Sim Domains: Structure and Function

S E C T I O N

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HYPOXIA-INDUCIBLE FACTOR

C H A P T E R

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Hypoxia-Inducible Factors Per/ARNT/ Sim Domains: Structure and Function Thomas H. Scheuermann, Jinsong Yang, Lei Zhang, Kevin H. Gardner, and Richard K. Bruick Contents 4 5 7 7 8 10 11 11 13 15 15 17 18 20 21

1. Introduction 2. Delineation of the HIF PAS Domains 3. Expression and Characterization of HIF PAS Domains 3.1. Recombinant protein expression 3.2. Limited proteolysis 3.3. Solution NMR spectroscopy 4. Assessing PAS Domain Protein–Protein Interactions 4.1. NMR spectroscopy 4.2. Reporter genes of HIF function 4.3. Preparation of nuclear lysates 4.4. Coimmunoprecipitation of HIF-a and HIF-b 4.5. Electrophoretic mobility shift assay 5. Discussion Acknowledgments References

Abstract Hypoxia-inducible factors (HIFs) are key transcriptional regulators of genes involved in cellular adaptation to reduced oxygen availability through effects on anaerobic metabolism, oxygen delivery, angiogenesis, and cellular survival and proliferation. As such, HIFs contribute to the pathogenesis of diseases in which oxygen availability is compromised, notably ischemia and tumorigenesis. Though tremendous progress has been made in elucidating the mechanisms underlying O2-dependent regulation of HIF by Fe(II)- and 2-oxoglutarate-dependent dioxygenases, HIF induction can be uncoupled from these modes of regulation in diseases such as cancer. Consequently, renewed interest has developed in understanding the structure/function relationships of individual Per/ARNT/Sim (PAS) domains that are important for maintaining transcriptionally active HIF

Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, Texas Methods in Enzymology, Volume 435 ISSN 0076-6879, DOI: 10.1016/S0076-6879(07)35001-5

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2007 Elsevier Inc. All rights reserved.

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complexes, regardless of the manner by which HIF is induced. This review highlights strategies for the biophysical and biochemical characterization of the PAS domains found within both HIF subunits and provides a platform for future efforts to exploit these domains in therapeutic settings.

1. Introduction Mammalian cells have a remarkable capacity to adapt to changes in their environment. One such pathway receiving considerable attention (Kaufman et al., 2004) is the hypoxic response pathway, which is induced in all mammalian cells in response to chronic O2 deprivation and is critical for many physiological and pathophysiological processes (Semenza, 2000). This response is based on changes in gene expression mediated through HIFs, heterodimers composed of HIF-a and HIF-b subunits (Wang et al., 1995). Oxygen-dependent regulation of HIF appears to be primarily mediated through posttranslational modifications of the HIF-a subunit (reviewed in Dann and Bruick, 2005). Under normoxic conditions, Fe(II)- and 2-oxoglutarate-dependent dioxygenases (Bruick and McKnight, 2001; Epstein et al., 2001; Hewitson et al., 2002; Ivan et al., 2002; Lando et al., 2002b) employ O2 to hydroxylate proline residues within the HIF-a oxygendependent degradation domain and an asparagine residue within the HIF-a C-terminal transactivation domain. These modifications promote HIF-a degradation by the proteasome (Ivan et al., 2001; Jaakkola et al., 2001) and interfere with coactivator recruitment (Lando et al., 2002a), respectively. Under low oxygen conditions, hydroxylation does not occur, and the HIF-a subunit binds to the O2-insensitive HIF-b subunit (also known as the aryl hydrocarbon nuclear receptor translocator, ARNT) and subsequently induces expression of genes that promote cellular adaptation to hypoxia (for examples, see Semenza, 2003). HIF can also be induced through O2-independent mechanisms, as revealed by genetic alterations that activate oncogenic-signaling pathways or inactivate tumor suppressor genes. Perhaps the best-studied examples are mutations that inactivate pVHL, the recognition subunit of the E3 ubiquitin ligase complex, which targets HIF-a for degradation under normoxia (Maxwell et al., 1999). Such mutations result in constitutive stabilization of the HIF-a subunit and contribute to the pathogenesis of a number of tumors (Maxwell, 2005). In contrast, activating mutations in several signaling pathways can induce HIF under normoxic conditions by increasing the rate of HIF-1a protein synthesis (Giaccia et al., 2003; Semenza, 2003). The ability of O2-dependent and independent mechanisms to induce HIF has prompted renewed interest in characterizing other domains relevant to HIF function, regardless of the manner in which it is induced.

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In addition to the O2-sensitive regions of HIF-a, both HIF subunits contain several O2-independent elements, including two PAS domains. The PAS domain family derives its name from three of the first proteins noted to contain this motif: period (Per; Huang et al., 1993), aryl hydrocarbon receptor nuclear translocator (ARNT; Hoffman et al., 1991), and single-minded (Sim; Nambu et al., 1991). These domains are found in a wide range of proteins, ranging from transcription factors to enzymes to ion channels (Taylor and Zhulin, 1999), where they typically mediate protein– protein interactions. While HIF PAS domains have been reported to contribute to HIF heterodimer formation (Chapman-Smith et al., 2004; Jiang et al., 1996), they also have been implicated as mediators of nuclear localization (Kallio et al., 1998) and HIF stabilization via heat shock protein 90 (Hsp90) association (Gradin et al., 2002; Isaacs et al., 2004; Kallio et al., 1998; Katschinski et al., 2004; Minet et al., 1999). While these studies provided some initial insight into the importance of PAS domains for HIF function, the interpretation of much of these data is complicated by the absence of structural information on PAS domains at the time and the incomplete biochemical characterization of reagents used in these studies. For example, the use of limited sequence data to define the boundaries of low homology PAS domains can result in the generation of inappropriate truncations within a structural domain, introducing unfolded regions within a protein instead of cleanly excising a single domain. These concerns may be compounded by over-reliance on exogenous protein overexpression (resulting in non-physiological interactions in the cell culture model system) or through the use of uncharacterized recombinant proteins that may not be properly folded. Here we describe efforts to avoid these issues in our studies of HIF PAS domains by using a combination of methods to characterize their structure and function, integrating data from in vitro and in-cell studies of the HIF heterodimer.

