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Mol Cell. Author manuscript; available in PMC Oct 8, 2008.
Published in final edited form as:
PMCID: PMC2563152
NIHMSID: NIHMS17236

RACK1 Competes with HSP90 for Binding to HIF-1α and is Required for O2-independent and HSP90 Inhibitor-induced Degradation of HIF-1α

Summary

Hypoxia-inducible factor 1 (HIF-1) regulates transcription in response to changes in O2 concentration. O2-dependent degradation of the HIF-1α subunit is mediated by prolylhydroxylase (PHD), the von Hippel-Lindau (VHL)/Elongin C/Elongin B E3 ubiquitin ligase complex, and the proteasome. Inhibition of heat shock protein 90 (HSP90) leads to O2/PHD/VHL-independent degradation of HIF-1α. We have identified the receptor of activated protein kinase C (RACK1) as a HIF-1α interacting protein that promotes PHD/VHL-independent proteasomal degradation of HIF-1α. RACK1 competes with HSP90 for binding to the PAS-A domain of HIF-1α in vitro and in human cells. HIF-1α degradation induced by the HSP90 inhibitor 17-allylaminogeldanamycin is abolished by RACK1 loss-of-function. RACK1 binds to Elongin-C and promotes ubiquitination of HIF-1α. Elongin-C binding sites in RACK1 and VHL show significant sequence similarity. Thus, RACK1 is an essential component of an O2/PHD/VHL-independent mechanism for regulating HIF-1α stability through competition with HSP90 and recruitment of the Elongin-C/B ubiquitin ligase complex.

Introduction

Oxygen homeostasis represents an essential organizing principle of metazoan evolution and biology. Hypoxia-inducible factor 1 (HIF-1) has been identified as a critical mediator of adaptive responses to reduced O2 availability in many developmental, physiological, and pathological contexts through its transcriptional regulation of genes that encode proteins required for tissue O2 delivery and energy metabolism (Hu et al. 2003; Manalo et al. 2005; Elvidge et al. 2006). HIF-1 is required for embryonic development in mice (Iyer et al. 1998; Ryan et al. 1998) and plays key roles in ischemic cardiovascular disease, stroke, and cancer (Melillo, 2004; Ran et al. 2005; Moeller and Dewhirst, 2006). Delineation of the mechanisms that regulate HIF-1 activity in these contexts has become a major challenge of contemporary molecular and cell biology.

HIF-1 is a heterodimeric transcription factor that consists of HIF-1α and HIF-1βsubunits (Wang et al., 1995). The amino-terminal half of each subunit consists of basic helix-loop-helix (bHLH) and PAS domains that mediate dimerization and DNA binding (Jiang et al., 1996a). HIF-1α protein is rapidly accumulated in hypoxia and degraded under non-hypoxic conditions (Wang et al. 1995; Jiang et al., 1996b; Salceda and Caro, 1997; Huang et al., 1998). The von Hippel-Lindau protein (VHL) binds both to HIF-1α and to Elongin-C, which in turn recruits Elongin-B and other subunits of an E3 ubiquitinligase, thus targeting HIF-1α for ubiquitination and degradation by the 26S proteasome (Salceda and Caro, 1997; Maxwell et al., 1999).

Hydroxylation of the 826-amino-acid human HIF-1α protein at proline residue 402 and/or 564 is required for VHL binding and subsequent degradation (Ivan et al., 3 2001; Jaakkola et al., 2001; Yu et al. 2001). Three prolyl hydroxylases (PHD1–3) were identified in mammalian cells and shown to utilize O2 and α-ketoglutarate as substrates to generate 4-hydroxyproline at P402 and/or P564 of HIF-1α (Bruick and McKnight 2001, Epstein et al., 2001). The HIF-1α transactivation domain is regulated by the binding of FIH-1 (factor inhibiting HIF-1; Mahon et al., 2001), which hydroxylates asparagine 803 to block binding of the coactivators CBP and p300 (Lando et al., 2002). HIF-2α, which is expressed in a more restricted distribution of cell types than HIF-1α, also heterodimerizes with HIF-1β and is regulated by O2-dependent asparagine and proline hydroxylation.

