. Exposure of nascent chains in vesicles prepared from ER membranes to proteinase K, a protease that cleaves nonspecifically, results in cleavage of cytoplasmically exposed domains while domains in the ER lumen are protected. The data for study of the amino terminus and HD1 to HD6 are presented in Fig. 2, A-G. Proteinase K treatment of homogenates from oocytes expressing full-length prolactin shows that membrane vesicles in the homogenate are intact as prolactin is fully protected from the protease (Fig. 2A). The intensity of the protected prolactin band is nearly identical to the intensity of the prolactin control band (Fig. 2A), indicating that the majority of vesicles in the Xenopus oocyte membrane homogenate are in the correct orientation. Prolactin is digested in the presence of proteinase K and a nonionic detergent (Fig. 2A) indicating that prolactin is not protease-insensitive. A protease-resistant band of 14-15 kDa was detected for the full-length prolactin as well as each of the GAT-1-prolactin fusions tested. This fragment corresponds to the predicted size of the prolactin tag and is not indicative of a particular membrane orientation. The GAT-1-prolactin chimera extracellular loop 1, EL1, was protected from proteinase K digestion in the absence but not the presence of a nonionic detergent (Fig. 2B). These data strongly suggest that the first hydrophobic domain (HD1) has been translocated into the ER lumen but that HD1 has not yet integrated into the membrane (schematized in Fig. 2B) since the amino terminus also has been protected from digestion. Cytoplasmic loop 1, CL1, was partially digested by proteinase K as indicated by the reduced size of the protected fragment in Fig. 2C (open arrow, lane 2). Comparison of the size of the untreated chimera with the protected fragment shows a difference of approximately 6 kDa. Partial digestion of CL1 indicates that HD1 has become associated with the membrane, and that a portion of CL1 slightly larger than the amino terminus has been digested. These data suggest that the amino terminus of CL1 is in the cytosol. No protected fragments were detected with EL2 (Fig. 2D), the chimera with the prolactin tag fused at the COOH terminus of the large loop connecting HD3 and HD4 and containing three canonical sites for N-linked glycosylation (Fig. 1A). This finding was surprising as several groups have shown that the sites for N-linked glycosylation in this loop are utilized, and therefore that this loop is extracellular (10-13). An 18-kDa fragment (approximately 2 kDa without the prolactin tag) was detected following protease treatment of CL2 (Fig. 2E, open arrow, lane 2). The approximate point of digestion of CL2 based on gel mobility is the NH2-terminal side of HD4 suggesting that the COOH-terminal part of the large loop connecting HD3 and HD4 extends into the membrane to the extent that it is accessible to protease. Protected fragments were detected following proteinase K treatment of EL3, whereas no protected fragments were detected following treatment of CL3 (Fig. 2, F and G). These data confirm that the loop connecting HD5 and HD6 is extracellular and that connecting HD6 and HD7 is intracellular as predicted by hydropathy analysis (Fig. 1A). The digested fragments from EL3 (Fig. 2F, open arrows, lane 2) are 7 and 17 kDa smaller than the full-length chimera indicating that the approximate points of digestion were HD1 and the NH2-terminal side of the loop connecting HD3 and HD4, respectively. The inaccessibility of the loop connecting HD4 and HD5 to protease in chimera EL3 suggests that this chimera has assumed a conformation where this loop is buried and thus inaccessible to protease.


Fig. 2. Proteolysis and immunoprecipitation of GAT-1-prolactin chimeras EL1 through EL3 and CL1 through CL3, and schematics of chimera transmembrane topology. Homogenates of Xenopus oocytes expressing chimeras were treated without protease (lane 1), with proteinase K (Prot. K) (lane 2), or with protease and Triton X-100 (TX-100) (lane 3), as indicated. Exposure to protease was terminated after 60 min, digested chimeras were immunoprecipitated with prolactin antisera, and immunoprecipitated proteins were analyzed by SDS-PAGE using 4-20% acrylamide gels. Closed arrows (lane 1) indicate untreated fusion proteins, open arrows (lane 2) indicate protected fragments, and the asterisk (A, lane 1) denotes the prolactin epitope. Units for molecular weight standards are kilodaltons. Three to five independent experiments were performed with each construct yielding highly reproducible results. Full-length (A) prolactin and (B) EL1 are protected from protease digestion, but digested in the presence of protease and detergent. The prolactin epitope is protected from protease in chimeras CL1 (C), CL2 (E), and EL3 (F) and digested in the presence of detergent. The prolactin epitope is susceptible to protease in chimeras EL2 (D) and CL3 (G) under nondetergent conditions. The transmembrane topology with respect to the ER membranes for each of these GAT-1-prolactin chimeras is diagrammed based on the protected status of the prolactin epitope after exposure to protease. Sites of protease cleavage for the two protected fragments for EL3 (F, a and b) have been denoted in the schematic.
[View Larger Version of this Image (40K GIF file)]

