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. Author manuscript; available in PMC: 2013 Nov 7.
Published in final edited form as: Cell Metab. 2012 Oct 25;16(5):645–657. doi: 10.1016/j.cmet.2012.10.001

mTOR Regulates Cellular Iron Homeostasis through Tristetraprolin

Marina Bayeva 1, Arineh Khechaduri 1, Sergi Puig 2, Hsiang-Chun Chang 1, Sonika Patial 3, Perry J Blackshear 3, Hossein Ardehali 1,*
PMCID: PMC3594686  NIHMSID: NIHMS418098  PMID: 23102618

SUMMARY

Iron is an essential cofactor with unique redox properties. Iron regulatory proteins 1 and 2 (IRP1/2) have been established as important regulators of cellular iron homeostasis, but little is known about the role of other pathways in this process. Here we report that the mammalian target of rapamycin (mTOR) regulates iron homeostasis by modulating transferrin receptor 1 (TfR1) stability and altering cellular iron flux. Mechanistic studies identify tristetraprolin (TTP), a protein involved in anti-inflammatory response, as the downstream target of mTOR that binds to and enhances degradation of TfR1 mRNA. We also show that TTP is strongly induced by iron chelation, promotes downregulation of iron-requiring genes in both mammalian and yeast cells, and modulates survival in low-iron states. Taken together, our data uncover a link between metabolic, inflammatory, and iron regulatory pathways, and point towards the existence of a yeast-like TTP-mediated iron conservation program in mammals.

INTRODUCTION

Iron deficiency anemia affects one quarter of the world’s population (WHO, 2008), yet the mechanisms of iron regulation remain largely unexplored. Iron is an essential micronutrient required for fundamental biological processes including oxygen delivery, protein synthesis, DNA replication and oxidative phosphorylation (Arredondo and Nunez, 2005). However, the ability of iron to easily gain and lose electrons also facilitates the production of reactive oxygen species (ROS) by Fenton reaction (Aisen et al., 2001). Thus, iron levels must be tightly controlled at both cellular and systemic levels and deregulation of iron homeostasis is associated with aging, metabolic disorders and cancer (Aisen et al., 2001; Altamura and Muckenthaler, 2009; Torti and Torti, 2011; Weinberg, 2010).

Maintenance of cellular iron homeostasis is dependent on RNA-binding iron regulatory proteins 1 and 2 (IRP1/2) which are activated under iron-deficient conditions. IRP1/2 function to restore iron levels through: 1) stabilization of transferrin receptor 1 (TfR1) mRNA and increased iron uptake, 2) mobilization of cellular iron stores, and 3) reduction in cellular iron export through the suppression of ferroportin 1 (Fpn1), the only cellular iron exporter identified to date (Hentze et al., 2010; Rouault, 2006; Wang and Pantopoulos, 2011). Additionally, an iron conservation response has been recently described in S. cerevisiae where coordinated expression of the tandem zinc finger (TZF) proteins Cth1p and Cth2p causes destabilization of mRNAs of nonessential iron-containing proteins and libration of iron for use in vital processes (Pedro-Segura et al., 2008; Puig et al., 2005; Puig et al., 2008). A mammalian homolog of Cth1p/Cth2p, tristetraprolin (TTP), has been extensively studied in the context of immunosuppression (Baou et al., 2009), but its role in iron regulation is not known.

Mammalian target of rapamycin (mTOR) is an important sensor of cellular energy state and a major hub for integration of environmental cues (Howell and Manning, 2011; Sengupta et al., 2010). The complex is activated under energetically favorable, low-stress states while adverse conditions, such as starvation, cytotoxic insults and DNA damage, act to inhibit mTOR signaling and promote survival through conservation of cellular resources (Dunlop and Tee, 2009; Soliman, 2005). Many of the processes governed by mTOR are dependent on iron as a co-factor and would require a steady supply of this metal. There appears to be a link between iron homeostasis and mTOR, as clinical use of rapamycin is associated with the development of microcytic anemia that is mechanistically distinct from the anemia of chronic disease (Kim et al., 2006; Maiorano et al., 2006; Sofroniadou et al., 2010). Here, we establish mTOR pathway as a regulator of iron homeostasis which acts through modulation of iron transporters and alteration of cellular iron flux. Moreover, we identify TTP as a target of mTOR which, at least in part, modulates its iron-regulatory effects through destabilization of TfR1 mRNA.

