FRINK Linked Data Fragments server

Matches in SemOpenAlex for { <https://semopenalex.org/work/W2137714098> ?p ?o ?g. }

• W2137714098 endingPage "553" @default.
• W2137714098 startingPage "544" @default.
• W2137714098 abstract "Article19 January 2006free access Iron-responsive degradation of iron-regulatory protein 1 does not require the Fe–S cluster Stephen L Clarke Stephen L Clarke Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USAPresent address: Department of Physiology, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd, Dallas, TX 75390-8854, USA Search for more papers by this author Aparna Vasanthakumar Aparna Vasanthakumar Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Sheila A Anderson Sheila A Anderson Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Corinne Pondarré Corinne Pondarré Department of Pathology, Children's Hospital and Harvard Medical School, Boston, MA, USA Search for more papers by this author Cheryl M Koh Cheryl M Koh Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Kathryn M Deck Kathryn M Deck Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Joseph S Pitula Joseph S Pitula Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Charles J Epstein Charles J Epstein Department of Pediatrics and Center for Human Genetics, University of California, San Francisco, CA, USA Search for more papers by this author Mark D Fleming Mark D Fleming Department of Pathology, Children's Hospital and Harvard Medical School, Boston, MA, USA Search for more papers by this author Richard S Eisenstein Corresponding Author Richard S Eisenstein Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Stephen L Clarke Stephen L Clarke Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USAPresent address: Department of Physiology, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd, Dallas, TX 75390-8854, USA Search for more papers by this author Aparna Vasanthakumar Aparna Vasanthakumar Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Sheila A Anderson Sheila A Anderson Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Corinne Pondarré Corinne Pondarré Department of Pathology, Children's Hospital and Harvard Medical School, Boston, MA, USA Search for more papers by this author Cheryl M Koh Cheryl M Koh Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Kathryn M Deck Kathryn M Deck Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Joseph S Pitula Joseph S Pitula Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Charles J Epstein Charles J Epstein Department of Pediatrics and Center for Human Genetics, University of California, San Francisco, CA, USA Search for more papers by this author Mark D Fleming Mark D Fleming Department of Pathology, Children's Hospital and Harvard Medical School, Boston, MA, USA Search for more papers by this author Richard S Eisenstein Corresponding Author Richard S Eisenstein Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA Search for more papers by this author Author Information Stephen L Clarke1, Aparna Vasanthakumar1, Sheila A Anderson1, Corinne Pondarré2, Cheryl M Koh1, Kathryn M Deck1, Joseph S Pitula1, Charles J Epstein3, Mark D Fleming2 and Richard S Eisenstein 1 1Department of Nutritional Sciences, University of Wisconsin, Madison, WI, USA 2Department of Pathology, Children's Hospital and Harvard Medical School, Boston, MA, USA 3Department of Pediatrics and Center for Human Genetics, University of California, San Francisco, CA, USA *Corresponding author. Department of Nutritional Sciences, University of Wisconsin-Madison, 1415 Linden Drive, Madison, WI 53706, USA. Tel.: +1 608 262 5830; Fax: +1 608 262 5860; E-mail: [email protected] The EMBO Journal (2006)25:544-553https://doi.org/10.1038/sj.emboj.7600954 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info The generally accepted role of iron-regulatory protein 1 (IRP1) in orchestrating the fate of iron-regulated mRNAs depends on the interconversion of its cytosolic aconitase and RNA-binding forms through assembly/disassembly of its Fe–S cluster, without altering protein abundance. Here, we show that IRP1 protein abundance can be iron-regulated. Modulation of IRP1 abundance by iron did not require assembly of the Fe–S cluster, since a mutant with all cluster-ligating cysteines mutated