|Home | About | Journals | Submit | Contact Us | Français|
Subunit assembly governs regulation of AMPA receptor (AMPA-R) synaptic delivery and determines biophysical parameters of the ion channel. However, little is known about the molecular pathways of this process. Here we present single particle electron microscopy (EM) 3D structures of dimeric biosynthetic intermediates of the GluA2 subunit of AMPA-Rs. Consistent with the structures of intact tetramers, the amino terminal domains of the biosynthetic intermediates form dimers. Transmembrane domains also dimerize despite the two ligand binding domains (LBD) being separated. A significant difference was detected between the dimeric structures of the wildtype and the L504Y mutant, a point mutation that blocks receptor trafficking and desensitization. In contrast to the wildtype, whose LBD is separated, the LBD of the L504Y mutant was detected as a single density. Our results provide direct structural evidence that separation of the LBD within the intact dimeric subunits is critical for efficient tetramerization in the endoplasmic reticulum and further trafficking of AMPA-Rs. The contribution of stargazin on the subunit assembly of AMPA-R was examined. Our data suggests that stargazin affects AMPA-R trafficking at a later stage of receptor maturation.
The majority of fast excitatory synaptic transmission in the brain is mediated by AMPA (α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid) receptors (AMPA-Rs), a subset of ligand-gated ion channels of the glutamate receptor family. Trafficking, anchoring, and gating of AMPA-Rs form the molecular basis for certain types of synaptic plasticity involved in learning and memory (Barry and Ziff, 2002; Malinow and Malenka, 2002; Nicoll et al., 2006). Dysfunction of AMPA-Rs is implicated in a variety of neurological and psychiatric disorders, including X-linked mental retardation, Alzheimer’s disease, amyotrophic lateral sclerosis, and Rasmussen’s encephalitis (Rogers et al., 1994; Shepherd and Huganir, 2007; Wu et al., 2007).
AMPA-R subunits are encoded by four different genes (GluA1-4). (Hollmann et al., 1989; Keinanen et al., 1990; Nakanishi et al., 1990). Each subunit consists of four domains (Fig 1A). The N-terminal domain (NTD) and ligand binding domain (LBD) are both extracellular. The LBD, made of S1 and S2 subdomains, undergoes conformational changes resulting in channel gating upon glutamate binding. The polypeptide chain forming the LBD is interrupted by the channel pore-forming transmembrane domain (TMD), which consists of three membrane spanning segments (M1, M3, and M4) and one re-entrant loop (M2) (Hollmann et al., 1994). A small C-terminal domain (CTD) extends into the cytoplasm, interacting with cytosolic proteins that regulate receptor anchoring and trafficking. (Scannevin and Huganir, 2000; Sheng and Lee, 2001; Malinow and Malenka, 2002; Ziff, 2007). Cumulative evidence suggests an AMPA-R subunit assembly model in which two dimers come together to form a tetramer, hence a dimer-of-dimer organization for mature tetrameric AMPA-Rs. (Armstrong et al., 1998; Gouaux, 2004; Mayer, 2006). In the brain, AMPA-R auxiliary subunits of the stargazin/TARP (transmembrane AMPA-R regulatory protein) family and the cornichon family are physically associated with the channel and regulate their trafficking and gating (Chen et al., 2000; Nakagawa et al., 2005; Nicoll et al., 2006; Ziff, 2007; Schwenk et al., 2009).
Trafficking of both newly synthesized and recycling AMPA-Rs is a critical component of synaptic plasticity (Malinow and Malenka, 2002; Ju et al., 2004; Park et al., 2004; Matsuo et al., 2008). The molecular anatomy of early phases of AMPA-R trafficking, including biosynthesis, is largely unknown. Studies have identified point mutations and splice variants that alter receptor trafficking and, together with the crystal structures of the mutated S1S2 domains, have provided insight into what might be happening at the ultrastructural level of the intact subunits during receptor assembly (Greger et al., 2002; Greger et al., 2003; Coleman et al., 2006; Greger et al., 2006). It is not clear how the domains are organized during the normal assembly of full-length subunits and how mutant subunits interfere with this process. In addition, how the auxiliary subunit stargazin influences biosynthesis of AMPA-R remains controversial.
Here we investigate AMPA-R subunit assemply and report single particle EM structures of newly synthesized AMPA-Rs in dimeric states. The study reveals that efficient subunit assembly requires a preferred conformation of AMPA-R biosynthetic intermediates and stargazin affects AMPA-R trafficking at the later stages of receptor maturation.
The GluA2 flop splice variant was used for all experiments. The L504Y mutation was introduced by in vitro mutagenesis using Quick change kit (Stratagene). The GFP-GluA2 fragment was a gift from Y.Hayashi and GFP was inserted immediately after the signal peptide, following the exact design as previously described (Hayashi et al., 2000). The FLAG epitope tag was inserted in the C-terminal domain of GluA2 (FATDYKDDDDKEGYNVYGIESVKI, where bold case indicates FLAG epitope) and placement preserves the original anti-GluA2CT epitope.
Wildtype HEK cells, GnTI(-)HEK cells, and the transformants created were maintained in a base media that consists of high glucose DMEM, 100 units/ml penicillin, 100 µg/ml streptomycin, and 10 % fetal calf serum. To isolate stable clones, we co-transfected a plasmid vector that expresses GluA2 under the CMV promoter and another plasmid vector that expresses a hygromycin resistant gene. Transfection was done by calcium phosphate methods and the selection of clones was done over two weeks in the presence of 160 µg/ml hygromycin. Isolated colonies were cultured until morphologically homogeneous cultures were established. Expression of GluA2-FLAG was tested for each clone using western blotting of the whole cell lysate by probing with custom made antibodies raised against the C-terminal peptide of GluA2 (EGYNVYGIESVKI) (Nakagawa et al., 2005). Through screening ~200 colonies we identified several clones that meet the criteria of optimal growth speed and expression. There was a tendency for highly expressing clones to be slow growing, consistent with toxicity to the host cell caused by overexpressing an ion channel. To assess stability, we kept culturing the established clones for seven months, and detected by immunofluorescence microscopy that 65% of the cells maintain expression of GluA2 (Supplemental Fig 1A). Thus the stable cell line we established can be used for large scale culture to produce recombinant GluA2 in large quantities. Typically a 1 liter culture of HEK cells was used for each purification in this study.