The protease protection assay data showed that each of the GAT-1-prolactin chimeras was translocated across the ER membrane, but did not address the membrane integration abilities of these proteins. Polypeptides were extracted from membranes in 0.1 M sodium carbonate, pH 11.5. Under these conditions peripheral membrane proteins and luminal polypeptides are extracted from the membranes while integral membrane proteins remain associated and readily sediment with the lipid bilayer (27). The secretory protein prolactin as well as EL1 are found in both the supernatant and pellet of Tris and carbonate-treated membranes (Fig. 3). In contrast, CL1 and CL2 are found in the Tris and carbonate pellets only (Fig. 3). EL2 and each of the remaining chimeras, EL3 through EL6 and CL3 through CL5, were found in the Tris and carbonate pellets following extraction (data not shown). The appearance of EL1 in the supernatant of the carbonate-extracted membranes confirms that HD1 has not completed integration into the membrane, while all of the chimeras with the exception of EL1 have integrated. The presence of prolactin and EL1 in the Tris supernatant is most likely due to lysis of some of the membrane vesicles in the procedure. The presence of some prolactin and EL1 in the carbonate pellet could be due to attachment of a portion of the nascent chains to ribosomes or to the presence of a fraction of sealed vesicles.


Fig. 3. Membrane integration status of PRO, and the GAT-1-prolactin chimeras EL1, CL1, and CL2. Xenopus oocyte homogenates expressing PRO, EL1, CL1, or CL2 were incubated either at pH 7.5 (Tris) or 11.5 (Carb). Membranes were pelleted by centrifugation and proteins immunoprecipitated from both the supernatant (S) and the pellet (P) with the prolactin antisera as described. The distribution of EL1 protein resembles that for the secretory protein prolactin whereas the chimeras CL1 and CL2 were found associated with the pellets only.
[View Larger Version of this Image (73K GIF file)]

The large loop between HD3 and HD4, containing sites for N-linked glycosylation, appears to be located in the cytosol based on data obtained in protease protection assays with EL2. If this loop is cytosolic then EL2 should not be glycosylated. To determine the glycosylation state of chimeras, homogenates were treated with PNGase F an enzyme that cleaves oligosaccharides between the innermost N-acetyl moiety and asparagine residues. Neither CL1, which has no potential sites for N-linked glycosylation, nor EL2 were affected by treatment with PNGase F (Fig. 4). In contrast, CL2 and EL3 (Fig. 4), and all remaining chimeras (data not shown), showed increased mobility on SDS-PAGE after treatment with PNGase F indicating that sugar moieties have been removed from these chimeras. These data indicate that a topogenic sequence in HD4, or in the loop connecting HD4 and HD5, is directing the extracellular orientation of the large loop connecting HD3 and HD4 such that it undergoes glycosylation. Taken together the PNGase F and protease protection data show that HD1 through HD6 attain a transmembrane orientation similar to that predicted by hydropathy analysis. However, these data point to potential differences in the membrane association of HD1 and the membrane association of the COOH-terminal region of the loop connecting HD3 and HD4.


Fig. 4. Glycosylation state of GAT-1-prolactin chimeras CL1, EL2, CL2, and EL3. Xenopus oocyte homogenates expressing GAT-1-prolactin chimeras were incubated in the absence or presence of PNGase F for 60 min and proteins immunoprecipitated with the prolactin antisera as described. PNGase F treatment did not change the migration of CL1 or EL2 as compared with untreated chimera. Mobilities of CL2 and EL3 were significantly increased following PNGase F treatment.
[View Larger Version of this Image (33K GIF file)]

Transmembrane Topology of GAT-1 HD7 through HD12 Resembles the Model Predicted by Hydropathy Analysis