to serine underwent iron-induced protein degradation. Phosphorylation of IRP1 at S138 favored the RNA-binding form and promoted iron-dependent degradation. However, phosphorylation at S138 was not required for degradation. Further, degradation of an S138 phosphomimetic mutant was not blocked by mutation of cluster-ligating cysteines. These findings were confirmed in mouse models with genetic defects in cytosolic Fe–S cluster assembly/disassembly. IRP1 RNA-binding activity was primarily regulated by IRP1 degradation in these animals. Our results reveal a mechanism for regulating IRP1 action relevant to the control of iron homeostasis during cell proliferation, inflammation, and in response to diseases altering cytosolic Fe–S cluster assembly or disassembly. Introduction Iron-regulatory protein 1 (IRP1) is an iron-regulated RNA-binding protein that controls cellular iron metabolism. IRPs bind to iron-responsive elements (IRE) in mRNAs, thereby controlling mRNA fate (Eisenstein, 2000; Hentze et al, 2004). Mutation of IRP1, its functional ortholog IRP2, or the IRE in specific mRNAs disrupts iron metabolism and can lead to iron-related abnormalities in humans and in mice (Beaumont et al, 1995; LaVaute et al, 2001; Fleming, 2002; Mok et al, 2004). Modulation of IRP1 RNA-binding activity is conventionally believed to occur as a result of insertion of a [4Fe–4S] cluster into the protein converting it to cytosolic aconitase (c-acon). Consequently, recent work has focused on identifying cytosolic proteins that directly modulate IRP1 Fe–S cluster assembly (Roy et al, 2003). However, it has been suggested that IRP1 RNA-binding activity is also controlled through regulated changes in protein degradation (Mascotti et al, 1995; Neonaki et al, 2001; Fillebeen et al, 2003). The role of these Fe–S cluster assembly/disassembly and protein degradation mechanisms in controlling IRP1 function is the focus of this investigation. Multiple lines of evidence suggest that there is a physiological need for an Fe–S cluster-independent mechanism to control accumulation of IRP1 RNA-binding activity. First, the c-acon form can be as much as 20–100-fold more abundant than the RNA-binding form of IRP1 (Chen et al, 1997; Meyron-Holtz et al, 2004a). This observation predicts that conditions that perturb the synthesis or enhance the destruction of Fe–S clusters might lead to maladaptive responses to changes in iron status due to IRP1 dysregulation. Furthermore, the activity of proteins required for cytosolic Fe–S cluster assembly and the level of reactive species (e.g. superoxide, O2•−) capable of disrupting Fe–S clusters differ between tissues (Tong and Rouault, 2000; Balk and Lill, 2004; Meyron-Holtz et al, 2004b). Consequently, a cluster-independent mechanism for iron regulation of IRP1 could provide an alternative pathway to normalize IRP1-binding activity in tissues or situations where the capacity for cytosolic Fe–S cluster assembly or disassembly differs. Despite the clear physiologic role of Fe–S cluster assembly and disassembly in controlling IRP1 RNA-binding activity, little is known of the effect of alterations in these pathways of Fe–S cluster metabolism on IRP1 function. In this regard, the Fe–S clusters of aconitases are susceptible to reactive oxygen and nitrogen species (i.e. O2•− and NO); yet, the extent to which such cluster perturbants dictate the mechanism of IRP1 regulation remains largely unexplored. For example, the pathogenesis of inherited amyotrophic lateral sclerosis due to mutations in superoxide dismutase 1 (SOD1) might, in part, involve disturbance of Fe–S cluster metabolism (Maier and Chan, 2002). Additionally, primary Fe–S biosynthetic defects contribute to the inherited neurodegenerative and hematologic disorders associated with abnormal cellular iron metabolism, including Friedreich's ataxia (Rotig et al, 1997; Stehling et al, 2004; Seznec et al, 2005) and X-linked sideroblastic anemia with ataxia (XLSA/A) (Pagon et al, 1985; Raskind et al, 1991; Hellier et al, 2001; Fleming, 2002). In the present study, we have specifically asked how assembly and disassembly of the Fe–S cluster in IRP1 contributes to the regulation of iron metabolism in normal and pathological states, and have uncovered a novel iron-dependent