A neomycin (G418) resistant TetON-HEK cell line (Clontech) has in its genome the expression module to produce rtTA (see Fig 2A). GluA2-FLAG, GluA2L504Y-FLAG, GFP-GluA2-FLAG, and GFP-GluA2L504Y-FLAG were subcloned into pTREtight vector (Clontech). TetON-HEK cells were co-transfected with a plasmid that expresses a hygromycin resistant gene and a GluA2 construct in pTREtight described above. Transfection was done by calcium phosphate and selection of clones was done over two weeks in the presence of 120 µg/ml hygromycin. The remaining procedure follows the generation of the stable HEK cell lines described above, except that we detected the expression of GluA2 using western blotting after inducing the isolated clones with 5 µg/ml DOX for 24 hours.
Stargazin-IRES-mCherry cassette was subcloned into pBOSS vector (a gift from Shigekazu Nagata and Hideki Sakahira) downstream of the elongation factor promoter. pBOSS-stg-IRES-mCherry vector and pCMVZeocin (Invitrogen) were co-transfected into the parental TetONGluA2 stable HEK cell and stable clones were isolated by selecting with antibiotics 125 µg/ml zeocin, 150 µg/ml hygromycin, and 125 µg/ml neomycin (G418). mCherry positive colonies were visually identified using an epi-fluorescent microscope, isolated, and subcultured. 80% of the mCherry positive clones also expressed stargazin as determined by Western blotting. DOX inducible GluA2 expression was also re-confirmed in all of the isolated cell lines.
Five confluent 15 cm dishes were taken from the CO2 incubator at a time. Media was aspirated off and 6 ml of ice-cold phosphate buffered saline (PBS) was added to each plate. The plates were tapped strongly from the side about 10 – 15 times to dislodge all cells from the bottom of the dish. Cells were pooled in a 250 ml centrifuge tube on ice. The dishes were further rinsed with 3 ml of PBS twice to collect all remaining cells. Cells collected from 20 plates fill up the 250 ml centrifuge tube. We centrifuged the cells at 1000 rpm at 4 °C for 10 min. After discarding the supernatant the pellet was resuspended in 50 ml of PBS and further centrifuged. After discarding the supernatant the cell pellet was flash frozen in liquid nitrogen and stored at −80 °C until use.
All purification procedures were conducted on ice or in the cold room to maintain specimen temperature below 4 °C. 1 liter of HEK cell culture (6 ml of cell pellet) was resuspended in 50 ml of buffer containing 50 mM K-HEPES pH7.4, 100 mM NaCl, 1mM Kynurenic acid, protease inhibitors (1 mM PMSF, 10 µg/ml leupeptin, atropinin, benzamidine, and pepstatin A). The cells were extracted with the detergent, DDM (0.25%) at 4 °C for three hours. Solubulization yield was above 90%, which is unsurprising given that AMPA-Rs from brain solubulize efficiently when synaptosomal fractions are extracted with mild detergents (Leonard et al., 1998). After clearing the lysate by ultracentrifugation (Beckman 45 Ti) at 45,000 rpm for 1 hour at 4 °C, the supernatant was applied to a column made of protein A sepharose beads (GE Amersham) crosslinked using DMP (Pierce) with anti-FLAG M2 monoclonal antibody (Sigma) at a concentration of 2 mg/ml. Following washing, bound proteins were released from the column using a buffer containing 0.5 mg/ml of FLAG epitope peptide. The peak fraction from the peptide elution was further separated by Superdex 200 gel filtration column (GE Amersham) in a buffer that contains 50 mM K-HEPES pH 7.4, 100 mM NaCl, and 0.1% DDM.
Purified GluA2 was resolved in 7.5% SDS-PAGE and Western blotting was done using the custom made anti-panTARP antibody as described (Nakagawa et al., 2005). In the case of the mass spectrometry, the gel was stained with CBB. All the bands between 30–60 kDa were cut out and digested with trypsin. The identity of each band was determined by liquid chromatography followed by tandem mass spectrometry (LC/MS/MS) at UCSD mass spectrometry facility.
400 mesh copper grids were coated with carbon to create a substrate for proteins to bind. 4 µl of protein solution was applied to a glow discharged grid and left for 30 sec to 5 min to allow the proteins to bind. The excess water was blotted on filter paper and the specimen was washed twice in water droplets to remove excess detergents. Purified proteins were negatively stained with 0.75% (w/v) uranyl formate as described (Ohi et al., 2004). Images were recorded using a FEI Sphera electron microscope equipped with a LaB6 filament operated at an acceleration voltage of 200 keV. Images were taken at a magnification of 50,000 X and defocus value = −1.5 µm. Specimens were imaged at 0° and 60° tilt for random conical tilt 3D reconstruction; the defocus value for 0° = −1.5 ~ −1.8 µm and 60° tilt = −2.0 ~ −2.2 µm. All images were recorded using SO-163 film and developed with a Kodak D-19 developer at full strength for 12 min at 20 °C. Particle images were taken at room temperature and under low dose conditions (20 e/Å2) to minimize radiation damage.
Fabs were purified using the Immunopure IgG1 F(ab’) and the F(ab’)2 Fab purification kit (Pierce) followed by gel filtration on a Superdex 200 column (Pharmacia). Anti-FLAG M2 monoclonal antibody (Sigma) was used as source. Labeling was performed by incubating dimeric AMPA-Rs with Fab fragments at a molar ratio of 1:4 to 1:8 overnight at 4°C in 50 mM HEPES, pH 7.4, 100 mM NaCl, 0.1% DDM.
