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By Neeraj Sharma, Chaitanya Patel, Marina Shenkman, Amit Kessel, Nir Ben-Tal, Gerardo Z. Lederkremer

Excerpt from the article published in Journal of Biological Chemistry, 297, 5, 101299, November 2021 DOI: https://doi.org/10.1016/j.jbc.2021.101299

Editor’s Highlights

• The Sigma-1 receptor (S1R) is a small transmembrane protein (25 KDa), which is localized predominantly in the endoplasmic reticulum (ER), preferentially at the mitochondrion-associated membranes (MAM).
• The S1R is expressed in many tissues and has important roles in modulating calcium signaling and mitochondrial function, with special importance in the central nervous system.
• Decreased levels of S1R or of its activity have been found associated with neurodegenerative diseases, and S1R activation by agonists is neuroprotective in several of these diseases.
• The S1R is an ER resident protein, not reaching the plasma membrane.
• The bulk of the S1R protein, spanning the C-terminal portion, is located in the ER lumen.
• The fact that the S1R ligand-binding domain is thus inside the ER lumen predicts the properties that are shared by S1R ligands, such as hydrophobicity.

Abstract

The Sigma-1 receptor (S1R) is a transmembrane protein with important roles in cellular homeostasis in normal physiology and in disease. Especially in neurodegenerative diseases, S1R activation has been shown to provide neuroprotection by modulating calcium signaling, mitochondrial function and reducing endoplasmic reticulum (ER) stress. S1R missense mutations are one of the causes of the neurodegenerative Amyotrophic Lateral Sclerosis and distal hereditary motor neuronopathies. Although the S1R has been studied intensively, basic aspects remain controversial, such as S1R topology and whether it reaches the plasma membrane. To address these questions, we have undertaken several approaches. C-terminal tagging with a small biotin-acceptor peptide and BirA biotinylation in cells suggested a type II membrane orientation (cytosolic N-terminus). However, N-terminal tagging gave an equal probability for both possible orientations. This might explain conflicting reports in the literature, as tags may affect the protein topology. Therefore, we studied untagged S1R using a protease protection assay and a glycosylation mapping approach, introducing N-glycosylation sites. Both methods provided unambiguous results showing that the S1R is a type II membrane protein with a short cytosolic N-terminal tail. Assessments of glycan processing, surface fluorescence-activated cell sorting, and cell surface biotinylation indicated ER retention, with insignificant exit to the plasma membrane, in the absence or presence of S1R agonists or of ER stress. These findings may have important implications for S1R-based therapeutic approaches.

The Sigma-1 receptor (S1R) is a small transmembrane protein (25 KDa), which is localized predominantly in the endoplasmic reticulum (ER), preferentially at the mitochondrion-associated membranes (MAM) (1, 2, 3, 4, 5). The S1R is expressed in many tissues and has important roles in modulating calcium signaling and mitochondrial function, with special importance in the central nervous system. Decreased levels of S1R or of its activity have been found associated with neurodegenerative diseases (6, 7, 8, 9, 10, 11), and S1R activation by agonists is neuroprotective in several of these diseases (12, 13, 14, 15). However, the basic topology of this protein remains controversial. Initial studies suggested the existence of two transmembrane domains (TMDs) (4). The determination of the crystal structure of the S1R points to only one TMD near the N-terminus of the protein, but still leaves open the possibility of two possible orientations in the membrane. There is no cleaved signal peptide, but the C-terminal bulk of the protein could be in the lumen of the ER (type II membrane protein) or in the cytosol (type III membrane protein).
Another important issue is to what extent the S1R reaches the cell surface. The S1R appears to be mainly localized at an ER subdomain that contacts mitochondria, the MAM (4). However, it has also been reported to be present on the plasma membrane (16) where it might be translocated upon activation with agonists (17). However, on closer inspection, the location seems to be not at the plasma membrane but at close subsurface areas (17, 18), perhaps at ER domains adjacent to the plasma membrane (19).
Given the biological and pharmacological importance of the S1R, a clear understanding of its topology relative to the membrane and of its presence or absence at the cell surface is essential. Prediction algorithms of S1R membrane topology give a wide array of possibilities, whereas the conflicting studies use tags that might affect S1R topology. Here we analyzed S1R topology using different approaches, some of them with untagged S1R. In one such approach, we introduced N-glycosylation sites at key residues and tested their glycosylation. These constructs also allowed us to follow their sugar chain processing as an indicator of ER exit.