mechanism for controlling IRP1 protein stability that can occur without participation of the Fe–S cluster. Results IRP1 undergoes iron-dependent protein turnover in the absence of cluster assembly Formation and loss of the Fe–S cluster in IRP1 has been widely considered to be the major mechanism through which iron influences IRP1 RNA-binding activity. Iron-dependent protein turnover may also control IRP1 function (Mascotti et al, 1995; Neonaki et al, 2001; Fillebeen et al, 2003). However, the circumstances under which a protein turnover mechanism is invoked, and whether or not it involves the Fe–S cluster, have not been determined. We reasoned that enhanced protein turnover of IRP1 could control accumulation of IRP1 RNA-binding activity, particularly under conditions when iron is elevated, but conversion of IRP1 to c-acon is not efficient. Therefore, we determined the effect of iron on wild-type IRP1 (IRP1WT) and an IRP1 mutant in which all cluster-ligating cysteines (residues 437, 503, and 506) were mutated to Ser (IRP13C>3S). To avoid potential toxicity from overexpression of IRP1, a tetracycline (tet) inducible system for expression of myc-tagged WT and mutant forms of IRP1 was used. Cells expressing IRP1WT or IRP13C>3S were treated for 24 h with hemin, an iron source, or desferal, an iron chelator. Hemin had little or no effect on the RNA-binding activity of IRP1WT at 24 h (Figure 1A and B), but with shorter exposure (8 h), RNA-binding activity of IRP1WT was reduced by 20–30% (results not shown). This is similar to the time-dependent effect of iron treatment on IRP1 in other cell types (Leibold and Munro, 1988). On the other hand, desferal-induced iron deficiency stimulated RNA binding by IRP1WT three-fold, consistent with recruitment of the RNA-binding form from the c-acon form (Figure 1A and B). As expected, in iron-sufficient cells the IRP13C>3S had RNA-binding activity similar to that observed for IRP1WT in iron-deficient conditions (Figure 1A, lanes 4 versus 3). In contrast, comparison of the activity of both proteins in iron-sufficient cells showed that IRP13C>3S had higher RNA-binding activity than IRP1WT (Figure 1A, lanes 4 versus 1). Furthermore, the RNA-binding activity of IRP13C>3S responded to iron even though this protein cannot form an Fe–S cluster; hemin decreased the RNA-binding activity of IRP13C>3S by 30%, while desferal increased binding by 45% (Figure 1A and B). Of note, iron excess and iron deficiency produced a similar fold change in the RNA-binding activity of IRP1WT and IRP13C>3S. However, for IRP13C>3S, the point at which this activity is maintained in untreated cells is more reflective of a response seen in iron deficiency or in proliferating cells. These results demonstrate that IRP1 RNA-binding activity responds to iron even when a Fe–S cluster cannot be inserted, and suggests that IRP1 can be iron-regulated by cluster-dependent and -independent means. Figure 1.Iron-dependent protein turnover of an IRP1 cluster mutant: Cells expressing IRP1WT or IRP13C>3S were grown without or with 100 μM hemin or 100 μM desferal for 24 h. (A) RNA-binding activity of IRP1WT or IRP13C>3S determined by EMSA. Control cells are indicated by (C), hemin-treated cells by (H) and desferal-treated cells by (D). A representative gel is shown. EndoIRP is endogenous IRP and NS refers to nonspecific band. (B) Quantified RNA-binding results for IRP1WT or IRP13C>3S incubated with no addition (Control), 100 μM hemin or 100 μM desferal. Results are expressed as percent of control value for each clone and are mean±s.e.m. (n=3 experiments). An asterisk indicates that desferal value is significantly different from control (P<0.05), while two asterisks indicate a hemin effect relative to control or desferal. (C) Representative immunoblot of IRP1WT and IRP13C>3S in lysates from cells incubated as in panels A and B. (D) Pulse-chase analysis of the half-life of IRP1WT; half-life of IRP1WT was 18 h (n=3) irrespective of iron status (details in Materials and methods). (E) The half-life of IRP13C>3S was 3.9±0.7 h in the presence of hemin, and was greater than 18 h in the presence of desferal (n=3) (details in Materials and methods). Representative decay curves are shown. Download figure Download PowerPoint We next determined