The levels of HIF-1α protein in normoxic tissues vary widely (Stroka et al., 2001), but the molecular basis for this regulation is unknown. HIF-1α degradation is regulated in an O2-independent manner by heat shock protein 90 (HSP90), a molecular chaperone that protects client proteins from misfolding and degradation (Neckers and Ivy, 2003; Whitesell and Lindquist, 2005). HSP90 binds to the HIF-1α PAS domain and the HSP90 inhibitors geldanamycin and 17-allylaminogeldanamycin (17-AAG) induce proteasomal degradation of HIF-1α even in renal carcinoma cells that lack functional VHL (Gradin et al. 1996; Isaacs et al. 2002, 2004; Mabjeesh et al. 2002).

In the present study, we identified the receptor for activated C-kinase 1 (RACK1) as a novel HIF-1α interacting protein through a proteomics-based screen. RACK1 was originally identified as an anchoring protein for activated protein kinase C (PKC) (Ron et al., 1994). However, RACK1 is now recognized as a multi-functional scaffold protein that plays an important role in diverse biological processes including intracellular signal transduction (McCahill et al., 2002) and assembly of the 80S ribosome from 40S and 60S subunits (Ceci et al., 2003). Here, we demonstrate that RACK1 competes with HSP90 for binding to HIF-1α, links HIF-1α to Elongin-C, and promotes HIF-1α degradation.

Results

We utilized a proteomics-based approach to identify proteins that bind to HIF-1α. HEK293 cell lysates were passed over a column containing Sepharose-4B covalently linked to glutathione-S-transferase (GST) or a fusion protein consisting of GST and residues 531–826 of human HIF-1α [GST-HIF-1α (531–826)]. Proteins that bound to GST or GST-HIF-1α (531–826) were eluted and labeled with the fluorescent dyes Cy3 and Cy5, respectively (Figure 1A). The labeled proteins were pooled and fractionated by 2-dimensional SDS-PAGE. The gel was scanned, 527 fluorescent spots were identified, and their Cy5/Cy3 ratios were calculated. A Cy5/Cy3 ratio ≥ 2.5 was chosen to distinguish specific binding to HIF-1α from non-specific binding to GST. Two spots that met this criterion were excised from the gel (Figure 1B). Matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometry was performed and the data were analyzed using a mass fingerprint search engine to query the NCBI Protein Database, which identified the two proteins as FIH-1 and RACK1 (Figure 1C). The identification of FIH-1, an established HIF-1α interacting protein (Mahon et al. 2001) validated this approach.

Figure 1
Identification of RACK1 as a HIF-1α Interacting Protein

In contrast to FIH-1, RACK1 was not previously known to interact with HIF-1α. To confirm the proteomics result, we performed in vitro binding assays. In vitro 5 transcribed and translated RACK1 bound to GST-HIF-1α (531–826), but not to GST alone (Figure 1D). GST-HIF-1α (531–826) also bound to endogenous RACK1 from HEK293 cell lysates (Figure 1E). Co-immunoprecipitation (co-IP) of RACK1 and endogenous HIF-1α from desferrioxamine-treated cells (Figure 1F) demonstrated that RACK1 also interacts with HIF-1α in human cells.

To investigate the functional consequences of RACK1 binding to HIF-1α, HEK293T cells were cotransfected with vectors encoding FLAG-tagged full-length HIF-1α and T7-tagged RACK1. FLAG-HIF-1α protein levels decreased in a dose-dependent manner as T7-RACK1 protein levels were increased (Figure 2A). The anti-RACK1 antibody (Ab) detected both T7-tagged RACK1 and endogenous RACK1, which also served as a loading control. Transfection of T7-RACK1 vector markedly decreased the levels of endogenous HIF-1α protein induced by exposure of HEK293T cells to 1% O2 (Figure 2B). HIF-2α protein levels were also decreased by T7-RACK1 under both 20% and 1% O2 (Figure 2C).

