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J. Biol. Chem., Vol. 282, Issue 31, 22865-22878, August 3, 2007
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Effects of the Isoform-specific Characteristics of ATF6
and ATF6
on Endoplasmic Reticulum Stress Response Gene Expression and Cell Viability*
From the San Diego State University Heart Institute and Department of Biology, San Diego State University, San Diego, California 92182
Received for publication, February 8, 2007 , and in revised form, May 22, 2007.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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The endoplasmic reticulum (ER)-transmembrane proteins, ATF6
and ATF6
, are cleaved during the ER stress response (ERSR). The resulting N-terminal fragments (N-ATF6 and N-ATF6) have conserved DNA-binding domains and divergent transcriptional activation domains. N-ATF6 and N-ATF6 translocate to the nucleus, bind to specific regulatory elements, and influence expression of ERSR genes, such as glucose-regulated protein 78 (GRP78), that contribute to resolving the ERSR, thus, enhancing cell viability. We previously showed that N-ATF6 is a rapidly degraded, strong transcriptional activator, whereas is a slowly degraded, weak activator. In this study we explored the molecular basis and functional impact of these isoform-specific characteristics in HeLa cells. Mutants in the transcriptional activation domain or DNA-binding domain of N-ATF6 exhibited loss of function and increased expression, the latter of which suggested decreased rates of degradation. Fusing N-ATF6 to the mutant estrogen receptor generated N-ATF6-MER, which, without tamoxifen exhibited loss-of-function and high expression, but in the presence of tamoxifen N-ATF6-MER exhibited gain-of-function and low expression. N-ATF6 conferred loss-of-function and high expression to N-ATF6, suggesting that ATF6 is an endogenous inhibitor of ATF6. In vitro DNA binding experiments showed that recombinant N-ATF6 inhibited the binding of recombinant N-ATF6 to an ERSR element from the GRP78 promoter. Moreover, siRNA-mediated knock-down of endogenous ATF6 increased GRP78 promoter activity and GRP78 gene expression, as well as augmenting cell viability. Thus, the relative levels of ATF6 and -, may contribute to regulating the strength and duration of ATF6-dependent ERSR gene induction and cell viability. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Stresses that alter the rough ER3 environment can impair folding of proteins synthesized by this organelle (1-4). Numerous proteins induced under such conditions are targeted to the ER, where they aid in nascent protein folding and thus, counteract the stress; this ER-initiated signaling process is known as the ER stress response (ERSR). ERSR elements (ERSEs) are located in the regulatory regions of many ERSR genes. One of the transcription factors that mediates ERSR gene induction via ERSEs is ATF6
, a 670-aa ER trans-membrane protein (5, 6) (Fig. 1A, ATF6). ER stress activates the proteolytic cleavage of 
400 aa from the N terminus of ATF6 (N-ATF6) (7), which translocates to the nucleus and activates numerous ERSR genes (8, 9). The transcriptional activation domain (TAD) of N-ATF6 resides in the N-terminal portion of the protein, whereas the basic leucine zipper (b-Zip) and nuclear localization domains reside in the C terminus (Fig. 1B, N-ATF6) (8, 10). N-ATF6 can bind directly to ATF6 binding sites (9), or it can combine with several other proteins to form a complex that binds to ERSEs and augments the induction of numerous ERSGs, such as the ER chaperone, glucose-regulated protein 78 kDa (GRP78) (8, 9, 11-13). N-ATF6 exhibits potent transcriptional activity, however, it is susceptible to proteasome-mediated degradation, and mutations in the TAD that reduce N-ATF6 transcriptional activity decrease degradation (14). Several other potent transcription factors that exert rapid, transient effects exhibit similar coupling of transcriptional activation and degradation (15), including the virally encoded protein, VP16 (16). An 8-aa domain in VP16, called VN8, confers strong transcriptional activity and susceptibility to degradation, and mutations in VN8 that reduce VP16 activity decrease degradation (17, 18). The TAD of ATF6 possesses a VN8-like sequence, and mutating it in ways known to decrease VP16 activity decrease ATF6 activity and degradation (14). To the best of our knowledge, the VN8 domain has not been found in any other mammalian transcription factor, including a second isoform of ATF6, ATF6.
Like ATF6, ATF6 is an ER-transmembrane protein (Fig. 1A, ATF6), and during ER stress proteolysis generates an N-terminal fragment of 400 aa (19). N-ATF6 and N-ATF6 possess highly conserved b-Zip domains, which allow them to bind to ERSEs as homo- or heterodimers (20); however, the N-terminal regions are divergent. For example, the region of N-ATF6 corresponding to the VN8 of N-ATF6 differs in 5 of 8 aa in ways predicted from studies with VP16 to diminish transcriptional activity (21) (Fig. 1B, N-ATF6). In support of this prediction were findings that ectopically expressed N-ATF6 is a poor ERSR gene inducer (6) that exhibits much greater stability than N-ATF6 (14). Accordingly, although they can bind to the same regulatory elements, N-ATF6 and - exhibit isoform-specific transcriptional activation and stability characteristics. Thus, N-ATF6 and - might function in a combinatorial fashion to fine-tune the strength of ERSR gene activation.