Electron micrographs were digitized with a CoolScan 9000 (Nikon) using a step size of 6.35 µm and 3 × 3 pixels were binned so the specimen level pixel size used was 3.81 Å. Projection averages were calculated from windowed small images of 100 × 100 pixels over 10 cycles of K-means classification and multi-reference alignment specifying 100 classes. For 3D reconstruction of GluA2wildtype dimer, 269 tilt pairs (total of 538 micrographs) were recorded on film from which 178 tilt pairs were selected based on image quality. A total of 13,345 particle pairs were interactively selected using WEB display program for SPIDER (Frank et al., 1996) and windowed, and the untilted particles were averaged into 100 classes as before. In the case of the GluA2L504Y dimer, 238 tilt pairs (total of 476 micrographs) were recorded from which 159 tilt pairs were selected based on image quality. A total of 9,000 particle pairs were selected and used for further analysis. Raw particle images were visually inspected after classification to make sure that tetramers were not mistakenly introduced into our dimer 3D reconstruction. Images of the tilted specimens for each class were used to calculate initial 3D reconstructions of individual classes by backprojection, backprojection refinement, and angular refinement (implemented in SPIDER). The final volume obtained by angular refinement with SPIDER was used as the input model for FREALIGN (Grigorieff, 2007); this was used for refinement of orientation parameters of individual particles and for individual image contrast transfer function correction based upon the defocus value. The tilt angles and defocus values of the center of each micrograph were determined with CTFTILT (Mindell and Grigorieff, 2003). The defocus of each particle was deduced from its position on the micrograph. Particles selected from tilted and untilted specimens were used for FREALIGN refinement (500 and 800 particles were used in the final reconstruction for wildtype and L504Y, respectively). To ensure that the final 3D reconstruction agreed with the raw data, the particle images were compared with reprojections.
Expression of GFP-GluA2 wildtype and GFP-GluA2L504Y was induced with 7.5 µg/ml DOX. 30 hr after induction cells were live labeled using an anti-GluA2 NTD monoclonal antibody for 15 min (Chemicon, MAB397). Cells were washed with warm DMEM and fixed with 4% formaldehyde in 0.1 M phosphate buffer pH 7.4. Surface GluA2 was detected using Alexa 568 conjugated anti-mouse IgG secondary antibody (Invitrogen). Total GluA2 was detected by the GFP fluorescence signal. Imaging was performed using an Olympus Fluoview confocal microscope using the 60X objective lens.
GFP-GluA2 expression was induced for 30 hrs with 7.5µg/ml DOX followed by labeling with 10 µg/ml anti-GluA2NTD monoclonal antibody (Chemicon, MAB397) in the CO2 incubator at for 1 hr. During the 1 hr incubation period, GFP-GluA2 labeled with the antibody undergoes endocytosis. Unbound antibodies were washed with DMEM (37 °C) and the cells were fixed with 4% formaldehyde. Alexa 568 conjugated anti-mouse IgG antibody (Invitrogen) was used as secondary antibody to label the primary antibodies. All the images were taken on an Olympus Fluoview confocal microscope, as described in the above section.
HEK cells were grown on a glass bottom dish for 24 hr before 7.5 µg/ml DOX was added. Time-lapse imaging occurred 30 hr after induction using an Olympus Fluoview confocal microscope mounted with a temperature (37 °C), CO2 (5%) and humidity controlled chamber was used equipped with a 60X objective lens (Olympus PLAPON, N.A. = 1.42) was used. 0.5 µm thick optical sections were obtained up to 4 µm from the bottom of the cell. Recording was done continuously, resulting in 20 sec time intervals between each Z-stack.
Primary cultures of rat hippocampal neurons were prepared from E18 rat embryos as previously described (Sala et al., 2003). HA-tagged GluA2 constructs were introduced into neurons at DIV14 using calcium phosphate transfection. Surface HA-GluA2 was stained while neurons were alive. Specifically, after labeling the DIV 18 neurons with anti-HA monoclonal antibody (HA.11, Covance), neurons were washed in DMEM to remove unbound antibodies and fixed with 4% formaldehyde in 0.1 M phosphate buffer pH 7.4. After washing with PBS, internal HA-GluA2 was labeled using anti-HA polyclonal antibody (Y-11, Santa Cruz) diluted in 1XGBD (0.2% gelatin, 0.6% TritonX-100, 33mM phosphate buffer pH 7.4, and 0.9M NaCl). Alexa 488 conjugated anti-mouse IgG (Invitrogen) and Alexa 568 conjugated anti-rabbit IgG (Invitrogen) were used as secondary antibodies. Neurons were imaged using an Olympus Fluoview confocal microscope using the 60X objective lens (Olympus PLAPON, N.A. = 1.42). Z-projections of confocal stacks are shown in Figure 3B.
Four clones (#2, 8, 9, and 10) of TetOnGluA2-stg cell lines and the parental TetONGluA2 cell were plated on the 6 well plates at a density of 0.6 × 106 cells/well, and incubated 24 hr to allow the cells to attach. At this point the cells were near confluent and 7.5 µg/ml DOX was added with 30 µM of NBQX to induce GluA2 expression. Cells were harvested 0, 6, 12, 18, and 24 hr after induction. Equal amounts of protein sample from each time point were loaded into each well of the SDS-PAGE gel (12.5% and 7.5% gels were used to resolve stargazin and GluA2, respectively). Western blotting was done using anti-panTARP and anti-GluA2C-terminal polyclonal antibodies (both generated in (Nakagawa et al., 2005)).
TetOnGluA2-stg Clone#10 cells were fixed as above 24hr after DOX induction. Anti-FLAG monoclonal (Sigma) and anti-panTARP polyclonal antibodies (Nakagawa et al., 2005) were used to detect GluA2 and stargazin, respectively. Cy5 conjugated anti-mouse IgG (Jackson ImmunoResearch laboratories) and Alexa 488 conjugated anti rabbit (Invitrogen) were used as secondary antibodies. Confocal Z-stack images of the cells were recorded as above.