Figure 2
RACK1 Inhibits HIF-1α Protein Expression by a Mechanism that is Independent of Prolyl Hydroxylation but Dependent on Proteasome Activity

HIF-1α is degraded in an O2-dependent manner through the activity of PHD2, which hydroxylates proline residues 402 and 564 (Bruick and McKnight, 2001; Epstein et al. 2001; Berra et al., 2003), and VHL, which binds to hydroxylated HIF-1α and promotes its ubiquitination and subsequent proteasomal degradation (Maxwell et al. 1999; Tanimoto et al. 2000). RCC4 renal carcinoma cells lack functional VHL and have high endogenous HIF-1α protein levels (Maxwell et al. 1999; Hu et al. 2003). In RCC4 cells that were transduced with a retrovirus encoding RACK1, HIF-1α protein levels were greatly decreased compared to cells transduced with a retrovirus encoding GFP or uninfected parental cells (Figure 2D).

To analyze the effect of RACK1 on HIF-1 transcriptional activity, cells were cotransfected with FLAG-HIF-1α expression vector and p2.1, a reporter plasmid that contains a hypoxia response element (HRE) upstream of SV40 promoter and firefly luciferase coding sequences (Semenza et al. 1996). Transcriptional activity mediated by FLAG-HIF-1α was also inhibited in a dose-dependent manner by cotransfection of T7-RACK1 vector (Figure 2E, middle columns). Therefore, RACK1 reduces HIF-1α protein levels and, by doing so, reduces HIF-1 transcriptional activity.

Full-length HIF-1α protein containing the proline-to-alanine substitutions P402A/P564A is resistant to PHD2-mediated hydroxylation and VHL-mediated ubiquitination/degradation. Similar to its effect on wild-type FLAG-HIF-1α (Figure 2F, lanes 2 and 3), RACK1 decreased FLAG-HIF-1α (P402A/P564A) protein levels (Figure 2F, lanes 6 and 7) and transcriptional activity mediated by the double mutant protein (Figure 2E, right columns). In contrast, PHD2 inhibited transcriptional activity mediated by wild-type FLAG-HIF-1α but not by FLAG-HIF-1α (P402A/P564A) (Figure 2E).

Expression vector encoding FLAG-tagged bacterial alkaline phosphatase was also transfected into the cells and the levels of FLAG-BAP were not affected by RACK1 (Figure 2F, middle panel). FLAG-BAP and FLAG-HIF-1αcoding sequences were inserted into the same plasmid backbone to control for any potential effects of RACK1 on transcription or translation of the tagged proteins, demonstrating that RACK1 specifically induces the degradation of FLAG-HIF-1α. In support of this conclusion, RACK1-induced degradation of FLAG-HIF-1α was blocked by addition of the proteasome inhibitor MG132 (Figure 2F, compare lanes 3 and 5). Taken together, the data in Figure 2 demonstrate that RACK1 induces degradation of HIF-1α that is independent of prolyl hydroxylation and VHL function but dependent on proteasome activity.

To complement our studies investigating the effect of RACK1 gain-of-function, we also performed an analysis of RACK1 loss-of-function by expressing a short hairpin RNA (shRNA) directed against RACK1 or a scrambled negative control (SNC) shRNA. In HEK293T cells, shRNA-RACK1 decreased levels of RACK1 mRNA (Figure 3E, 3F) and protein (Figure 3A, 3C). shRNA-RACK1 increased FLAG-HIF-1α protein levels in a dose-dependent manner and there was a remarkable negative correlation between the levels of RACK1 and FLAG-HIF-1α in multiple experiments (Figure 3A and data not shown). shRNA-RACK1 also increased endogenous HIF-1α protein levels at both 20% and 1% O2 (Figure 3C). Transcription of the HRE reporter gene mediated by overexpressed FLAG-HIF-1α (Figure 3B) or endogenous HIF-1α (Figure 3D) was increased in cells co-transfected with shRNA-RACK1 at both 20% and 1% O2. HIF-1α mRNA levels were not affected by shRNA-RACK1, whereas the levels of mRNAs encoded by the HIF-1 target genes VEGF and GLUT1 were increased by shRNA-RACK1 but not by shRNA-SNC or empty vector, as determined by conventional RT-PCR (Figure 3E) and quantitative real-time RT-PCR (Figure 3F). The data demonstrate that RACK1 loss-of-function increases endogenous HIF-1α protein levels and the expression of HIF-1 target genes.