RESULTS

mTOR regulates cellular iron homeostasis

mTOR activation stimulates several important anabolic processes that depend on the presence of iron as a cofactor; therefore, we hypothesized that mTOR may play a role in the regulation of cellular iron availability and utilization. To test this, we modulated mTOR signaling in wild type (WT) mouse embryonic fibroblasts (MEFs) and H9c2 cardiac myoblasts followed by quantification of heme and non-heme iron content. Rapamycin effectively inhibited mTOR signaling in all cell lines (Figure 1A), as assessed by the ablation of pS6 expression. Treatment with rapamycin led to a significant increase in both non-heme and heme iron content in MEFs (Figure 1B and S1A) and H9c2 cells (Figure S1B,C). Consistent with elevation in iron levels, protein content of ferritin light chain, an iron storage molecule, was also significantly increased by rapamycin treatment in WT MEFs (Figure S1D). Increased heme levels were likely due to inhibition of heme degradation, as the mRNA levels of heme oxygenase 1 (HMOX1) were significantly decreased, while the expression of key heme synthetic enzymes, δ-aminolevulinic acid synthase 1 (ALAS1) and ferrochelatase, remained unchanged (Figure S1E). To activate mTOR, we used TSC2 knockout MEFs in which the upstream inhibitor of the pathway is genetically deleted and pS6 levels are increased (Figure 1A). The knockout MEFs were transfected with either an empty pEF6 vector (referred to as TSC2 KO-V MEFs) or with pEF6/TSC2 vector to reconstitute TSC2 expression in the knockout MEFs (TSC2 KO-TSC2 MEFs), followed by selection with blasticidin to generate stable transgenic cell lines as described (Zhang et al., 2003). Constitutive activation of mTOR in TSC2 KO-V MEFs decreased heme and non-heme iron, and rapamycin restored their levels to that of control (Figure 1C and S1F), indicating that the observed changes were due to mTOR modulation.

Figure 1. mTOR regulates cellular iron content and uptake.

Figure 1

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(A) mTOR modulation in MEFs (left) and H9c2 (right) cells assessed by Western blot of phosphorylated S6 protein. Densitometry analysis is shown below the Western blot (n=3). TSC2 protein levels (right) in TSC2 KO-V and TSC2 KO-TSC2 MEFs. (B) Non-heme iron content in WT MEFs treated with 20nM rapamycin for 48 hrs normalized to protein concentration (n=12). All subsequent experiments were carried out with 20nM rapamycin treatment for 48 hrs in all mammalian cell lines, unless noted otherwise. (C) Non-heme iron content in TSC2 KO-V and TSC2 KO-TSC2 MEFs with or without rapamycin treatment (n=6–12). (D) Radioactive iron uptake over 1 hour by WT MEFs with rapamycin or vehicle control treatment (n=12). (E) Radioactive iron uptake over 1 hour by TSC2 KO-V and TSC2 KO-TSC2 MEFs with or without rapamycin (n=6–12). (F) Cellular content of 55Fe in WT MEFs treated with rapamycin or vehicle control following the 24-hour incubation with radioactive iron (n=12). (G) Cellular content of 55Fe in TSC2 KO-V and TSC2 KO-TSC2 MEFs with or without rapamycin following the 24-hour incubation with radioactive iron (n=6). Data are presented as mean ± SEM. * p<0.05 vs. control, # p<0.05 vs. TSC2 KO-V MEFs. See also Figure S1.

To determine if changes in cellular iron resulted from alterations in iron import, we measured the uptake of radioactive iron (55Fe) after mTOR modulation. Unexpectedly, rapamycin treatment significantly reduced iron uptake, while TSC2 KO-V MEFs or TSC2 siRNA-treated H9c2 myoblasts displayed enhancement of iron uptake after 60 minutes of incubation (Figure 1D,E and S1G,H). The differences persisted even after 24 hours of growth with 55Fe in all cell lines (Figure 1F,G and S1I,J). These results suggest that inhibition of mTOR results in cellular iron accumulation despite reduction in iron uptake, while mTOR activation has the opposite effect.