With the exception of EL4 and CL4, protease protection assays with chimeras designed to test loops connecting HD9 through HD12 yielded data consistent with the predicted topology for GAT-1. There were no detectable fragments on SDS-PAGE following proteinase K digestion of EL4 (Fig. 5A), suggesting a cytosolic placement for this loop. However, protease treatment of CL4 resulted in the protection of a fragment of approximately 20 kDa (4 kDa without the prolactin tag) (Fig. 5B) indicating that HD7 has integrated into the membrane and that the COOH-terminal portion of the loop connecting HD7 and HD8 is accessible to protease. Proteinase K treatment of CL5 resulted in no detectable fragments (Fig. 5D) which suggests a cytosolic placement of the loop connecting HD10 and HD11. Several protected fragments were detected for both EL5 and EL6 (Fig. 5, C and E) suggesting that the nascent chain goes through various conformational states in acquiring the appropriate topology and as a result not all cytosolic loops are accessible to the protease. The sizes of the protected fragments reveal sites in the chimera which are accessible to protease. Protected fragments of 29, 27, and 21 kDa were detected for EL5 (Fig. 5C). The approximate sites of cleavage for the 29 and 27 kDa (13 kDa and 11 kDa without the prolactin tag) are in the loop connecting HD7 and HD8, while the approximate site of cleavage for the 21-kDa fragment (5 kDa without the prolactin tag) is in the loop connecting HD8 and HD9. These data confirm the protease accessibility of the loop connecting HD7 and HD8 that was detected with the CL4 chimera. In addition, these data place the loop connecting HD8 and HD9 in the cytosol, consistent with the predicted transmembrane topology of GAT-1. Despite the presence of a single mutation in HD8 (Ile to Met, as described under "Experimental Procedures") the protease protection data obtained with EL5 is confirmed by data obtained with EL6 suggesting that the mutation in EL5 has not had an effect on transmembrane topology. Four fragments were detected following treatment of EL6 with protease (34, 31, 28, and 19 kDa) (Fig. 5E). Approximate sites of cleavage for the 34- and 31-kDa fragments (18 and 16 kDa without the prolactin tag) are in the loop connecting HD7 and HD8 and in the NH2 terminus of HD8, respectively, confirming that at least part of this loop is accessible to protease. The 28- and 19-kDa fragments (12 and 4 kDa without the prolactin tag) are generated by cleavage of the loop connecting HD8 and HD9, and cleavage of the NH2 terminus of HD11, respectively. These data confirm the placement of the loops connecting HD8 and HD9 and HD10 and HD11 in the cytosol. Together these data confirm the predicted transmembrane orientation of GAT-1 HD7 through HD12 and reveal that part of the loop connecting HD7 and HD8 extends into the membrane such that it is accessible to protease.


Fig. 5. Proteolysis and immunoprecipitation of GAT-1-prolactin chimeras EL4 through EL6 and CL4 through CL5, and schematics of chimera transmembrane topology. Homogenates of Xenopus oocytes expressing chimeras were treated as described in the legend for Fig. 2 and samples were analyzed on 8-16% acrylamide gels. Closed arrows (lane 1) indicate untreated fusion proteins and open arrows (lane 2) indicate protected fragments. Three to five independent experiments were performed with each construct yielding similar results. The prolactin epitope is susceptible to protease in chimeras EL4 (A) and CL5 (D) under nondetergent conditions. The prolactin epitope is protected from protease in chimeras CL4 (B), EL5 (C), and EL6 (E) and digested in the presence of detergent. The transmembrane topology with respect to the ER membranes for each of these GAT-1-prolactin chimeras is diagrammed based on the protected status of the prolactin epitope after exposure to protease. Sites of protease cleavage for the protected fragments for EL5 (C, lane 2, a, b, and c) and EL6 (E, lane 2, a, b, c, and d) have been denoted in the schematic.
[View Larger Version of this Image (52K GIF file)]

Amino and Carboxyl Termini of GAT-1 Are Located in the Cytosol

The transmembrane location of GAT-1 amino and carboxyl termini was studied using immunocytochemistry of GAT-1 with the FLAG epitope fused at the amino (FLAGA-N) and carboxyl (FLAGA-C) termini. Transport of GABA by the FLAG-GAT-1 chimeras transiently expressed in CV-1 cells was equivalent to that of wild-type GAT-1 (data not shown). Fig. 6 shows the results of immunocytochemistry of the FLAG-GAT-1 chimeras with the FLAG M2 monoclonal antibody in the absence and presence of detergent. There is no immunofluorescence in the absence of detergent (Fig. 6, A and C) showing that the epitopes are not accessible to the antibody. In contrast, permeabilization of the cells makes the epitopes readily accessible to the antibody and results in robust immunofluorescence (Fig. 6, B and D). These data are consistent with a cytosolic localization of both the amino and carboxyl termini of GAT-1.


