Iron-dependent degradation of IRP2 requires its C-terminal region and IRP structural integrity
© Wang et al; licensee BioMed Central Ltd. 2008
Received: 18 September 2007
Accepted: 28 January 2008
Published: 28 January 2008
Iron regulatory protein 2 (IRP2), a post-transcriptional regulator of cellular iron metabolism, undergoes iron-dependent degradation via the ubiquitin-proteasome pathway. A stretch of 73 amino acids within the N-terminal domain 1 of the protein was reported to function as an iron sensor. However, mutants lacking this fragment remain sensitive to degradation in iron-replete cells.
To identify elements within IRP2 involved in the control of its stability, we undertook a systematic mutagenesis approach. Truncated versions of IRP2 were expressed in H1299 cells and analyzed for their response to iron. Deletion mutants lacking the entire C-terminal domain 4 (amino acids 719–963) of IRP2 remained stable following iron treatments. Moreover, the replacement of domain 4 of IRP1 with the corresponding region of IRP2 sensitized the chimerical IRP11–3/IRP24 protein to iron-dependent degradation, while the reverse manipulation gave rise to a stable chimerical IRP21–3/IRP14 protein. The deletion of just 26 or 34 C-terminal amino acids stabilized IRP2 against iron. However, the fusion of C-terminal IRP2 fragments to luciferase failed to sensitize the indicator protein for degradation in iron-loaded cells.
Our data suggest that the C-terminus of IRP2 contains elements that are necessary but not sufficient for iron-dependent degradation. The functionality of these elements depends upon the overall IRP structure.
Iron regulatory proteins, IRP1 and IRP2, post-transcriptionally control the expression of several mRNAs bearing iron responsive elements (IREs). In iron-deficient cells, IRE/IRP interactions account for the stabilization of transferrin receptor 1 (TfR1) mRNA and the translational inhibition of ferritin (H- and L-) mRNAs, resulting in increased uptake and reduced sequestration of iron . IRPs regulate the expression of additional IRE-containing transcripts, such as those encoding erythroid aminolevulinate synthase (ALAS2), mitochondrial aconitase, the iron transporter ferroportin 1, myotonic dystrophy kinase-related Cdc42-binding kinase α (MRCK α), hypoxia inducible factor 2 α (HIF2α), and splice variants of the divalent metal transporter DMT1 and the kinase Cdc14A [2–4]. Experiments with IRP1-/- and IRP2-/- cells and animals revealed that IRP2 exerts a dominant regulatory function in vivo .
Both IRP1 and IRP2 share significant sequence similarity [1, 2, 5]. A major difference in their primary structure is that IRP2 contains a unique insertion of 73 amino acids close to its N-terminus (referred to hereafter as 73d). In iron-replete cells, IRP1 binds a cubane 4Fe-4S cluster, which precludes IRE-binding, renders the protein to a cytosolic aconitase and maintains it in a closed conformation [6, 7]. Under these conditions, IRP2 undergoes rapid ubiquitination and degradation by the proteasome [1, 2, 5]. Phosphorylation or defects in Fe-S cluster assembly may also sensitize IRP1 to iron-dependent proteasomal degradation, albeit with slower kinetics compared to IRP2 [8–10].
The mechanism for IRP2 degradation is far from being understood. It has been proposed that the 73d functions as an "iron-dependent degradation domain". One model postulates that the iron-sensing capacity of the 73d is based on site-specific oxidation of conserved cysteine residues upon direct iron binding . Another model suggests that IRP2 degradation is triggered by oxidative modification following high affinity binding of heme within the 73d [12, 13]. Nevertheless, experiments in cultured cells showed that IRP2 deletion mutants lacking the entire 73d remain as sensitive to iron as wild type IRP2 [14–16]. Moreover, the 73d failed to destabilize GFP fusion indicator constructs in iron-loaded cells , casting further doubt on its proposed function as a necessary and sufficient regulatory element for IRP2 degradation. Recent results showed that 73d is sensitive to proteolytic cleavage and that heme binding only occurs in its truncated form .