We next studied the mechanism for these observations by measuring the expression of genes involved in iron regulation in MEFs and H9c2 (Anderson and Vulpe, 2009; Hentze et al., 2010). Consistent with the decreased iron uptake, rapamycin significantly reduced the mRNA and protein levels of transferrin receptor 1 (TfR1) required for the uptake of transferrin-bound iron (Figure 2A,C). Additionally, TfR1 levels were reduced in MEFs and H9c2 treated with Torin-1, a more specific inhibitor of the mTOR pathway (Figure S2A). However, mRNA and protein levels of an iron exporter, Fpn1, were also reduced by rapamycin and Torin-1 (Figure 2B,C, Figure S2A). Consistently, mTOR activation increased the expression of TfR1 and Fpn1, while rapamycin suppressed this effect (Figure 2D). These results suggest that mTOR inhibition leads to coordinated reduction in the expression of TfR1 and Fpn1, resulting in a net accumulation of cellular iron, while mTOR activation has the opposite effect.

Figure 2. mTOR regulates expression of iron transporters.

Figure 2

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Protein levels of TfR1 in WT MEFs (A) and Fpn1 in mouse hearts (B) following rapamycin treatment. Densitometry analyses are presented below the Western blots. (C) mRNA levels of iron transporters, TfR1 and Fpn1, as determined by qRT-PCR, in WT MEFs with or without rapamycin treatment (n=6–12). (D) mRNA levels of iron transporters in TSC2 KO-V and TSC2 KO-TSC2 MEFs with or without rapamycin treatment (n=6–12). (E) mRNA levels of genes unrelated to iron homeostasis in WT MEFs treated with rapamycin or vehicle control (n=6). Data are presented as mean ± SEM. * p<0.05 vs. control, # p<0.05 vs. TSC2 KO-V MEFs. See also Figure S2.

In addition to its effects on TfR1 and Fpn1, rapamycin led to a global suppression of many iron-regulatory genes in MEFs and H9c2, consistent with a reduction in cellular iron flux, while the opposite was observed with activation of mTOR (Figure S2B–E). On contrary, we found no downregulation of several genes not involved in regulation of cellular iron upon rapamycin treatment of WT MEFs (Figure 2E), suggesting that its effects on iron regulatory network are specific, and do not represent an artifact of a generalized transcriptional repression.

TfR1 is a target of mTOR

Concurrent decrease in the expression of a protein mediating iron import (TfR1) and an iron exporter (Fpn1) by rapamycin raises two possibilities. First, mTOR inhibition may negatively regulate the expression of TfR1, leading to reduced iron import and net iron loss from the cell. The resulting deficiency would activate IRP1/2 with subsequent inhibition of Fpn1 to prevent further iron loss (Figure S3A). Alternatively, rapamycin may primarily target Fpn1 leading to cellular iron accumulation due to decreased export. This would suppress IRP1/2 activity and secondarily reduce TfR1 levels to stop iron gain through the import (Figure S3B). To distinguish between these two possibilities, we assessed the time-course of TfR1 and Fpn1 expression during the first 24 hours of rapamycin treatment. We observed a steady decrease in TfR1 mRNA levels with rapamycin, while Fpn1 expression was initially upregulated at 2 and 4 hours, followed by a decline (Figure 3A). These findings point towards the first possibility of mTOR primarily regulating TfR1 expression, and the changes in Fpn1 being the secondary effect (Figure S3A). Consequently, we focused our efforts on studying the regulation of TfR1 by mTOR.

Figure 3. TTP is functionally similar to yeast Cth1p/2p.

Figure 3

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(A) Time-course of TfR1 and Fpn1 mRNA regulation by rapamycin in WT MEFs over 24 hrs (n=12–18). (B) TfR1 mRNA stability in WT MEFs with and without rapamycin pre-treatment as assessed by qRT-PCR after incubation with transcriptional inhibitor actinomycin D (act D), normalized TfR1 mRNA levels at time point zero in each respective group (n=3). (C) IRP1 mRNA levels in IRP2 KO MEFs following treatment with IRP1 siRNA (n=6). (D) TfR1 mRNA levels in IRP1/2 KD/KO MEFs following rapamycin treatment (n=6). TTP mRNA (E) and TTP protein (F) levels in WT MEFs after DFO treatment for 16 hours (n=3–6). Densitometry analysis is presented below the Western blot. (G) TTP mRNA levels in H9c2 cells after 16-hour incubation with 2,2-BPD or FAC for iron chelation and overload, respectively (n=6). (H) mRNA levels of iron-containing proteins in WT MEFs treated with 150µM DFO for 16 hours (n=6). Unless noted otherwise, all subsequent studies were performed using this protocol for iron chelation with DFO. (I) mRNA levels of iron-containing proteins in TTP WT and KO MEFs (n=12–18). (J) mRNA levels of iron-containing proteins in TTP KO and WT MEFs in the presence and absence of DFO (n=6–12). (K) mRNA levels of iron-containing Cth2p targets in cth1Δcth2Δ yeast with ectopic expression of human wild-type and TZF mutant TTP. Data are presented as mean ± SEM. * p<0.05 vs. control, # p<0.05 vs. control at 180 minutes. See also Figure S3.