Results

 In vivo biotinylation of BAP-tagged S1R

The crystal structure of the human sigma-1 receptor indicated the existence of only one TMD near the N-terminus of the protein (20). Previous reports had suggested one or two TMDs (4, 21, 22, 23). Furthermore, analyses of the S1R protein sequence (Fig. 1A) by different topology prediction methods estimate a surprisingly wide range of topologies, with one to three TMDs, and both possible orientations of the N-terminus, toward the ER lumen or toward the cytosol (Fig. S1).

If indeed the S1R has a single TMD, given the lack of a signal peptide, it could adopt either a type II orientation (C-terminus in the ER lumen) or a type III orientation (N-terminus in the ER lumen). Moreover, both the prediction published with the 3D structure (20) and the combined prediction of the CCTOP server (24) (Fig. S1) suggest a type III orientation. To further inspect the S1R orientation, we first used a system that allows in vivolabeling in cells and sensitive detection of protein epitopes in the ER lumen or in the cytosol. This system uses a specific biotinylation reaction by the Escherichia coli-derived biotin-ligase BirA (25) on a 15 aa long biotin acceptor peptide (BAP), which is linked to the protein of interest. BirA can be expressed in the cytosol in mammalian cells, reacting only with BAP exposed to the cytosol (26). In contrast, a luminally expressed BirA (sec-BirA) labels BAP when exposed to the ER lumen. We have made constructs expressing BAP-tagged S1R, with the tag on the N-terminus (BAP-S1R) or C-terminus (S1R-BAP). If the protein has a type II orientation, S1R-BAP should be biotinylated by sec-BirA and not by BirA and vice versa for BAP-S1R. Conversely, if it has a type III orientation, S1R-BAP should be biotinylated by BirA and not by sec-BirA and vice versa for BAP-S1R (Fig. 1B). An SV5 tag was added next to the BAP tag, to detect the protein independently of biotinylation. The tagged proteins were expressed in HEK293 cells together with BirA or sec-BirA, and the cells were incubated with biotin. Cell lysates (prepared in the presence of N-ethylmaleimide to prevent postlysis biotinylation) were immunoblotted with anti-V5 to detect total levels of the proteins, or with Streptavidin linked to HRP (Strep-HRP) to detect only biotinylated molecules. The results show biotinylation of S1R-BAP by sec-BirA and only to a minor extent (∼25% of total) by BirA, suggesting a mostly type II orientation (Fig. 1, C and E). This was similar to the results obtained with a BAP-tagged construct made with a protein known to have a type II orientation, asialoglycoprotein receptor H2a (27) (Fig. 1, D and E). The small amount of labeling by BirA probably reflects molecules that have undergone retrotranslocation toward ERAD ((26, 28) and our unpublished results). In contrast to S1R-BAP, BAP-S1R was biotinylated to a similar extent by both BirA and sec-BirA (Fig. 1, C and E). This result was surprising and might be due to the N-terminal tag affecting the membrane orientation of the protein, giving equal probability for insertion in the two opposite orientations. This effect of the tag might explain the conflicting results in other studies that used S1R with other tags (4, 21, 29, 30). The orientation of H2a with a BAP-tag in the N-terminus (BAP-H2a) was affected to some degree, but still showed a predominant type II orientation (63% of total BAP-H2a) (Fig. 1, D and E).
We analyzed whether the tags affect the oligomerization of the S1R. For this, we separated Triton-insoluble species (higher order oligomers (31)) and kept the lysates at 4 °C before SDS-PAGE, which prevents dissociation of S1R dimers (32). To avoid the formation of mixed oligomers, we performed these experiments by expressing the constructs in HEK293 cells where S1R was deleted using CRISPR/Cas9 technology (S1R−/−) (15). Whereas C-terminal BAP tagging caused a small increase in dimerization and did not affect association into higher-order oligomers, N-terminal tagging reduced significantly the presence of dimers (Fig. 2, A and C) and Triton-insoluble species (Fig. 2, B and C) compared with WT untagged S1R.