IRP2 is stabilized in response to hypoxia [14, 18, 19], by analogy to HIF α subunits that play a crucial role in cellular adaptation to low oxygen levels . Under normoxic conditions, HIF α subunits undergo post-translational modification by the prolyl-hydroxylases PHD1–3, which tag them for ubiquitination by the E3 ubiquitin ligase VHL and degradation by the proteasome . These enzymes, as well as other 2-oxoglutarate-dependent dioxygenases, catalyze the hydroxylation of protein substrates by using 2-oxoglutarate. The reaction yields a hydroxylated amino acid, succinate and carbon dioxide, and proceeds via an iron-oxo intermediate . The availability of ferrous iron, oxygen and ascorbate (presumably to maintain iron in a reduced state) is critical for catalysis.
Experimental evidence supports a mechanism for IRP2 degradation via 2-oxoglutarate-dependent dioxygenases. Thus, dimethyl-oxalyl-glycine (DMOG), a substrate analogue of 2-oxoglutarate, protected IRP2 against iron-dependent degradation [14, 15]. Furthermore, ascorbate and other antioxidants accelerated the degradation of IRP2 . Nevertheless, the protective effect of DMOG was restricted to cells that were previously depleted from iron by the chelator desferrioxamine, indicating that the iron-dependent degradation of IRP2 is mediated by multiple mechanisms. One of them appears to interfere with the heme biosynthetic pathway, as succinyl-acetone, an inhibitor of endogenous heme synthesis, stabilized IRP2 against iron ; this result has independently been reproduced in various laboratories [13, 16, 19, 24]. The E3 ubiquitin ligases VHL  and HOIL-1  are not required for IRP2 degradation in response to iron.
To identify structural elements within IRP2 that may play a role in iron sensing and in the regulation of its stability, we undertook a systematic deletion analysis. Our data suggest that C-terminal sequences are necessary for the iron-dependent degradation of IRP2. However, these sequences only operate in a context of IRP structure and do not have intrinsic iron sensing properties.
The C-terminal domain of IRP2 is necessary for its iron-dependent degradation
The C-terminal domain 4 of IRP2 sensitizes IRP11–3-IRP24 chimeras for iron-dependent degradation
The swapping of domains 4 between IRP1 and IRP2 impinges on the IRE-binding properties of the chimeras
Truncation of the C-terminal domain 4 of IRP2 to identify minimal iron-sensing sequences
Even though the peptides encompassing the 60 C-terminal amino acids of IRP1 and IRP2 are largely conserved (Additional File 1A), they display notable differences and cluster separately in a phylogenetic dendrogram (Additional File 1B). For example, the IRP2 C-terminus is serine-rich, whereas serines are completely absent from the respective segment of mammalian IRP1s. We considered IRP2-specific amino acids in this region as potential regulatory sites and hypothesized that their replacement with IRP1-specific counterparts might confer an IRP1-like phenotype (stabilization in iron-loaded cells). Thus, we focused on non-conserved amino acids that differ significantly (in terms of chemical side chains) between IRP2 and IRP1 and generated IRP2 variants with S929D, S939R and L948Y IRP1-like point mutations. However, none of these manipulations resulted in stabilization of the IRP2 mutants against iron (Additional File 1C).
The sensitivity of IRP2 to iron has been known for years . Nevertheless, the mechanism for its iron-dependent degradation remains unresolved and, moreover, little progress has been made towards identifying and characterizing IRP2 sequences with iron-sensing properties. The IRP2-specific 73d has been proposed to function as an iron sensor either via direct binding of iron to C168, C174 and C178 , or binding of heme to C168  or C201 . Nevertheless, several labs have demonstrated that the above cysteine residues and the entire 73d are dispensable for IRP2 degradation in cultured cells [14, 15, 19], even though published data with opposing views exist . We reasoned that an unbiased systematic deletion analysis of IRP2 might shed light onto this issue and facilitate the identification of segments of the protein with iron-sensing capacity in vivo. To this end, we generated several IRP2 deletion mutants and transfected them into H1299 cells for tetracycline-inducible expression. To avoid saturation effects related to overexpression , we selected and analyzed multiple clones with variable IRP2 expression levels for their response to iron treatment. We were unable to isolate stable clones expressing the N-terminal deletion mutant ΔD1; the reason for this is unclear.