To uncover the mechanism by which mTOR regulates TfR1, we first assessed the stability of TfR1 mRNA after rapamycin treatment. Incubation of rapamycin-treated MEFs with a transcriptional inhibitor actinomycin D led to a significant reduction in TfR1 mRNA stability relative to the control group (Figure 3B), suggestive of post-transcriptional regulation. The key known post-transcriptional regulators of TfR1 are IRP1/2 proteins, which, we hypothesized, may mediate the mTOR-dependent changes in TfR1. To determine if IRP1/2 pathway mediates the suppression of TfR1 by rapamycin, we downregulated IRP1 expression in IRP2 KO MEFs using siRNA approach (IRP1/2 KD/KO MEFs), which resulted in a significant knockdown of IRP1 expression (Figure 3C) and a reduction in TfR1 mRNA, consistent with a functional inhibition of the IRP1/2 pathway (Figure S3C). However, the repression of TfR1 by rapamycin was intact in the IRP1/2 KD/KO MEFs (Figure 3D), indicating that regulation of TfR1 by mTOR is independent of the IRP1/2 pathway.

TTP is functionally similar to iron-regulating yeast Cth1p/Cth2p proteins

Recent studies in S. cerevisiae found TZF proteins, Cth1p and Cth2p, to regulate cellular iron homeostasis by binding to AU-rich elements (AREs) and destabilizing the mRNAs of iron-requiring proteins thus salvaging iron for use in essential functions (Puig et al., 2005). Since we showed that IRP1/2 proteins do not mediate the effects of mTOR on TfR1, we hypothesized that TZF proteins may regulate TfR1 and cellular iron homeostasis in mammals. Three TZF proteins with homology to Cth1p and Cth2p, TTP (ZFP36), CMG1 (Zfp36l1) and TIS11D (Zfp36l2), have been identified in mammals (Baou et al., 2009) and studied in the context of cytokine signaling (Carrick et al., 2004). To determine whether these proteins modulate iron homeostasis similar to Cth1p/Cth2p in yeast, we first assessed their regulation by cellular iron levels. Iron chelation with DFO resulted in a significant and dose-dependent upregulation of TTP mRNA (Figure 3E) and protein expression (Figure 3F). Treatment with 2,2–bipyridyl (2,2-BPD), a lipophilic iron chelator that is mechanistically distinct from DFO, also induced TTP expression (Figure 3G), suggesting that the observed effect is indeed due to a reduction in cellular iron and is not restricted to DFO. Moreover, iron overload with ferric ammonium citrate (FAC) suppressed TTP expression (Figure 3G). The mRNA levels of the other two TZF proteins were also increased by DFO in MEFs and H9c2 cells (Figure S3D,E), however, they were excluded from subsequent studies due to the lack of regulation by mTOR (Figure S3F,G). Taken together, our results show that, similar to Cth1p/Cth2p in yeast, TTP is regulated by cellular iron status in mammals.

To further confirm the functional similarity between Cth1p/Cth2p and TTP, we studied the ability of TTP to reduce the expression of ARE-containing iron-requiring proteins (ABCE1, a Fe/S cluster protein involved in ribosome biogenesis and Lias, lipoic acid synthase) (Figure S3H). Similar to yeast, iron chelation reduced the expression of ABCE1 and Lias in WT MEFs (Figure 3H), but did not suppress mRNA levels of most other proteins that do not require iron for their function (Figure S3I). Moreover, mRNA levels of ABCE1 and Lias were increased in TTP KO MEFs (Figure 3I and S3J), while ABCE1 levels were reduced by TTP overexpression in WT MEFs (Figure S3J,K), confirming the inhibitory function of TTP. DFO treatment of TTP KO MEFs not only failed to decrease, but instead led to an exaggerated increase in the expression of ABCE1 and Lias (Figure 3J), suggesting that TTP is critical for the repression of iron-containing proteins by iron chelation. Finally, to determine whether mammalian TTP can complement the Cth1p/Cth2p deletion in yeast, we expressed the WT and TZF-mutant allele of human TTP (C124R) (Lai et al., 1999) under the control of the iron-regulated CTH2 promoter in yeast cells. Similar to MEFs, treatment of Cth1ΔCth2Δ yeast with an iron chelator bathophenanthroline (BPS) led to an increase in the mRNA levels of two known Cth1p/Cth2p targets, succinate dehydrogenase subunit 4 (SDH4) and aconitase (ACO1), while transfection of these cells with Cth2 plasmid restored the suppression of iron-containing genes by iron deficiency (Figure 3K). Moreover, overexpression of WT human TTP, but not the C124R construct, reduced the mRNA levels of SDH4 and ACO1 under iron-limiting conditions, suggestive of a functional conservation between yeast Cth1p/Cth2p and mammalian TTP (Figure 3K). Taken together, our findings suggest that the role of TTP in regulation of cellular iron homeostasis is similar to that performed by yeast Cth1p/Cth2p, namely suppressing the expression of iron-containing proteins and potentially optimizing iron utilization in low-iron states.