To obtain pure intact AMPA-Rs for EM analyses, we stably expressed FLAG epitope tagged GluA2 flop (Fig 1A) in HEK cells. Recombinant GluA2 was solubilized in dodecylmaltoside (DDM) in the presence of 1 mM kynurenic acid and purified by affinity chromatography using a sepharose column conjugated with anti-FLAG M2 monoclonal antibody (Fig 1C). The purity was further improved by gel filtration chromatography. The majority of purified GluA2 existed as tetramers (Supplementary Fig 1C). Because of glycosylation, the purified protein migrated in the SDS-PAGE as a doublet (Fig 1C). Only the tetrameric species were glycosylated (Supplementary Fig 1B–D). Based on the yield of purification we estimate that at least 15,000~20,000 GluA2 tetramers are expressed per cell.
Recombinant GluA2 tetramers were homogeneous in shape and size when imaged by negative stain EM (Fig 1D, left panel). The projection structures (class averages) of GluA2 tetramers were very similar to those of native AMPA-Rs from rat brain (compare Fig 1B and 1D). The NTD, LBD and TMD were clearly identified as distinct features (Fig 1E). The TMD of native AMPA-Rs is larger than that of recombinant GluA2, a difference caused by the presence or absence of auxiliary stargazin/TARP subunits (Chen et al., 2000). Consistently we did not detect stargazin/TARP protein in the purified recombinant GluA2 fraction when examined by western blotting or by protein identification using liquid chromatography followed by tandem mass spectroscopy (LC/MS/MS, data not shown). Glycosylation did not affect the overall structure of tetrameric GluA2. The shapes of negative stained glycosylated GluA2 tetramers were very similar to GluA2 tetramers purified from the GnTI(-) HEK cell line, a line defective in complex mannose glycosylation (Reeves et al., 2002) (Supplementary Fig 1E). Altogether, our stable HEK cell line is a simple, robust and highly reproducible system to study AMPA-Rs using single particle EM.
To enrich biosynthetic intermediates that have a shorter lifetime than mature and targeted proteins, we induced expression of GluA2-FLAG in HEK cells and purified them at an early time point. In this system, HEK cells stably express a reverse Tet transcriptional activator that enhances transcription from the minimal CMV promoter upstream of the GluA2-FLAG transgene only in the presence of doxycycline (DOX) (Fig 2A). We named this cell line TetONGluA2. GluA2 protein expression reached a maximum 24 hr after DOX application (Fig 2B). The magnitude of GluA2 expression in DOX inducible system was similar to that of the constitutive expression when the expression was induced for 24 hr at 7.5 µg/ml DOX. Enrichment of the glycosylated band became prominent 12 to 24 hr after transcription initiation (emergence of the upper band in Fig 2B).
When GluA2 was purified 20 hr after addition of DOX and resolved by gel filtration, equal amounts of tetramers and dimers were detected (Fig 2C solid line). The tetramer peak increased relative to the dimer peak 20 hr to 24 hr after DOX induction. At earlier time points (15 hr post induction) the majority of GluA2 existed as dimers (Supplementary Fig 2). Collectively, this suggests that the dimer population precedes the formation of the tetramer population, and thus represents a biosynthetic intermediate of pre-assembled GluA2 tetramers.
Interestingly, even at 15 hr after DOX induction we were unable to detect a distinct population that represents the monomeric subunits (Supplementary Fig 2). Because the appearance of the monomers should in theory precede the dimers, we interpret that at a given moment, the monomer population is much lower relative to the dimer population. It is thus difficult to detect monomers in our experimental system. Such an interpretation is consistent with an idea that the dimer-to-tetramer transition is the rate-limiting step compared to the monomer-to-dimer transition.
When GluA2 biosynthetic dimer intermediates were imaged by negative stain EM, the particles were homogeneous in size but existed in a variety of shapes (Fig 2D, upper panel). Approximately 7,000 particles were analyzed by multivariate statistical analysis, classification and multi reference alignment. Representative class averages are shown in Fig 2D, lower panels. There is an elongated bipartite density at the top of the particle, and a featureless globular density at the bottom of the particle. In some class averages the upper bipartite density appears as a squarish density with a weaker density in the center (Fig 2D). Between these structures two small round densities are positioned at both sides. The arrangement of the bipartite density relative to the small round densities on the sides differs significantly between class averages, and as a result, the heights of the particles vary between class averages.
The projection structures were interpreted by molecular labeling. To immunolabel the C-terminal FLAG epitope, antigen binding fragments (Fabs) were generated by proteolytic digest from IgG of anti-FLAG monoclonal antibody (Fig 2E left). Anti-FLAG Fab consistently labeled the bottom featureless density of the particles (Fig 2E right panels). Consistent with our interpretation that these particles are dimers, some particles were labeled with two Fabs (Fig 2E, far right). Because the CTD is small and attached to the TMD, we interpret that the bottom round density represents the TMD and CTD (Fig 2G).
In order to GFP label the NTD, we purified particles from a DOX inducible HEK cell line that expresses a GluA2 construct whose amino terminal end of the NTD is fused to GFP (Fig 2F). The timecourse of GFP-GluA2 expression after DOX induction and the elution profile of purified GFP-GluA2 in gel filtration chromatography were similar to those without the GFP tag (data not shown), suggesting that GFP does not affect overall processing. In the class averages of GFP-GluA2 dimers, two GFP densities were always attached to each side of the bipartite density (Fig 2F). The small round densities between the NTD and the TMD were neither labeled by anti-FLAG Fab or GFP, so we interpret that they are the LBDs (Fig 2G).
Maturation and trafficking are tightly coupled processes in membrane proteins. To understand the functional significance of the dimer structure, we decided to study the structure of the L504Y mutant, which has trafficking and maturation deficits (Fig 3A). This mutation within the LBD was originally identified in the GluR3 subunit because it blocks receptor desensitization (Stern-Bach et al., 1998). L504Y mutation is located in helix D in the S1 subdomain and the mutated tyrosine interacts with helix J in the S2 subdomain of the adjacent subunit (Sun et al., 2002). This interaction stabilizes the LBD dimer interface, and thus blocks the receptor from entering into the desensitized state. Recently it was found that surface expression of the GluA2L504Y mutant is impaired (Greger et al., 2006). Consistently, we observe significantly reduced surface expression of GluA2L504Y compared to GluA2 wildtype when expressed for 4 days starting from DIV14 in primary rat hippocampal neurons (Fig 3B).