2. Delineation of the HIF PAS Domains Since its initial description in 1993 (Huang et al., 1993), the concept of a ‘‘PAS domain’’ has undergone several revisions as additional sequence and structural information have become available. As originally detailed by sequence comparisons among the Per, ARNT, Sim, and aryl hydrocarbon receptor (AHR) proteins, the PAS domain was thought to be roughly 270 amino acids in length and to contain two copies of a tandem repeat sequence. Subsequent characterization of a larger number of PAS-containing proteins revealed that these proteins actually contain two separate PAS domains (designated PAS-A and PAS-B). This led to several suggestions in which PAS domains actually contain two sequence homology elements,

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a 50- to 60-residue N-terminal S1 box (Zhulin et al., 1997) or PAS (Ponting and Aravind, 1997) motif that is frequently accompanied by a C-terminal 45- to 50-residue S2 box or PAS C-terminal (PAC) motif. Efforts to identify PAS-containing sequences are complicated by poor sequence similarity and the variable length of loops connecting several conserved secondary structure elements, particularly in the S2/PAC motif. These sources of primary structure diversity are likely of functional importance, perhaps reflecting the broad range of PAS domain interactions with proteins, cofactors, and ligands (Pellequer et al., 1999). The tertiary structures of several PAS domains reveal a conserved structural unit comprised of a mixed a/b fold in which several a-helices pack against a common face of a five-stranded antiparallel b-sheet (Fig. 1.1) (Amezcua et al., 2002; Card et al., 2005; Erbel et al., 2003; Morais Cabral et al., 1998). These structural studies confirmed earlier suspicions that the S1/PAS and S2/PAC motifs are integral to the PAS domain tertiary structure, with the former encompassing the Ab-Fa secondary structure elements and the latter containing the last three b-strands of the PAS b-sheet (Hefti et al., 2004) (see Fig. 1.1). Unfortunately, the PAS/PAC nomenclature is still occasionally encountered and has led to studies of 70- to 80-residue PAS domain fragments that undoubtedly lack the last three strands of the PAS b-sheet. As a general rule, minimal PAS domains that lack extensive loop structures contain approximately 100 amino acid residues, and we suggest that shorter constructs are likely missing elements required for a stably folded, biologically relevant protein. It should be noted that some PAS structures display additional structural elements N- or

A

B

Ca

Da

Bb N

Ca

C

Ib

Ab Da

Hb Ea

Ab Hb

Bb Ea

Gb

Gb

Ib

Fa Fa

Figure 1.1 Per/ARNT/Sim (PAS) domain topology and structure. (A) Schematic of a typical arrangement of secondary structure elements in PAS domains. (B) Ribbon diagram of the solution structure of the hypoxia-inducible factors (HIF)-b PAS-B domain (Card et al., 2005). In both panels, the PAS motif/S1 box (red) and PAC motif/S2 box (blue) sequence homology elements are indicated, and secondary structure elements are labeled using the nomenclature initially established for FixL (Gong et al., 1998).

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C-terminal to the core PAS fold, and that these non-canonical structural features can associate with the core a/b fold in structurally and functionally essential ways (e.g., Per [Yildiz et al., 2005], photoactive yellow protein [PYP] [Craven et al., 2000], and phototropin1 [phot1] kinase [Harper et al., 2003, 2004]). We have routinely used a combination of computational and biophysical tools detailed elsewhere (Card and Gardner, 2005) to aid in the identification of PAS domain boundaries that provide well-folded proteins amenable to biophysical and functional characterization. To identify preliminary domain boundaries, the secondary structure of a PAS-containing protein can be predicted with the web-accessible program JPred (Cuff et al., 1998). These predictions are then compared with known PAS tertiary structures, paying particular attention to identify any potential secondary structures adjacent to the minimal a/b PAS fold. Using this information, we design a series of protein constructs with variable N- and C-termini, each of which are evaluated for recombinant expression levels, solubility, structural integrity, and oligomeric state. Such analyses performed on the PAS domains from the three human HIF-a subunits, HIF-1a, HIF-2a, and HIF-3a (Ema et al., 1997; Gu et al., 1998; Tian et al., 1997), and HIF-b have led to the predicted sequence boundaries defining each HIF PAS domain shown in Fig. 1.2.