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In the present study, we examined the molecular mechanism and function of the isoform-specific characteristics of N-ATF6
and -, addressing the following hypotheses: 1) the isoform-specific characteristics of N-ATF6 and - are conferred by their divergent N-terminal TADs; 2) N-ATF6-mediated transcriptional activation and rapid degradation are coordinated processes, and 3) the relative levels of N-ATF6 and - impact ERSR gene induction and cell viability in ways consistent with roles of N-ATF6 as a transcriptional repressor of N-ATF6. |
EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Methods
Replicates and Statistical Analysis—Unless otherwise stated in the legends or figures, each treatment was performed on three identical cultures, and each experiment was repeated at least three times. Representative experiments are shown. Statistical analyses were performed using a one-way ANOVA followed by the Student-Newman-Keul post-hoc analysis. *, #, or
= p < 0.05; **, ##, , or 
= p < 0.01.
Cell Culture—HeLa Cells were maintained in Dulbecco's modified Eagle's medium containing 10% fetal calf serum. For transfection experiments, HeLa cells were resuspended at 5-9 x 106 cells per 400 µl of cold Dulbecco's phosphate-buffered saline and electroporated in a 0.4-cm gap electroporation cuvette at 250 V and 950 microfarads using a GenePulser II Electroporator (Bio-Rad). The cells were then plated at a density of 0.5 x 106 per 24-mm well for experiments involving luciferase and -galactosidase enzyme assays, or 1.5 x 106 per 35-mm well for experiments involving immunoblotting. Reporter assays and immunoblotting were carried out as previously described (22).
Plasmids
CMV-Galactosidase—CMV--galactosidase, which codes for a -galactosidase reporter driven by the CMV promoter, was used to normalize for transfection efficiency.
N-ATF6-VN8 Mutations—Mutations were introduced into 3x FLAG-ATF6-(1-392) to mimic the VN8 region of ATF6. Accordingly, the following changes were introduced into ATF6-VN8-M1: V65F, G66D, and M67L; whereas the following changes were introduced into ATF6-VN8-M2: V65F, G66D, M67L, V69L, and S70L. ATF6-VN8-M1 and ATF6-VN8-M2 were created by PCR, using QuikChange from Stratagene and the relevant primers required to introduce the desired amino acid substitutions.
N-ATF6 and N-ATF6 Chimeras—Constructs encoding chimeric proteins composed of various portions of N-ATF6 and - were designed by aligning the sequences of N-ATF6 and - and selecting homologous regions for domain swapping studies. Appropriate PCR fragments were generated from 3x FLAG-N-ATF6 and -, so that an XhoI restriction site was introduced at the ATF6 and - junction in each chimera. The amino acid sequence used to name each chimera refers to the original, full-length sequence of either ATF6 and -. The constructs generated are: ATF6-(1-114)/ATF6-(116-392), ATF6-(1-180)/ATF6-(191-392), and ATF6-(1-302)/ATF6-(322-392), which are named constructs 3, 4, and 5, respectively, in Fig. 3A.
N-ATF6 and N-ATF6 Gal4 DBD Fusion Proteins—A construct encoding the Gal4 DBD fused to the N terminus of ATF6-(1-114) (i.e. Gal4 DBD-ATF6-(1-114)) was created by PCR, using 3x FLAG-ATF6-(1-373) as the template, and the appropriate primers, to create an amplicon with a BamH1 site on the 5'-end, and a termination codon and SacI site on the 3'-end. This PCR product was then cloned into Gal4 DBD (pSG424, GenBankTM accession number X85976
[GenBank]
). Clones Gal4 DBD-M (L32P) and Gal4 DBD-M (L32P)-ATF6-(1-114) were generated using QuikChange from Stratagene and the appropriate primers.
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N-ATF6
-MER Fusion Protein—A PCR product composed of the nucleotides encoding aa 281-599 of the mouse mutated (G525R) estrogen receptor (MER, a gift from Dr. Michael Reth, Max-Plank-Institute, Freiburg, Germany) was cloned into the NotI/EcoR1 site of pCDNA3.1-3x FLAG vector to create 3x FLAG-MER. Subsequently, a PCR product of N-ATF6 was generated, which introduced a NotI site and removed the termination site after aa 373, was cloned into the XhoI/NotI site of 3x FLAG-MER to create 3x FLAG-N-ATF6-MER.
3x HA-ATF6—3x HA-ATF6 was generated by subcloning N-ATF6 from 3x FLAG-N-ATF6 into pCDNA3.1-3x HA, which has been described elsewhere (22).