A protease protection assay and the introduction of N-glycosylation sites in untagged S1R reveal a type II orientation

Given the confounding effect of N-terminal tagging, we decided to determine the topology of untagged S1R. We applied a classical protease protection assay on microsomes prepared from S1R−/− HEK293 cells transfected with WT S1R and compared with S1R-BAP and H2a-BAP. The microsomes were treated with proteinase K after treatment or not with 1% SDS. All three proteins were completely digested in the presence of SDS and were protected from the protease in the absence of SDS (Fig. 3A). Both S1R and S1R-BAP showed a small shift, using anti-S1R or anti-V5 antibodies for the latter, indicating a cleavage by proteinase K of a peptide of ∼1.3 kD, which is consistent with digestion of the small cytosolic tail in a type II orientation (Fig. 3B). H2a-BAP showed a larger cleavage of ∼6 kD, which is the size of its cytosolic tail. As controls, a luminal protein, BiP-RFP showed complete protection in the absence of SDS and calnexin (CNX), a type I membrane protein, was no longer recognized by an anti-C terminal antibody, in the absence or presence of SDS (Fig. 3A).

To discard any changes that might take place during the manipulation needed for the preparation of microsomes, we used another approach to reveal the topology of untagged S1R in intact cells. S1R is a nonglycosylated protein. To analyze the orientation of untagged S1R, we made different constructs where individual N-glycosylation sites were introduced and expressed them in S1R−/− HEK293 cells. As N-glycosylation takes place only in the ER lumen, these constructs should reveal the orientation of the protein. N-glycosylation does not affect the normal topology of the protein as the N-glycans are added only after translocation into the ER lumen of a given segment containing an N-glycosylation sequon. N-glycosylation sites were introduced at exposed hydrophilic regions and with minimal effects on polarity (Fig. 4A). A viability assay was used to test if the constructs expressed functional S1R, which would protect cells from ER stress-induced cytotoxicity caused by long-term incubation with thapsigargin (TG). S1R functionality would reflect a native conformation. All constructs protected the cells to a similar or slightly higher extent than WT S1R, except for S1R L214N, which showed a lower, although still significant protection (Fig. 4B). Constructs with N-glycosylation sites near the N-terminus, S1R A4N and S1R T32N, resulted in proteins with no change in migration in SDS-PAGE compared with WT S1R, suggesting no glycosylation at these sites (Fig. 4C). In contrast, a construct with a more C-terminal N-glycosylation site, S1R Q44N, and one near the C-terminus, S1R L214N, resulted in a slower migration in SDS-PAGE compared with WT S1R, consistent with N-glycosylation at these sites. We observed the same differences between these proteins when expressed in S1R+/+ or S1R1−/− cells (Fig. 4C). Only a minority of S1R Q44N (14%) and S1R L214N molecules (18%) were not glycosylated, measured in S1R1−/− cells. These results suggest that most of the protein, including its C-terminus, is in the ER lumen. The T32N site is in the lumen, but too close to the TMD (residues 10–31) to be glycosylated, as a stretch of a minimum of 12 residues C-terminal to a membrane span is needed for N-glycosylation to take place (33) (Fig. 4D). An opposite orientation, with the N-terminus in the ER lumen would have resulted in no glycosylation for any of the constructs, as the S1R A4N is also too close to the membrane span. The results suggest a type II transmembrane protein topology.

Neither S1R agonists nor ER stress changes the topology or the ER exit of S1R constructs

To test whether the topology or orientation of the S1R constructs was affected by an S1R agonist, we incubated S1R−/− HEK293 cells expressing WT S1R or the constructs with added N-glycosylation sites with the S1R agonist Pre-084 (34). There was no change in the migration in SDS-PAGE for any of the constructs with or without Pre-084 incubation (Fig. 5A). Similar results were obtained with the S1R agonist pridopidine (35, 36) (Fig. 5B).