Fig. 6. Immunofluorescence microscopy of FLAGA-N and FLAGA-C expressed in CV-1 cells. At 72 h post-transfection, transiently transfected CV-1 cells were fixed and incubated with monoclonal FLAG-M2 antibody in the absence (A and C) or presence (B and D) of detergent. Both FLAGA-N (A and B) and FLAGA-C (C and D) exhibited immunofluorescence in detergent permeabilized cells.
[View Larger Version of this Image (126K GIF file)]


DISCUSSION

Until recently, studies of Na+- and Cl--dependent transporter structure and function have been based on the transporter model predicted by hydropathy analyses (Fig. 1A). GAT-1 was chosen as a representative of this transporter family in the study described here, which is among the first attempts to examine experimentally the transmembrane topology of these proteins in detail. Using protease protection of defined GAT-1 reporter epitopes our data reveal novel differences in the topology of some of the loops connecting HDs, while the topology of GAT-1 HDs is not significantly different from the predicted model. Specifically, the loops connecting HD3 and HD4, and HD7 and HD8 are accessible to protease in the cytosol suggesting that regions of these loops extend into the membrane as depicted in Fig. 7. In addition to elucidating the transmembrane topology of GAT-1, these methods reveal processes governing the transmembrane assembly of this transporter. Although the presence of topogenic sequences throughout GAT-1 makes interpretation of these data difficult and limits the utility of this methodology, many approaches have been taken to study the topology of integral membrane proteins, and each method displays certain limitations. Immunocytochemical localization of peptide epitopes is a preferred method for studying transmembrane topology because it is possible to study the full-length unmodified, and functional protein. Yet it is not always possible to generate antibodies to epitopes of interest and it is difficult to distinguish between buried extracellular epitopes and intracellular ones. Another approach involves the detection of post-translational modification of residues as an indication of membrane sidedness. Several integral membrane proteins such as the cystic fibrosis transmembrane regulator (30), GluR1 glutamate receptor (31), Glut 1 glucose transporter (32), the SGLT1 glucose transporter (33), and most recently GAT-1 and the GLYT1 glycine transporter (14, 15) have been studied using glycosylation site insertion. The insertion of such sites throughout a protein has the potential of altering the native topology. In addition, the insertion of glycosylation sites into the loops of some proteins can be detrimental to function as has been shown for the Glut 1 glucose transporter (32), GAT-1 (14), and GLYT1 (15).


Fig. 7. Proposed GAT-1 transmembrane topology. Tree-like structures denote potential sites for N-linked glycosylation. Barrels represent hydrophobic domains and darkened areas of loops indicate regions of extracellular loops accessible to protease.
[View Larger Version of this Image (19K GIF file)]

The approach taken in this study to examine the transmembrane topology of GAT-1 has been used by several groups to study other integral membrane proteins including the human P-glycoprotein (19-21), the nicotinic acetylcholine receptor subunits (23), and the GluR3 glutamate receptor (22). In this method the COOH-terminal 144 amino acid residues of prolactin are fused at defined locations to the GAT-1 transporter. In a membrane vesicle preparation from X. laevis oocytes expressing GAT-1-prolactin chimeras, the protease sensitivity of the epitope reveals the cellular location of the hydrophilic loop being examined. Protection of the epitope from protease digestion indicates that the loop is located in the ER lumen and therefore becomes ultimately extracellular, whereas, digestion of the epitope would indicate that the loop is cytosolic. In using this approach the assumption has been made that truncation of GAT-1 and fusion of the prolactin epitope does not affect the native transmembrane topology. This assumption is supported by several lines of evidence. First, the prolactin epitope has no topogenic sequences itself and has been shown to be permissive to the actions of defined polytopic integral transmembrane proteins (17). Second, the transmembrane topologies of some proteins studied with prolactin fusions have been confirmed through the use of epitope-directed antibodies (20, 23). Finally, fusion of the prolactin epitope to the COOH terminus of GAT-1 had no effect on transport function (data not shown). The effect of truncation and prolactin fusion on the native transmembrane topology of EL1 through EL6 and CL1 through CL5 could not be tested, however. It is evident from the limitations described for each of the methods that convergent data obtained using different methods will prove most convincing in determining the transmembrane topology of the Na+- and Cl--dependent transporters.