We created stable HEK cell lines in which expression of GFP-GluA2 wildtype or GFP-GluA2L504Y is DOX inducible. In both cell lines, the timecourses of protein expression after DOX induction were similar to those without the GFP tag (data not shown), suggesting that the GFP has minimal effect on protein turnover. At 30 hr after DOX induction, there was significantly less GFP-GluA2L504Y on the cell surface compared to GFP-GluA2 wildtype, while the total expression levels of each protein were the same (Fig 3C). This suggests that the defect in surface expression of GluA2L504Y can be replicated in our simplified HEK cell system.
To gain further insight into the differential dynamics of newly synthesized GFP-tagged GluA2 wildtype and L504Y, we conducted time-lapse confocal imaging of HEK cells. We imaged a thin layer of cytoplasm between the bottom of the nucleus and the bottom of the cell that provides clear images of receptor trafficking (yellow volume in Fig 3D). For the first 24 hours following DOX induction, GluA2 wildtype and GluA2L504Y showed similar subcellular distribution, localized predominantly in the endoplasmic reticulum (ER) (data not shown). After 24 hours dynamic punctate structures emerged in the GluA2 wildtype cells but not in the GluA2L504Y cells (compare Fig 3E and F). The vesicles appeared at time points after the majority of receptor maturation was complete. More than 50 % of the punctate structures in GluA2 wildtype cells were dynamic and translocated rapidly (Fig 3G and supplementary movies). These punctate structures were absent from GluA2L504Y cells even 48 hrs after DOX induction, suggestive of differential vesicle trafficking patterns caused by the mutation (Supplementary Fig 3). Double staining GFP-GluA2 puncta with the known organelle markers revealed no co-localization with EEA1 (an early endosomal marker), transferrin (a recycling endosomal marker), nor lysotracker (a lysosomal marker). However, a subpopulation of GFP-GluA2 vesicles co-localized with rab6 (a small GTPase localized to a subset of the post Golgi vesicles), GM130 (a Golgi marker), and PDI (a ER marker) (Supplemental Fig 4). In addition, the GFP-GluA2 puncta partially co-localized with a population of GFP-GluA2 that underwent endocytosis while they were live-labeled for 1 hr with anti-GluA2NTD antibody. These data suggest that the GFP-GluA2 puncta are a mixture of vesicles that belong to the ER or Golgi apparatus, or are involved in post-Golgi trafficking and endocytic pathways (Supplemental Fig 4). The time lag between tetramer formation and the appearance of puncta of GFP-GluA2 wildtype may also suggest the possibility that tetrameric GFP-GluA2 is not readily competent to exit the ER. Collectively, these results suggest that vesicle trafficking is severely reduced in GluA2L504Y cells compared to wildtype. Next, we tested if there is also a maturation deficit in this mutant.
The timecourse of GluA2L504Y protein expression following DOX induction was similar to that of GluA2 wildtype. However, unlike GluA2 wildtype, GluA2L504Y was detected as a single band by SDS-PAGE when purified 24 hr after induction (Fig 4A and B). Furthermore, the mobility of GluA2L504Y was identical to that of GluA2 wildtype purified from GnTI(-)HEK cells deficient in complex mannose glycosylation (data not shown), indicating that the majority of GluA2L504Y is not glycosylated within 24 hr of expression.
Next, we tested if tetramerization was affected by the L504Y mutation. Interestingly, even 24 hour after induction, the majority of GluA2L504Y remained as dimers when resolved by gel filtration chromatography (Fig 4C blue). In contrast, GluA2 wildtype exists primarily as tetramers 24 hr after DOX induction (Fig 4C compare red and blue). Western blotting of the gel filtration fractions suggests that the glycosylated species of GluA2 wildtype are tetramers whereas the non-glycosylated species are primarily enriched as dimers (Fig 4D). In contrast to GluA2 wildtype, GluA2L504Y exists mostly as dimers, although a small quantity is detected as tetramers. These results are in keeping with the observation that GluA2L504Y mutant has high degree of aberrant multimerization of subunits including limited tetramer formation in HEK cells (Penn et al., 2008).
The projection structure of the negative stained tetrameric GluA2L504Y is very similar to that of GluA2 wildtype (Supplementary Fig 5A and B). The yield of these particles was extremely low, so despite our efforts, it was not feasible to obtain 3D structures of GluA2L504Y tetramers. We considered the possibility that the yield was low because tetramers in the non-desensitizing state might be less stable in detergent. However, this is unlikely because our previous study showed that brain derived AMPA-R locked in the non-desensitized state by treatment with 330 µM cyclothiazide (CTZ) and 1 mM glutamate had similar structures to the untreated receptors (Nakagawa et al., 2005). Collectively, these results suggest that the GluA2L504Y mutation causes defects in the dimer-to-tetramer transition.
To gain further insight into the difference in tetramerization between GluA2 wildtype and L504Y, EM images were taken from negative stained GluA2L504Y dimers. GluA2L504Y dimer particles were homogeneous in size, but had less conformational heterogeneity and were more elongated than wildtype dimers (Fig 5A). Representative class averages show three layers of domains stacked on top of each other. Similar to the GluA2 wildtype dimer structures, the top domain of GluA2L504Y was bipartite and elongated or appeared squarish with a weaker density in the center in some class averages. (Fig 5A, bottom right panels). The bottom domain was a featureless globular density. In contrast to the wildtype dimer, in most class averages, the middle portion existed as a single density, and was split into two in only a minority of the class averages (Fig 5A, bottom row, second from right).
Fab labeling and GFP tagging were used for domain assignment. Similar to GluA2 wildtype dimers, anti-FLAG epitope Fabs consistently decorated the bottom round featureless density of the particles (Fig 5B) and two Fabs sometimes bound to a single particle (Fig 5B, right panel). Moreover, two extra densities representing N-terminal GFP tags were detected at the top of the elongated bipartite density of GFP-GluA2L504Y dimers. From these experiments we interpret the bottom density as the TMD and CTD dimer, the middle density as the LBD dimer, and the top elongated bipartite domain as the NTD dimer (Fig 5D).