3. Expression and Characterization of HIF PAS Domains 3.1. Recombinant protein expression Numerous PAS domains have proven amenable to heterologous expression in Escherichia coli, which, as of January 2007, has provided the quantities of protein required to determine the high-resolution structures of at least 16 different domains or their complexes. In our work, we employ a parallel cloning approach with a series of bacterial expression plasmids (Sheffield et al., 1999), each of which provides expression of a different N-terminal fusion protein tag while using a common multiple cloning site. We have extended this system, generating expression plasmids with N-terminal Gb1 or His6-Gb1 tags (Amezcua et al., 2002; Harper et al., 2003). The latter tag affords the benefits of affinity chromatography while avoiding the poor expression and/or solubility demonstrated by some His6-tagged proteins (Hammarstrom et al., 2002). While this approach is sufficient for many PAS domains, allowing us to generate large amounts (>50 mg/l) of folded protein in certain cases, more challenging expression targets may demand alternative expression strategies. Such problems may sometimes be addressed by the use of different bacterial expression vectors or host cell lines to address

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Ab

Bb

Ca

Da

Ea

Fa

HIF1a HIF2a HIF3a HIFb

91 90 86 167

FYLKALDGFVMVLT-DDGDMIYISDNVNKYMGLTQFELTGHSVFDFTHPCDHEEMREML-THRNG LYLKALEGFIAVVT-QDGDMIFLSENISKFMGLTQVELTGHSIFDFTHPCDHEEIRENL-SLKNG CYLKALEGFVMVLT-AEGDMAYLSENVSKHLGLSQLELIGHSIFDFIHPCDQEELQDAL-TPQQT LILEAADGFLFIVSCETGRVVYVSDSVTPVLNQPQSEWFGSTLYDQVHPDDVDKLREQLSTSENA

HIF1a HIF2a HIF3a HIFb

154 153 149 232

--LVKKGKEQN------------------TQRSFFLRMKCTLTS------------------RGR SGFGKKSKDMS------------------TERDFFMRMKCTVTN------------------RGR --LSRRKVEAP------------------TERCFSLRMKSTLTS------------------RGR --LTGRILDLKTGTVKKEGQQSSMRMCMGSRRSFICRMRCGSSSVDPVSVNRLSFVRNRCRNGLG

HIF1a HIF2a HIF3a HIFb

181 182 176 295

TMNIKSATWKVLHCTGHIHVYDT-NSNQPQCGY--KKPPMT-CLVLICEPIPHPSNIEIPLDSKTVNLKSATWKVLHCTGQVKVYNNCPPHNSLCGY--KEPLLS-CLIIMCEPIQHPSHMDIPLD--TLNLKAATWKVLNCSGHMRAYKP-PAQTSPAGSPDSEPPLQ-CLVLICEAIPHPGSLEPPLGRGA SVKDGEPHFVVVHCTGYIKAWPP-AGVSLPDDDPEAGQGSKFCLVAIGRLQVTSSPNCTDMSNVC

PAS-A

Gb

Ib

Hb

Bb

PAS-B

Ab

Ca

Da

Linker

Fa

Ea

HIF1a HIF2a HIF3a HIFb

238 240 235 359

DSKTFLSRHSLDMKFSYCDERITELMGYEPEELLGRSIYEYYHALDSDHLTKTHHDM DSKTFLSRHSMDMKFTYCDDRITELIGYHPEELLGRSAYEFYHALDSENMTKSHQNL GRGAFLSRHSLDMKFTYCDDRIAEVAGYSPDDLIGCSAYEYIHALDSDAVSKSIHTL QPTEFISRHNIEGIFTFVDHRCVATVGYQPQELLGKNIVEFCHPEDQQLLRDSFQQV

HIF1a HIF2a HIF3a HIFb

295 297 292 416

FT-KGQVTTGQYRMLAKRGGYVWVETQATVIYNTKNSQPQCIVCVNYVVSGIIQHD CT-KGQVVSGQYRMLAKHGGYVWLETQGTVIYNPRNLQPQCIMCVNYVLSEIEKND LS-KGQAVTGQYRFLARSGGYLWTQTQATVVSGGRGPQSESIVCVHFLISQVEETG VKLKGQVLSVMFRFRSKNQEWLWMRTSSFTFQNPYSDEIEYIICTNTNVKNSSQEP

Gb

Hb

Ib

Figure 1.2 Sequence and structural elements of the hypoxia-inducible factors (HIF) Per/ARNT/Sim (PAS) domains. Multiple sequence alignments of PAS-A and -B fragments from human HIF-b (P27540) and three human HIF-a isoforms (Q16665, Q99814, and Q9Y2N7) are shown. The secondary structures plotted above the PAS-B sequences are those observed in the structure of HIF-2a PAS-B (Erbel et al., 2003). For the HIF PAS-A domain, the plotted secondary structures are those predicted by use of the JPred algorithm (Cuff et al.,1998) in combination with manual placement of the D- and E-helices and the F-helix boundaries. All accession numbers given correspond to the sequences deposited in the UniProt Knowledgebase.

proteolysis, redox, codon bias, or toxicity issues. In addition, PAS domains that are poorly soluble and that demonstrate large unstructured regions, such as the 18 and 23 kDa PAS-A domains of HIF-a and HIF-b proteins, may benefit from coexpression approaches, as has been reported for full-length bHLH-PAS-A and -B fragments of the HIF heterodimer (Chachami et al., 2005; Chapman-Smith et al., 2004; Kikuchi et al., 2003).

3.2. Limited proteolysis The structural integrity of recombinantly expressed proteins is of utmost concern and must be experimentally validated. Using relatively dilute (10–100 mM) protein solutions, techniques such as size exclusion chromatography and circular dichroism spectroscopy can quickly ascertain that the