Figure 3
RACK1 Knockdown by RNA Interference Increases HIF-1α Protein Levels and HIF-1 Transcriptional Activity

RACK1 was originally identified as a platform to bring together activated PKC and its substrates (Ron et al. 1994; McCahill et al. 2002). To investigate whether PKC was involved in the RACK1-mediated inhibition of HIF-1α, we used RO-32-0432, which inhibits all known PKC isoforms with an IC50 ≤ 0.1 μM (Wilkinson at al., 1993). Exposure of HEK293T cells to 5 μM RO-32-0432 had no effect on the RACK1-mediated degradation of HIF-1α protein (data not shown). Although PKC-ε was reported to interact with HIF-1α in mouse heart (Ping et al. 2001), we found that expression of constitutively active PKC-ε had no effect on HIF-1α protein levels and HIF-1 transcriptional activity in HEK293T cells (data not shown).

Exposure of cells to O2 induces PHD/VHL-dependent degradation of HIF-1α. In contrast, the HSP90 inhibitors geldanamycin and 17-allylaminogeldanamycin (17-AAG) induce degradation of HIF-1α that is independent of PHD/VHL activity. HSP90 binding stabilizes HIF-1α and disruption of HSP90-HIF-1α interaction by geldanamycin or 17-AAG results in HIF-1α degradation even in cells without functional VHL (Isaacs et al., 2002, 2004; Mabjeesh et al., 2002). We analyzed purified GST-HIF-1α fusion proteins that contain different domains of HIF-1α for their binding to endogenous RACK1 and HSP90 present in HEK293T cell lysate. GST-HIF-1α (1–329) bound strongly both to HSP90 and RACK1, whereas GST-HIF-1α (429–608) bound to neither HSP90 nor RACK1, and both GST-HIF-1α(575–786) and (786–826) bound weakly to RACK1 (Figure 4A). Thus, although our initial identification of RACK1 was based on its interaction with HIF-1α residues 531–826, RACK1 shows strongest interaction with residues 1–329. GST-RACK1 did not bind to HSP90 (Figure 4B, lane 1), but it inhibited the binding of HSP90 to GST-HIF-1α (1–329) in a dose-dependent manner (lanes 2–4).

Figure 4
RACK1 Competes with HSP90 for Binding to HIF-1α

If RACK1 competes with HSP90 for binding to GST-HIF-1α(1–329), then a truncated form of HIF-1α containing only residues 1–329 may be subject to RACK1-mediated degradation. T7-RACK1 did in fact decrease FLAG-HIF-1α (1–329) protein levels in cotransfected cells (Figure 4C, compare lanes 2 and 3). The proteasome inhibitor MG132 blocked RACK1-mediated degradation of FLAG-HIF-1α(1–329) (compare lanes 3 and 5). The levels of FLAG-HIF-1α (1–329) protein were not regulated in response to changes in the cellular O2 concentration (lanes 1 and 2), which is mediated through HIF-1α residues 400–600 (Huang et al. 1998). Thus, RACK1 competes with HSP90 for binding to HIF-1α (1–329) and this competition is sufficient to destabilize HIF-1α in an O2-independent manner.

To further investigate the competition between RACK1 and HSP90, we divided GST-HIF-1α (1–329) into smaller domains and studied their binding. HIF-1α residues 81–200 and 81–329 bound to both RACK1 and HSP90, whereas residues 1–80 and 201–329 bound to neither RACK1 nor HSP90 (Figure 4D). Thus, residues 81–200 represent the minimal sequence analyzed that binds to RACK1 and HSP90. GST-HIF-1α (81–200) bound to HSP90 from HEK293T cell lysate and the addition of GST-RACK1 decreased HSP90 binding in a dose dependent manner (Figure 4E). Reciprocally, the addition of GST-HSP90 dose-dependently decreased RACK1 binding to GST-HIF-1α 81–200) (Figure 4F). These data indicate that RACK1 and HSP90 compete for binding to residues 81–200, which encompass the PAS-A subdomain of HIF-1α (Wang et al. 1995).