if protein level contributed to the responses of IRP1WT and IRP13C>3S to hemin (Figure 1C). As expected, IRP1WT protein was not altered by hemin or desferal. In contrast, IRP13C>3S protein level was inversely related to iron status, and was closely related to changes in its RNA-binding activity. When treated with hemin, the abundance of IRP13C>3S decreased by 30%, while after desferal treatment it increased 80%. To determine if this change in protein level was due to post-translational regulation, we determined IRP1 protein half-life (Figure 1D and E). We found that IRP13C>3S had a half-life of 4 h in hemin-treated cells that increased to greater than 18 h in the presence of desferal. In contrast, IRP1WT did not exhibit this response. Hence, in the absence of Fe–S cluster assembly, IRP1 RNA-binding activity is substantially regulated through protein degradation. S138 phosphomimetic mutants of IRP1 exhibit cluster-independent regulation of IRP1 protein stability Previous studies have shown that S138 phosphomimetic mutants of IRP1 (e.g. IRP1S138E) can be converted to c-acon, but have markedly unstable Fe–S clusters (Brown et al, 1998). Since this would predict preferential accumulation of IRP1S138E in the RNA-binding form, we determined if this phosphomimetic mutant was also subject to cluster-independent regulation. We found that in transfected cells the fraction of IRP1S138E in the RNA-binding form was five-fold more than IRP1WT or nonphosphorylatable IRP1S138A (Supplementary Figure 1). In fact, IRP1S138E displayed RNA-binding characteristics similar to the IRP13C>3S cluster mutant. Hemin treatment reduced the RNA-binding activity of IRP1S138E by 60%, while desferal increased RNA binding by 30% (Figure 2A). In contrast, IRP1S138A behaved like its WT counterpart, showing no or little response to hemin at 24 h, but a two-fold increase in RNA-binding activity after desferal treatment (compare Figures 2A and 1B). Since IRP1S138E can be an aconitase (Brown et al, 1998) (Supplementary Figure 1), we determined if the Fe–S cluster was required for iron regulation of RNA binding by creating the IRP1S138E/3C>3S quadruple mutant. We found that IRP1S138E/3C>3S displayed changes in RNA-binding activity similar to IRP1S138E after hemin or desferal treatment, indicating that the Fe–S cluster is not required for iron regulation of IRP1S138E (Figure 2A). Figure 2.Altered iron regulation and protein degradation of the S138E phosphomimetic mutant of IRP1. Cells expressing IRP1S138A, IRP1S138E or IRP1S138E/3C>3S were grown without or with 100 μM hemin or 100 μM desferal for 24 h. (A) RNA-binding activity of IRP1S138A, IRP1S138E or IRP1S138E/3C>3S determined by quantitative EMSA. Cells were incubated with no addition (Control), 100 μM hemin or 100 μM desferal for 24 h. Results are expressed as percent of control value for each clone and are mean±s.e.m. (n=3). An asterisk indicates that desferal value is significantly different from control (P<0.05), while two asterisks indicate a hemin effect relative to control or desferal. (B) Representative immunoblot of IRP1S138A, IRP1S138E, and IRP1S138E/3C>3S in lysates from cells incubated as in panel A. Treatments are indicated by Control (C); hemin (H); and desferal (D). (C) The half-life of IRP1S138E was 4.1±0.3 h in hemin, but greater than 18 h in desferal (n=3) (details in Materials and methods). (D) The half-life of IRP1S138E/3C>3S was 5.1±1.1 h in the presence of hemin, and was greater than 18 h in the presence of desferal (n=3) (details in Materials and methods). Representative decay curves are shown. Download figure Download PowerPoint To extend the parallelism with IRP13C>3S, we examined the protein level of the IRP1S138E and IRP1S138E/3C>3S mutants, and found that their protein levels were reduced by 40 and 30%, respectively, after hemin treatment (Figure 2B). Furthermore, desferal increased the protein level of both mutants by 60%. Like IRP1WT, the protein level of nonphosphorylatable IRP1S138A was not affected by hemin or desferal treatment. Similar to IRP13C>3S, IRP1S138E had a half-life of 4 h in hemin-treated cells, and desferal treatment stabilized the protein, resulting in a half-life of >18 h (Figure 2C); similar results have been observed by others (Fillebeen et al, 