JVI Accepts, published online ahead of print on 20 February 2008 J. Virol. doi:10.1128/JVI.02054-07 Copyright 2008, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.
XBP-1, a novel HTLV-1 Tax binding protein, activates HTLV-1 basal and Tax-activated transcription
Sebastian C. Y. Ku1, Jialing Lee1, Joanne Lau1, Meera Gurumurthy1, Raymond Ng1, Siew Hui Lwa1, Joseph Lee1, Zachary Klase2, Fatah Kashanchi2, and Sheng-Hao Chao1*
Expression Engineering Group, Bioprocessing Technology Institute, 20 Biopolis Way, #06-01 Centros, Singapore 138668, Singapore
Department of Biochemistry and Molecular Biology, the George Washington University School of Medicine, Washington DC 20037, USA.
Email: jimmy_chao@bti.a-star.edu.sg

Fax: (65) 6478-9561

Running title: Activation of HTLV-1 LTR by XBP-1
To whom correspondence should be addressed:

T P E C

Word count for abstract: 195. Word count for text: 5223.
Abstract X-box binding protein 1 (XBP-1), a basic leucine zipper transcription factor, plays a key role in cellular unfolded protein response (UPR). There are two XBP-1 isoforms in cells, the spliced XBP-1S and the unspliced XBP-1U. XBP-1U has been shown to bind to the 21-bp Tax-responsive element 1 repeat of human T-lymphotropic virus type 1 (HTLV-1) long terminal repeat (LTR) in vitro and transactivate HTLV-1 transcription. Here we identify XBP-1S as a transcription activator of HTLV-1.
Compared to XBP-1U, XBP-1S demonstrates stronger activating effects on both basal
and Tax-activated HTLV-1 transcription in cells. Our results show that both XBP-1S and XBP-1U interact with Tax and bind to HTLV-1 LTR in vivo. In addition, elevated mRNA levels of XBP-1 and several UPR genes are detected in the HTLV-1-infected T cell lines, C10/MJ and MT2 cell lines, suggesting that HTLV-1 infection may trigger UPR in the host cells. We also identify Tax as a positive regulator of XBP-1 gene expression. Activation of UPR by tunicamycin shows no effect on HTLV-1 LTR, suggesting that HTLV-1 transcription is specifically regulated by XBP-1. Collectively, our study demonstrates a novel host-viral interaction between a cellular factor XBP-1 and transcriptional regulation of HTLV-1.
Introduction Human T-lymphotropic virus type 1 (HTLV-1) is the causative agent of adult Tcell leukemia/lymphoma and the neurological disorder, HTLV-1-associated myelopathy/tropical spastic paraparesis (14,33,34,49). The HTLV-1 transactivator, Tax, activates viral transcription through three 21-bp repeats, which are known as Tax-
responsive element (TRE), located within the HTLV-1 long terminal repeat (LTR) (5,20). Each 21-bp TRE repeat contains a cyclic AMP response element (CRE) recognized by
members of the CRE binding protein/activating transcription factor (CREB/ATF) family of proteins. All CREB/ATF proteins contain a basic-region leucine zipper (bZIP)
domain, which is involved in DNA binding and gene regulation (15). Tax does not bind the TRE repeats directly but interacts with CREB (or other CREB/ATF members) to form a protein complex that associates with the DNA (1,3,12). The Tax-CREB complex serves as a binding site for the recruitment of cellular transcriptional co-activators, including CREB binding protein (CBP), p300, and p300/CBP-associated factor (PCAF), resulting in the activation of viral transcription (16,22,26). CREB1 (previously known as CREB), CREB2 (also known as TREB7), ATF-1 (also known as TREB36), and ATF-2 have been identified as the cellular Tax-binding proteins, suggesting that these CREB/ATF play a key role in the transcriptional regulation of HTLV-1 (12,35,45,53). X-box binding protein 1 (XBP-1) is a bZIP protein belonging to the CREB/ATF family of transcription factors. XBP-1 plays a major role in the cellular unfolded protein response (UPR), which is triggered by accumulation of unfolded or malfolded proteins in the endoplasmic reticulum (ER) (6). There are two protein isoforms of XBP-1, XBP-1U and XBP-1S. XBP-1U (also known as TREB5), which consists of 261 amino acids (a.a.),
is translated from the unspliced mRNA of XBP-1 and is the dominant isoform under nonstress conditions. It has been reported that XBP-1U is transcriptionally inactive (46). Activation of UPR induces the endoribonuclease activity of inositol requiring enzyme 1, an ER transmembrane protein, resulting in the excision of 26 bases (between the nucleotides 531 and 556 of XBP-1 mRNA) from the XBP-1 transcript. Splicing of the 26 nucleotides leads to a frame shift at a.a. 165 and the generation of a longer and

expression vector by deleting the 26 nucleotides between the positions 531 and 556 of
human XBP-1 mRNA (46). The coding regions of XBP-1 (1-74), XBP-1U (134-261),
and XBP-1S (134-378) (which contain XBP-1 a.a. 1-74, XBP-1U a.a. 134-261, and XBP1S a.a. 134-378, respectively) were amplified by PCR using wild-type XBP-1U or XBP1S vectors as the templates. The amplified fragments were then subcloned into pcDNA6 plasmid (Invitrogen) to generate the XBP-1 mutant vectors. Tax expression vectors containing wild-type Tax (pcTax), the M22 mutation, and the M47 mutation were generous gifts from Dr. Warner C. Greene (37,39). The coding region of HTLV-1 Tax was amplified by PCR using pcTax plasmid as the template and subcloned into the cytomegalovirus (CMV)-enhanced green fluorescent protein (EGFP) vector (kindly provided by Dr. Zhiwei Song, Bioprocessing Technology Institute, Singapore) to generate CMV-EGFP-Tax. The expression of EGFP-Tax was under the control of human CMV major immediate early (MIE) promoter. HTLV-1-LUR-GL3 firefly luciferase (HTLV-1-LTR-F-Luc) was a kind gift from Dr. Arnold Rabson (29). The vectors BiP-Luc and ATF6(1-373) were provided by Dr. Kazutoshi Mori (47). The firefly luciferase plasmids, CMV-Luc, MLV-LTR-Luc, and HIV-LTR-Luc, in which the expression of luciferase was driven by HCMV major immediate-early (MIE) promoter,
Moloney murine leukemia virus (MLV) LTR, and human immunodeficiency (HIV) LTR, respectively, were previously described (8,9,32). To generate the HTLV-1-LTR-F-Luc stable clones, we first constructed the HTLV-1-LTR-GL2 firefly luciferase vector [HTLV-1-LTR-F-Luc (GL2)], which contains the selective marker blasticidin for mammalian stable cells. The CMV MIE
promoter in the CMV-F-Luc plasmid (8) was replaced with HTLV-1 LTR to generate
HTLV-1-LTR-F-Luc (GL2). HeLa cells were then transfected with the HTLV-1-LTR-FLuc (GL2) plasmid using Fugene 6 (Roche) as described in the manufacturers manual.
Selection was performed using 12 g/ml blasticidin (Invitrogen) to generate a stable pool of HTLV-1-LTR-F-Luc cells. Single clones displaying XBP-1S- and Tax-inducible expression of luciferase were selected.
adherent cells, HEK293, 293T, HeLa, and HTLV-1-LTR-F-Luc stable cells, were performed using Fugene 6 (Roche). To perform the luciferase-based assays, the cells were grown to 50-80% confluence in 96-well plates. Transfection of Jurkat cells were carried out using Amaxa electroporation system (Amaxa). Cells were co-transfected with an indicated expression plasmid, a firefly luciferase reporter, and a Renilla luciferase plasmid, pRL-RSV (Promega). pRL-RSV was used to normalize transfection efficiency. Firefly and Renilla luciferase activities were measured 48 hours post-transfection using the Dual-Glo assay system (Promega) and the activities were determined using an Infinite 200 multiplate reader (Tecan). Chromatin immunoprecipitation (ChIP). ChIP assay was performed as described previously (28). The HTLV-1-LTR-F-Luc stable cells were transfected with