GRP78-Promoter-luc—Reporter constructs encoding the GRP78 promoter from -284 to +7or -284 to +221 driving luciferase have been described elsewhere (23).
Small Interfering RNAs
The use of small interfering (si) RNA targeted against human ATF6 and - has been described elsewhere (22). Briefly, HeLa cells were plated on 6-well plates at 400 K cells per well, then transfected with 50 ng of the relevant dicer siRNAs, using Lipofectamine 2000TM. After 24 h, cells were treated with or without tunicamycin (2 µg/ml), and then examined by real-time quantitative PCR (see below), or, before tunicamycin treatment, they were removed from the plate with TripLE (Invitrogen), and re-plated in 96-well plates at 10 K cells per well in preparation for viability assays. To examine viability, cells in 96-well plates were treated with or without tunicamycin (2 µg/ml) and 2-deoxyglucose (3 mM) in serum-free media for 32 h. Cell viability was then assessed using an MTT Cell Proliferation Kit according to the manufacturer's protocol (Roche Applied Science). Samples were read at 570 nm in a VersaMax microplate reader (Molecular Devices, Downingtown, PA).
Real-time Quantitative PCR
HeLa cells were transfected with the appropriate siRNA, as described above, then after treatments, they were lysed and RNA was extracted using an RNeasy kit (Qiagen). cDNA was generated by reverse transcription using a Superscript III kit (Invitrogen). Real-time quantitative PCR was performed on cDNA using the Quanti-Tect SYBR Green PCR kit (Qiagen) on an ABI Prism 7000 (Applied Biosystems, Foster City, CA). The relative abundance of GRP78 RNA was calculated using the 
Ct method, as previously described (24). Primers (see below) were designed using primer express version 2.0 (Applied Biosystems). All primers were determined to be 90% to 110% efficient, and all exhibited only one dissociation peak as follows: GRP78: (+) CCACCTCAGTCTCCCAGCTAA; (-) GCCGAGCATGGTGGTAACA; ATF6:(+) CACAGCTCCCTAATCACGTGG; (-) ACTGGGCTATTCGCTGAAGG; ATF6:(+) CAGCCATCAGCCACAACAAG; (-) GGCATCACCAGGGACATCTT; and glyceraldehyde-3-phosphate dehydrogenase: (+) GCCACATCGCTCAGACACC; (-) CAAATCCGTTGACTCCGACC.
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Electrophoretic Mobility Shift Assays
Electrophoretic mobility shift assays (EMSAs) were carried out essentially as previously described (25). Double-stranded synthetic oligonucleotides were used as 32P-labeled probes or competitors, as described in the figure legends. The GRP78 ERSE-1 sequence was: CCGGGAGGGCCTTCACCAATCGGCGGCCTCCACGACGGGGCTGGC (underlined nucleotides are a consensus ERSE), and the GRP78 ERSE1 MM sequence was CCGGGAGGGCCTTCAgactaCGGCGGCCTgatgtACGGGGCTGGC (mutated nucleotides indicated by lowercase). Nuclear extracts, which provide the source of other proteins (e.g. NF-Y, YY1, and TFII-I) needed for ATF6 binding to ERSEs, were prepared as previously described (25). Binding reactions were carried out in a solution composed of 20 mM HEPES (pH 7.9), 10% glycerol, 1 mM MgCl2, 1 mM 2-mercaptoethanol, 0.1% Tween 20, 0.2 µg of poly(dI-dC), 6 µg of HeLa nuclear extract, 2 µl of in vitro translated ATF6
-(1-373), ATF6-(115-373), or ATF6-(1-392), prepared as previously described (10), and 10,000 cpm of 32P-labeled probe. Reactions were incubated at room temperature for 20 min, and then fractionated on a 5% polyacrylamide gel at 200 V for 150 min in 0.5x TBE buffer (45 mM Tris borate, 1 mM EDTA). For supershift EMSAs, 1 µl of anti-ATF6 (Santa Cruz Biotechnology, sc-22799), 1 µl of anti-ATF6 (Santa Cruz Biotechnology, sc-30596), or 1 µl of non-immune mouse antisera was added 15 min prior to probe addition. For oligonucleotide specificity assessment, 250-fold excess of unlabeled double-stranded oligonucleotide was added 15 min prior to addition of probe. |
RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Isoform-specific Characteristics of N-ATF6