To determine whether the competitive binding of RACK1 and HSP90 to HIF-1α was relevant to the mechanism of action of 17-AAG, we analyzed protein-protein interactions in 17-AAG-treated cells. The 17-AAG-induced degradation of FLAG-HIF-1α (P402A/P564A) was blocked by the proteasome inhibitor MG132 (Figure 5A, left panel). Although FLAG-HIF-1α (P402A/P564A) levels were not affected by 17-AAG in the presence of MG132, co-IP of HSP90 by anti-FLAG Ab was decreased in 17-AAG-10 treated cells and co-IP of RACK1 was increased (Figure 5A, right panel). Decreased HSP90 and increased RACK1 binding to HIF-1α were observed when 17-AAG was added directly to cell lysate in vitro followed by IP of endogenous HIF-1α (Figure 5B).

Figure 5
HIF-1α Degradation Induced by 17-AAG is Dependent on RACK1 Binding

The increased interaction of RACK1 and HIF-1α in the presence of 17-AAG may simply reflect the loss of competition by HSP90 for binding to HIF-1α. Alternatively, RACK1 binding may be required for HIF-1α degradation in 17-AAG-treated cells. To distinguish between these models, HEK293T cells were cotransfected with FLAG-HIF-1α (P402A/P564A) and shRNA-RACK1 or shRNA-SNC. 17-AAG decreased FLAG-HIF-1α (P402A/P564A), but not HIF-1βor β-actin, protein levels when shRNA-SNC was used (Figure 5C, lane 2). However, the degradation of FLAG-HIF-1α (P402A/P564A) by 17-AAG was abolished in the presence of shRNA-RACK1 (lane 4). These results indicate that the mechanism of action of 17-AAG is dependent upon RACK1 activity.

To delineate the mechanism of RACK1-mediated degradation of HIF-1α, we investigated the connection between RACK1 and HIF-1α E3 ubiquitin ligase. VHL is the substrate recognition subunit that recruits HIF-1α to Elongin-C/B (Kamura et al. 2002; Min et al., 2002). Both GST-VHL and GST-RACK1 bound to in vitro translated Elongin-C (Figure 6A). Interaction of endogenous RACK1 and Elongin-C in cell lysates was demonstrated by co-IP (Figure 6B). GST-RACK1 increased the co-IP of FLAG-HIF-1α (1–329) and Elongin-C in a dose dependent manner (Figure 6C). In vitro translated Elongin-C recruited Elongin-B to RACK1 dose dependently (Figure 6D). These data indicate that RACK1 recruits HIF-1α to the Elongin-C/B subunits of E3 ubiquitin ligase. In further support of this model, ubiquitination of HIF-1α was increased by RACK1 in the presence of the proteasomal inhibitor MG132, similar to the effect of overexpressed PHD2 (Figure 6E, bottom panel).

Figure 6
RACK1 Promotes Elongin-C Binding to HIF-1α and Ubiquitination of HIF-1α

RACK1 is composed of 7 Trp-Asp-40 (WD-40) repeats and forms a homodimer through WD-40 repeat 4 (WD4; Thornton et al, 2004). Next we determined which WD-40 repeats in RACK1 bind to Elongin-C and HIF-1α. Both GST-RACK1-WD1-7, which contains all 7 WD-40 repeats, and GST-RACK1-WD567, which contains only the last 3 WD-40 repeats, bound to FLAG-HIF-1α (1–329) as well as to Elongin-C (Figure 6F). RACK1-WD1-7 also bound to HIF-2α (Figure S1). Further deletion revealed that WD56 or WD6 alone bound to both FLAG-HIF-1α (1–329) and Elongin-C (Figure 6G). Because RACK1 is a dimer, it may be able to co-recruit HIF-1α and Elongin-C even though they both bind to the WD6 repeat. Alternatively, HIF-1α and Elongin-C may bind to 2 distinct sequences within the 48-amino-acid WD6 domain, such that a RACK1 monomer can interact with both proteins simultaneously.

Since the WD6 domain of RACK1 and the α domain of VHL (Stebbins et al, 1999) both bind to Elongin-C, we aligned their sequences from multiple vertebrate species and identified a conserved domain of 22–24 residues with significant similarity (Figure 7A). These results provide a molecular basis for the observed ability of both VHL and RACK1 to recruit Elongin-C to HIF-1α.