Transient transfection and luciferase assays. Transient transfections of the
the XBP-1U or XBP-1S expression vectors. The chromatins (which were sonicated to the DNA sizes between 300 and 1000 bp) of the transfected HTLV-1-LTR-F-Luc stable cells were isolated 48 hours post-transfection and were used for ChIP analyses with normal rabbit IgG (Upstate Biotechnology), anti-CREB1 antibody (Upstate Biotechnology), or anti-XBP-1 antibody (Santa Cruz Biotechnology).
Immunoprecipitated DNA was analyzed by PCR by using specific primers to the HTLV1 LTR (5-AAGGTCAGGGCCCAGACTAAG-3 and 5-
GAGGTGAGGGGTTGTCGTCAA -3) and the luciferase coding region (5-
GTTACAACACCCCAACATCTT-3 and 5-ATTTGGACTTTCCGCCCTTCT-3).
Co-immunoprecipitation (Co-IP) and western blotting. HEK293 cells were co-transfected with XBP-1S/Tax or XBP-1U/Tax expression plasmids. Lysates prepared from the transfected cells were utilized for Co-IP. Co-IP was performed using an immunoprecipitation kit according to manufacturers manual (Roche) and the anti-XBP-1 and -Tax antibodies (52). SDS-PAGE and western blotting were carried according to the standard protocols.
circular cover slips in 6-well plates, one day before transfection and transfected with the indicated expression vectors (i.e. EGFP-Tax, XBP-1S, or XBP-1U). The cells were fixed with 4% paraformaldehyde and permeabilized using 0.1% triton X-100 in PBS 48 hours post-transfection. The samples were then incubated with anti-XBP-1 antibody. After washing with PBT (PBS + 0.1% Tween 20), the cells were incubated with the anti-rabbit Alexa Fluor 594 antibody (Invitrogen) for 1 hour. The samples were finally mounted with 4,6-diamidino-2-phenylindole (DAPI) mixed ProLong Gold Antifade Reagent
Immunofluorescence and fluorescent microscopy. HeLa cells were seeded on

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(Invitrogen). Fluorescent microscopy was carried out using Zeiss Axio Imager.Z1 microscope. Quantitative RT PCR. RNAs of the T cell lines and the transfected cells were isolated using TRIZOL Reagent (Invitrogen). Improm II Reverse Transcription System (Promega) and SYBR Green PCR Master Mix (Applied Biosystems) were utilized to carry out reverse transcription and real time PCR, respectively. Amplification and
detection of specific mRNAs were performed using ABI Prism 7000 Thermal-Cycler
(Applied Biosystems). GAPDH was used as an endogenous control. To normalize the
data for RNA loading, threshold cycle (CT) value of GAPDH was subtracted from that of each gene in the respective samples.
Results XBP-1S activates basal and Tax-dependent HTLV-1 transcription. It was reported more than 11 years ago that XBP-1U bound to the 21-bp repeats in the HTLV-1 LTR and transactivated the viral transcription (10,30,50). Since the discovery of XBP-1S in 2001, most studies demonstrated that XBP-1S is a transcription activator while XBP-

1U is inactive (46). However, to date, the effect of XBP-1S on HTLV-1 transcription is still unknown.
We first performed cell-based reporter assays to investigate the effect of XBP-1S on HTLV-1 LTR. To determine the influence of XBP-1S specifically, we generated a
XBP-1S expression vector by deleting the 26 nucleotides located between the positions 531 and 556 of human XBP-1 mRNA. The 26-bp deletion results in a shift in the open reading frame of XBP-1 mRNA, generating only the XBP-1S isoform (46). HEK293, 293T, and HeLa cells were transiently transfected with HTLV-1-LTR-F-Luc and an indicated XBP-1 expression vector (i.e. XBP-1S or XBP-1U). Human ATF6(1-373), a cleaved form of ATF6 (containing amino acids 1-373 of ATF6) and a known transcription activator for ER chaperon genes and the XBP-1 gene, was used as a control (47). Both XBP-1S and ATF6(1-373) have been shown to activate the transcription of an ER chaperon, BiP (47). 5- to 35-fold increases in HTLV-1-LTR-dependent transcription were detected when XBP-1S was overexpressed in HEK293, 293T, and HeLa cells (Fig. 1A-C). Overexpression of XBP-1U resulted in a significantly lower level of activation in HTLV-1 transcription in HEK293 and 293T cells, and had no effect in HeLa cells (Fig. 1A-C). In all cell types, ATF6(1-373) did not cause any detectable changes in HTLV-1 LTR-driven expression (Fig. 1A-C). To further confirm the
activating effect of XBP-1S on HTLV-1 LTR, a similar set of cell-based assays were carried out by titrating the amounts of XBP-1S expression plasmids. A positive correlation between the fold change of HTLV-1 LTR activation and the dosage of XBP1S was observed in HEK293, 293T, and HeLa cells (Fig. 1D-F). Notably, a greater than 100-fold increase in HTLV-1 LTR transactivation was detected in 293T cells (Fig. 1E). We next examined the effects of XBP-1 on other viral promoters, including
HCMV MIE promoter, Moloney MLV LTR, and HIV LTR. While XBP-1S and XBP1U stimulated 7.4- and 2.0-fold increase on HTLV-1 LTR-dependent expression,
respectively, little or no effects were detected on other viral promoters (Fig. 2). These results demonstrated the specific activation of HTLV-1 LTR by XBP-1.
Our results showed that XBP-1S and ATF6(1-373) exhibited different effects on the HTLV-1 LTR (Fig. 1). Since both XBP-1S and ATF6(1-373) are known to be key regulators in UPR, we thus further investigated the effect of UPR activation on HTLV-1 transcription. Tunicamycin was utilized to elicit UPR. As UPR is known to stimulate BiP transcription (23,24,46), BiP-Luc, a reporter plasmid in which the expression of firefly luciferase is driven by BiP promoter, was utilized as a control (46). Cells were transiently transfected with BiP-Luc or HTLV-1-LTR-Luc in the presence of tunicamycin. Treatment with tunicamycin significantly stimulated BiP-dependent transcription as expected (Fig. 3). In contrast, no activation on HTLV-1-mediated expression was detected (Fig. 3). Similar results were obtained when several HTLV-1LTR-F-Luc stable cell lines were incubated with tunicamycin (data not shown). The influence of XBP-1S and XBP-1U on HTLV-1 LTR-mediated transcription was examined in a T cell line, the Jurkat cell. Cells were separately co-transfected with

the luciferase coding region (i.e. Luc, Fig. 5), indicating the in vivo association between XBP-1S, XBP-1U, and the HTLV-1 LTR.
transactivated HTLV-1 Tax-dependent transcription (Fig. 4B) and bound to the HTLV-1 LTR region containing the 21-bp repeats in vivo (Fig. 5). It is known that Tax can not bind to HTLV-1 promoter by itself. Tax requires an additional Tax-binding protein which can bind to the TRE repeat and recruit Tax to the HTLV-1 LTR to activate HTLV1 transcription. Therefore, it is possible that XBP-1 may be a Tax interacting partner which facilitates Tax transactivation of HTLV-1 LTR. Co-IP was carried out using the lysates of the cells co-transfected with Tax and one of XBP-1 isoforms. To ensure the expression of XBP-1S and XBP-1U in the transfected cells (Fig. 6, CL input), in vitro synthesized XBP-1S and XBP-1U proteins were used as references (Fig. 6, TNT). IP was carried out using an anti-XBP-1 antibody which recognizes both XBP-1 isoforms, and normal IgG was used as a negative control.
Both XBP-1S and XBP-1U interact with Tax. We have shown that XBP-1