Due to the presence of topogenic sequences throughout GAT-1 it was not possible to infer the transmembrane topology of a loop based on a single GAT-1-prolactin fusion protein. Protease protection studies to determine the topology of both prokaryotic and eukaryotic integral membrane proteins have shown that data on individual fusion proteins may be misleading (21, 34). Transmembrane assembly of GAT-1 must require the cooperative actions of several topogenic sequences. Therefore, consideration of data collected from studying all the constructs was necessary to provide a two-dimensional picture of this polytopic protein. Cooperative actions were necessary for the assembly of HD1 through HD4 as membrane integration for each of these HDs did not occur until the next downstream HD was present. HD1 did not become membrane integrated until HD2 was present, as evidenced by extraction of EL1 from the membranes at high pH. In addition, the digestion of the amino terminus when CL1 was treated with protease suggests that HD1 has become integrated. As revealed by the difference in the glycosylation states of EL2 and CL2, HD3 did not become inserted in the membrane until HD4 was added. The extracellular placement of the loop connecting HD3 and HD4 is consistent with data from many studies which have shown that the sites for N-linked glycosylation in this loop are utilized (10-13). Immunocytochemical data reveal that this loop is extracellular in the norepinephrine transporter (9). However, EL3 data showed that in this fusion protein the NH2-terminal portion of the large loop connecting HD3 and HD4 is accessible to protease. These data may be unique to the conformation of this particular construct or may reveal that this region of the transporter is topologically different from the predicted model. A different approach will be necessary to confirm that this change exists in the full-length functional transporter. Furthermore, in EL4 and CL4, HD7 and HD8 did not insert in the membrane in the predicted manner, yet they do become integrated as indicated by EL5 and EL6 chimera data. Protected digestion products from both EL5 and EL6 show that the loop connecting HD8 and HD9 is accessible to protease and therefore located in the cytoplasm, consistent with the predicted model for GAT-1. The loop connecting HD7 and HD8 in NET has been shown to be extracellular by immunocytochemistry with epitope-specific peptide antibodies (9). Due to the significant homology between GAT-1 and NET sequences and therefore the expected similarity in transmembrane topology of the proteins, it is assumed that the majority of the loop connecting HD7 and HD8 in GAT-1 is also extracellular. Our immunocytochemical analysis of the amino and carboxyl termini of GAT-1 indicates that both are located in the cytoplasm. This is consistent with data published for the norepinephrine and glycine transporters (9, 11, 35, 36).

While the transmembrane orientation of the HDs is consistent with that determined by hydropathy analysis, accessibility of residues in the loops connecting HD3 and HD4 and HD7 and HD8 was not predicted. Determination of cleavage sites in GAT-1 chimeras was based on gel mobility of cleaved fragments. Since hydrophobic proteins such as transporters can migrate anomolously on SDS gels, these sites of cleavage must be considered approximate until confirmed by mapping. While the specific sites of protease cleavage remain to be defined, the data indicate that regions within the loops connecting HD3 and HD4 and HD7 and HD8, loops which are thought to be extracellular, were accessible to protease in the cytosol. Data from the assay of CL2 indicate that the COOH-terminal portion of the large loop connecting HD3 and HD4 is accessible to protease, and therefore must extend to some degree into the membrane (Fig. 7). Similarly, data from assay of CL4, EL5, and EL6 show that a portion of the loop connecting HD7 and HD8 is accessible to protease (Fig. 7). Prolines and short stretches of hydrophobic residues in the COOH-terminal regions of both of these loops may participate in the formation of loop-type structures that extend into the membrane, or into a channel-like opening, that are accessible from the cytoplasm (Fig. 7). Such pore loop structures have been identified in a number of ion channels (37). Pore loops in the voltage-gated potassium channel are thought to extend into the central pore from the extracellular side of the membrane forming an ion selectivity filter (37). Putative pore loops in GAT-1 may function in substrate selectivity. In fact, Tamura et al. (38) have suggested that the loop connecting HD7 and HD8 in the GABA transporter is involved in forming a pocket to which substrate binds. These authors provide evidence that the loop connecting HD7 and HD8 is involved in determining the GABA binding affinities for different GABA transporters.