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recombinant protein adopts a compact, folded structure. Limited proteolysis experiments can complement these methods by assessing structural integrity, working under the assumption that most potential proteolysis sites are inaccessible in a well-folded protein (Cohen et al., 1995). For limited proteolysis experiments, we generally prepare separate reactions containing a 100:1 w/w ratio of protein to trypsin, chymotrypsin, or V-8 proteases, which cleave proteins after basic, aromatic, or acidic residues, respectively. For small PAS domains on the order of 15 kDa molecular mass, reactions typically consist of 1 mg of purified PAS domain and 10 mg of protease in a 500-ml reaction volume. Following protease addition to the protein solution, we withdraw 20-ml aliquots at time points ranging from 5 min to 16 h and quench the reaction by boiling in the presence of an equivalent volume of denaturing buffer. Samples are subsequently analyzed by sodium dodecyl phosphate–polyacrylamide gel electrophoresis (SDS-PAGE). To precisely determine protease-accessible sites in the primary structure, electrospray ionization (ESI) mass spectrometry–compatible samples may be prepared by quenching with a 1% final concentration of trifluoroacetic acid (in a chemical safety hood), freezing or holding quenched samples on ice, and purifying peptide/protein components by reverse phase high-performance liquid chromatography (HPLC). Given accurate masses of proteolytic fragments, we routinely use the MS-Digest component of the ProteinProspector suite of proteomic tools (http://prospector.ucsf.edu) to identify cleavage sites in the primary structure. It must be emphasized that limited proteolysis experiments do not directly assay the folded state of a protein, but rather reveal the accessibility of potential protease sites. The most easily interpreted data is collected from protein samples representing the extremes of completely disordered or compact, well-folded polypeptides. The former sample would be completely proteolyzed during initial time points, while the latter sample is expected to show little to no degradation over the course of the experiment. There exists a potentially confusing middle ground where well-folded protein domains demonstrate degradation on account of one or more extended loops. For example, the HIF-2a PAS-B domain has a single, highly-accessible trypsin site within the extended loop connecting the Hb and Ib strands (HI loop; see Fig. 1.1) that leads to accumulation of a large protein fragment followed by complete degradation over several hours. Likewise, limited chymotrypsin proteolysis of a PAS-A fragment of PAS kinase (Amezcua et al., 2002) revealed the Gb/Hb loop to be similarly protease-accessible. Without high-resolution structures, these data might be misinterpreted as indicating domain boundaries at sites that actually reside within well-folded domains. Attempts to trim these two PAS domains to more minimal amino acid sequences that express only the large proteolytic fragment would excise residues composing internal strand(s) of the PAS b-sheet and almost certainly disrupt the PAS fold. Therefore, one must

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interpret such data in light of known PAS domain structures and discriminate between the rapid proteolysis of entirely unstructured proteins and the slower degradation of folded proteins.

3.3. Solution NMR spectroscopy Optimal domain boundaries may also be refined by analysis of twodimensional (2D) 15N/1H heteronuclear spin quantum coherence (HSQC) spectra (Kay et al., 1992), a solution nuclear magnetic resonance (NMR) experiment commonly used to study protein structure and dynamics. Collected from uniformly 15N-isotopically enriched protein, either purified or in clarified bacterial lysates (Gronenborn and Clore, 1996), a single 15N/1H HSQC experiment can qualitatively describe the extent to which a protein is structurally well-ordered. In these spectra, each cross-peak corresponds to a specific pair of proton and nitrogen nuclei separated by a single bond, as found in amide groups in the protein backbone. The chemical shifts of these peaks are sensitive to their local magnetic environment, which will significantly change depending on the state of protein folding near each site. Unstructured proteins, where amide groups largely interact with solvent, typically exhibit 15N/1H HSQC spectra with most peaks clustered in the middle of the proton dimension (
7.9–8.5 ppm) (Fig. 1.3A). In contrast, spectra from well-folded proteins have peaks dispersed over a broader range of 1H frequencies (
5.5–11 ppm) and demonstrate uniform peak intensities and line widths (see Fig. 1.3B). Comparing these spectra collected from protein constructs with different domain boundaries may identify a minimal, well-folded protein construct. Shorter protein constructs that truncate amino acids required for folding will exhibit 15N/1H HSQC spectra with poor amide proton chemical shift dispersion. Larger constructs that merely add unstructured amino acids to the folded domain provide spectra with narrow and intense peaks in the 1H
7.9- to 8.5-ppm range in addition to the well-dispersed spectral peaks corresponding to the folded domain. Various NMR-based measurements of rotational correlation time or translational diffusion coefficients may also be used to confirm the oligomeric state of recombinant protein fragments (Altieri et al., 1995; Kay et al., 1989), potentially in combination with analytical ultracentrifugation or size exclusion chromatography methods. By these approaches, boundaries have been identified for the C-terminal PAS (PAS-B) domains of HIF-1a (residues 238–349), HIF-2a (residues 240–350) and HIF-b (residues 350–470) and subsequently used for the solution structure determinations of the isolated HIF-2a and HIF-b domains (Card et al., 2005; Erbel et al., 2003) and to calculate an NMR data-guided model of an HIF-2a:HIF-b PAS-B heterodimer (Card et al., 2005).

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HIF PAS Domains

A

B 107 111

C257S

C336S

-

15N

ppm -

115 119 123 127 131 10

9

8

7

6 10 - 1H ppm -

9

8

7

6

Figure 1.3 Use of nuclear magnetic resonance (NMR) spectroscopy to validate the structures of recombinant proteins. Displayed are 15N/1H heteronuclear spin quantum coherence (HSQC) spectra collected from two different Gb1-HIF-2a PAS-B fusion proteins bearing single amino acid substitutions in the Per/ARNT/Sim (PAS) domain. (A) Hypoxia-inducible factors (HIF)-2a PAS-B (C257S). In this spectrum, the majority of peaks are found within the 1H
7.9- to 8.5-ppm (denoted by blue lines) boundaries associated with poorly folded or disordered protein elements. The poor signal intensity of this spectrum also implies that this construct is aggregation prone, a function of NMR spectroscopy’s sensitivity to macromolecular size. (B) HIF-2a PAS-B (C336S). A 15N/1H HSQC spectrum of this construct indicates that it is a well-folded protein, as demonstrated by the presence of a large number of well-dispersed peaks showing similar line widths. The spectrum collected from the Gb1-fused mutant PAS domain is given in black, while a reference Gb1-only spectrum is given in red.This example underscores the importance of confirming the folded state of recombinantly expressed proteins. While neither of these conservative substitutions was expected to disrupt the HIF-2a PAS-B structure, qualitative analysis of these spectra provided early insight regarding the viability of these reagents for subsequent studies.