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Tax was present in the immunoprecipitated complexes of both XBP-1S/Tax and XBP1U/Tax co-expressing cells. Stronger interaction between XBP-1S and Tax was observed (Fig. 6, IP XBP-1). Results of reciprocal IP analyses reconfirmed the interaction between XBP-1S/Tax and XBP-1U/Tax. In contrast to the previous observation, the binding affinities between Tax and the two XBP-1 isoforms were similar (Fig. 6, IP
Tax). A small amount of Tax was observed in the negative control for XBP-1S/Tax
expressing cells (Fig. 6, IP IgG), but this is insignificant when compared to the amounts of Tax proteins that were found in the immunoprecipitates using anti-XBP-1 or anti-Tax antibodies (Fig 6, IP XBP-1 and IP Tax, XBP-1S/Tax co-expressing cells).
Nuclear co-localization of XBP-1 and Tax in cells. We further examined subcellular localization of XBP-1S, XBP-1U, and Tax by immunofluorescence. Cells were co-transfected with an EGFP-Tax vector and an indicated XBP-1 plasmid (i.e. XBP-1S or XBP-1U expression vectors). XBP-1U and XBP-1S were immunostained with antiXBP-1 antibody. XBP-1U was evenly distributed within the nucleus and a small portion of XBP-1U was detected in the cytoplasm (Fig. 7). The cytoplasmic localization of XBP1U has been reported (48). XBP-1S was also localized in the nucleus with stronger labeling observed on the nuclear periphery (Fig. 7). Tax showed a predominant nuclear localization but was also detected in the cytoplasm (Fig. 7). The cytoplasmic presence of Tax was detected in all EGFP-Tax transfected cells (data not shown). Cytoplasmic localization of Tax was expected since its localization in ER and Golgi complex was previously reported (2). In the overlaid images, XBP-1S/Tax and XBP-1U/Tax were found to be co-localized with the nuclei (Fig. 7).

The stimulating effect of XBP-1U on the HTLV-1 LTR was reported by two
is possible that XBP-1U requires additional cellular co-factors to regulate the activity of HTLV-1 LTR and the expression of the XBP-1U-interacting factors is cell-specific. However, it is still unknown why XBP-1U can activate HTLV-1 transcription. The mechanism of XBP-1U activation requires further investigation. XBP-1S demonstrates stronger activating effects on both basal and Tax-
dependent HTLV-1 transcription than XBP-1U (Fig. 1 and 4). This is not surprising
since XBP-1S has been shown to have higher transcriptional activity (46). Because of
the UPR-induced splicing, the C-terminus of XBP-1S, but not that of XBP-1U, contains a transactivation domain (46,48). We also compared the activating effect of XBP-1S and XBP-1U to two known positive regulators of HTLV-1 Tax-activated transcription, CREB1 and CREB2. XBP-1S induced a 5-fold increase in the Tax-dependent transcription, while less than 2-fold activation was caused by CREB1 and CREB2 (Fig. 4B). Both CREB1 and CREB2 had little or no effect on basal HTLV-1 transcription, while XBP-1S and XBP-1U resulted in 9- and 5-fold activation of HTLV-1 LTR, respectively (Fig. 4A). Previously, elevated expression of XBP-1 mRNA was detected in the HTLV-1-infected T cell (i.e. HuT 102) (50). Compared with the uninfected T cell (i.e. Jurkat cell), transcriptional levels of CREB1 and CREB2 genes was unchanged or significantly lower in HuT 102 cells (50). In our study, we also detected higher levels of XBP-1 mRNA in two HTLV-1 infected cell lines (Table 1). Collectively, XBP-1 may play a more important role than CREB1 and CREB2 in transcriptional regulation of HTLV-1. Interaction between Tax and XBP-1 proteins was examined by IP experiments. Compared to XBP-1U, more Tax was precipitated with XBP-1S, suggesting a stronger
interaction between XBP-1S and Tax (Fig. 6, IP XBP-1). However, in reciprocal IP assays, the amounts of XBP-1S and XBP-1U co-immunoprecipitated by Tax were similar (Fig. 6, IP Tax). One possible explanation is that the epitope recognized by anti-XBP-1 antibody is not required for Tax-XBP-1S interaction and remains accessible to the antibody when associated with Tax. Therefore, anti-XBP-1 antibody may precipitate most of the Tax-XBP-1S complexes. However, the epitope recognized by anti-Tax
antibody may be involved in the protein-protein interaction between XBP-1S and Tax or
become inaccessible to the antibody when complexing with XBP-1S. Therefore, the antiTax antibody may only pull down a small portion of Tax-XBP-1S complexes.
Quantitative RT-PCR analyses of the HTLV-1 infected cells reveal that HTLV-1 infection may trigger UPR in the host cells (Table 1). We examined the effects of UPR on the HTLV-1 LTR promoter by overexpressing an UPR regulator, ATF6(1-373) or by tunicamycin treatment. Although ATF6(1-373) and tunicamycin up-regulate XBP-1 transcription and/or induce the generation of XBP-1S, neither of them cause any significant changes on HTLV-1 transcription (Fig. 1 and 3). Besides the activation of XBP-1, ATF6(1-373) and tunicamycin have profound effects on UPR-inducible genes and UPR signaling pathways,. It is possible that other signaling pathways or gene expression induced by ATF6(1-373) or tunicamycin may directly or indirectly repress the transactivation of HTLV-1 LTR. Besides the nuclear localization, Tax has been found to be present in the medium of Tax-transfected cells and localize in the organelles associated with protein secretion, including ER and Golgi complex (2). It is possible that Tax may trigger UPR, resulting in up-regulation of XBP-1 expression due to its presence in the ER. Another possibility

is that Tax may activate XBP-1 transcription directly. Several cellular target genes of Tax, such as c-myc and human proliferating cell nuclear antigen, have been reported (11,36). The domains of Tax and XBP-1 required for functional Tax-XBP-1 interaction were investigated. Since XBP-1 belongs to CREB/ATF protein family, it is possible that XBP-1 may utilize the same molecular mechanism in binding to Tax and stimulating HTLV-1 Tax transactivation as other Tax-binding CREB proteins. As expected, the CREB-deficient Tax, M47, failed to activate the XBP-1S-mediated HTLV-1 LTR
transactivation (Fig. 8A). In addition, it has been shown that the bZIP domains of CREB
proteins are essential for CREB-Tax interaction (13,45). Using the bZIP deletion mutants of XBP-1, our results demonstrated the involvement of bZIP domain of XBP-1 in HTLV1 LTR transactivation (Fig. 8B and C). However, XBP-1S (134-378) still could induce a 70% increase in Tax transactivation (Fig. 8C). Study shows that the C-terminus of XBP1S, which is unique to the XBP-1S isoform, contains a transcriptional activation domain (46). It is possible that the C-terminus of XBP-1S may induce a specific set of cellular genes which directly or indirectly regulate HTLV-1 transcription without interacting with Tax.
Although Tax M22 and M47 demonstrated different effects on XBP-1-dependent HTLV-1 Tax transactivation, both mutants and the wild-type Tax showed similar levels of activation on XBP-1 transcription (Fig. 10). The results exclude the requirement of NF-B and CREB proteins in regulating XBP-1 transcription. It is known that Tax can not bind to DNA directly. Therefore, it is clear that Tax requires an unidentified DNAbinding cellular factor to bind to the promoter and up-regulate the expression of XBP-1.