Recent studies of GAT-1 and GLYT1 (glycine transporter) transmembrane topology using glycosylation site insertion, cysteine mutagenesis, and gene fusion methods (14, 15) suggest that significant revisions of the amino-terminal portion of the transporter model are needed. In these authors' revised model, the first HD extends into the membrane from the cytosolic face as a loop; HD2 is the first true HD; CL1 is located on the extracellular face of the membrane; and an additional HD is found in the amino-terminal portion of EL2. The transmembrane topology of HD4 through HD12 in these authors' model is in agreement with the predicted hydropathy model and the model presented in the current study. Due to the incremental mode of insertion and integration of the transporter prolactin chimeras, our study does not confirm or disprove the revisions suggested by these authors. However, data obtained with EL3 did indicate that the amino-terminal portion of EL2 may be accessible to protease, which would support the presence of an additional HD in this region.

The particular approach chosen here to study GAT-1 transmembrane topology has yielded data that begins to elucidate the unusual transmembrane assembly of this protein. Translocation of a nascent polypeptide across the ER membrane depends upon the presence of a signal sequence. More specifically, signal anchor sequences are responsible for membrane targeting, translocation, and membrane integration of nascent chains. Partial translocation of a nascent chain across a membrane requires the presence of a second type of topogenic sequence, a stop-transfer sequence. It has been proposed that proteins which span the membrane several times acquire their unique topology by a series of alternating signal and stop-transfer sequences (39, 40). Study of the membrane translocation of bovine opsin suggests that alternating signal and stop-transfer sequences are responsible for directing the assembly of this protein (41). Assembly of the carboxyl-terminal portion of GAT-1, including HD5, HD6, and HD9-HD12 appears to occur through an alternating pattern of signal and stop-transfer sequences similar to that observed for bovine opsin (41). My data are confirmed by Olivares et al. (15) studying individual and paired GLYT1 HDs. However, amino-terminal GAT-1 assembly of HD1-HD4 as well as HD7 and HD8 seems complex. Based on the predicted hydropathy model for GAT-1, HD1 should possess signal anchor sequence properties which would direct translocation of the prolactin epitope into the ER lumen and integrate the nascent polypeptide into the ER membrane such that the amino terminus extends into the cytoplasm. However, protease protection data reveal that HD1 does not become integrated into the membrane until HD2 is also present, and HD2, which is expected to have stop-transfer ability based on the predicted topology of GAT-1, does not exhibit this property in the CL1 fusion protein. The membrane integration of HD3 and HD4, as well as that of HD7 and HD8, also reveal the coordinate action of topogenic sequences other than a series of alternating signal and stop-transfer sequences. Similarly, the topogenic properties of GLYT1 individual and paired HD1 and HD2 and individual HD4 do not traverse the membrane in the predicted manner (15).

During synthesis of apolipoprotein B, translocation is paused resulting in nonintegrated, transmembrane intermediates which become fully translocated over time (42, 43). The pause in translocation is thought to be triggered by a pause-transfer sequence that halts the nascent protein in an aqueous channel until translocation and integration is resumed. Pause-transfer sequences have also been identified in the prion protein, and are thought to mediate the unusual translocation of this protein (43). Pause-transfer sequences from these two proteins have been identified and are strikingly similar (KPKTNMKHMA, apolipoprotein B B'; KKTKNSEEFA, prion protein STE) (43). Analysis of the GAT-1 amino acid sequence reveals the presence of a stretch of residues (KPKTLVVKVQKK) in the amino terminus with remarkable similarity to the pause-transfer sequences identified in apolipoprotein B and the prion protein. Pause-transfer sequences and other unidentified unconventional topogenic sequences may mediate the unusual translocation and integration of GAT-1. More detailed analyses are necessary to elucidate the topogenic sequences responsible for GAT-1 transmembrane assembly and may aid in understanding the transmembrane assembly of other complex polytopic integral membrane proteins.

In the absence of x-ray crystallography data of complex integral membrane proteins such as GAT-1, one must infer the transmembrane topology of such proteins using methods such as those described above. While each of these approaches has limitations, valuable information may be obtained as long as results are interpreted with caution. Given the significant sequence homology between the members of this transporter family, it is expected that the model of GAT-1 proposed here is representative of all Na+- and Cl--dependent transporters. These results provide a foundation for future topology studies of this transporter family using alternative methodologies.