4. Assessing PAS Domain Protein–Protein Interactions 4.1. NMR spectroscopy Investigations of many PAS-containing proteins suggest that they function as protein–protein interaction domains, often using the solvent-exposed b-sheet surface as an interaction interface. Crystal structures of the isolated PAS domains from E. coli Dos heme domain EcDOS (Fig. 1.4A) (Kurokawa

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Figure 1.4 Use of the Per/ARNT/Sim (PAS) b-sheet as a dimerization interface. (A) The crystallographic structure of the EcDOS (heme-regulated phosphodiesterase from Escherichia coli) PAS domain (Kurokawa et al., 2004; Park et al., 2004) (bound heme omitted for clarity), in which the b-sheet does not directly act as a dimerization surface, but instead hosts interactions with a non-canonical N-terminal helix that serves as part of the interface. (B) The crystallographic structure of tandem PAS domains from the Per protein (Yildiz et al., 2005) reveals a non-canonical helix from the C-terminal PAS domain of one subunit (gray) interacting extensively with the b-sheet of the N-terminal PAS domain (black) from the other subunit. (C) The NMR data-guided model of the hypoxia-inducible factors (HIF)-2a PAS-B (gray) and HIF-b (black) heterodimer, in which the two domains adopt an antiparallel relative orientation with respect to each other. HIF-2a PAS-B amino acids making direct contact with the HIF-b PAS-B domain are indicated by spheres.

et al., 2004; Park et al., 2004) and a tandem PAS fragment from Per (see Fig. 1.4B) (Yildiz et al., 2005) reveal that the solvent-exposed surface of the PAS b-sheet is often used as a homodimerization interface. Photosensory PAS domains likewise employ the b-sheet as an intramolecular interaction interface that can be regulated by internally bound cofactors (Craven et al., 2000; Harper et al., 2003). Combined with the established role of PAS domains in protein–protein interactions, it was clear from the

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initial identification of HIF that these domains could play a potential role in heterodimer assembly (Wang and Semenza, 1995). To characterize these interactions, we have used solution NMR spectroscopy to take advantage of its strengths in characterizing protein–protein interactions in vitro, particularly in its ability to quantitatively evaluate relatively weak complexes (low mM–low mM dissociation constants), which may not be accessible to other techniques, and to provide atomic resolution information. For the HIF-2a:HIF-b PAS-B complex, 15N/1H HSQC spectra were collected from samples containing 200 mM 15N-isotopically enriched HIF-2a PAS-B and 0 to 800 mM HIF-b PAS-B with natural abundance 15N (0.37%) (Erbel et al., 2003). Taking advantage of the tendency of NMR peaks to broaden and become less intense with increasing molecular size, diminishing 15N HIF-2a PAS-B peak heights were plotted as a function of HIF-b PAS-B concentration and fit to a model of single-site binding, providing a KD ¼ 30 mM for the two isolated domains. Other 15N/1H HSQC peaks shifted position during the course of the titration on account of these nuclei experiencing different magnetic environments in the monomeric and heterodimeric states. Having assigned the chemical shifts of monomeric HIF-2a PAS-B nuclei as a prelude to structure determination, it was possible to correlate changes in cross peak locations to sites that were perturbed upon complex formation. These chemical shift changes mapped to the solvent-exposed b-sheet surface of HIF-2a PAS-B. The reverse titration experiment with unlabeled HIF-2a PAS-B titrated into 15N-HIF-b PAS-B revealed that HIF-b similarly uses its b-sheet as an interaction interface. These findings were supported by analysis of PAS mutants characterized in vitro and in the context of living cells (Erbel et al., 2003; Yang et al., 2005). Together, these data were used to calculate the mixed NMR-docking structure of the HIF-2a PAS B:HIF-b PAS B heterodimer (see Fig. 1.4C) (Card et al., 2005). This model shows the two domains packing together via their solvent-exposed sheets, arranged in an antiparallel manner with the N- and C-termini of each domain on opposite sides of the complex. This model was subsequently validated by obtaining ˚ ) distance restraints using paramagnetic relaxation long range (
25 A enhancements to measure intersubunit distances between a nitroxide spin label covalently attached to one subunit of the heterodimeric complex and NMR-active nuclei in the other partner (Card et al., 2005). These data were fully consistent with the antiparallel orientation identified by modeling.

4.2. Reporter genes of HIF function To functionally validate such models, one can employ a combination of domain deletions and point mutations to probe the involvement of domains and their surfaces in protein–protein interactions in vitro and within living cells. Choices underlying the nature and location of mutations benefit from the