2003). Likewise, IRP1S138E/3C>3S had a half-life of 5 h in hemin-treated cells and >18 h when cells were exposed to desferal (Figure 2D). Hence, iron-mediated degradation independent of the Fe–S cluster accounts for the alterations in protein abundance and RNA-binding activity of IRP13C>3S as well as the IRP1S138E and IRP1S138E/3C>3S mutants. S138 phosphorylation targets the RNA-binding form of IRP1 for iron-dependent degradation Phosphorylation state-specific antibodies were developed and used to demonstrate that IRP1 is phosphorylated at S138 in control HEK cells and the level of S138 phosphorylation increased after treatment of the cells with the protein kinase C (PKC) activator phorbol 12-myristate 13-acetate (PMA) (Supplementary Figures 2 and 3). The S138 site was also recognized by these antibodies after incubation of purified IRP1WT, but not IRP1S138A, with PKC (Supplementary Figure 2). These antibodies were used to determine the properties of S138-phosphorylated IRP1 in HEK cells. In order to determine the fraction of IRP1 that is phosphorylated in cells, two-dimensional gel electrophoresis was employed. Cells expressing IRP1WT were labeled with [35S]Met/Cys or [32P]orthophosphate in the presence or absence of PMA, and IRP1 was collected by immunoprecipitation. In control cells, two species of [35S]IRP1 were observed, with the more acidic form (form 2) present in lower abundance (Figure 3A). After PMA treatment, this acidic isoform (form 2) increased in relative abundance by about 20% and two additional more acidic isoforms (forms 3 and 4) of [35S]IRP1 were observed (compare Figure 3A with B). When labeled with [32P], form 3 and possibly form 4 were detected in control cells (Figure 3C). After PMA treatment, the amount of [32P]IRP1 increased substantially and [32P] was detected in all acidic isoforms (forms 2–4) (Figure 3D). Hence, forms 2–4 correspond to phosphorylated IRP1. The S138 phosphospecific antibody recognized forms 2–4 in PMA-treated cells (Figure 3E); S138 phosphorylated IRP1 was present at a lower level in control cells (Supplementary Figure 3). Since IRP1 can also be phosphorylated at S711 (Pitula et al, 2004), the phosphorylated forms 2 and 3 likely represent mono- and diphosphorylated IRP1. Form 4 may represent IRP1 phosphorylated at sites in addition to S138 and S711 as additional phosphorylation sites have been observed in cultured cells and with purified IRP1 protein (S Anderson and R Eisenstein, data not shown). On the basis of the intensity of the 35S-labeled protein species, approximately 2% of IRP1 is phosphorylated in control cells, while in PMA-treated cells phosphorylation increased six-fold such that 12% of the protein was phosphorylated. Our previous work using purified IRP1 demonstrated that the free apoprotein form is the preferred substrate for phosphorylation; the c-acon and the RNA-bound forms of IRP1 are poor substrates for phosphorylation (Schalinske et al, 1997). The level of IRP1 phosphorylation observed in HEK cells suggests that this is the case in vivo. In agreement with this, desferal treatment of cells stimulated phosphorylation of IRP1 at S138 several fold, and IRP13C>3S is more highly phosphorylated than IRP1WT in untreated cells (A Vasanthakumar and R Eisenstein, data not shown). Figure 3.Abundance and iron-dependent loss of S138 phosphorylated IRP1. Panels (A) and (B): HEK cells were incubated with 100 μCi/ml of [35S]Met/Cys for 2 h before the addition of vehicle (DMSO, control) or PMA (1 μM) for an additional 2 h. Myc-tagged IRP1 was immunoprecipitated and analyzed by 2-D gel electrophoresis. (C) HEK cells were incubated with 0.75 mCi/ml of [32P]orthophosphate for 4 h. (D) HEK cells were incubated with 0.75 mCi/ml of [32P]orthophosphate for 2 h before addition of PMA and incubation for an additional 2 h. Immunoprecipitated myc-tagged IRP1 was analyzed in panels C and D. (E) Myc-tagged IRP1 was immunoprecipitated from unlabeled HEK cells treated with 1 μM PMA for 2 h. After 2D gel separation, S138 phosphorylated IRP1 was detected by phosphoblotting. (F) HEK cells were treated with 1 μM PMA for 2 h. The media was removed and replaced with normal media with or without 100 μM hemin. After 2 or 4 h, myc-tagged IRP1 was immunoprecipitated from cell lysates and