and - Are Conferred by Their Divergent N-terminal TADs—Because we previously showed the importance of the VN8 sequence for the transcriptional activity and rapid degradation of ATF6 (22), we assessed whether the lack of a VN8 sequence in ATF6 is responsible for its low transcriptional activity and high stability. Upon sequence alignment of N-ATF6 and -, we found that residues 64-71 of correspond to residues 61-68 of , the VN8-like region (Fig. 1B). Accordingly, residues 64-67 of N-ATF6 (i.e. ATF6-(1-392)) were mutated to the same residues found in the VN8-like region of ATF6, which possess the Phe and Leu known to be required for optimal activity (Fig. 2A, construct 3, ATF6-VN8-M1). We also prepared a mutation that converted the entire 64-71 region N-ATF6 to be identical to the VN8 in ATF6 (Fig. 2A, construct 4, ATF6-VN8-M2). The abilities of native N-ATF6 or the VN8 mutations to activate the promoter of the prototypical ERSR gene, GRP78, in HeLa cells were compared with N-ATF6 (i.e. ATF6-(1-373)). As previously seen (22), N-ATF6 exerted strong GRP78 promoter activation, whereas native N-ATF6 exhibited weak effects (Fig. 2B, constructs 1 versus 2). Among the ATF6 VN8 mutations, only ATF6-VN8-M2 exhibited detectible GRP78 promoter activation, although it amounted to only 8% of N-ATF6 (Fig. 2B, construct 4). Previous studies showed that the relative levels of ectopically expressed ATF6 and - are proportional to their half-lives (22). N-ATF6 was expressed in very low quantities (Fig. 2C, construct 1), consistent with its known short half-life, whereas all forms of N-ATF6 were expressed at much higher levels (Fig. 2C, constructs 2-4), suggesting that they exhibited relatively long half-lives, as previously shown for native N-ATF6 (22). Quantification demonstrated that N-ATF6, N-ATF6-VN8-M1, and N-ATF6-VN8-M2 were expressed at 40-, 29-, and 20-fold higher levels than N-ATF6, respectively (Fig. 2C). Thus, although their expression levels and apparent half-lives decreased in coordination with the minor increases in activity, it was apparent that the low transcriptional activity and high stability of N-ATF6 were not due entirely to the lack of a consensus VN8 sequence, but that larger portions of ATF6 must be required to confer these isoform-specific characteristics.
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To examine the effects of mutating larger portions of ATF6
and -, a series of domain-swap mutations were generated where the N terminus of N-ATF6 was replaced with progressively larger portions of the corresponding sequences from N-ATF6 (Fig. 3A). As expected, native N-ATF6 was a strong activator of the GRP78 promoter, whereas native N-ATF6 was much weaker (Fig. 3B, constructs 1 versus 2). However, when the N-terminal 115 or 190 aa of N-ATF6 were replaced with corresponding sequences from , transcriptional activity increased progressively (Fig. 3B, constructs 3 and 4). Finally, when the N-terminal 321 aa of N-ATF6, representing all but the 71-aa b-Zip domain, were replaced by corresponding sequences from N-ATF6, GRP78 promoter activity increased to about the same level as that observed using native N-ATF6 (Fig. 3B, construct 5), suggesting that the b-Zip domains of N-ATF6 and - were interchangeable. As expected, the level of expression of N-ATF6 was 80-fold greater than N-ATF6 (Fig. 3C, constructs 1 and 2); moreover, expression levels of the chimeras declined coordinately as more sequences from N-ATF6 replaced corresponding sequences in (Fig. 3C, constructs 3-5), consistent with the hypothesis that the degradation rate of ATF6 coordinates with its transcriptional activity. When GRP78 promoter activity was normalized to the levels of ectopic N-ATF6 or - protein expression, the only domain-swap mutant exhibiting activity approximating that of native N-ATF6 was construct 5 (Fig. 3D). Accordingly, these data suggested that, although the b-Zip domain of ATF6 can substitute for the b-Zip domain of ATF6 without much loss of function, most of the sequences lying to the N terminus of the b-Zip domain of ATF6 are necessary to confer the full transcriptional activity and rapid degradation characteristic of this ATF6 isoform.