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availability of structural models and are dictated by the particular question(s) being asked. In every case, consideration should be given to the consequences of deletions/mutations with respect to the potential for impinging upon adjacent functional elements or unintentionally compromising global folding. Techniques previously described are useful for assessing the effects of mutations in vitro on domain folding/integrity and on protein–protein interactions using recombinant protein preparations. As with any such model, it is important to correlate effects observed in the context of isolated domains with effects on full-length transcription factors in functional settings. To this end, a number of functional assays for HIF function can be performed. For example, HIF activity can be reported as a function of target gene induction. Several suitable assays are available, employing surrogate reporter constructs (such as luciferase) under the control of HIF response elements (for example, see Bruick, 2000) or the examination of endogenous HIF target genes by Northern blot analysis or quantitative reverse transcriptase polymerase chain reaction (RT-PCR). Interpretation of such experiments must be taken with care, as HIF activity can be influenced by a number of PAS-independent regulatory mechanisms. Furthermore, exogenous overexpression of HIF subunits containing mutations or deletions may overwhelm relevant endogenous regulatory factors leading to artifactual behaviors. Fortunately, a number of cell lines exist in which one of the two HIF subunits has been inactivated (Hoffman et al., 1991; Wood et al., 1998). Such cell lines are well-suited for examining the activity of transiently- or stably-transfected HIF constructs expressed at physiologically relevant levels in a background absent of endogenous HIF. A variety of mechanisms exist by which mutations to HIF subunits can lead to a decrease in transcriptional activity in cultured cells including, but not limited to, decreased protein accumulation, improper localization, impaired coactivator recruitment, heterodimer disruption, or decreased ability to bind DNA. While many assays are available to assess each of these functions, we have found two to be particularly useful; coimmunoprecipitation (co-IP) assays, which monitor HIF heterodimerization, and electrophoretic mobility shift assay (EMSA), which monitors HIF DNA binding activity, are described here. In our experience, the effects of PAS domain mutations on the function of full-length HIF proteins in these assays strongly correlates with the effects of those same mutations on individual PAS domains (Yang et al., 2005). Though these assays can be performed with recombinantly expressed fragments of HIF, we typically use HIFcontaining nuclear lysates, presuming that HIF PAS domains are most likely to be folded properly in this context.

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4.3. Preparation of nuclear lysates Following a modified version of protocols described in other texts (Wang and Semenza, 1996), cells are grown to near confluency in complete medium in 150-mm plates. For HeLa cells, typically 5  106 cells are inoculated/plate and grown for 2 days at 37 in an incubator maintained under ambient atmosphere plus 5% CO2. To induce HIF accumulation, cells are then incubated under hypoxic conditions (1% O2, 5% CO2, balance N2) in a 37 incubator for 12 h. If a hypoxic incubator is not available, HIF accumulation can be induced upon addition of 100 mM desferrioxamine mesylate, an iron chelator that blocks prolyl hydroxylasemediated degradation of HIF-a under normoxic conditions. Following HIF induction, the cells are washed with cold phosphate buffered saline (PBS), followed by addition of cold PBS plus 1 mM Na3VO4, after which cells are harvested by scraping. After pelleting the cells for 2 min at 3500g and carefully removing all the PBS by aspiration, the cell pellet is resuspended in 1 ml buffer containing 20 mM Hepes (pH 7.5), 0.1 mM EDTA, 5 mM NaF, and 10 mM Na2MoO4 and incubated on ice for 10 min. Nonidet P-40 is added to a final concentration of 0.5%, followed by vigorous vortexing of the sample for 10 s. To recover the cell nuclei, the sample is pelleted at 13,000 rpm in a microfuge for 1 min at 4 , after which the supernatant is removed by aspiration. To extract nuclear proteins, resuspend the pelleted nuclei in extraction buffer (420 mM KCl, 20 mM Tris-Cl (pH 7.5), 1.5 mM MgCl2, 20% glycerol, 2 mM DTT, 1 mM Na3VO4, 0.3 mM PMSF; add 1:200 protease inhibitor cocktail [PIC; P8340, Sigma-Aldrich, St. Louis, MO] immediately prior to addition). For HeLa cells, 400 ml extraction buffer is added for every 150-mm plate of cells harvested. Incubate for 30 min at 4 with slow mixing. Pellet the insoluble debris at 13,000 rpm in a refrigerated microfuge for 10 min at 4 and recover the supernatant. The nuclear extract can generally be stored at –80 for several months. Prior to storage or usage, the nuclear lysate can be dialyzed into a low salt buffer. However, dialysis is often unnecessary for the assays described here given that the final protein concentration of the lysate is
1.5 to 2.5 mg protein/ml, as is typically observed when using HeLa cells. Yields will vary among different cell lines.

4.4. Coimmunoprecipitation of HIF-a and HIF-b Co-IP assays are particularly useful for assessing HIF heterodimerization in vitro. Briefly, an antibody specific for one of the HIF subunits is used to immunoprecipitate (IP) that protein from a sample. The IP’ed material is then resolved by SDS-PAGE, and amounts of the IP’ed protein and co-IP’ed–associated proteins can be ascertained by Western blot analysis using appropriate antibodies. However, success of this approach requires