The particles were quantified based on LBD separation. Approximately 7,000 particles from negative stained EM images were classified into 100 classes by multivariate statistical analysis and multi-reference alignment. We assigned each class average (and the particles in these classes) either to “LBD separated” or “LBD fused”. All classes that could not unambiguously be assigned to either were termed “unclassifiable”. In GluA2 wildtype dimers, 71.3% had a separated LBD, 2.3% had a fused LBD, and 26.4% were unclassifiable. In contrast, of GluA2L504Y dimers, 66 % had a fused LBD, 20% had a separated LBD, and 14% were unclassifiable. Among the 66% of GluA2L504Y dimers that adopted fused LBD, 33.3% had shapes exhibiting two-fold symmetry, while 66.7% had asymmetric shapes. Among the classifiable particles of GluA2 wildtype and L504Y dimers, 7.8% and 12.1% adopted squarish NTDs, respectively, indicating that the particles that adopt squarish NTDs are the minority. These results demonstrate a robust difference in the molecular shapes between dimers of GluA2 wildtype and L504Y (Fig 2D, Fig 5A, and Fig 6). Furthermore, the difference in molecular shapes is correlated with their contrasting ability to transition from dimer-to-tetramer (Fig 4).
To confirm that the structural differences seen in the projection structures indeed reflect structural differences in 3D, the 3D density maps of the GluA2 wildtype and L504Y dimers were calculated using random conical tilt reconstruction (Frank, 1996; Frank et al., 1996). For this purpose, particle images were recorded as tilt pairs at specimen tilt angles of 0° and 60°. Raw particle images recorded at 0° were analyzed by multivariate statistical analysis, classification and multi reference alignment. Well aligned and highly represented class averages were chosen (Fig 6A and C, left box) and corresponding tilted images of particles in the selected class were used to calculate the 3D structure. Refinement was done in three steps using backprojection refinement, angular refinement (implemented in SPIDER) (Frank et al., 1996) and FREALIGN refinement (Grigorieff, 2007). Three different views of the final reconstruction of wildtype and L504Y dimers are shown in Fig 6A and C, respectively. The resolution of the final reconstruction was 35Å (wildtype) and 34 Å (L504Y) at FSC = 0.5, and with the less conservative criteria 29Å (wildtype) and 28Å (L504Y) (FSC = 0.142) (Rosenthal and Henderson, 2003). The images shown in Fig 6 are filtered at the resolution determined by FSC = 0.5, the more conservative resolution criteria. In addition, to demonstrate the structural contrast between immature and mature receptors, the 3D maps of GluA2 wildtype dimer and AMPA-R tetramer are compared side-by-side in Fig 6E.
The 3D maps of wildtype and L504Y dimers were significantly different. The projection structures and the 3D EM maps viewed from the front correspond well for both wildtype and L504Y dimers (Fig 6A and C). We assigned each globular feature in the 3D maps to individual domains of the subunit based on the domain labeling experiments (Fig 2G and Fig 5D). The wildtype dimer has the bipartite NTD dimer at the top and two smaller globular LBD densities attached at both sides of the NTD dimer. The following dimensions are the maximal distances. The height of the particle is 14.5 nm and the width is 14.6 nm. The central empty cavity of the particle suggests a clear physical separation of the two LBDs. The dimensions of the cavity are 3.7 nm in height and 5.6 nm in width. In contrast, the height and width of the L504Y dimer is 17.0 nm and 8.7 nm, and the L504Y dimer has no LBD separation.
Similar to the class averages (Fig 6A and C), the overall 3D structures did not reveal global two fold symmetry, despite that they represent homodimeric subunits of GluA2 wildtype or L504Y. The linker between the NTD and LBD consists of 16 amino acids, and we predict it has structural flexibility to accommodate twist that may exist between the NTD and LBD dimers. Consistently, the NTD was observed as an elongated bipartite density in some class averages, whereas in the others it appeared as a squarish density (Fig 2D and Fig 5A). We believe that the different appearance of the NTD in different class averages results from viewing the NTD dimer from different angles. A variety of orientations of the NTD relative to LBD can be explained by the structural flexibility present in the linker connecting these domains.
To interpret our 3D density map, we used known crystal structures of AMPA-R subunit domains. The crystal structure of the NTD of GluA2 (Jin et al., 2009) fits nicely in the NTD density of our EM structure (Fig 6B, and D). Additionally, the two separated small globular densities of the wildtype dimer can each accommodate the crystal structure of the GluA2 LBD monomer (Armstrong et al., 1998), while the density that corresponds to the LBD in the L504Y dimer is consistent with the dimeric crystal structure of the LBD (Fig 6B and D) (Sun et al., 2002). Because of the limited resolution of our EM map, we did not refine the position of the crystal structures using computational algorithms. However, placing crystal structures into our 3D reconstruction demonstrates that the size and shape of the densities of the extracellular domains of dimeric GluA2 wildtype and L504Y are compatible with known crystal structures. Thus, we conclude that the structural differences of the class averages between GluA2 wildtype and GluA2L504Y do not reflect different views of an identical 3D structure but represent true structural differences. Specifically, the structural difference between the wildtype and L504Y dimers can be attributed to the different global arrangement of the LBDs.
Stargazin/TARPs are auxiliary subunits of native AMPA-Rs in the brain. Because endogenous stargazin/TARPs were not detected in HEK cells, we investigated the effect of stargazin on subunit assembly of GluA2 by introducing stargazin into the parental stable HEK cell that DOX-dependently expresses FLAG-tagged GluA2 flop (TetONGluA2 cell, Fig 2A). The new stable HEK cell lines, which we named TetONGluA2-stg cells, constitutively express stargazin-IRES-mCherry using the elongation factor (EF) promoter, and DOX-dependently express GluA2 flop (Fig 7A and B). The mCherry was co-expressed (not as a fusion protein) with stargazin to facilitate the visual isolation of stable clones by fluorescent microscopy.