Figure 7
Structural and Functional Similarities Between RACK1 and VHL

Discussion

RACK1 contains WD-40 repeats that form a 7-bladed propeller structure (Steele et al., 2001). RACK1 was originally discovered as a protein that can activate and translocate PKC (Ron et al., 1994). However, the multiple WD-40 repeats provide a molecular scaffold for the combinatorial interaction of various repeats with many different partners through which RACK1 influences a wide variety of cellular processes (McCahill et al., 2002). In this study, we demonstrated that RACK1 binds to HIF-1α both in vitro and in human cells. Overexpression of RACK1 leads to increased proteasomal degradation of HIF-1α protein that is independent of O2, proline hydroxylation, or VHL binding. RACK1 loss-of-function by RNA interference increases HIF-1α protein levels and promotes HIF-1 downstream target gene expression. Although we initially identified RACK1 through its interaction with residues 531–826, its strongest interaction is with the amino terminal half of HIF-1α and this interaction is sufficient to promote HIF-1α degradation. Further studies are required to determine whether RACK1 regulates other aspects of HIF-1α function through its binding to residues 531–826.

RACK1 competes with HSP90 for the binding to residues 81–200 of HIF-1α, which encompass the PAS-A domain (Wang et al. 1995). RACK1 recruits Elongin-C to HIF-1α through the WD6 domain, which has significant sequence similarity to the Elongin-C binding domain of VHL (Figure 7A). Elongin-C, in turn, recruits Elongin-B. Based on the extensive prior analyses of the VHL/Elongin-C/B complex (Stebbins et al., 1999; Kamura et al., 2002), it is likely that other subunits of the E3 ubiquitin ligase complex are also recruited, leading to the observed increase in ubiquitination and proteasomal degradation of HIF-1α in response to 17-AAG treatment or RACK1 over expression. Thus, by competing with HSP90 for binding to HIF-1α, RACK1 mediates a degradation pathway that is mechanistically similar to the VHL pathway with the critical difference that it is O2 independent (Figure 7B). In vitro assays of ubiquitination using purified proteins are required to determine whether RACK1 is sufficient to substitute for VHL within the previously defined (Kamura et al., 2002) E3 ubiquitin ligase that modifies HIF-1α. Alternatively, RACK1 may recruit additional proteins that are required for ubiquitin ligase activity.

Basal levels of HIF-1α vary among tissue and cell types (Stroka et al., 2001). Our results implicate RACK1 as a major determinant of the basal rate of HIF-1α protein degradation. Responses to changes in cellular O2 concentration that are mediated by the PHDs and VHL are superimposed upon this basal rate. Thus, competition between HSP90 and RACK1 for interaction with HIF-1α may contribute to establishment of the HIF-1α “set-point” in each cell type. Further studies are required to investigate whether specific developmental or physiological signals lead to qualitative or quantitative changes in HSP90, RACK1, or Elongin-C, which in turn alter the degradation of HIF-1α in an O2-independent manner. In particular, additional studies are required to determine whether the RACK-HIF-1α interaction is regulated by post-translational modification of either protein. We have demonstrated effects of RACK1 gain- and loss-of-function under both non-hypoxic and hypoxic conditions, suggesting that both VHL and RACK1 may contribute to the degradation of HIF-1α to varying degrees at physiological concentrations of O2. These results are consistent with the finding that the HSP90 inhibitor 17-AAG reduces HIF-1α levels in both non-hypoxic and hypoxic cells.