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antibodies capable of immunoprecipitating a protein without disrupting its association with other factors. We tested several commercially available and custom antibodies raised against each HIF subunit for their ability to IP the HIF heterodimer. Most commercial antibodies we tested were unable to IP an intact HIF heterodimer, though we did find that rabbit polyclonal antibodies raised against human HIF-b (ARNT) residues 1 to 140 or various human HIF-2a epitopes in the C-terminal of the PAS domains worked for this application (Dan Peet, personal communication). Alternatively, one can transfect cells with HIF-a or -b expression constructs in which the HIF subunits are tagged with epitopes (i.e., FLAG or myc) for which commercial antibodies are available. However, care must be taken when using such tagged proteins to correlate HIF activity with heterodimerization via mutational analysis, as the presence of multiple C-terminal epitope tags can independently compromise HIF activity observed upon transfection ( J. Garcia, personal communication). Exact conditions for the co-IP assay will vary depending on the antibodies and cell lines used and should be optimized on a case-by-case basis. As a starting point, we recommend diluting 50 to 100 mg of total protein from the nuclear lysate to a final volume of 400 ml with IP buffer (150 mM KCl, 10 mM Tris-Cl [pH 7.5], 2 mM EDTA [ethylenediamine tetraacetic acid], 0.1% Triton X-100, 0.5 mM PMSF [phenylmethylsulfony fluoride], 1% skim milk powder). Add 15 to 30 ml of a 50% protein G agarose slurry (Cat# 1 243 233; Roche Applied Science, Indianapolis, IN) pre-equilibrated with IP buffer and an experimentally determined amount of the IP antibody. Protein A–agarose beads also can be used, though we have found that protein G beads give more consistent results. The reaction mixture is then incubated for 2 h at 4 with gentle rotation. Antibodybound beads are pelleted for 1 min at 2000 rpm in a microfuge, and the supernatant is removed by aspiration. The beads are then washed once with 1.25 ml of IP buffer lacking PMSF. To elute proteins, 50 ml of 2 protein loading buffer (125 mM Tris-Cl [pH 6.8], 4% SDS, 20% glycerol, 10% 2-mercaptoethanol) is added to the beads, followed by boiling the sample for 5 min. After a brief centrifugation to pellet the beads, the soluble sample can be resolved in a 10% SDS-PAGE gel. Proteins from the SDS-PAGE gel are electro-transferred to Hybond-C Extra nitrocellulose (Amersham, Piscataway, NJ) for Western blot analysis. As we typically employ rabbit polyclonal antibodies for immunoprecipitation, mouse monoclonal antibodies are used for Western blot analysis. Several commercially available antibodies perform well in Westerns, including an anti-HIF-1a mouse monoclonal antibody (Cat# 610959; BD Biosciences, San Jose, CA), an anti-EPAS ([endothelial PAS domain protein] HIF-2a) mouse monoclonal antibody (EP190b, NB 100–132; Novus Biological, Littleton, CO), and an anti-ARNT (HIF-1b) mouse monoclonal antibody (H1beta234, NB 100–124; Novus Biological).

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Peroxidase-conjugated AffiniPure donkey anti-mouse immunoglobulin (Ig) G (H þ L) from Jackson ImmunoResearch (West Grove, PA; Cat# 715–035–150) is used as the secondary antibody because it displays minimal species cross-reactivity with the large amount of rabbit heavy chain present on the blot from the IP antibody. Together, these precautions minimize background signal on the blot following addition of the chemiluminescent peroxidase substrate (Cat# 34080; Pierce, Rockford, IL).

4.5. Electrophoretic mobility shift assay Complementing the ability of co-IP assays to probe protein–protein interactions in HIF, EMSA assays can be used to assess the DNA-binding activity. Because assembly of the HIF heterodimer is a prerequisite for DNA binding, these assays can also be used to indirectly probe protein– protein interactions between the HIF subunits. Briefly, a radiolabeled double-stranded DNA probe encompassing an HIF binding site is incubated with protein. The binding of HIF to the DNA probe is reflected as a shift in mobility of the HIF:DNA complex with respect to the unbound DNA in a nondenaturing PAGE gel, as observed by autoradiography. The sequence of DNA probes is typically derived from hypoxia-response elements (HREs) of endogenous HIF promoters. To detect HIF-1 and -2 DNA binding, we typically employ a sequence from the vascular endothelial growth factor (VEGF) promoter, 50 -TACACAGTGCATACGTGGGTTTCCACAGGTCGTCT-30 (Ema et al., 1997). Other sequences, such as the W18 probe derived from the erythropoietin (EPO) promoter (50 -GATCGCCCTACGTGCTGTCTCA-30 ) (Wang and Semenza, 1995), work better for HIF-1 than HIF-2. To prepare radiolabeled double-stranded DNA probes, equal amounts of these oligonucleotides and their complements are annealed in STE buffer (10 mM Tris, pH 8.0, 50 mM NaCl, 1 mM EDTA) by heating to 95 followed by slow cooling to room temperature over
45 min. The DNA is radiolabeled upon incubation with T4 DNA polynucleotide kinase and [g-32P] ATP (adenosine triphosphate) and purified by extraction in phenol/chloroform/ isoamyl alcohol (25:24:1) followed by desalting using a ProbeQuant G-50 Micro column (Amersham, Piscataway, NJ). Approximately 0.2 pmol of the radiolabeled probe is incubated with 10 mg of nuclear lysate in a 20-ml reaction mixture containing 10 mM Tris-Cl (pH 7.5), 1 mM MgCl2, 1 mM EDTA, 2 mM DTT (dithiothreitol), 1:200 PIC, and 0.05 mg/ml herring sperm DNA (Invitrogen, Carlsbad, CA) for 20 min at 4 room temperature. The samples are then resolved in a 5% nondenaturing polyacrylamide gel. To verify that changes in probe mobility are due to binding by HIF, antibodies to one of the subunits can be added to the reaction mixture to form a larger complex with a greater mobility shift (supershift).

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The co-IP and EMSA assays can be performed with recombinant protein preparations, though the latter requires the presence of the bHLH domains for DNA-binding; however, we caution that it is important to thoroughly characterize recombinant PAS domain-containing proteins with respect to folding. Careful examination of several HIF-1a and -2a PAS-A–containing constructs expressed in E. coli has revealed that HIF-a PAS-A domains, while soluble, are often unfolded (unpublished data). If the HIF PAS domains do act cooperatively, as discussed later, an unfolded PAS-A domain may complicate interpretation of PAS-A and PAS-B function in vitro. Tellingly, we have found that consequences of deletions/mutations to recombinant protein preparations in the co-IP and EMSA assays often do not correlate well with the effects of those same deletions/mutations assayed in the context of HIF proteins residing in nuclear lysates. Because data obtained with HIF from nuclear lysates better predict functional consequences in intact cells, one should proceed with caution when using poorly characterized recombinant protein preparations that may not accurately recapitulate biologically meaningful interactions.