We then analyzed the extent of ER exit of the proteins by measuring their resistance to endo-β-N-glycosaminidase H (endo H), which cleaves high-mannose type N-glycans. N-glycans are processed in the Golgi to complex-type sugar chains, acquiring resistance to endo H. Endo H treatment caused no change in the migration of WT, S1R A4N, and S1R T32N, as they are not glycosylated. In contrast, endo H treatment of S1R Q44N and S1R L214N changed their migration to a position similar to the WT S1R (Fig. 6A). There were few if any S1R Q44N and S1R L214N molecules that were resistant to endo H. To discard the possibility that even though they exit to the Golgi, these proteins carry high mannose N-glycans that are not converted to complex-type, cells were treated with brefeldin A (BFA). BFA causes ER-Golgi fusion and therefore delivers Golgi enzymes to the ER. When cells expressing S1R L214N were treated with BFA, the glycoprotein became partially (34 %) endo H resistant (Fig. 6A, rightmost lane). This result indicates that the S1R L214N N-glycan is capable of undergoing processing by Golgi enzymes to an endo H-resistant form, but is normally totally endo H-sensitive, suggesting complete ER retention.

As S1R agonists were reported to translocate the protein to or near the plasma membrane (17), we tested whether they cause any change in the endo H sensitivity. Neither Pre-084 nor pridopidine changed the endo H sensitivity of S1R Q44N or S1R L214N (Fig. 6, B and C). We also tested whether ER stress causes any changes, because the S1R is activated by ER stress (37). Incubation of cells with TG for 16 h did not change the endo H sensitivity of the glycosylated S1R constructs (Fig. 6D).
Although the results suggest that no S1R exits the ER, we wanted to test whether any molecules reach the cell surface. With this aim we first used fluorescence activated cell sorting (FACS) on nonpermeabilized S1R−/− cells expressing S1R or endogenous S1R in HEK293 cells. As a positive control, we measured the surface expression of an unassembled receptor subunit, asialoglycoprotein receptor H1, which can exit the ER and reach the cell surface but only to a modest extent (27). A very minor cell surface signal, although statistically significant, was observed for both transfected and endogenous S1R using anti-S1R antibodies (which recognize an epitope that should be exposed if S1R reaches the cell surface (Fig. 7A and Fig. S2)). In contrast, H1 showed strong surface fluorescence. When analyzing permeabilized cells, the signals obtained for S1R and H1 were of similar magnitude (Fig. 7B). The ratio of cell surface (nonpermeabilized cells) to total (permeabilized cells) was ∼30% for H1 and only ∼4% for S1R. Cell incubation with Pre-084 caused only a small increase in this ratio. We also analyzed the cell surface expression of BAP-S1R, reasoning that the altered topology caused by an N-terminal tag might affect the ER retention, but it also gave a very minor cell surface signal in the absence or presence of the agonist (Fig. S2). Therefore, only a minor fraction of S1R, if any, appears to reach the cell surface (Fig. 7C).

We then used another approach and applied a sensitive and specific surface biotinylation approach, using the membrane-impermeant reagent sulfosuccinimidyl-6-(biotinamido)hexanoate (sulfo-NHS-LC-Biotin). Biotinylated cell surface proteins from S1R−/− cells expressing WT S1R or S1R L214N were precipitated from cell lysates using streptavidin-agarose beads and immunoblotted with anti-S1R compared with total S1R. A weak signal of biotinylated S1R could be observed only upon overexposure of the blot and accounted for 2.2% of total WT S1R and 2.8% of total S1R L214N (Fig. 8, A and D and Fig. S3). There was no change when cells were treated with Pre-084. H1, when expressed in S1R−/− cells showed 12.7% of cell surface expression (Fig. 8, B and D). We also analyzed H1 expression in S1R+/+ cells, where it showed similar levels of cell surface biotinylated protein. To evaluate the significance of the low percent of surface S1R, we immunoblotted the same biotinylated samples with anti-CNX antibodies, as this is an ER-retained chaperone. Cell surface-biotinylated molecules of CNX were 1.7% of the total CNX in WT S1R expressing cells and 3.2% in S1R L214N expressing cells, not significantly different than the cell surface labeling obtained for the S1R (Fig. 8, C and D). Altogether, these results suggest that the S1R is an ER resident protein, not exiting the ER to any meaningful extent.