Preclinical studies have demonstrated that HIF-1 plays important roles in tumor angiogenesis, metabolism, and invasion/metastasis and clinical studies have demonstrated that increased HIF-1α protein levels in tumor biopsies are associated with increased patient mortality (Melillo, 2004). These findings have sparked interest in the identification of HIF-1 inhibitors, several of which have been shown to inhibit tumor xenograft growth in vivo, including 17-AAG. HSP90 inhibitors may be particularly potent anti-cancer agents because HSP90 is also required to prevent the degradation of many activated or overexpressed oncoproteins (Neckers and Ivy, 2003; Whitesell and Lindquist, 2005; Solit and Rosen, 2006). The HSP90 chaperone complex in cancer cells binds HSP90 inhibitors with 100-fold greater avidity relative to the HSP90 complex in non-transformed cells, which provides a therapeutic window to selectively target cancer cells (Kamal et al., 2003). The anti-cancer efficacy of 17-AAG is under investigation in phase II clinical trials (Sharp and Workman, 2006). Our data indicate that RACK1 is required for the action of 17-AAG as a HIF-1α inhibitor and RACK1 levels in cancer cells may be an important determinant of their sensitivity to HSP90 inhibitors.

The common binding of Elongin-C by VHL and RACK1 has uncovered a molecular mechanism by which an E3 ubiquitin ligase complex may be recruited to HIF-1α by O2-dependent and O2-independent specificity factors, respectively. Our studies have revealed a novel role for RACK1 in promoting the ubiquitination and degradation of HIF-1α and, likely, HIF-2α. Interaction with RACK1 is also required for proteasomal degradation of the transcription factor ΔNp63α in response to DNA damage (Fomenkov et al., 2004), suggesting that RACK1 may also recruit the Elongin-C/B ubiquitin ligase complex to ΔNp63α.

Experimental Procedures

Tissue Culture

HEK293, HEK293T, and RCC4 cells were cultured as described (Semenza et al. 1996; Hu et al. 2003). Cells were maintained at 37°C in a 5% CO2, 95% air incubator. For hypoxic exposures, cells were placed in a modulator incubator chamber (Billups-Rothenberg) flushed with 1% O2/5% CO2/balance N2 and incubated at 37°C.

Two Dimensional PAGE

HEK293 cells were lysed in 50 mM Tris-Cl (pH7.5), 150 mM NaCl, 0.1% NP40 with protease inhibitor cocktail (Roche). 112 mg of lysate was precleared with 1 ml of GST-coupled CNBr-activated Sepharose-4B column (GE Healthcare). The flow-through was divided in half and loaded onto 1-ml columns of GST- and GST-HIF-1α(531–826)-coupled Sepharose-4B, washed with 10 ml of lysis buffer, and eluted with 2 ml of 50 mM Tris-Cl (pH 7.5), 1 M NaCl, 8 M Urea. The eluate was concentrated using a Microcon 10K filter (Millipore), precipitated with trichloroacetic acid and acetone, and resuspended in 30 mM Tris-Cl (pH 8.5), 7 M urea, 2 M thiourea, 4% CHAPS. 50 μg of eluted protein from each column was labeled with 400 pmol of Cy3 or Cy5. 50 μg of labeled protein and 85 μg of unlabeled protein from each column were pooled together and loaded onto a 24-cm, pH 3–10 IPG strip (GE Healthcare), which was rehydrated for 10 hr actively at 50V. IEF was run overnight for 75 kVh. The IPG strip was reduced, alkylated, equilibrated in Laemmli buffer, and second-dimension SDS-PAGE (24-cm, 12% polyacrylamide gel in Tris-Glycine buffer) was performed at 2W overnight. Cy3 and Cy5 fluorescence images were obtained with a Typhoon scanner, processed with DeCyder 5.1 DIA software (GE Healthcare), and Cy5/Cy3 ratios were calculated. The gel was silver stained and spots of interest were excised.

Protein Identification by MALDI-TOF Mass Spectrometry

In gel digestion of spots excised from the gel was performed as described (Shevchenko et al., 1996; Gharahdaghi et al., 1999). Trypsin-digested peptides were mixed with 2,5-dihydroxybenzoic acid and loaded on a MALDI plate. Spectra were obtained using a Voyager DE-STR MALDI–TOF mass spectrometer (Applied Biosystems). The data were analyzed with Data Explorer 5.1 (Applied Biosystems). The mass lists were searched against protein database NCBInr.2006.02.16 using Protein Prospector MS-Fit v4.0.6 (http://prospector.ucsf.edu) with 50 ppm mass tolerance.