N-ATF6-mediated Transcriptional Activation and Rapid Degradation Are Coordinated Processes—It is not known whether it is the sequences in the TAD of ATF6 that confer strong transcriptional activation and rapid degradation, or whether rapid degradation is a function of the engagement of ATF6 in a productive transcription complex. If the latter is true, then mutating the basic region of the b-Zip domain to disrupt binding of N-ATF6 to ERSEs should decrease transcriptional activation and decrease degradation. Consistent with this hypothesis was our finding that mutating the basic region of N-ATF6 (Fig. 4A, construct 2) to disrupt the binding of N-ATF6 to ERSEs (9) resulted in decreased GRP78 promoter activation (Fig. 4B, constructs 1 and 2) and increased N-ATF6 expression of >3-fold (Fig. 4C, FLAG blot, constructs 1 and 2). To test this hypothesis in a heterologous gene expression system, we used a truncated form of the yeast transcription factor Gal4, Gal4-(1-147), composed of the Gal4 DBD, which does not possess a TAD. The binding of the Gal4 DBD to appropriate DNA sequences was assessed using a luciferase reporter driven by a neutral promoter flanked by tandem repeats of the Gal4 binding element. A mutation known to block the binding of Gal4-(1-147) to the Gal4 binding element (26) was introduced into a construct featuring the TAD of ATF6 without the ATF6 DBD, i.e. ATF6-(1-114), fused to the Gal4 DBD (Fig. 4A, constructs 5 and 6). As expected, the ATF6(1-114)/Gal4 DBD fusion protein without the DBD mutation exhibited robust transcriptional activation, compared with the Gal4 DBD alone (Fig. 4B, constructs 3 and 5); however, the ATF6(1-114)/Gal4 DBD fusion protein harboring the DBD mutation exhibited no transcriptional activation (Fig. 4B, construct 6). Moreover, the level of expression of ATF6/Gal4 DBD-M was 2-fold greater than that of ATF6/Gal4 DBD (Fig. 4C, Gal4 blot, constructs 5 and 6), whereas the level of expression of Gal4 DBD-M was actually somewhat lower than that of Gal4 DBD (Fig. 4C, Gal4 blot, constructs 3 and 4). These results are consistent with the hypothesis that rapid degradation of N-ATF6 requires its engagement in transcriptional activation.
To examine the relationship between transcriptional engagement and ATF6 degradation in a different model system, we designed a method for conditionally activating N-ATF6 in a ligand-dependent manner. For this purpose, we generated a construct encoding N-ATF6 fused to a fragment of the MER, which has no TAD or DBD, but features a tamoxifen-ligand-binding domain replacing the estrogen-binding domain. By analogy to the way MER affects other proteins to which is has been fused (27), we reasoned that, in the absence of tamoxifen, the MER would attract other cellular components, e.g. HSP90, which would block functional domains of ATF6, but that, upon tamoxifen binding, release of HSP90, among others, would reveal functional domains and allow full engagement of ATF6 in transcription (Fig. 5A). Accordingly, constructs encoding FLAG-MER or FLAG-N-ATF6-MER, where MER is fused to the C terminus of FLAG-N-ATF6, were prepared (Fig. 5B), and the abilities of each to activate the GRP78 promoter were examined. As expected, FLAG-MER exhibited essentially no activity (Fig. 5C, construct 1), whereas FLAG-N-ATF6 exhibited high activity that was affected very little by tamoxifen (Fig. 5C, construct 3). In contrast, FLAG-N-ATF6-MER exhibited little activity in the absence of tamoxifen, but, upon tamoxifen addition, activity increased 3-fold, nearly equal to that of FLAG-N-ATF6 (Fig. 5C, construct 2). The protein levels of FLAG-MER were relatively high in the absence of tamoxifen (Fig. 5D, lanes 1 and 2), and actually increased by 1.6-fold in the presence of tamoxifen (Fig. 5D, lanes 3 and 4), which was somewhat expected, because tamoxifen stabilizes MER. The levels of FLAG-N-ATF6 were low and unchanged by tamoxifen (Fig. 5D, lanes 9-12). However, the levels of FLAG-N-ATF6-MER were decreased by 5-fold in tamoxifen-treated cells (Fig. 5D, lanes 5-8), suggesting coordination of tamoxifen-activated transcription and rapid degradation of FLAG-ATF6-MER.
Relative Levels of N-ATF6 and - Impact ERSR Gene Induction and Cell Viability in Ways Consistent with Roles of N-ATF6 as a Transcriptional Repressor of N-ATF6—The ATF6 loss-of-function mutations in this and previous studies exhibit decreases in degradation. We previously showed that N-ATF6 mimicked ATF6 loss-of-function mutations in terms of inhibiting N-ATF6-mediated transcription (22), but its effect on ATF6 expression level and degradation is not known. Accordingly, a construct encoding FLAG-N-ATF6 was used to distinguish it from HA-N-ATF6 on immunoblots, and the ratios of ectopically expressed FLAG-N-ATF6 and HA-N-ATF6 were varied by transfecting HeLa cells with different amounts of the appropriate plasmids.