analyzed for S138-phosphorylated or total IRP1 by immunoblotting. Graph (i) represents the 4-h time point. Graph (ii) represents the 4-h time point from an experiment where cells were treated with PMA for 2 h and then hemin for 4 h. In one set of plates, the protease inhibitor ALLN (10 μM) (N-acetyl-leucyl-leucyl-norleucinal, Calbiochem) was added with the PMA and hemin. Results in graphs i and ii are mean±s.e.m. for three separate experiments. (G) Cells expressing IRP1S138A/3C>3S were treated without or with 100 μM hemin or 100 μM desferal for 24 h. Representative immunoblot of myc-tagged IRP1. The graph shows quantified results for n=3 independent pools of clones (mean±s.e.m.). Panels F and G: an asterisk indicates significant difference (P<0.05). Download figure Download PowerPoint To further establish the physiological relevance of phosphorylation, we determined the effect of hemin treatment on the abundance of S138-phosphorylated IRP1. To do so, cells were treated with PMA for 2 h and then incubated in the presence or absence of hemin for another 2 or 4 h. Addition of hemin reduced the level of S138-phosphorylated IRP1 by 45% at 4 h (Figure 3F, graph i). In contrast, the level of S138-phosphorylated and total IRP1 did not change in cells that were not hemin-treated. In the presence of the protease inhibitor ALLN (plus hemin), the level of S138-phosphorylated IRP1 protein was 52% greater than in the presence of hemin alone (Figure 3F, graph ii). This shows that S138-phosphorylated IRP1 undergoes iron-dependent degradation, consistent with the notion that IRP1 undergoes cluster-independent regulation when phosphorylated at S138. Phosphorylation at S138 is not required for iron regulation of protein stability of IRP1 cluster mutants We next determined whether phosphorylation is required for, or merely induces, cluster-independent regulation of IRP1. The requirement for S138 phosphorylation for turnover of the IRP1 cluster mutants was evaluated by creating the nonphosphorylatable IRP1S138A/3C>3S mutant. Hemin treatment reduced the level of IRP1S138A/3C>3S protein by 25%, whereas desferal promoted a 150% increase in its abundance (Figure 3G). Hence, S138 phosphorylation is not required for iron regulation of the turnover of IRP1 cluster mutants. Nonetheless, these findings do suggest that IRP1 S138 phosphorylation is one of the physiological mechanisms that impairs the accumulation of the c-acon form, thereby rendering IRP1 susceptible to iron-dependent turnover. Overall, these results indicate that S138 phosphorylation promotes, but is not required for, iron-dependent control of IRP1 protein turnover, and that other scenarios where cluster accumulation is impaired are likely to promote a similar mode of regulation. Sod1−/− mice exhibit iron-dependent regulation of IRP1 protein levels To establish the relevance of our findings in vivo, we used two animal disease models with perturbed cytosolic Fe–S cluster metabolism. As the Fe–S cluster in aconitases is known to be sensitive to superoxide, we first examined the effect of loss of the Fe–S cluster in vivo in mice lacking the copper-zinc superoxide dismutase, SOD1 (Huang et al, 1997; Elchuri et al, 2005). SOD1 deficiency is well established to promote cytosolic oxidative stress, which can damage Fe–S clusters, including that of c-acon (Strain et al, 1998; Missirilis et al, 2003; Starzynski et al, 2005). Iron-replete Sod1−/− mice had 50% more IRP1 RNA-binding activity in liver compared to their WT littermates (Figure 4A). Treatment with 2-mercaptoethanol (2-ME) activates RNA binding by c-acon and other inactive forms of IRP1, allowing an indirect assessment of total IRP1 protein. In WT mouse liver, IRP1 RNA-binding activity was increased 260-fold after 2-ME treatment, whereas in Sod1−/− mice a 15-fold increase was observed (Figure 4B). In addition, the total level of 2-ME inducible RNA-binding activity was lower in Sod1−/− liver. In agreement with these observations, IRP1 protein abundance was reduced by 80% (Figure 4C) and c-acon activity was not detectable in Sod1−/− liver (Figure 4D); others have obtained similar results (Starzynski et al, 2005). Mitochondrial aconitase activity was not reduced in liver of Sod1−/− mice (Figure 4D). To determine if this effect