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However, we could not rule out the possibility that Tax may induce XBP-1 transcription indirectly. Based on our study, we propose a model for the host-viral interaction between XBP-1 and HTLV-1 transcription (Fig. 11). Higher expression of XBP-1 mRNA was detected in HTLV-1 infected cells, resulting in the generation of XBP-1S and XBP-1U.
We did not detect a significant increase in XBP-1 splicing in the HTLV-1 infected cells
by RT-PCR (data not shown). It was technically difficult to determine the expression of XBP-1S mRNA quantitatively. However, higher mRNA levels of EDEM (ER

degradation-enhancing alpha-mannosidase-like protein), a XBP-1S target gene (27), were detected in HTLV-1 infected cells using quantitative RT-PCR (data not shown), suggesting that HTLV-1 infection might induce the splicing of XBP-1 as well. Higher expression of XBP-1 would be expected then since XBP-1 has been identified as a target gene of XBP-1S (27). We identified Tax as a transcription activator of XBP-1, suggesting that XBP-1 could be a cellular target gene of Tax (Fig. 11). XBP-1S and XBP-1U were found to interact with Tax and bind to the HTLV-1 LTR in vivo (Fig. 5 and 6). Both XBP-1 isoforms stimulated HTLV-1 basal as well as Tax-mediated transcription through the interaction with Tax (Fig. 1, 2, 4, and 6). Future work will be required to investigate the involvement of XBP-1 in viral replication of HTLV-1.

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Acknowledgement We would like to thank Dr. Kazutoshi Mori, Dr. Arnold Rabson, and Dr. Zhiwei Song for providing plasmids, Dr. Peter Nissom for critical review of the manuscript, and Dr. Yong Xiao and Grant Tan for expert technical assistance. This work is supported by the Agency for Science, Technology and Research (Singapore), and the National Institutes of Health (U.S.) grant AI043894 to F.K.
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43. Tirosh, B., N. N. Iwakoshi, B. N. Lilley, A. H. Lee, L. H. Glimcher, and H. L. Ploegh. 2005. Human cytomegalovirus protein US11 provokes an unfolded protein response that may facilitate the degradation of class I major histocompatibility complex products. J.Virol. 79:2768-2779. 44. Twizere, J. C., L. Lefebvre, D. Collete, C. Debacq, P. Urbain, H. Heremans, J. C. Jauniaux, A. Burny, L. Willems, and R. Kettmann. 2005. The homeobox protein MSX2 interacts with tax oncoproteins and represses their transactivation activity. J.Biol.Chem. 280:29804-29811.
45. Yin, M. J., E. J. Paulssen, J. S. Seeler, and R. B. Gaynor. 1995. Protein domains involved in both in vivo and in vitro interactions between human T-cell leukemia virus type I tax and CREB. J.Virol. 69:3420-3432. 46. Yoshida, H., T. Matsui, A. Yamamoto, T. Okada, and K. Mori. 2001. XBP1 mRNA is induced by ATF6 and spliced by IRE1 in response to ER stress to produce a highly active transcription factor. Cell 107:881-891.
47. Yoshida, H., T. Okada, K. Haze, H. Yanagi, T. Yura, M. Negishi, and K.
Mori. 2000. ATF6 activated by proteolysis binds in the presence of NF-Y (CBF)
directly to the cis-acting element responsible for the mammalian unfolded protein response. Mol.Cell Biol. 20:6755-6767.

48. Yoshida, H., M. Oku, M. Suzuki, and K. Mori. 2006. pXBP1(U) encoded in
XBP1 pre-mRNA negatively regulates unfolded protein response activator pXBP1(S) in mammalian ER stress response. J.Cell Biol. 172:565-575.
49. Yoshida, M., I. Miyoshi, and Y. Hinuma. 1982. Isolation and characterization of retrovirus from cell lines of human adult T-cell leukemia and its implication in the disease. Proc.Natl.Acad.Sci.U.S.A 79:2031-2035.
50. Yoshimura, T., J. Fujisawa, and M. Yoshida. 1990. Multiple cDNA clones encoding nuclear proteins that bind to the tax-dependent enhancer of HTLV-1: all contain a leucine zipper structure and basic amino acid domain. EMBO J. 9:2537-
51. Yu, C. Y., Y. W. Hsu, C. L. Liao, and Y. L. Lin. 2006. Flavivirus infection activates the XBP1 pathway of the unfolded protein response to cope with endoplasmic reticulum stress. J.Virol. 80:11868-11880.

665 666

52. Zhang, W., J. W. Nisbet, B. Albrecht, W. Ding, F. Kashanchi, J. T. Bartoe, and M. D. Lairmore. 2001. Human T-lymphotropic virus type 1 p30(II) regulates gene transcription by binding CREB binding protein/p300. J.Virol. 75:9885-9895. 53. Zhao, L. J. and C. Z. Giam. 1992. Human T-cell lymphotropic virus type I
(HTLV-I) transcriptional activator, Tax, enhances CREB binding to HTLV-I 21base-pair repeats by protein-protein interaction. Proc.Natl.Acad.Sci.U.S.A 89:7070-7074.

671 672

674 675

678 679

684 685
Table 1: Expression of UPR genes in different T-cell lines analyzed by quantitative RTPCR
Fold Change* in mRNA of UPR Genes

Cell lines

HTLV-1-free cells: HuT 78#
HTLV-1-infected cells: C10/MJ

1.0 + 0.1$

1.3 + 0.2

1.0 + 0.1

1.5 + 0.1
*: Fold change for a particular gene is calculated using the equation Fold change (relative to HuT78) = 2-CT where CT = CT(sample) CT(HuT78)

3.5 + 0.7

2.8 + 0.4

0.9 + 0.2

3.0 + 0.4

2.9 + 0.2

1.9 + 0.3

3.7 + 0.5

2.5 + 0.2

1.6 + 0.1

1.0 + 0.2

1.5 + 0.2

7.0 + 1.8

2.7 + 1.0

0.3 + 0.2

2.6 + 0.5

: The mRNA levels of the indicated UPR genes in HuT 78 cells were used as the control

for comparison.

: Data represent the average of triplicates with standard deviations.
Figure legends Fig. 1. Effects of XBP-1 on HTLV-1 LTR-dependent transcription in HEK293, 293T, and HeLa cells. HEK293 (A), 293T (B), and HeLa (C) cells were transiently transfected with HTLV-1-LTR-F-Luc and an expression vector [i.e. ATF6 (1-373), XBP-1S, or XBP-1U]. The empty plasmid (i.e. Mock) was used as a negative control. In a similar set of experiments, HEK293 (D), 293T (E), and HeLa (F) cells were transiently transfected with HTLV-1-LTR-F-Luc with the titration of XBP-1S plasmids (at 2-fold increment). Fold changes were determined by comparing with the reading of the negative control.