Discussion

Previous studies of the S1R suggested a wide variety of topologies, including two TMDs, with both N- and C- termini facing the cytosol (21) or with both N- and C- termini facing the lumen (4). Studies of S1R truncation mutants suggest that the second hydrophobic segment, toward the C-terminus, may be peripherally attached, not being a TMD (22, 23). The recent determination of the crystal structure of the S1R indicates the existence of only one transmembrane span near the N-terminus. The probability that other hydrophobic segments would cross the membrane, as suggested by some of the bioinformatic predictions and by other studies, is extremely low.
Regarding the orientation of the S1R in the membrane, the studies also conflict. The crystal structure indicated one TMD; therefore the N- and C- termini of the protein must be on opposite sides of the membrane. However, the C-terminus was suggested to be in the cytosol in that study, from bioinformatic prediction (20). A recent study, based on BRET and ligand binding, confirmed that the N- and C- termini are on opposite sides, but was inconclusive as to the orientation (30). Our results with the BAP-tagged S1R, compared with those obtained with the untagged protein, suggest that at least N-terminal tagging can affect the orientation of the protein in the membrane. This is likely due to the short N-terminal tail, and thus the proximity of the tag to the TMD. N-terminal tagging was also reported to affect S1R oligomerization (32), which we confirmed with BAP-S1R. A study using an ascorbate peroxidase 2 (APEX2) tagging approach indicated a type II orientation, thus in this case the tagging did not seem to affect the orientation of the protein (29). Our study suggests that untagged S1R is in a type II orientation, with a short 9 amino acid N-terminal cytosolic tail, a TMD of 22 residues (10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21, 22, 23, 24, 25, 26, 27, 28, 29, 30, 31), and a C-terminal portion, amino acids 32–223 in the lumen of the ER (Figs. 3 and 4). The classical protease protection assay indicates this clearly (Fig. 3).
The N-glycosylation mapping approach also yielded clear-cut results, with the site near the amino-terminus not being glycosylated, whereas the sites carboxy-terminal to the TMD are glycosylated, except for S1R T32N, which is too close to the membrane. This indicates that the S1R is a transmembrane protein with a signal anchor and a type II orientation, which is consistent with the positive-inside rule (38), as there are more positive residues in the amino-terminal, cytosolic side of the TMD (Fig. 4D). Only the TMHMM method gave this prediction (Fig. S1). The glycosylation results would also be compatible with Prodiv, but this would need 3 TMDs, which as mentioned above, is very improbable and is not in line with the crystal structure.
The minor cell surface biotinylation obtained for CNX (Fig. 8) could be due to a technical limitation of the experiment or to real cell surface expression of a small population of molecules of the ER chaperone, as has been reported previously in some cell types (39). Therefore, the similar minor biotinylation of the S1R suggests either no cell surface expression or only to a very minor extent (∼2%), comparable to that of an ER chaperone. Although this might be influenced by the fact that there is only one lysine in the S1R exposed domain that could be biotinylated, the FACS experiment confirms the finding. The results of the FACS experiment also yielded very low cell surface expression (∼4%). This suggests that the S1R cannot exit the ER to a significant extent by itself, although we cannot exclude that in some cell types it might exit the ER in complex with other protein(s).
The ER localization and type II orientation are consistent with S1R association with the ER luminal chaperone BiP and S1R activation upon dissociation from BiP (4). It is also in line with the main S1R function, modulation of the activity of the ER-localized inositol 1,4,5-triphosphate receptor (IP3R) (4, 40, 41).
S1R activation could hypothetically affect its topology, and it has been reported to translocate to the plasma membrane upon activation with agonists (17). However, our results indicate that neither the widely used S1R agonist Pre-084 (13, 34, 42) nor the high-affinity S1R ligand pridopidine (35, 36) affect the topology or ER exit of the S1R (Figure 5, Figure 6, Figure 7, Figure 8). Likewise, ER stress, which also activates the S1R (4), did not alter its ER retention (Fig. 6). It is possible that the report of plasma membrane location might have been due to technical shortcomings, and it might have been rather in internal membranes close to the plasma membrane, as mentioned above, likely at ER domains adjacent to the plasma membrane, as has also been described (19).
In conclusion, our results suggest that the S1R is an ER resident protein, not reaching the plasma membrane. The bulk of the S1R protein, spanning the C-terminal portion, is located in the ER lumen. This outcome concerning the S1R transmembrane orientation is the opposite of that predicted by some web-based algorithms and by the report on the crystal structure of the S1R (20). The fact that the S1R ligand-binding domain is thus inside the ER lumen predicts the properties that are shared by S1R ligands, such as hydrophobicity. Given the notable protective activity of the S1R in diseases, especially neurodegenerative diseases (12, 13, 14), our results could have important consequences in informing target-based therapies.