Acknowledgments

I thank M. Brownstein for generous support and encouragement, R. Seal and T. Usdin for helpful discussions, J. Northup and W. Clark for helpful discussions and critical review of the manuscript, Z. Hall for critical review of the manuscript, and C. Smith for assistance with the laser scanning microscope. V. Lingappa and W. Skach generously provided pSPSp+1L.ST.gG.pT, BPI, and many helpful suggestions.


FOOTNOTES

*   This work was supported in part by the National Institute of Mental Health Intramural Research Program and a Pharamacology Research Associate Training (PRAT) award from the National Institute of General Medical Sciences, National Institutes of Health.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.
Dagger    Recipient of the PRAT Award (National Institute of General Medical Sciences) and an Intramural Research Training Award (National Institute of Mental Health). Tel.: 301-496-2290; Fax: 301-402-1748; E-mail: janet{at}codon.nih.gov.
1   The abbreviations used are: GABA, gamma -aminobutyric acid; NET, norepinephrine transporter; HD, hydrophobic domain(s); ER, endoplasmic reticulum; PNGase F, peptide:N-glycosidase F; Endo H, endoglycosidase H. 

REFERENCES

  1. Amara, S. G., and Kuhar, M. J. (1993) Annu. Rev. Neurosci. 16, 73-93 [CrossRef][Medline] [Order article via Infotrieve]
  2. Clark, J. A., and Amara, S. G. (1993) Bioessays 15, 323-332 [Medline] [Order article via Infotrieve]
  3. Rudnick, G., and Clark, J. A. (1993) Biochim. Biophys. Acta 1144, 249-263 [Medline] [Order article via Infotrieve]
  4. Guastella, J., Nelson, N., Nelson, H., Czyzyk, L., Keynan, S., Miedel, M. C., Davidson, N., Lester, H., and Kanner, B. (1990) Science 249, 1303-1306 [Medline] [Order article via Infotrieve]
  5. Pacholczyk, T., Blakely, R. D., and Amara, S. G. (1991) Nature 350, 350-354 [CrossRef][Medline] [Order article via Infotrieve]
  6. Mestikawy, S. E., Giros, B., Pohl, M., Hamon, M., Kingsmore, S. F., Seldin, M. F., and Caron, M. G. (1994) J. Neurochem. 62, 445-455 [Medline] [Order article via Infotrieve]
  7. Liu, Q.-R., Mandiyan, S., López-Corcuera, B., Nelson, H., and Nelson, N. (1993) FEBS Lett. 315, 114-118 [CrossRef][Medline] [Order article via Infotrieve]
  8. Uhl, G. R., Kitayama, S., Gregor, P., Nanthakumar, E., Persico, A., and Shimada, S. (1992) Mol. Brain Res. 16, 353-359 [Medline] [Order article via Infotrieve]
  9. Brüss, M., Hammermann, R., Brimijoin, S., and Bönisch, H. (1995) J. Biol. Chem. 270, 9197-9201 [Abstract/Free Full Text]
  10. Tate, C. G., and Blakely, R. D. (1994) J. Biol. Chem. 269, 26303-26310 [Abstract/Free Full Text]
  11. Olivares, L., Aragón, C., Giménez, C., and Zafra, F. (1995) J. Biol. Chem. 270, 9437-9442 [Abstract/Free Full Text]
  12. Melikian, H. E., McDonald, J. K., Gu, H., Rudnick, G., Moore, K. R., and Blakely, R. D. (1994) J. Biol. Chem. 269, 12290-12297 [Abstract/Free Full Text]
  13. Melikian, H. E., Ramamoorthy, S., Tate, C. G., and Blakely, R. D. (1996) Mol. Pharmacol. 50, 266-276 [Abstract]
  14. Bennett, E. R., and Kanner, B. I. (1997) J. Biol. Chem. 272, 1203-1210 [Abstract/Free Full Text]
  15. Olivares, L., Aragón, C., Giménez, C., and Zafra, F. (1997) J. Biol. Chem. 272, 1211-1217 [Abstract/Free Full Text]