The four stable TetONGluA2-stg cell lines (clone #2, 8, 9, and 10), express stargazin at different levels. Interestingly, these cells started to die after 24 hr of DOX induction, suggesting that the expression of GluA2 induced cytotoxicity in the presence of stargazin. The cytotoxicity was stronger in the clones that express higher levels of stargazin and, as a result, all the cells died within 48 hr in clone#10 and 8. The cytotoxicity was suppressed when the cells were cultured with 30 µM of NBQX, an antagonist of AMPA-Rs. Because the same level of GluA2 expression was not cytotoxic in TetONGluA2 cells, we interpret that the glutamate in the media caused cell death by activating AMPA-Rs whose function was enhanced by stargazin. These observations suggest that both GluA2 and stargazin are functional in these cells.
First, we investigated if stargazin alters the biosynthesis of GluA2. All the TetONGluA2-stg cell lines expressed similar levels of GluA2 24hr after DOX induction. To assess the effect of stargazin on GluA2 maturation, we induced GluA2 expression in various TetONGluA2-stg cells and compared the timecourse of expression with the parental TetONGluA2 cells that lack stargazin. We were unable to detect any significant difference in the rate of GluA2 expression (Fig 7B). Thus, in HEK cells stargazin is not the rate limiting molecular chaperone for GluA2 biosynthesis.
When GluA2 was purified from clone #10, 12 hr after DOX application, no detectable amount of stargazin was co-purified. However, when purification was done 24 hr after DOX induction, stargazin co-purified with GluA2 and associated only with the tetrameric forms of GluA2, as determined by Superdex 200 gel filtration (Fig 7C). Consistently both proteins co-localized at the cell periphery when clone#10 cells were double stained 24 hr after induction (Fig 7D).
This study establishes a new approach to investigate subunit assembly of AMPA-Rs using a combination of genetic engineering and single particle EM. We utilize recombinant GluA2 tetramers obtained from HEK cells whose structures (at the resolution of our current study) are indistinguishable from those purified from rat brain (Nakagawa et al., 2005; Nakagawa et al., 2006) (Fig 1). This system has several advantages because the subunit composition can be precisely controlled, genetic manipulation is feasible, and the protocol is simple and highly reproducible. While AMPA-Rs form heterotetramers in the brain (Wenthold et al., 1996), it has been established that homotetrameric receptors are functional ion channels when expressed in HEK cells (Swanson et al., 1997). Many structural studies of AMPA-Rs are done using the GluA2 subunit. In addition, this subunit renders AMPA-Rs impermeable to calcium, therefore they are less toxic when overexpressed in cells. Insights obtained from homomeric GluA2 may therefore also apply to the heteromeric AMPA-Rs.
Recombinant AMPA-Rs were structurally more homogeneous than the AMPA-R particles purified from the brain (Nakagawa et al., 2005; Nakagawa et al., 2006). Because application of glutamate to the purified AMPA-R induces conformational changes of the NTD, we speculate that the structural heterogeneity observed in brain derived AMPA-Rs is caused by exposure of the particles to the endogenous glutamate during purification. Consistently, it is critical to include antagonist kynurenic acid (or NBQX) during the detergent solubulization and subsequent immunoaffinity chromatography to obtain structurally homogeneous recombinant AMPA-R tetramers.
The AMPA-R auxiliary subunit stargazin/TARP was not detected in HEK cells, nor from fractions of purified GluA2 from HEK cells that should in principal enrich endogenous stargazin/TARPs if they are present (Note, however, that two proteins co-purified when stargazin was introduced into the system (Fig 7C)). We estimate that there is very little, if any, and that possible endogenous stargazin/TARPs in HEK cells contribute little to tetrameric assembly of AMPA-R subunits. Consistently, stargazin/TARPs form a stable complex with GluA2 tetramers but not with the dimer intermediates during biogenesis (Fig 7C). In addition, the timecourse of expression of the newly synthesized GluA2 was not affected by the presence of stargazin in these cell lines. Stargazin functions as a molecular chaperone for AMPA-Rs in the presence of the ER stress response (Vandenberghe et al., 2005a). Because our expression level of GluA2 was modest, it is unlikely that the ER stress response was elicited. Taken together, it is likely that stargazin functions as a chaperone only when there is a high demand to synthesize AMPA-Rs. The lack of accumulation of AMPA-R dimers in the stargazer mutant mice is likely due to the fact that the ER is unlikely to be in a stress state (Vandenberghe et al., 2005b). Cornichons are newly identified auxiliary subunits of AMPA-Rs that modulate receptor trafficking (Schwenk et al., 2009). This suggests the possibility that they are also involved in the assembly process of AMPA-Rs. However, no definitive data is currently available.
Trafficking intermediates of membrane receptors are typically not easily accessible. By harvesting GluA2 within a short period after induction of transcription, we were able to enrich the biosynthetic intermediates. GluA2 protein expression reached a plateau more than 24 hr after DOX induction in HEK cells. Interestingly, the half-life of an AMPA-R subunit is 18 hr as determined by radioisotope metabolic labeling in cortical neurons (O'Brien et al., 1998). As the duration of protein turnover and molecular structures are consistent between AMPA-Rs expressed in HEK cells and endogenous receptors in neurons, HEK cells likely have the necessary molecular chaperones to correctly assemble AMPA-Rs.
The ratio of tetramer to dimer increased as a function of time after GluA2 induction by DOX. When determined by negative stain EM, purified AMPA-R tetramers remained in tetrameric form up to 5 days in buffer containing 0.1% DDM, suggesting that the tetrameric complexes are relatively stable (data not shown). Thus, it is unlikely that the dimers we purified are an artifact of detergent solubilization. Dimeric GluA2, a biogenic trafficking intermediate, is a previously uncharacterized molecular species of AMPA-Rs from a structural point of view. We speculate that stable dimers are common biosynthetic intermediates of AMPA-Rs, regardless of subunit composition. Such an idea is consistent with the preferential interaction of GluA1 with GluA1 NTD relative to GluA2 NTD (Leuschner and Hoch, 1999).