of loss of SOD1 on IRP1 protein accumulation was a phenomenon applicable to other cytosolic Fe–S proteins, xanthine oxidase (XO) was examined. In contrast to IRP1, XO protein levels were not altered in Sod1−/− mouse liver (Figure 4C), even though XO activity was reduced by 74% (Figure 4D). Hence, the reduction in IRP1 protein in Sod1−/− liver reflects regulation of the stability of only certain cytosolic Fe–S proteins. Figure 4.Iron regulates IRP1 protein abundance in the liver of Sod1−/− mice. Liver IRP1 RNA-binding activity, enzyme activities and protein levels were determined in the liver of wild-type (WT) or Sod1−/− mice. (A) IRP1 RNA-binding activity was determined by EMSA of liver cytosol from 8-week-old mice (n=4). IRP1 RNA-binding activity was 0.072 pmol/mg protein (WT) and 0.11 pmol/mg protein (Sod1−/−). (B) RNA binding after treatment of cytosol with 4% 2-ME. IRP1 RNA-binding activity was 19 pmol/mg protein in WT liver and 1.6 pmol/mg protein in Sod1−/− liver. (C) Immunoblot for IRP1 and XO in the liver of WT or Sod1−/− mice fed a diet containing 50 ppm iron. (D) c-Acon, XO, and mitochondrial aconitase activity in WT and Sod1−/− liver. Unit activity is μmol cis-aconitate produced/min for aconitases and is μmol Amplex-Red consumed per min for XO. Results for n=3 animals/group. (E) IRP1 RNA-binding activity and protein level (immunoblot) in WT or Sod1−/− mice fed an iron-replete (50 ppm Fe) or iron-deficient (<2 ppm Fe) diet for 3 weeks. Immunoblot of IRP1 or α-tubulin in the liver cytosol of WT or Sod1−/− mice fed either the iron-sufficient or -deficient diet for 3 weeks. For all panels an asterisk indicates that Sod1−/− is significantly different from WT (P<0.05). In panel E graph, iron-deficient values are different from iron-sufficient values for both genotypes (P<0.05). Download figure Download PowerPoint The iron-dependent regulation of IRP1 protein abundance observed with the cluster mutants (IRP13C>3S, IRP1S138/3C>3S) in cultured cells was recapitulated in Sod1−/− mice. Iron-deficient WT mice had three-fold more IRP1-binding activity in liver than did iron-replete WT animals (Figure 4E) and, as expected, there was no change in IRP1 protein level. However, there was five-fold more IRP1-binding activity in iron-deficient versus iron-replete Sod1−/− mice. Furthermore, and unexpectedly, iron deficiency resulted in a 2.5-fold increase in IRP1 protein levels in mutant animals, compared to iron-sufficient mutant mice. No such effect was seen in wild-type mice. Thus, one-half of the increase in IRP1 RNA-binding activity in the liver of iron-deficient Sod1−/− mice can be attributed to an increase in IRP1 protein. Impaired assembly of cytosolic Fe–S clusters in Abcb7lv/Y mice leads to iron regulation of IRP1 protein abundance To confirm and extend the findings in Sod1−/− mice, an alternative method predicted to impair accumulation of c-acon from IRP1 was used. Here, the expression of Abcb7, a protein implicated in cytosolic Fe–S cluster assembly, that is defective in the human disorder X-linked sideroblastic anemia with ataxia (XLSA/A), was specifically ablated in liver using a tissue-specific deletion strategy (Pondarré et al, submitted). Male mice containing the Abcb7 conditional allele and a liver-specific Cre recombinase transgene are hereafter referred to" @default.
• W2137714098 created "2016-06-24" @default.
• W2137714098 creator A5001926930 @default.
• W2137714098 creator A5003554057 @default.
• W2137714098 creator A5006370557 @default.
• W2137714098 creator A5013266798 @default.
• W2137714098 creator A5013945791 @default.
• W2137714098 creator A5016817792 @default.
• W2137714098 creator A5024941834 @default.
• W2137714098 creator A5033261280 @default.
• W2137714098 creator A5074830346 @default.
• W2137714098 creator A5081882131 @default.
• W2137714098 date "2006-01-19" @default.
• W2137714098 modified "2023-10-03" @default.
• W2137714098 title "Iron-responsive degradation of iron-regulatory protein 1 does not require the Fe–S cluster" @default.
• W2137714098 cites W1530657831 @default.
• W2137714098 cites W1535109335 @default.
• W2137714098 cites W1538728268 @default.
• W2137714098 cites W1547848353 @default.
• W2137714098 cites W1568912715 @default.
• W2137714098 cites W1592463355 @default.