752 753

HTLV-1-LTR-F-Luc and an indicated Tax expression vector [i.e. wild-type Tax (Tax WT), NF-B-deficient Tax mutant (Tax M22), or CREB-deficient Tax mutant (Tax M47)] in the absence or presence of a XBP-1S plasmid. Cells transfected with HTLV-1LTR-F-Luc and an empty vector (i.e. no Tax and XBP-1S plasmids included) were used as a negative control. Fold changes were determined by comparing with the reading of
the control. (B) Schematic representation of truncated XBP-1 mutants. The basic-region leucine zipper domain (black box) and the C-terminal regions unique to XBP-1S and XBP-1U due to the UPR-induced splicing are indicated (striped and dotted boxes,
respectively). (C) Jurkat cells were co-transfected with HTLV-1-LTR-F-Luc and an
indicated XBP-1 expression vector with or without a wild-type Tax expression plasmid. Fold changes were determined by comparing with the reading of the negative control (i.e. no Tax and XBP-1S plasmids transfected).
transfected with a Tax-producing plasmid at the indicated amount (0.01, 0.05, 0.1, 0.5, and 1 g). The empty vector was used as a negative control. Tunicamycin treatment (4 g/ml tunicamycin) and an ATF6 (1-373) expression vector (1 g) were used as positive controls. RNAs isolated from the cells were analyzed by quantitative RT-PCR to examine the expression of XBP-1 and NF-YC genes. Fig. 10. Effects of Tax mutants on XBP-1 expression. HEK293 cells were transiently transfected with 1 g of an indicated plasmid to express wild-type Tax (Tax WT), NF-B-deficient Tax mutant (Tax M22), or CREB-deficient Tax mutant (Tax M47) expression vector. The empty vector was used as a negative control while an ATF6 (1-
Fig. 9. Tax stimulates XBP-1 transcription. HEK293 cells were transiently

756 757

373) expression vector (1 g) were used as a positive control. RNAs isolated from the cells were analyzed by quantitative RT-PCR to examine the expression of XBP-1. Fig. 11. A model for cellular-viral interaction between XBP-1 and HTLV-1 transcription.

MOLECULAR AND CELLULAR BIOLOGY, Sept. 2002, p. 63216335 0270-7306/02/$04.000 DOI: 10.1128/MCB.22.18.63216335.2002

Vol. 22, No. 18

MINIREVIEW
Classication of Human B-ZIP Proteins Based on Dimerization Properties
Charles Vinson,* Max Myakishev, Asha Acharya, Alain A. Mir, Jonathan R. Moll, and Maria Bonovich
Laboratory of Metabolism, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892 B-ZIP transcription factors (98) are exclusively eukaryotic proteins that bind to sequence-specic double-stranded DNA as homodimers or heterodimers to either activate or repress gene transcription (34). We have examined both of the recently published DNA sequences of the human genome (51, 95) and identied 56 genes that contain the B-ZIP motif. Three sequences were identical, giving a total of 53 unique B-ZIP domains with the potential to form 2,809 dimers. This creates the possibility for a tremendous range of transcriptional control (23, 50, 52). While signicant effort has been directed at identifying dimerization partners of B-ZIP proteins, the full complement of dimerization partners remains to be elucidated. This review highlights two topics: (i) the known structural rules that regulate leucine zipper dimerization specicity and (ii) experimental data addressing mammalian B-ZIP dimerization partners. We have annotated the leucine zippers of all human B-ZIP domains, highlighting amino acids in the a, d, e, and g positions that appear critical for leucine zipper dimerization specicity. These data were used to group B-ZIP proteins into 12 families with similar dimerization properties: (i) those that strongly favor homodimerization within the family (PAR, CREB, Oasis, and ATF6), (ii) those that have the ability to both homodimerize and heterodimerize with similar afnities (C/EBP, ATF4, ATF2, JUN, and the small MAFs), and (iii) those that favor heterodimerization with other families (FOS, CNC, and large MAFs). BACKGROUND In the late 1980s, several mammalian B-ZIP proteins were puried by double-stranded DNA afnity chromatography, and the genes encoding these proteins were cloned. Among the rst cloned were the AP-1 (c-FOS and c-JUN) heterodimer, (4), the CREB homodimer (65), and the C/EBP homodimer (37, 53). These newly isolated genes were used as probes in low-stringency DNA hybridizations to identify new sequencerelated B-ZIP proteins (5, 78, 101). In addition, new B-ZIP proteins (25, 36, 102) were isolated by screening lambda phage protein expression libraries with radiolabeled DNA binding elements (29, 85, 97). These functional DNA binding assays successfully isolated B-ZIP proteins because the B-ZIP motif is compact and refolds easily (87). The wealth of new sequences led to a confusing nomenclature, because multiple groups independently isolated and named the same B-ZIP proteins. Moreover, initial classication into families was often based on apparent DNA binding activity, resulting in grouping of proteins with different dimerization properties. Several reviews have helped clarify these issues, including a comprehensive review by Hurst in 1995 (34), one focusing on ATF proteins (24), and another focusing on FOS and JUN proteins (6). We hope this review will further contribute to a systematic B-ZIP classication. B-ZIP STRUCTURE Amino acid alignment of B-ZIP proteins allowed the identication of the B-ZIP motif, a long bipartite -helix that is 60 to 80 amino acids long (98). The N-terminal half contains two clusters of basic amino acids responsible for sequence-specic DNA binding, while the C-terminal half contains an amphipathic protein sequence of variable length with a leucine every seven amino acids. The shorter leucine zippers have less protein sequence exibility, because amino acids must be optimized for dimerization stability. Longer leucine zippers allow better regulation of dimerization specicity, because they can contain amino acids that are suboptimal for stability but favor interaction with a particular partner. This amphipathic sequence, termed the leucine zipper (52), mediates homo- and heterodimerization of B-ZIP proteins (3, 20, 41, 44, 54, 77, 86, 91). Figure 1 shows the X-ray crystal structure of the B-ZIP domain from yeast GCN4 bound to DNA (15). B-ZIP DNA binding stabilizes the basic region inducing the random coil to form an -helical extension of the leucine zipper (73, 84). Several B-ZIP proteins, including the small and large MAF proteins (13, 42), contain additional DNA binding elements N terminal to the basic region that increase the number of specic DNA bases that can be bound. The leucine zipper dimerization domain forms a parallel coiled coil (75) that consists of four to ve heptads, in which each heptad is composed of two -helical turns or seven amino acids, labeled a, b, c, d, e, f, and g (61). Amino acids in the a, d, e, and g positions regulate leucine zipper oligomerization, dimerization stability, and dimerization specicity. Amino acids in the a and d positions are on the same surface of the -helix and are typically hydrophobic. The a and d amino acids from one monomer interact with the complemen6321

* Corresponding author. Mailing address: Building 37, Room 2D24, Laboratory of Metabolism, National Cancer Institute, National Institutes of Health, Bethesda, MD 20892. Phone: (301) 496-8753. Fax: (301) 496-8419. E-mail: Vinsonc@dc37a.nci.nih.gov.

MOL. CELL. BIOL.