In the first series of experiments, the level of FLAG-N-ATF6-encoding plasmid was held constant, while the level of the HA-N-ATF6-encoding plasmid was varied. As expected, GRP78 promoter activation by FLAG-N-ATF6 was inhibited as the level of HA-N-ATF6 was increased (Fig. 6A, transfections 1-3). FLAG and HA immunoblots showed that the quantity of HA-N-ATF6 increased as a function of increased plasmid, as expected (Fig. 6B, HA-ATF6); interestingly, the levels of FLAG-N-ATF6 also increased, even though each culture had been transfected with the same quantity of the FLAG-N-ATF6 plasmid (Fig. 6B, FLAG-ATF6). These results suggested that HA-N-ATF6 not only inhibited the ability of FLAG-N-ATF6 to activate the GRP78 promoter, but also increased its half-life. We examined degradation of FLAG-N-ATF6 using cycloheximide (CHX) to inhibit new protein synthesis, as previously described (28). The apparent degradation of FLAG-N-ATF6 was extremely rapid when no HA-N-ATF6 was co-expressed. Within 9 min of CHX addition, only 12% of the FLAG-N-ATF6 originally present remained (Fig. 6C, transfection 1, Blot A, versus transfection 1, Blot B). In contrast, the degradation rate of FLAG-N-ATF6 was reduced in the presence of HA-N-ATF6; moreover, as the level of HA-N-ATF6 was increased, degradation rate of FLAG-N-ATF6 decreased. For example, at intermediate or high levels of HA-FLAG-N-ATF6, 66 and 82% of the original FLAG-N-ATF6 remained after 9 min of CHX treatment (Fig. 6C, transfections 2 and 3, Blot A versus Blot B).
In the second series of experiments, the FLAG-N-ATF6-encoding plasmid was varied, while the HA-N-ATF6-encoding plasmid was held constant. As expected, HA-N-ATF6 alone conferred very little GRP78 promoter activation (Fig. 6D, transfection 4), whereas, increasing the levels of FLAG-N-ATF6 increased GRP78 promoter activity (Fig. 6D, transfections 5 and 6). FLAG and HA immunoblots showed that the level of FLAG-N-ATF6 increased as more plasmid was transfected, as expected (Fig. 6E, FLAG-ATF6, transfections 4-6); however, surprisingly, the levels of HA-N-ATF6 decreased, even though each culture had been transfected with the same quantity of that plasmid (Fig. 6E, HA-ATF6, transfections 5 and 6). These results suggested that FLAG-N-ATF6 can increase the degradation rate of HA-N-ATF6. Consistent with this hypothesis was the finding that, in the absence of FLAG-N-ATF6, 58% of the original HA-N-ATF6 was still present following 17 min of CHX treatment (Fig. 6F, transfection 4, Blot A versus B). In contrast, the degradation rate of HA-N-ATF6 was increased in the presence of FLAG-N-ATF6; moreover, as FLAG-N-ATF6 was increased, the degradation rate of HA-N-ATF6 increased. For example, at intermediate and high levels of FLAG-N-ATF6, 32 and 26% of the original HA-N-ATF6 was still present 17 min after CHX treatment (Fig. 6F, transfections 5 and 6, Blot A versus B). Taken together, the results of the experiments shown in Fig. 6 indicate that ATF6 and - can influence each other, such that the isoform-specific transcriptional and degradation characteristics of each are dependent upon their relative levels. This finding is consistent with a mechanism whereby N-ATF6 and - can regulate ERSR gene expression and cellular function in a combinatorial fashion.
Because ATF6 and - can both bind to ERSEs (20), EMSAs were performed to assess the abilities of recombinant ATF6 and - to compete for binding to an ERSE in the GRP78 gene. Incubation of nuclear extract from untreated HeLa cells with a labeled oligonucleotide that replicates ERSE-1 in the GRP78 gene resulted in the formation of complex 1 (Fig. 7A, lane 1). Formation of complex 1 has previously been shown to be due to binding of other nuclear proteins (e.g. NF-Y A, B, and C) to the ERSE in the absence of ATF6 (25). Adding recombinant ATF6-(1-373) or ATF6-(1-392) to the nuclear extract resulted in the formation of complexes 3 and 4, respectively, which migrated with relative mobilities consistent with the sizes of each form of ATF6 that was added (Fig. 7A, lanes 2 and 3). Adding a shortened form of ATF6, ATF6-(115-373), which should retain ERSE-binding ability, also exhibited a complex, complex 2, the mobility of which was consistent with the size of ATF6-(115-373) relative to the other forms of ATF6 used in this analysis (Fig. 7A, lane 4). When an excess of unlabeled wild-type GRP78 ERSE-1 oligonucleotide was added to the incubation, all of the complexes disappeared (Fig. 7A, lanes 5-8), as expected. However, excess unlabeled mutated GRP78 ERSE-1 was unable to compete for labeled oligonucleotide binding (Fig. 7A, lanes 9-12). These results demonstrate the dependence of each complex on the presence of the native GRP78 ERSE-1.
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To verify the presence of ATF6 isoforms in complexes 2-4, supershift EMSA experiments were carried out. Addition of preimmune antiserum to the nuclear extract did not alter the EMSA profile (Fig. 7B, lanes 1-4). However, addition of an antiserum specific for ATF6
altered the mobility of complexes 2 and 3, only (Fig. 7B, compare lanes 6 and 8 to lanes 2 and 4, respectively), whereas addition of an antiserum specific for ATF6 altered the mobility of complex 4, only (Fig. 7B, compare lane 11 with lane 3). These results verify that complex 1 does not contain either form of ATF6, whereas complexes 2-4 contain ATF6-(115-373), ATF6-(1-373), and ATF6-(1-392), respectively.