mTOR regulates TTP expression

Next, we assessed whether mTOR regulates the expression of TTP. Rapamycin treatment of MEFs and H9c2 cells induced expression of TTP mRNA and protein (Figure 4A,B and S4A). Significant upregulation of TTP mRNA was also observed in Torin-1 treated MEFs and H9c2 cells (Figure S4B,C). TTP levels were suppressed by mTOR activation in TSC2 KO-V MEFs, and rapamycin reversed these effects (Figure 4C). Moreover, rapamycin treatment of WT MEFs resulted in a significant decrease in the levels and stability of TTP targets ABCE1 and Lias, consistent with TTP induction (Figure S4D,E). Treatment of cells with both rapamycin and DFO led to a greater induction of TTP expression than either agent alone (Figure S4F), suggesting an additive effect. Importantly, the downstream effects of TTP and rapamycin on iron regulation were independent of the inflammatory response, as we did not detect TNFα mRNA expression in MEFs or H9c2 cells (Figure S4G,H). Finally, to determine whether the regulation of TZF proteins by mTOR is conserved between eukaryotic organisms, we treated WT yeast strain with rapamycin and observed a significant induction of Cth1 and Cth2 mRNA (Figure 4D), consistent with our observations in the mammalian cells.

Figure 4. mTOR regulates TTP expression.

Figure 4

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TTP mRNA (A) and TTP protein (B) levels in WT MEFs treated with rapamycin (n=6–8). (C) TTP mRNA levels in TSC2 KO-V and TSC2 KO-TSC2 MEFs with and without rapamycin treatment (n=6–12). (D) Cth1 and Cth2 mRNA levels in WT yeast cells treated with 1µg/mL rapamycin for 24 hours (n=6). mRNA levels of TTP in ARNT KO MEFs treated with rapamycin (E) or DFO (F) (n=15). TTP mRNA levels in IRP1/2 KD/KO MEFs treated with rapamycin (G) or DFO (H) (n=6). (I) TfR1 mRNA levels in TTP KO and control MEFs (n=15). (J) TfR1 mRNA levels in WT MEFs transfected with TTP- or GFP-containing (control) vectors (n=6). (K) TfR1 mRNA stability in TTP WT and KO MEFs treated with the transcriptional inhibitor act D (n=6). Data are presented as mean ± SEM. * p<0.05 vs. control. See also Figure S4.

Since both mTOR and cellular iron status are known to regulate hypoxia-inducible factor (HIF) signaling, we assessed TTP regulation in aryl hydrocarbon receptor nuclear translocator (ARNT) KO MEFs that lack the obligate binding partner of HIF1α and HIF2α, thus disrupting HIF pathway (Patel and Simon, 2008). We found TTP levels to be increased both with DFO and rapamycin treatment in ARNT KO MEFs (Figure 4E,F), indicating that the observed regulation is HIF-independent. Moreover, we observed significant induction of TTP expression in IRP1/2 KD/KO MEFs (Figure 4G,H), suggesting that the activity of IRP1/2 system does not mediate the upregulation of TTP expression by rapamycin and DFO. In summary, we found TTP to be negatively regulated by mTOR pathway in a HIF- and IRP1/2-independent manner.