FIG. 1. X-ray structure of the yeast B-ZIP homodimer, GCN4 (blue -helices) bound to DNA (red helices). The N-terminal DNA recognition helix lies in the major groove of the DNA. An almost invariant leucine present every two turns of the C-terminal -helix (at the d position) is shown in gray.
tary a and d amino acid positions in the opposite monomer ( refers to the second -helix in the dimer). This interaction creates a hydrophobic core essential for dimer stability (87). The g and e positions typically contain charged amino acids (8, 96). X-ray crystallography reveals g7e interhelical interactions between amino acids in the g position and oppositely charged amino acids in the e, which is ve amino acids C terminal (2, 15, 19, 21, 75, 80). Electrostatic interactions between the amino acids in the g7e pair can be either attractive or repulsive and thus can regulate both homodimerization and heterodimerization. Furthermore, Van der Waal interactions between the g and e methylene groups and the underlying a and d amino acids contribute to stability (2). LIST OF ALL HUMAN B-ZIP PROTEINS GROUPED BY DIMERIZATION PROPERTIES We have examined two versions of the DNA sequence of the human genome (51, 95) and identied 56 genes that contain the B-ZIP motif. Three of these motifs were identical, resulting in 53 unique B-ZIP domains. Four of the proteins were found only in the Celera database. Table 1 provides the chromosomal location, unique database search identiers, and the number of amino acids found N terminal and C terminal of the B-ZIP domain. We have generated three dendrograms that examine the relatedness of the amino acid sequences of the 53 human B-ZIP domains. One dendrogram is based on the entire B-ZIP
domain, one is based on the basic region that is critical for DNA binding, and the last is based on the leucine zipper region that regulates dimerization specicity (Fig. 2). The three dendrograms are similar. The differences are interesting, because they reveal whether similarities are based on DNA binding properties or dimerization specicities. For example, the basic region dendrogram places the CREB, ATF6, and Oasis families together, reecting their binding to the CRE DNA sequence (5-TGACGTCA-3). In contrast, the leucine zipper dendrogram places the Oasis family separately from the CREB and ATF6 families, reecting their different dimerization properties. It is possible that these families could compete for binding to the CRE DNA sequence to give a range of transcriptional control. Another example is the XBP protein, which is not related to any sequence when its basic region is examined, but clusters with the ATF6 proteins when its leucine zipper is examined. To achieve a functional classication of the B-ZIP proteins, we have considered the dimerization properties of the B-ZIP domains. We have examined the amino acids in the g, a, d, and e positions of the leucine zipper region of the B-ZIP domains to rationalize the known and predict the unknown dimerization properties of the 53 human B-ZIP domains whose amino acid sequences are presented in Fig. 3. Using this analysis, we have grouped B-ZIP proteins with similar dimerization properties into 12 families. Our grouping is consistent with the

VOL. 22, 2002 TABLE 1. Human B-ZIP proteinsa
Family and protein Alternate name Chromosomal no.b Accession no.c
Tail length (no. of amino acids)d N-terminal C-terminal
Homodimer PAR TEF TEF paralog DBP HLF NFIL3 CREB CREB ATF1 CREM Oasis Oasis CREB-H CREB3 hCP201085 hCP1698600 ATF6 ATF6 CREBL1 XBP1 Homo- and heterodimer C/EBP CEBPA CEBPB CEBPD CEBPE CAA60698 CEBPG CHOP10 ATF4 ATF4 ATF4 paralog hCP1709392 ATF5 ATF2 ATF2 ATF7 CRE-BPa JUN JUND JUN JUNB S-MAF MAFK MAFG MAFF Heterodimer FOS c-FOS FOSB FRA1 FRA2 hCP34067 ATF3 JDP2 JDP1 BATF CNC BACH1 NRF1 NFE2 BACH2 NRF2
DABP NFIL3A, E4BP4 CREB1 HCREM-1, ICER1 BBF-2 LZIP, Luman
NP_003207.1 hCP1717043 NP_001343 NP_002117.1 NP_005375 NP_004370.1 NP_005162.1 AAC60617.2 XP_054856.1 NP_115996 NP_006359 hCP201085 hCP1698600 BAA34722 XP_004202.3 NP_005071

320 68

313 126
CEBP NFIL6, IL6DBP, LAP, TCF5 CRP3 CRP1 HP8 peptide GCSF DDIT3, GADD153, GA15 CREB2, TAXREB67 ATFX CREBP1, (CRE82) ATFA

17 X 17 22

XP_009180.2 XP_009510.1 NP_005186.1 NP_001796.1 CAA60698 S26300 NP_004074 NP_001666 hCP1640238M hCP1709392 NP_036200 XP_002767.2 NP_006847.1 XP_004938.3 NP_005345.2 NP_002219.1 NP_002220.1 NP_002351.1 NP_002350.1 NP_036455.1

AP1 NFE2U, p18

FOS FOSL1 FOSL2 LRF-1, LRG-21, CRG-5 p21SNFT, SNFT SFA-2 NFE2L1, TCF11, LCR-F1 p45 NFE2L2
NP_005243 XP_009143.2 NP_005429.1 NP_005244.1 hCP34067 NP_001665 AAF21148.1 NP_061134 NP_006390.1 NP_001177.1 NP_003195.1 XP_028656 NP_068585.1 XP_002548.1

644 479

130 43
Continued on following page
MINIREVIEW TABLE 1Continued

Family and protein

Alternate name

Chromosomal no.b

Accession no.c
NFE2L3 NFE2L3 paralog L-MAF NRL MAF-B C-MAF HCF
AAC09039.1 hCP46847 NP_006168.1 XP_009665.2 AAC27037 NP_067035

286 127

MAFB, KRML MAF ZF, C1, VCAF, CFF
NCBI LocusLink gene names are given in boldface (http://www.ncbi.nlm.nih.gov/Locuslink). b Chromosomal number at which the B-ZIP protein is located. c Accession number to uniquely identify the B-Zip domain in the NCBI or Celera database. d Number of amino acids N terminal and C terminal of the B-ZIP domain.

dendrograms and agrees particularly with the dendrogram based on the leucine zipper sequence. An examination of the human genome by Green and colleagues (90) grouped B-ZIP proteins into eight previously identied families, (PAR, CREB, C/EBP, FOS, JUN, Maf, CNC, and ATF6) based on amino acid similarity throughout the B-ZIP domain. Our analysis divides their FOS family into FOS, ATF2, and ATF4; the MAF family into the small MAFs and large MAFs; and the CREB family into the CREB and Oasis families. We also reassigned two sequences, XBP1 was moved from the FOS family to the ATF6 family, and NFLI3 was moved from the C/EBP to the PAR family. We divide these 12 families into three general groups: (i) those that strongly favor homodimerization within the family (PAR, CREB, Oasis, and ATF6), (ii) those that homodimerize and heterodimerize (C/EBP, ATF4, ATF2, JUN, and the small MAFs), and (iii) those that strongly favor heterodimerization with other families (FOS, CNC, and large MAFs). Amino acids in the leucine zipper region of each human B-ZIP domain that regulate attractive and repulsive interactions are color-coded in Fig. 3 and reveal similar interaction patterns within each family. Figure 4 presents a helical wheel representation of homo- and heterodimers to help explain the color code used in Fig. 3. A similar annotation for Drosophila B-ZIP proteins indicates that the 12 families we have identied are conserved in the insects (17). An examination of the g7e pairs in the rst four heptads of human B-ZIP proteins results in several general observations. Thirty percent are attractive, with acidic-basic pairs (orange) predominating; 23% are repulsive with acidic-acidic (red) pairs predominating; and 30% contain a single charged amino acid that can stabilize either homodimerization or heterodimerization. The d position of the hydrophobic interface typically contains leucine with very few polar and no charged amino acids. The 2nd heptad a position typically contains asparagine. The a position in other heptads typically contains hydrophobic amino acids, but asparagine and basic amino acids are occasionally observed, which are critical for regulating dimerization specicity. The surprisingly limited diversity of amino acids in the a, d, e, and g positions suggests that a limited set of rules might regulate leucine zipper dimerization specicity.