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Additional EMSAs were performed to determine the effects of ATF6
and ATF6 together on complex formation. Addition of ATF6-(1-373) and ATF6-(1-392) resulted in a decrease in the intensities of complexes 3 and 4 (Fig. 7C, compare lanes 2 and 3 to lane 4). Moreover, adding ATF6-(115-373) and ATF6-(1-392) demonstrated a decrease in the intensities of complexes 2 and 4 (Fig. 7C, compare lanes 7 to lanes 5 and 6). Taken together, the results of the EMSA studies shown in Fig. 7 support the hypothesis that ATF6 and - compete for binding to the canonical ERSE-1 in the GRP78 gene.
To examine the cellular effects of altering the relative levels of endogenous ATF6 and -, we used an siRNA approach that was previously shown by immunoblotting to selectively reduce the quantity of each ATF6 isoform in HeLa cells (22). The selectivity of the siRNA reagents was verified here by quantitative reverse transcription-PCR assessment of ATF6 and - mRNA in extracts from cells treated with siRNA targeted to green fluorescent protein (control), ATF6, ATF6, or another ERSR gene that is not the focus of this study, XBP1. Validating the specificity of the siRNAs was the finding that the ATF6-targeted siRNA reagent decreased the level of ATF6 and not ATF6 or XBP1 mRNA (Fig. 8A), whereas the ATF6-targeted siRNA decreased the level of ATF6 and not ATF6 or XBP1 mRNA (Fig. 8B). Knocking down ATF6 decreased basal and tunicamycin (TM)-stimulated GRP78 promoter activity by 2- to 3-fold (Fig. 9A, bars 1 versus 2 and 4 versus 5). In contrast, knocking down endogenous ATF6 had little effect on basal GRP78 promoter activity, but it increased TM-induced GRP78 luciferase by 2-fold (Fig. 9A, bars 4 versus 6). Coordinate with these results were the findings that knockdown of ATF6 decreased TM-induced GRP78 mRNA by 2-fold (Fig. 9B, bar 4 versus 5), whereas knockdown of ATF6 increased TM-induced GRP78 mRNA by 1.5-fold (Fig. 9B, bar 6). Because many ERSR genes, including GRP78, encode proteins that foster protection, we examined the effects of knocking down endogenous ATF6 or - on HeLa cell viability. We found that, although 32 h of TM treatment conferred no change in viability in cells treated with control siRNA (Fig. 9C; bars 1 versus 4), knockdown of ATF6 significantly decreased viability with or without TM (Fig. 9C; bars 1 versus 2, and 4 versus 5), although knockdown of ATF6 significantly increased viability with or without TM (Fig. 9C; bars 1 versus 3 and 4 versus 6). These findings indicate that the isoform-specific characteristics of ATF6 and - can influence TM-stimulated GRP78 expression, as well as viability of HeLa cells in ways that are consistent with the protective aspects of N-ATF6 and the putative abilities of N-ATF6 to serve as an endogenous repressor of ATF6. Moreover, because knocking down ATF6 or altered viability-TM, it is apparent that even in the absence of TM-mediated ER stress, ERSR genes, such as GRP78, must contribute to cell viability.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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In this study we examined the structural features underlying the isoform-specific characteristics of ATF6
and -, the coordination and mechanism of ATF6 transcriptional activation and rapid degradation, and whether the relative levels of ATF6 and - affect their binding to ERSEs and regulate ERSR gene induction and cell viability. Our findings showed that there is structural information spanning most of the N-terminal 300 aa of ATF6 and - that is required for isoform-specific characteristics. We also found that the rapid degradation of ATF6 is coordinate with its engagement in an active transcription complex, the latter of which can evidently be modulated by ATF6. Lastly, we determined the ratio of ATF6 and - that modulates ERSR gene induction, as well as cell viability, in a manner consistent with the hypothesis that ATF6 is a strong but labile transcriptional activator, whereas ATF6 is a weak, stable transcriptional activator.
In addition to transcriptional activity and the rate of degradation, the timing of ATF6 and - activation following ER stress is likely to be another important, albeit, not thoroughly studied isoform-specific characteristic. Although it is well known that both ATF6 isoforms are cleaved upon ER stress, to our knowledge, only one study showed that, depending on the stress, activation of ATF6 can occur earlier than that of ATF6 (19). Combined with their isoform-specific characteristics, sequential activation of the ATF6 isoforms (Fig. 10A) is consistent with the possibility that their relative levels could change as a function of time after ER stress, such that there is an initial, strong activation of ATF6-mediated ERSR gene induction, followed by modulation toward weak activation (Fig. 10B). One potential mechanism by which ATF6 and - could regulate the strength of ER stress involves how these isoforms bind to ERSR genes. ATF6 and - bind to ERSEs, and possibly other elements, as dimers, which, interact with the C subunit of the NF-Y A, B, and C trimer (19), as well as with other proteins, e.g. SRF (5), TFII-I (12), and perhaps YY1 (13). Together, these proteins evidently facilitate ERSR gene induction. Thus, it is conceivable that, as a result of isoform-specific rates of generation and degradation, the relative levels of ATF6 and - in transcriptional complexes change during progression of the ERSR and that, as a result of differences in their transcriptional activities, ERSR gene induction is finely tuned, as shown in Fig. 10C. The results of the gel shift experiments in this study showed that ATF6 and - can compete with each other for binding to the GRP78 ERSE (Fig. 7C), which lends further support to this hypothesis.