The 3D structure of dimeric GluA2 revealed that the LBDs are spatially separated, a novel feature that was not previously observed in AMPA-R structures. The NTD and TMD form dimers but the LBD remains separated. This observation provides a structural basis for a previous model derived from electrophysiological experiments in which the NTD and TMD are critical determinants for the formation of functional channels (Ayalon and Stern-Bach, 2001). Our structural finding suggests that dimerization of both the NTD and TMD precedes tetramerization. This agrees with the interruption of the polypeptide sequence of the LBD by the TMD, which implies that folding of the TMD coordinates with that of the LBD during translation.
To study the functional significance of the structure of GluA2 wildtype dimers, we identified GluA2L504Y as a mutant defective in tetramerization, determined the structure of the dimers of this mutant, and compared the structure with the wildtype. GluA2L504Y and analogous mutants in other AMPA-R subunits have been studied extensively as non-desensitizing mutants (Rosenmund et al., 1998; Stern-Bach et al., 1998; Robert et al., 2001; Sun et al., 2002). The tetramerization defect, however, was not observed when GluA2L504Y was exogenously expressed in neurons (Greger et al., 2006). This can be explained by the possible co-assembly of exogenous mutant GluA2L504Y subunits with endogenous wildtype AMPA-R subunits. However it remains unclear why heterotetramers that contain GluA2L504Y did not exit the ER. Additional mechanisms such as sampling of gating motion in the ER may regulate this process (Penn et al., 2008).
In the dimers of GluA2L504Y, all three domains (NTD, LBD, and TMD) appear to be dimerized or at least in close proximity. The dimeric modules are arranged linearly, thus making the overall shape of the complex elongated compared to wildtype structures (compare Fig 2, Fig 5, and Fig 6). The compact structure of the LBD in dimeric GluA2L504Y is consistent with the high affinity dimer formed by the S1S2 construct of the LBD that carries the same mutation (Sun et al., 2002). Our data is also consistent with previous results indicating that native tetrameric AMPA-Rs treated with 1 mM glutamate have more compact structures in the presence of 330 µM cyclothiazide, an allosteric inhibitor of desensitization (Nakagawa et al., 2005).
The two LBDs are separated in the GluA2 wildtype dimers and therefore, intermolecular dimerization of the LBDs can potentially occur between two GluA2 wildtype dimers during assembly of a tetramer. In contrast, the LBDs are fused in the GluA2L504Y dimers. Since this mutation causes a defect in dimer-to-tetramer transition, it is conceivable that the separation of LBD dimers is required to drive tetramerization (Fig 8A). Thus, our results support a new model for the subunit assembly pathway of AMPA-Rs in which the dimer-to-tetramer transition accompanies formation of two new LBD dimers between the two molecular dimers of subunits (Fig 8B). This suggests an unexpected domain arrangement in tetrameric AMPA-Rs, in which the NTD and LBD of each subunit forms a dimer with a different neighboring subunit.
As with any structural approaches, technical artifacts must always be considered. The carbon support and the negative stain can potentially introduce a small distortion in the molecular structure. Even if we assume the presence of a small distortion, the robust structural differences between the GluA2 wildtype dimer and L504Y dimer were detected when both specimens were prepared under identical experimental procedures. Therefore the observed structural differences of the particles on EM grid should reflect the structural differences in solution. In addition, when carefully inspected, the peak elution volume of wildtype and L504Y dimers are different (Fig 4C and D). Because the molecular weights of the wildtype and the L504Y mutant are nearly identical, this small difference in the peak elution volume suggests a structural difference between the two molecular forms. The direct structural inspection by single particle EM and the hydrodynamic properties obtained from quantitative gel filtration chromatography both support the presence of a structural difference between the wildtype and L504Y dimers, thus experimental artifacts are an unlikely explanation for the observed gross structural differences.
In both AMPA-Rs and kainite receptors, point mutations that prevent receptor desensitization also result in decreased surface delivery (Greger et al., 2006; Priel et al., 2006). Accordingly, it has been proposed that glutamate receptors sense glutamate prior to being delivered to the cell surface, which can prevent non-desensitizing and non-functional subunits from going to the surface (Mah et al., 2005; Valluru et al., 2005) (Penn et al., 2008). Because globular LBDs are clearly detected in immature AMPA receptors, glutamate might be used as a tool to facilitate progression through the subunit assembly pathway of AMPA-Rs as well. All in all we have created a system that has allowed us to study molecular details of AMPA-R assembly. Comparison of wildtype and mutant receptors has provided insight into the domain arrangements of the dimeric intermediates which are conducive to the receptor transitioning into mature tetrameric form.
We thank Peter Seeburg and Yasunori Hayashi for rat GluA2 flop, and GFP-GluA2flop cDNA clones. We acknowledge the use of the UCSD Cryo-Electron Microscopy Facility which was supported by NIH grants 1S10RR20016 and GM033050 to Dr. Timothy S. Baker and a gift from the Agouron Institute to UCSD. We thank Anirvan Ghosh for giving us access to their epifluorecent microscope during the initial phase of this study. We also thank Palmer Taylor for HEK wt and HEK GnTI(-) cell lines and Toni Koller for conducting LC/MS/MS. We thank Thomas Walz for discussion. We thank Hiro Hakozaki, Ohkyung Kwon, Guido Gaietta, and Tom Deerinck for their help at NCMIR. We thank Mark Elliott for transfecting and immuno staining neurons. This work is supported by grants from John Merck Fund, Hellman Foundation (to T.N.) and NIH P41004050 (to M.H.E.). N.F.S is supported by the NIH Molecular Biophysics Training Grant (NIH GM08326). A.N.F is supported by the NIH CMG Training Grant (T32 GM007240-34). T.N. is a recipient of NARSAD Young Investigator Award.
Note added in proof:
While this manuscript was being reviewed, a paper reporting the X-ray crystal structure of GluA2cryst was published (Sobolevsky et al., 2009). Despite the fact that 6 out of 16 amino acids in the wildtype GluA2 were deleted from the linker that connects the NTD and LBD, the domain arrangement reported for the X-ray crystal structure of GluA2cryst is consistent with our model described in Figure 8.
Sobolevsky AI, Rosconi MP, Gouaux E (2009) X-ray structure, symmetry and mechanism of an AMPA-subtype glutamate receptor. Nature 462:745–756.