The truncation of IRP2 at its C-terminus was sufficient for the stabilization of the protein in iron-loaded cells, independently of the presence or absence of 73d (Fig. 1). These findings not only confirmed that 73d fails to function as an iron sensor in vivo, but also shifted our focus for the identification of iron-sensing sequences to the C-terminal domain 4 of IRP2. Further support for the significance of this region in the control of IRP2 stability was provided by experiments with IRP1/IRP2 chimeras, generated by swapping the domains 4 between these proteins (Figs. 2, 3, 4). Thus, domain 4 of IRP2 accelerated the decay of the IRP11-3-IRP24 chimera in iron-loaded cells (Fig. 3C). The stability of ΔD4 IRP1 against iron (Fig. 2C) denotes that this effect is not due to merely the loss of the IRP1 C-terminal region. Along these lines, domain 4 of IRP1 abolished the iron responsiveness of the IRP21–3-IRP14 chimera (Fig. 3D). Collectively, these data suggest that the C-terminus of IRP2 is necessary for its iron-dependent degradation. It should, however, be noticed that the response of the IRP11–3-IRP24 chimera to iron was modest, compared to wild type IRP2 (Fig. 3B), indicating the involvement of additional IRP2 sequences, outside domain 4, in controlling the rate of its degradation.
The iron-sensitive IRP11–3-IRP24 chimera retained RNA-binding activity (Fig. 4A) and was functional in regulating downstream IRE-containing mRNA targets, as shown by the increase in TfR1 expression (Fig. 4B). By contrast, the apparently labile IRP21–3-IRP14 chimera was resistant to iron and failed to bind RNA (Fig. 4C), raising the possibility that the iron responsiveness may be linked to the RNA-binding capacity. We did not directly address this issue; nevertheless, the recent description of other iron-sensitive IRP1/IRP2 chimeras that are inactive in RNA-binding suggests that this is not the case . Considering that the 4Fe-4S cluster is necessary to maintain IRP1 in a compact structure [6, 7] and a failure to assemble it sensitizes the protein for slow iron-dependent degradation [8–10], it will be of interest to examine whether the chimerical proteins retain a capacity to assemble a 4Fe-4S cluster, and whether this correlates with their responsiveness to iron.
In an attempt to narrow down the putative iron-sensing region of IRP2, we further truncated domain 4 of the protein and established that the deletion of just small C-terminal segments abolished iron-dependent degradation of IRP2 (Fig. 5). However, the fusion of 60 or 168 C-terminal amino acids of IRP2 to luciferase failed to promote the degradation of this chimerical protein in iron-loaded cells (Fig. 6).
Conceivably, the observed stabilization of the IRP2 deletion mutants is related to a disruption of the overall IRP2 structure that may prevent the exposure of iron-sensing regions. Alternatively, the C-terminal domain of IRP2 may contribute to an iron-dependent rearrangement of the protein structure, which would render it more sensitive to proteolysis. In this scenario, the actual iron sensor could be another factor, such as a proteolytic enzyme, that may attack the destabilized protein upon activation by iron.
Our results suggest that sequences within the C-terminus of IRP2 are necessary, but not sufficient for iron-dependent degradation, at least outside an IRP context. Thus, the C-terminus of IRP2 does not possess the properties of a bona fide iron sensor but its regulatory function requires IRP structural integrity and very likely additional elements within other domains of IRP2 and/or other factors.
Hemin and ferric ammonium citrate were purchased from Sigma (St. Louis, MI).