• W2137714098 cites W1594389718 @default.
• W2137714098 cites W1606423922 @default.
• W2137714098 cites W1671669579 @default.
• W2137714098 cites W167888359 @default.
• W2137714098 cites W1910176080 @default.
• W2137714098 cites W1943372705 @default.
• W2137714098 cites W1967036581 @default.
• W2137714098 cites W1969490516 @default.
• W2137714098 cites W1969496526 @default.
• W2137714098 cites W1972537129 @default.
• W2137714098 cites W1975944453 @default.
• W2137714098 cites W1985841747 @default.
• W2137714098 cites W1986700855 @default.
• W2137714098 cites W1987287339 @default.
• W2137714098 cites W1987697140 @default.
• W2137714098 cites W1990354671 @default.
• W2137714098 cites W1992193197 @default.
• W2137714098 cites W2006178088 @default.
• W2137714098 cites W2009029742 @default.
• W2137714098 cites W2015796740 @default.
• W2137714098 cites W2024089487 @default.
• W2137714098 cites W2025403494 @default.
• W2137714098 cites W2029584806 @default.
• W2137714098 cites W2041575942 @default.
• W2137714098 cites W2042637324 @default.
• W2137714098 cites W2045864586 @default.
• W2137714098 cites W2047898325 @default.
• W2137714098 cites W2057404038 @default.
• W2137714098 cites W2062253763 @default.
• W2137714098 cites W2065999538 @default.
• W2137714098 cites W2070141684 @default.
• W2137714098 cites W2082798387 @default.
• W2137714098 cites W2084432680 @default.
• W2137714098 cites W2085866164 @default.
• W2137714098 cites W2088360596 @default.
• W2137714098 cites W2107845441 @default.
• W2137714098 cites W2111599402 @default.
• W2137714098 cites W2120374147 @default.
• W2137714098 cites W2133155560 @default.
• W2137714098 cites W2136827920 @default.
• W2137714098 cites W2147701752 @default.
• W2137714098 cites W2154439380 @default.
• W2137714098 cites W2155506096 @default.
• W2137714098 cites W2163557814 @default.
• W2137714098 cites W2166595673 @default.
• W2137714098 cites W2167860376 @default.
• W2137714098 cites W2176933263 @default.
• W2137714098 doi "https://doi.org/10.1038/sj.emboj.7600954" @default.
• W2137714098 hasPubMedCentralId "https://www.ncbi.nlm.nih.gov/pmc/articles/1383537" @default.
• W2137714098 hasPubMedId "https://pubmed.ncbi.nlm.nih.gov/16424901" @default.
• W2137714098 hasPublicationYear "2006" @default.
• W2137714098 type Work @default.
• W2137714098 sameAs 2137714098 @default.
• W2137714098 citedByCount "92" @default.
• W2137714098 countsByYear W21377140982012 @default.
• W2137714098 countsByYear W21377140982013 @default.
• W2137714098 countsByYear W21377140982014 @default.
• W2137714098 countsByYear W21377140982015 @default.
• W2137714098 countsByYear W21377140982016 @default.
• W2137714098 countsByYear W21377140982017 @default.
• W2137714098 countsByYear W21377140982018 @default.
• W2137714098 countsByYear W21377140982019 @default.
• W2137714098 countsByYear W21377140982021 @default.
• W2137714098 countsByYear W21377140982022 @default.
• W2137714098 countsByYear W21377140982023 @default.
• W2137714098 crossrefType "journal-article" @default.
• W2137714098 hasAuthorship W2137714098A5001926930 @default.
• W2137714098 hasAuthorship W2137714098A5003554057 @default.
• W2137714098 hasAuthorship W2137714098A5006370557 @default.
• W2137714098 hasAuthorship W2137714098A5013266798 @default.
• W2137714098 hasAuthorship W2137714098A5013945791 @default.
• W2137714098 hasAuthorship W2137714098A5016817792 @default.
• W2137714098 hasAuthorship W2137714098A5024941834 @default.
• W2137714098 hasAuthorship W2137714098A5033261280 @default.
• W2137714098 hasAuthorship W2137714098A5074830346 @default.
• W2137714098 hasAuthorship W2137714098A5081882131 @default.
• W2137714098 hasBestOaLocation W21377140981 @default.

• next