  16. Katz, F. N., Rothman, J. E., Lingappa, V. R., Blobel, G., and Lodish, H. F. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 3278-3282 [Abstract]
  17. Rothman, R. E., Andrews, D. W., Calayag, M. C., and Lingappa, V. R. (1988) J. Biol. Chem. 263, 10470-10480 [Abstract/Free Full Text]
  18. Wessels, H. P., and Spiess, M. (1988) Cell 55, 61-70 [CrossRef][Medline] [Order article via Infotrieve]
  19. Skach, W. R., and Lingappa, V. R. (1994) Cancer Res. 54, 3202-3209 [Abstract]
  20. Skach, W. R., Calayag, M. C., and Lingappa, V. R. (1993) J. Biol. Chem. 268, 6903-6908 [Abstract/Free Full Text]
  21. Skach, W. R., and Lingappa, V. R. (1993) J. Biol. Chem. 268, 23552-23561 [Abstract/Free Full Text]
  22. Bennett, J. A., and Dingledine, R. (1995) Neuron 14, 373-384 [Medline] [Order article via Infotrieve]
  23. Chavez, R. A., and Hall, Z. W. (1992) J. Cell Biol. 116, 385-393 [Abstract]
  24. Blakely, R. D., Clark, J. A., Rudnick, G., and Amara, S. G. (1991) Anal. Biochem. 194, 302-308 [Medline] [Order article via Infotrieve]
  25. Graminski, G. F., Jayawickreme, C. K., Potenza, M. N., and Lerner, M. R. (1993) J. Biol. Chem. 268, 5957-5964 [Abstract/Free Full Text]
  26. Blakely, R. D., Clark, J. A., Pacholczyk, T., and Amara, S. G. (1991) J. Neurochem. 56, 860-871 [Medline] [Order article via Infotrieve]
  27. Fujiki, Y., Hubbard, A. L., Fowler, S., and Lazarow, P. B. (1982) J. Cell Biol. 93, 97-102 [Abstract]
  28. Laemmli, U. K. (1970) Nature 227, 680-685 [Medline] [Order article via Infotrieve]
  29. Corey, J. L., Davidson, N., Lester, H. A., Brecha, N., and Quick, M. W. (1994) J. Biol. Chem. 269, 14759-14767 [Abstract/Free Full Text]
  30. Chang, X.-B., Hou, Y.-X., Jensen, T. J., and Riordan, J. R. (1994) J. Biol. Chem. 269, 18572-18575 [Abstract/Free Full Text]
  31. Hollmann, M., Maron, C., and Heinemann, S. (1994) Neuron 13, 1331-1343 [Medline] [Order article via Infotrieve]
  32. Hresko, R. C., Kruse, M., Strube, M., and Mueckler, M. (1994) J. Biol. Chem. 269, 20482-20488 [Abstract/Free Full Text]
  33. Turk, E., Kerner, C. J., Lostao, M. P., and Wright, E., M. (1996) J. Biol. Chem. 271, 1925-1934 [Abstract/Free Full Text]
  34. Hennessey, E. S., and Broome-Smith, J. K. (1993) Curr. Opin. Struct. Biol. 3, 524-531 [CrossRef]
  35. Zafra, F., Aragón, C., Olivares, L., Danbolt, N. C., Giménez, C., and Storm-Mathisen, J. (1995) J. Neurosci. 15, 3952-3969 [Abstract]
  36. Olivares, L., Aragón, C., Giménez, C., and Zafra, F. (1994) J. Biol. Chem. 269, 28400-28404 [Abstract/Free Full Text]
  37. MacKinnon, R. (1995) Neuron 14, 889-892 [Medline] [Order article via Infotrieve]
  38. Tamura, S., Nelson, H., Tamura, A., and Nelson, N. (1995) J. Biol. Chem. 270, 28712-28715 [Abstract/Free Full Text]
  39. Lingappa, V. R., Lingappa, J. R., and Blobel, G. (1979) Nature 281, 117-121 [Medline] [Order article via Infotrieve]
  40. Blobel, G. (1980) Proc. Natl. Acad. Sci. U. S. A. 77, 1496-1500 [Abstract]
  41. Audigier, Y., Friedlander, M., and Blobel, G. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 5783-5787 [Abstract]
  42. Chuck, S. L., and Lingappa, V. R. (1992) Cell 68, 9-21 [Medline] [Order article via Infotrieve]
  43. Nakahara, D. H., Lingappa, V. R., and Chuck, S. L. (1994) J. Biol. Chem. 269, 7617-7622 [Abstract/Free Full Text]

©1997 by The American Society for Biochemistry and Molecular Biology, Inc.