follows: E7R, 1.3 kcal/mol; E7A, 0.1 kcal/mol; and A7R, 0.7 kcal/mol. The additional energy of the E7R pair compared to the A7R and E7A pairs is 0.5 kcal/mol/saltbridge [E7R (A7R E7A) coupling energy] [1.3 (0.7 0.1) 0.5] and represents the energy of interaction (coupling energy) between E and R. Table 2 lists the stability of each g7e pair relative to A7A, and Table 3 gives their coupling energy. The larger coupling energy for the E7R pair (0.5 kcal/ mol) than that of the E7K pair (0.3 kcal/mol) indicates that the E7R pair contributes more to dimerization specicity than E7K. The like-charged E7E, K7K, and R7R pairs all have destabilizing coupling energies (0.7, 0.6, and 0.8 kcal/mol, respectively) that are larger than the E7R and E7K attractive coupling energies (Table 3). This suggests that preventing a repulsive g7e pair is more important for driving dimerization specicity than forming an attractive pair. The energetic basis of the coupling energy for g7e pairs is a subject of ongoing debate in the literature (47, 56, 59), with some studies suggesting the measured coupling energy does not have a strong electrostatic component.
The suggestion that coupling energy may not be driven by charge interactions is highlighted by the polar glutamine that has a repulsive coupling energy in pairs with either acidic E or basic K (Table 3). Because of the positive calculated coupling energies, we have color-coded E7Q and Q7E pairs in red as depicting repulsive acidic pairs and K7Q, R7Q, and Q7K in blue as depicting repulsive basic pairs (Fig. 3). Many B-ZIP families, including JUN, CNC, and C/EBP, have only one charged amino acid in the g7e pair. These charged amino acids contribute to the stability of the homodimer, as seen in the A7R and E7A pairs in the double-mutant thermodynamic analysis. Heterodimers, however, may be preferred if they form an attractive g7e pair, because of the coupling energy. Thus, incomplete g7e pairs can stabilize both homodimer and heterodimeric interactions. The a and d positions. Amino acids in the a and d positions of the leucine zipper are typically hydrophobic, with a variety of amino acids in the a positions and leucine in 84% of the d positions. An exception is the 2nd heptad a position, which contains asparagine in most homodimerizing B-ZIP proteins.
MOL. CELL. BIOL. TABLE 3. Coupling energy of interaction for g7e pairs (Gint in kcal mol1/pair) a

g e E Q R K

E Q R K

0.7 0.2 1.1 0.9

0.2 0.0 0.4 0.3

0.5 0.3 0.8 0.6

0.3 0.3 0.8 0.6
Values were calculated from Table 2.
FIG. 5. Schematic describing the four proteins used for a doublemutant thermodynamic cycle. The top panel depicts two alanines in the g and e positions of a g7e. The second panel shows that an E7A pair is 0.1 kcal/mol more stable than an A7A pair. The third panel shows that a A7R pair is 0.7 kcal/mol more stable than an A7A pair. The fourth panel shows a E7R pair. Instead of being 0.8 kcal/mol more stable than A7A, as would be expected if the two amino acids did not interact, E7R is 1.3 kcal/mol more stable. The additional 0.5 kcal/mol is described as the coupling energy indicative of an physical interaction between the E and R side chains.

FIG. 6. Schematic of the A-ZIP dominant negative. (A) A model of a B-ZIP dimer with the basic region (blue) unstructured. (B) A B-ZIP dimer bound to DNA with the basic region now -helical. (C) A B-ZIP|A-ZIP heterodimer with the protein-protein interface extending N terminally into the basic region. The basic region (blue) and the designed acidic amphipathic region (red) interact as -helices to extend the coiled-coil domain.
large MAF, and small MAF B-ZIP families use basic amino acids in the a position to create heterodimerizing B-ZIP families REPULSION BETWEEN LEUCINE ZIPPERS: USING AZIP DOMINANT NEGATIVES A problem with analyzing leucine zipper interactions by CD spectroscopy is that the stability of heterodimers can only be measured if the heterodimer is signicantly more stable than either homodimer. To gain further insights into dimerization specicity, we have developed an A-ZIP protein consisting of a leucine zipper and a designed amphipathic acidic -helical sequence that replaces the B-ZIP basic region (Fig. 6). This acidic extension forms a coiled-coil structure with the basic region in the B-ZIP|A-ZIP heterodimer and stabilizes the complex by up to 8 kcal/mol (1, 22, 49, 64, 71). The B-ZIP|AZIP heterodimer drives interactions between weakly attractive or even somewhat repulsive leucine zippers and gives us access to measuring a range of dimerization afnities. This is important because in vivo dimerization is often driven by DNA binding. The B-ZIP|A-ZIP heterodimer is more stable than the B-ZIP protein bound to DNA, so that the A-ZIPs specifically prevent B-ZIP DNA binding at equimolar concentrations. Because the acidic extension interacts with all basic regions, the specicity of interaction between A-ZIP and B-ZIP domains is primarily leucine zipper dependent (1, 22, 64, 70). The inhibition of DNA binding provides an assay for dimerization. Competition between A-ZIP protein and DNA for interac-
tion with a B-ZIP protein can be analyzed with the gel shift assay. Figure 6 shows the FOS|JUND heterodimer and the C/EBP, CREB, and PAR homodimers binding to their canonical DNA binding sites. One molar equivalent of A-PAR inhibits the DNA binding of PAR; it does not inhibit the DNA binding of FOS|JUND, C/EBP, or CREB, even at 100 molar equivalences (Fig. 7, top panel). Similarly, 1 molar equivalent of A-CREB, A-C/EBP, and A-FOS specically inhibit the DNA binding of CREB, C/EBP, and FOS|JUND, respectively. In contrast, A-ATF4 has somewhat promiscuous dimerization properties, inhibiting the DNA binding of FOS|JUND and C/EBP at 1 molar equivalent (Fig. 7E). We can rationalize the specicity seen in Fig. 7 based on the leucine zipper sequence of each protein. The homodimerizing PAR, CREB, and C/EBP families have similar a and d positions with an asparagine in the a position of the 2nd heptad. The families differ in their attractive g7e pairs, PAR has four attractive g7e pairs, while CREB and C/EBP each have two attractive g7e pairs that are subsets of the PAR pattern. Dimerization between PAR and CREB or PAR and C/EBP is prevented, because PAR preferentially homodimerizes due to the 4.0-kcal/mol/dimer of coupling energy from the eight attractive g7e pairs in the four heptads. Therefore, CREB or C/EBP remain to homodimerize. C/EBP and CREB do not interact because their g7e pairs are in different heptads. To test the validity of this idea, we mutated only three amino acids in the g and e positions of the C/EBP leucine zipper to confer the PAR pattern of g7e pairs. The mutated C/EBP displays dimerization properties similar to those of PAR (64).