Construction of IRP2 mutants
IRP2 deletion or point mutants were generated from either the pUHD-IRP2 or the pUHD-IRP2Δ73 plasmids  with the ExSite mutagenesis kit (Stratagene). The forward primer for PCR amplification to yield all C-terminal deletion mutants was 5'-TACCCATACG ATGTTCCAGA TT-3'. The reverse primers were: i) for ΔD4 and ΔD4/-73, 5'-TAAGTCCCAT GGAAACAAAA CT-3'; ii) for ΔD3,4 and ΔD3,4/-73: 5'-TATTGAATTC AGATTAATCT GG-3'; iii) for ΔD2,3,4: 5'-TAAAGTAAGA GAAACTGGCA GA-3'; iv) for ΔC168: 5'-TGTCATTACA GCATCATTAC CT-3'; v) for ΔC60: 5'-GCCCAAGGAA TCTGCATTTT CT-3'; and vi) for ΔC34: 5'-GCTTGTCTGT ATATTCAATG TA-3'. The ΔD1 deletion mutant was generated with 5'-GAGGGATCCA CCATGGGCCC AGAGGTGGTT GGATGT-3' (forward) and 5'-GTGGCACGAA AATTCTCATA CCCATACGAT GTTCCAGATT ACGCTTAGTA AGGATCCTAC G-3' (reverse) primers. The ΔC26 deletion mutant was generated with 5'-ACTGGAAAAG TATTCAGCGT GA-3' (forward) and 5'-GCCCAAGGAA TCTGCATTTT CT-3' (reverse) primers. IRP2 point mutants were generated with following sets of primers: i) for S929D: 5'-AATATACAGA CAGATACTGG AAAAGTA-3' (forward) and 5'-TACTTTTCCA GTATCTGTCT GTATATT-3' (reverse); ii) for S939R: 5'-AGCGTGATTG CTCGATTTGA AGATGAT-3' (forward) and 5'-ATCATCTTCA AATCGAGCAA TCACGCT-3' (reverse); and iii) for L948Y: 5'-TGGAAATAAC ATACTACAAA CATGGA-3' (forward) and 5'-TCCATGTTTG TAGTATGTTA TTTCCA-3' (reverse). To generate the ΔD4 IRP1 deletion mutant, pUHD-IRP1  was digested with MscI/SnaBI (thereby excising domain 4) and religated.
Construction of IRP1/IRP2 chimeras
A hybrid molecule in which the domain 4 of IRP1 was replaced by that of IRP2 (IRP11–3IRP24) was generated as follows: The region spanning domains 1–3 of IRP1 was PCR-amplified from pUHD-IRP1  with 5'-AGTGTGGGAT CCTGTACAAC CAGG-3' (forward) and 5'-AGTGTGCTCG AGTGCCTGGA TCTCGTCTCT AGT-3' (reverse) primers. Domain 4 of IRP2 was PCR-amplified from pUHD-IRP2 with 5'-AGTGTGCTCG AGGAAGAACA TGTTATA-3' (forward) and 5'-CGTAGGATCC TTACTAAGCG TAATCTGGAA CATCGTATGG GTATGAGAAT TTTCGTGCCA C-3' (reverse) primers. Finally, the two fragments were cloned into the BamH1 site of pUHD-IRP1. Likewise, a hybrid molecule in which the domain 4 of IRP2 was replaced by that of IRP1 (IRP21–3IRP14) was generated by PCR-amplification of the region spanning domains 1–3 of IRP2 from pUHD-IRP2 with 5'-GCTCGAGGAT CCCATGGACG CCCC AAA-3' (forward) and 5'-CTGACGCTCG AGTGCATGAA CTTCTTCTCG ACT-3' (reverse) primers. Domain 4 of IRP1 was PCR-amplified from pUHD-IRP1 with 5'-CAGGCACTCG AGCGTCAGTA TGTCATC-3' (forward) and 5'-AGCGTAGGAT CCTTACTAAG CGTAATCTGG AACATCGTAT GGGTACTTGG CCATCTTGCG GAT-3' (reverse), and the two fragments were cloned into the BamH1 site of the pUHD10-3 vector .