5. Discussion Over the past decade, much has been learned regarding the structure and function of various PAS domains. Structural studies of PAS-containing proteins have typically emphasized isolated PAS domains for a number of reasons, apart from a limited number of structures that detail PAS:PAS or PAS:protein interactions. In contrast, in vitro and cellular function studies have focused on intact proteins or larger constructs. By using approaches such as those already described, a framework for understanding how PAS domains contribute to HIF function is emerging. These approaches rely on integrated studies of the molecular mechanisms of PAS function, relating structural observations to in vitro properties to behaviors in the context of multi-domain proteins within intact cells. The solvent-exposed surface of the central b-sheet appears to be frequently used by PAS domains in other proteins to mediate protein– protein interactions. Indeed, the PAS domains within HIF-a and -b serve this function, providing interactions between subunits that are critical for the formation of active heterodimers in vitro and in living cells (Erbel et al., 2003; Yang et al., 2005). Structural studies of PAS domains from HIF-a and -b indicate that these interactions involve residues on an exposed b-sheet surface, commonly used among many PAS domains from different proteins (Card et al., 2005; Erbel et al., 2003). Interestingly, the data reveal an antiparallel arrangement of the HIF PAS-B domains, likely maintained by electrostatic complementarity between the subunits (Erbel et al., 2003).

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A variety of models are compatible with this antiparallel arrangement maintained within an HIF heterodimer, particularly given the flexibility afforded by the linkers between the two PAS domains in each subunit (see Fig. 1.2). Furthermore, based on precedents observed in recent structural data of complexes involving multiple PAS domains from the Per and LuxQ proteins (Neiditch et al., 2006; Yildiz et al., 2005), there may in fact be a wide range of intra- and intermolecular interactions among PAS domains in the HIF-a and HIF-b subunits. We view it as increasingly likely that all four PAS domains form a cooperative, intermolecular assembly (Fig. 1.5) necessary for HIF heterodimerization. This model is compatible with recent fluorescence resonant energy transfer (FRET) measurements collected from HIF proteins in living cells (Wotzlaw et al., 2006), suggesting a compact tertiary structure for the intact heterodimer. This model has important implications as well for PAS surfaces that mediate interactions between HIF and other proteins, such as coactivators and corepressors (Beischlag et al., 2004; Sadek et al., 2000; To et al., 2006). C-term C-term

HIFa PAS-B

HIFb PAS-B 17 residues

16 residues

PAS-A PAS-A 20 residues 22 residues

bHLH bHLH

Figure 1.5 Schematic model of hypoxia-inducible factors (HIF) domain interactions within the DNA-bound HIF heterodimer. Shown are the bHLH:DNA interactions, represented by the crystal structure of DNA-bound Max protein (Ferre-D’Amare et al., 1993), and the antiparallel heterodimer of HIF-a (gray) and HIF-b (black) Per/ARNT/ Sim (PAS)-B domains. As the structures of the HIF PAS-A domains have yet to be determined, they are represented schematically. (Reprinted from Card et al., 2005, with permission.)

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Lastly, given the absolute dependence on these PAS domain interactions for HIF function, it is appropriate to consider strategies that would target these domains for therapeutic purposes. For example, increased levels of HIF have been associated with tumor aggressiveness, resistance to conventional therapies, and mortality for a variety of human cancers through the induction of genes that promote metabolic adaptation, angiogenesis, changes in cell proliferation/survival, and cellular invasion and metastasis (Semenza, 2003). While a number of agents have been shown to antagonize HIF (Giaccia et al., 2003; Semenza, 2003), almost all are believed to act indirectly. As a consequence, their utility is often dependent on the mechanism of HIF induction while their therapeutic effects mediated through HIF can be difficult to distinguish from effects on other downstream pathways. Inspiration for a possible route to regulate PAS function in HIF comes from the observations that protein–protein interactions of a wide range of PAS domains are regulated by environmental stimuli, including light, oxygen, and redox state (Taylor and Zhulin, 1999). Sensitization to such diverse stimuli is achieved via small organic cofactors bound within the PAS domain, using changes in the configuration or occupancy of these ligands to alter protein structure and dynamics in ways that modulate function. Examples of this regulation are provided by photosensory PAS domains, which use light-driven bond formation or isomerization to break interactions between the sensory PAS domain and an effector helix (Harper et al., 2003; Rubinstenn et al., 1998). Such observations have led us to speculate that small organic compounds might be used to analogously disrupt an HIF heterodimeric complex. Although high-resolution HIF PAS-B structures have not revealed any internally-bound cofactors or preformed ligandbinding pockets, prior data from other PAS domains (e.g., hPASK [Amezcua et al., 2002]) indicate that this does not preclude ligand binding. Though protein–protein interactions are notoriously difficult to directly disrupt by small molecules, one can imagine that HIF PAS domains might bind small molecules within their cores, leading to conformational changes that propagate to the domain surface and disrupt association with other protein domains. Again, an experimental framework that relates domain structure to in vitro activity to in vivo function is crucial in identifying and evaluating potential PAS ligands (Park et al., 2006).

ACKNOWLEDGMENTS This work was supported by grants from the NIH (CA115962 to R.K.B.; CA90601 to K.H.G.; CA95471 to K.H.G. and R.K.B.) and the Robert A. Welch Foundation (I-1568 to R.K.B.). T.H.S. was supported by the American Cancer Society High Plains Division–North Texas Postdoctoral Fellowship. R.K.B. is the Michael L. Rosenberg Scholar in Medical Research and is supported by a Career Award in the Biomedical Sciences from the Burroughs Wellcome Fund.

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