R7K (blue) pair in the 1st heptad found in JUN, a repulsive E7E (red) pair in the 2nd heptad found in FOS, and a histidine in the 5th d position, as found in JUN, FOS, and ATF2. (ii) ATF4. The ATF4 family has three members, ATF4, ATF5, and hcp1709392. Besides homodimerizing, ATF4 heterodimerizes with C/EBP (93, 96), FOS (23), and NRF2 (27). This family is noteworthy in having acidic and basic repulsive g7e pairs as well as attractive g7e pairs, which may explain their promiscuous dimerization properties. The interface contains the 2nd heptad asparagine in the a position found in homodimerizing B-ZIP proteins. (iii) ATF2 family. The ATF2 family contains three proteins: ATF2, ATF7, and CRE-BPa (68). These proteins contain an asparagine in the 2nd heptad a position and attractive g7e pairs (orange) in the 3rd and 4th heptads, structural features that favor homodimerization. The 5th heptad d position contains a histidine that is also found in the FOS and JUN families and may be important for interaction with both of these families. ATF2 has been reported to heterodimerize with JUN (23, 58), FOS (28), and C/EBP (83) family members. Heterodimerization with FOS could be driven by an incomplete g7e pair in the 1st heptad of C/EBP interacting with the E7E pair in the 1st heptad of FOS to form an attractive K7E pair. Heterodimerization between C/EBP and ATF2 has been observed on a chimeric DNA sequence composed of a C/EBP half site and an ATF2 half site (83). CEBP and ATF2 homodimers have four attractive interhelical salt bridges and no repulsive g7e pairs, while a ATF2|C/EBP heterodimer has two attractive and one repulsive g7e pair, suggesting that these two proteins would prefer to homodimerize. The formation of heterodimerization on a chimeric site demonstrates the importance of DNA sequence in modulating dimerization specicity. (iv) JUN family. The JUN family is comprised of three proteins, c-JUN, JUND, and JUNB. The best-studied JUN partner is FOS. JUN and FOS heterodimerize to form the AP-1 transcription factor, originally isolated as a biochemical activity, that binds the 5-TGAGTCA-3 DNA sequence, termed the TRE (12-O-tetradecanoylphorbol-13-acetate response element) (79, 82). Many other proteins have been reported to heterodimerize with JUN family members, among them CNC, ATF2 (58, 68), ATF3, and c-Maf. JUN family members can also homodimerize, but these complexes bind DNA poorly, bringing into question their biological function (70). The amino acid sequence of the JUN leucine zipper is consistent with the experimentally observed promiscuous dimerization. Properties that drive heterodimerization are repulsive K7K (blue) and Q7K pairs (blue) in the 1st and 4th heptads, respectively. An incomplete g7e pair in the 3rd heptad also promotes promiscuous heterodimerization. In contrast, the asparagine in the 2nd heptad a position is commonly found in homodimerizing B-ZIP proteins. Heterodimerization with ATF2 creates a canonical interface and attractive g7e pairs. An elegant study (94) showed that by changing amino acids in the e and g positions, the promiscuous dimerization of JUN could be restricted to either FOS or ATF2 with a corresponding change in the biological activity of the JUN mutants. The large number of basic amino acids in the g and e positions

encourages heterodimerization with acidic proteins such as FOS. A histidine (H) in the 5th heptad d position is conserved among the JUN, FOS, and ATF2 proteins and contributes to dimer stability (9), but its contribution to dimerization specicity has yet to be elucidated. (v) Small MAF family. There are three small MAF (S-MAF) musculoaponeurotic brosarcome proteins, MafF, MafG, and MafK. The S-MAFs homodimerize, but do not contain, a transactivation domain and thus repress transcription. However, they also heterodimerize with CNC and FOS family members to activate gene expression (35, 40, 43). The S-MAF leucine zipper contains attractive E7R (orange) in the 3rd and 4th heptads that favor homodimerization (also observed in the ATF2 family) and an asparagine in the 3rd a heptad. Features that promote heterodimerization include a lysine in the 1st heptad a position (also found in the L-MAFs) and an incomplete glutamic acid g7e pair (red) in the 2nd heptad. The heterodimerizing B-ZIP families. The heterodimerizing B-ZIP families are FOS, CNC, and the large Maf. Three general properties are apparent for these proteins: (i) repulsive g7e pairs that inhibit homodimerization; (ii) amino acids in the a positions include lysine or arginine, which discourages homodimerization; (iii) incomplete g7e pairs that promote promiscuous heterodimerization. (i) FOS family. There are nine FOS family proteins: c-Fos, FosB, Fra1, Fra2, hcp34067, ATF3, JDP2, SNFT, and BATF. FOS family members heterodimerize with the JUN, CNC, and small Maf families (reviewed in reference 6). The FOS family has acidic amino acids in the g and e positions and heterodimerize with basic JUN zippers (74). Specically, FOS dimerization properties can be divided into three sets of characteristics that are slightly variable. Five of the proteins (c-Fos, FosB, Fra1, Fra2, and hcp34067) contain conserved repulsive E7E or Q7E pairs (red) in the 1st and 4th heptads and repulsive E7Q or E7E pairs (red) in the 2nd and 3rd heptads. The hydrophobic interface is composed of a threonine in the 1st a position, lysines in the 2nd and 4th a positions, and a histidine in the 5th heptad. The lysines in the a position inhibit homodimerization and drive heterodimerization. The remaining FOS family members, ATF3, JDP2, SNFT, and BATF, are not as acidic and do not contain as many repulsive lysines in the a position as the prototypical FOS leucine zipper. They can be further divided into two groups: SNFT and BATF, which do not contain the repulsive 4th heptad a position basic amino acid found in ATF3 and JDP2. (ii) CNC family. The second acidic leucine zipper family is named after the founding member, capncollar (CNC), a Drosophila protein (62). There are six members: BACH1, CNC1/ NRF1, NF-E2, BACH2, CNC2/NRF2, and NF-E2L3. These proteins heterodimerize with the S-MAF family (35, 66). These proteins have either a repulsive acidic g7e pair or an acidic incomplete g7e pair in the 1st heptad. The presence of multiple incomplete g7e pairs is expected to confer promiscuous dimerization properties. Like the FOS family, the CNC family has lysines in the a positions that drive heterodimerization. However, in contrast to the FOS family, where the lysines are in the 2nd and 4th heptad a positions, CNC proteins contain a basic amino acid in the 2nd and 3rd heptad a positions. This arrangement should alter the heterodimerization partners for the CNC proteins compared to FOS proteins. Interestingly, the

CNC|S-MAF heterodimer forms a 3rd heptad a position interaction between N and K, similar to the 2nd heptad interaction found between FOS and JUN in the 2nd heptad. (iii) Large MAF family. The large MAF (L-MAF) family contains three members: NRL, c-MAF, and MAF-B. These proteins can homodimerize and heterodimerize with FOS and JUN proteins (39, 43). L-MAF family members form heterodimers driven by the repulsive Q7K (blue) or Q7R pairs (blue) in the 2nd heptad, while homodimerization is mediated through attractive E7K pairs (orange) in the 4th heptad. Along with the small MAF proteins, these are the only B-ZIP proteins with aliphatic amino acids in the 2nd a position. The 1st and 4th heptad a position lysine or arginine discourages homodimerization. Features that promote promiscuous heterodimerization include incomplete g7e pairs composed of glutamic acid (red) in the 1st and 3rd heptads. (iv) HCF family. The HCF (host cell factor) protein, also known as C1 (45), VCAF (76), or CFF (38), exists in human cells as a family of proteolytic cleavage products of the primary HCF protein (46). HCF has a unique pattern of interactions that suggest it will homodimerize. This protein contains attractive g7e pairs (orange) in the 1st and 4th heptads. The a positions contain asparagine in the 1st and 2nd heptad a positions and serine (a small polar amino acid similar to asparagine) in the 4th heptad. HCF was found to interact with Luman (18, 55). SUMMARY We have reviewed the literature on the dimerization properties of mammalian B-ZIP proteins and made predictions about the amino acids in the a, d, e, and g positions of the leucine zipper that mediate their known dimerization specicities. We have extended these predictions to all the identied B-ZIP proteins in the human genome. These predictions appear more robust for the homodimerizing proteins than for the heterodimerizing proteins. This type of analysis will be valuable in predicting the dimerization properties of B-ZIP proteins from newly sequenced genomes for which much less experimental data exists. Additional experimental data is needed to quantify the attraction and repulsion between different B-ZIP leucine zippers to gain insight into which dimers may form in vivo. In addition, the contribution of DNA binding to B-ZIP stability needs to be quantied to gain insight into how DNA sequences can regulate B-ZIP dimer partner choice.
ACKNOWLEDGMENTS We thank Tsonwin Hai, Jon Shuman, and an anonymous reviewer for comments on the manuscript.

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