Construction of luciferase fusion indicators
A common reverse primer was employed to amplify C-terminal sequences of IRP2, including the hemagglutinin (HA) tag and a stop codon, from pUHD-IRP2: 5'-AGTGTGCTCG AGCTAAGCGT AATCTGGAAC-3'. The forward primers were: i) for C168: 5'-AGTGTGGAAT TCAGAGGCAC TTTTGCAAAT-3' and for C60: 5'-AGTGTGGAAT TCCTCTCCGG TAGAGAAACA-3'. The primers introduce EcoR1 and Xho1 restriction sites. The resulting fragments were subcloned into the respective sites of a firefly luciferase cDNA in the pcDNA3.1 vector.
H1299 lung cancer cells were maintained in Dulbecco's modified Eagle medium (DMEM) supplemented with 10% fetal bovine serum, 2 mM glutamine, 100 U/ml penicillin, and 0.1 mg/ml streptomycin. Stable H1299 clones for tetracycline-inducible expression of all IRP2 mutants and IRP1/IRP2 chimeras and for non-inducible expression of luciferase fusion constructs were obtained as in [15, 29]; the clones expressing wild type IRP1 [8–10] and IRP2  were earlier described. All inducible clones were maintained in DMEM containing 2 μg/ml tetracycline, 2 μg/ml puromycin and 250 μg/ml G418. The cells were plated for 24–48 h in tetracycline-free media to allow expression of transfected proteins. Tetracycline (2 μg/ml) was then added back to shut off transcription of the inducible promoters.
Cells were washed twice in phosphate-buffered saline (PBS) and lysed in "cytoplasmic lysis buffer" (25 mM Tris-Cl pH 7.4, 40 mM KCl and 1% Triton X-100). Cell debris was cleared by centrifugation and protein concentration was determined with the Bradford reagent (BioRad). Cell lysates (30 μg) were resolved by SDS-PAGE on 8% gels and proteins were transferred onto nitrocellulose filters. The blots were saturated with 10% non-fat milk in PBS and probed with FLAG (M2-FLAG, Sigma), HA (Santa Cruz), TfR1 (Zymed), luciferase (Promega) or β-actin (Sigma) antibodies, diluted 1:1000 (or 1:500 for luciferase) in PBS containing 5% non-fat milk and 0.5% Tween-20 (PBST). Following wash with PBST, the blots with monoclonal FLAG or TfR1 antibodies were incubated with peroxidase-coupled rabbit anti-mouse IgG (1:4000 dilution). The blots with polyclonal HA, β-actin or luciferase antibodies were incubated with peroxidase-coupled goat anti-rabbit IgG or donkey anti-goat IgG (1:5000 dilution). Detection was performed with the enhanced chemiluminescence (ECL) method (Amersham). The immunoreactive bands were quantified by densitometric scanning.
Pulse chase and immunoprecipitation
Cells were metabolically labeled for 2 h with 50 μCi/ml Trans-[35S]-label, a mixture of 70:30 35S-methionine/cysteine (ICN). The radioactive medium was then removed and the cells were chased in cold media. The chase was terminated by wash in PBS. The cells were lysed in a buffer containing 50 mM Tris-Cl pH 7.4, 300 mM NaCl and 1% Triton X-100. Cell debris was cleared by centrifugation and cell lysates were subjected to quantitative immunoprecipitation with the HA or FLAG antibodies. Immunoprecipitated proteins were analyzed by SDS-PAGE. Radioactive bands were visualized by autoradiography and quantified by phosphorimaging.
Electrophoretic mobility shift assay (EMSA)
Cytoplasmic lysates were analyzed for IRE-binding activity by an electrophoretic mobility shift assay with a 32P-labeled IRE probe . Supershifts were performed by the addition of HA antibody to the binding reaction .
The work was supported by a grant from the Canadian Institutes for Health Research (CIHR). JW was recipient of a fellowship from the Fonds de la recherche en santé du Quebéc (FRSQ). KP holds a senior career award from FRSQ.
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