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The mechanism governing the rate of degradation of ATF6 during transcription is not known. However, a great deal is known about the coupling of transcriptional activation and rapid degradation of other labile transcription factors, e.g. c-Myc, Gal4, VP16, SMAD2, STAT, and Hac1p (29). In those cases, the most active transcription factors are also very susceptible to ubiquitination and proteasome-mediated degradation, both of which evidently take place during engagement in transcription (15, 30). This "unstable when active" phenomenon is thought to allow tight control over transcription by ensuring that the activation of target genes is linked to the ongoing synthesis of the transcriptional regulator (30). Accordingly, such transcription factors are usually potent target gene activators, which must effect their function in a transient manner, although, rapid degradation of transcription factors has been linked to modulation as well as augmentation of activity (31, 32). Because ubiquitinligase/proteasome machinery exists in the nucleus and is intimately associated with active transcription complexes, it has been proposed that proteasome-mediated degradation of certain transcription factors occurs as they become engaged in transcriptional activation and may actually require polymerase II. The polymerase II requirement might contribute to distinguishing the half-lives of N-ATF6
and -; presumably, because N-ATF6 is a poor transcriptional activator, it does not engage in a transcriptional complex that efficiently recruits active polymerase II. It was recently shown in yeast that the ER stress-activated transcription factor, Hac1p, which is homologous to another mammalian ER stress-activated transcription factor, XBP1, is degraded rapidly upon ER stress; moreover, Hac1p degradation requires nuclear localization and was impaired by Hac1p mutations that forced its nuclear exclusion (33). These findings suggest that, like ATF6, transcriptional induction by Hac1p in yeast, and perhaps by XBP1 in mammalian cells, is engineered to be rapid and transient. The sequence responsible for Hac1p rapid degradation was localized using PESTFind (34), to a PEST motif, i.e. a stretch of the protein that is enriched in proline, glutamine, serine and threonine. This motif has been found in numerous other rapidly degraded proteins that are degraded in a conditional manner (35). Using PESTFind, we identified a potential PEST sequence in N-ATF6 that exhibits a similar PESTFind score as that found in Hac1p, and resides in a region, suggested by the domain-swap mutations carried out in this study, to be critical for transcriptional induction and rapid degradation of ATF6. Thus, it will be of interest to examine whether this potential PEST sequence contributes to the rapid degradation of N-ATF6 upon transcriptional engagement.
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ERSR gene expression by ATF6 is apparently regulated, in part, by the isoform-specific characteristics of ATF6
and -. The results presented in this study suggest that it is possible that these isoform-specific characteristics contribute to ATF6-mediated gene induction in subtle ways that fine-tune this aspect of the ERSR. Future studies examining the impact of ATF6 and - in various cells and tissues subjected to ER stress will be required to fully appreciate the roles of these ATF6 isoforms in this complex process. |
FOOTNOTES |
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* This work was supported in part by National Institutes of Health Grants HL75573 and NS025037 (to C. C. G.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 Funded by a San Diego State University Heart Institute/Rees-Sealy Research Foundation graduate fellowship, and a graduate fellowship from the San Diego Chapter of the Achievment Rewards for College Scientists Foundation.
2 To whom correspondence should be addressed: Dept. of Biology, San Diego State University, 5300 Campanile Drive, San Diego CA 92182. Tel.: 619-594-2959; Fax: 619-594-5676; E-mail: cglembotski{at}sciences.sdsu.edu.
3 The abbreviations used are: ER, endoplasmic reticulum; ERSR, ER stress response; aa, amino acid(s); N-ATF6, N-terminal ATF6; TAD, transcriptional activation domain; b-Zip, basic leucine zipper; GRP78, glucose-regulated protein 78; MER, mutant estrogen receptor; ANOVA, analysis of variance; CMV, cytomegalovirus; DBD, DNA-binding domain; siRNA, small interfering RNA; EMSA, electrophoretic mobility shift assay; ERSE, ERSR element; CHX, cycloheximide; TM, tunicamycin; STAT, signal transducers and activators of transcription; XBP1, X-box-binding protein 1.
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