GST Pull-Down Assays

GST fusion protein was purified as described (Baek et al., 2005). [35S]-methionine-labeled proteins were generated in reticulocyte lysates using a T7-coupled transcription/translation system (Promega). Whole cell lysates (WCL) were prepared in 50 mM Tris-Cl (pH 7.5), 150 mM NaCl, 0.1% NP40 and protease inhibitor cocktail. Unless otherwise specified, 5 μg of GST-HIF-1α fusion protein and 800 μg WCL were incubated at 4°C for 3 hr. 20 μl of glutathione-Sepharose-4B beads were added to the samples and incubated at 4°C for 1 hr to capture the GST fusion proteins. After washing with lysis buffer three times, the proteins were eluted in Laemmli buffer and analyzed by SDS-PAGE followed by immunoblot (IB) assay.

Transfection Assays

HEK293T cells were seeded onto 48-well plates and transfected with plasmid DNA using Fugene-6 (Roche). Control reporter pSV-Renilla (1 ng), HRE reporter p2.1 (10 ng), and other expression vectors were used. Cells were lysed and luciferase activities were determined by multi-well luminescence reader (PerkinElmer), using the Dual-Luciferase Reporter Assay System (Promega). Three independent transfections were performed. For IB assays, HEK293T cells were seeded onto 6-cm or 10-cm culture dishes. The following day, the cells were cotransfected with expression vectors using Fugene-6. 24 hr after transfection, the cells were exposed to 20% or 1% O2 in the presence or absence of 10 μM MG132 for 4 hr and lysed in RIPA buffer with protease inhibitor cocktail (Roche).

IP and IB Assays

Cells were lysed in 50 mM Tris-Cl (pH 7.5), 150 mM NaCl, 0.1% NP40, 1 mM DTT, protease inhibitor cocktail, sodium orthovanadate, and sodium fluoride. 2 μg of Ab or control IgG was incubated with 1 mg of WCL overnight at 4°C. To stabilize HSP90 protein interactions, 20 mM sodium molybdate was added. 20 μl of protein G-Sepharose (GE Healthcare) was added to the samples and incubated for 2 hr at 4°C to capture IgG. Beads were washed 3 times with lysis buffer. Proteins were eluted in Laemmli sample buffer and analyzed by SDS-PAGE. Antibodies used in IB assays were: FLAG (Sigma); RACK1 (BD); GST (GE Healthcare); Ubiquitin (MBL); β-actin, Elongin-C, and HSP90 (Santa Cruz); and HIF-1α, HIF-1β, HIF-2α, and PHD2 (Novus Biologicals).

ShRNA Assays

The vector pSR.retro.GFP.Neo.circular.stuffer (OligoEngine) was used for expression of shRNA in HEK293T cells. The shRNA-RACK1 insert was a previously published 19-nucleotide sequence (Lopez-Bergami et al., 2005). shRNA-SNC was previously described (Baek et al., 2005). Oligonucleotides were annealed and ligated into BglII- and HindIII-digested vector.

RT-PCR Assays

Total RNA was extracted from HEK293T cells using Trizol (Invitrogen) and treated with DNase I (Ambion). One microgram of total RNA was used for first-strand synthesis with iScript cDNA Synthesis system (BioRad). Real-time PCR was performed by using IQ SYBR Green Supermix and the iCycler Real-time PCR Detection System (BioRad). Expression of RACK1 mRNA relative to 18S rRNA was calculated based on the threshold cycle (CT) for amplification as 2−(ΔCT), where ΔCT=CT,RACK1 − CT,18S.

Statistical analysis

Data are presented as mean ± SEM. Differences between experimental conditions were analyzed for statistical significance (P < 0.05) by Student’s t-test.

Supplementary Material

Acknowledgments

We thank Karen Padgett (Novus Biologicals) for providing antibodies against HIF-2α and PHD2; Alexey Fomenkov and Edward Ratovitski (Johns Hopkins University) for RACK1 expression vector; Peipei Ping (UCLA) for PKC-ε vector; Jaime Caro (Thomas Jefferson University) and Peter Ratcliffe (Oxford University) for PHD2 vector; and Joan Conaway (Stowers Institute for Medical Research) for Elongin-C and -B vectors.

Footnotes

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