TTP associates with and regulates TfR1 mRNA

Since TTP modulates cellular iron homeostasis and is a downstream target of mTOR, we hypothesized that TTP may regulate TfR1. Computational analysis of the 3’UTR of TfR1 revealed multiple putative AREs, some of which were found in a close proximity to or overlapped with IREs (Figure S5A). Consistently, we found TfR1 mRNA levels to be increased in TTP KO MEFs (Figure 4I) and with TTP siRNA in H9c2 cells (Figure S5B), while TTP overexpression reduced TfR1 levels in MEFs (Figure 4J). Moreover, TfR1 mRNA was significantly stabilized in TTP KO MEFs in the presence of actinomycin D (Figure 4K), confirming that TTP regulates TfR1 expression on the level of mRNA stability.

To determine whether TTP regulates TfR1 mRNA on its 3’UTR, we cloned the full-length 3’UTR of mouse TfR1 downstream of the firefly luciferase gene (TfR1-3’UTR-Luc, Figure 5A), and used renilla luciferase construct to normalize for transfection efficiency. Consistent with TTP suppression of TfR1 on its 3’UTR, transfection of the TfR1-3’UTR-Luc construct into WT MEFs overexpressing TTP led to a significant reduction in the luminescence signal (Figure 5B), while luminescence was increased in TTP KO MEFs (Fig 5C).

Figure 5. mTOR regulates TfR1 through TTP.

Figure 5

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(A) Schematic representation of TfR1-3’UTR-Luc construct with IREs indicated in white and putative AREs in grey. Luciferase assay of TfR1-3’UTR activity in WT MEFs overexpressing TTP (B) or TTP KO and WT MEFs (C), normalized to renilla luciferase (n=6). (D) mRNA levels of TfR1 in total RNA co-precipitated with TTP and IgG (control) antibodies in HEK293 cells. Four primer sets targeting different regions near the 3’UTR of TfR1 gene were used in the RNA co-IP assay and normalized to 18S as an internal control. Data are presented as fold enrichment over the IgG control (n=3). (E) mRNA levels of TfR1 in TTP WT and KO MEFs treated with rapamycin or vehicle control (n=12). (F) mRNA levels of ABCE1 and Lias in TTP WT and KO MEFs with and without rapamycin treatment (n=12). (G) TfR1 mRNA stability after 3-hour incubation with act D in WT and TTP KO MEFs in the presence or absence of rapamycin. Data are presented as fold change over TfR1 mRNA levels at time point zero in each respective group (n=3–6). (H) TfR1 mRNA levels in TTP WT and KO MEFs with and without DFO treatment (n=6). (I) Cell death as assessed by propidium iodine (PI) and Annexin V labeling and flow cytometry in TTP WT and KO MEFs treated with 250µM DFO or 250µM 2,2-BPD for 40 hours and normalized to the vehicle-treated control (n=4). (J) Cell death in TTP KO MEFs with or without 16-hour pre-treatment with rapamycin incubated with 250µM DFO or 250µM 2,2-BPD for 40 hours with or without rapamycin (n=4). Data are presented as mean ± SEM. * p<0.05 vs. control, # p<0.05 vs. rapamycin pre-treated control group. Rapa, rapamycin. See also Figure S5.

Finally, to assess the interaction between TTP and TfR1 mRNA, we performed an RNA co-immunoprecipitation (RNA Co-IP) experiment using the anti-TTP or control IgG antibody as bait and analyzing mRNA targets of TTP by qRT-PCR (Table 1), as described previously (Emmons et al., 2008). Since the monoclonal antibody used in the studies was raised against human TTP, the pull-down experiments were performed in human HEK293 cells. We designed four distinct primer sets for TfR1 to target regions near its 3’UTR, and two primer sets for each of the positive controls. We observed enrichment in the known targets of TTP (VEGF (Essafi-Benkhadir et al., 2007) and Pitx-2 (Briata et al., 2003)) in the TTP antibody group compared to the IgG, while no enrichment was observed for any of the negative controls tested (Table 1). TfR1 mRNA levels were significantly enriched in the TTP group compared to the IgG control (Table 1, Figure 5D), indicative of an interaction between TTP and TfR1 mRNA. Taken together, our results show that TTP is induced by rapamycin and regulates TfR1 expression at the posttranscriptional level.

Table 1. Interaction between TTP and TfR1 mRNA.

RNA-CoIP experiment using a specific TTP antibody or control IgG antibody as bait establishes physical interaction between TTP and TfR1 mRNA. Four distinct sets of primers were designed to target regions of TfR1 mRNA adjacent to its 3’UTR. Relative mRNA levels were assessed by qRT-PCR and are displayed as fold enrichment over IgG control. VEGF1 and Pitx-2 are used as positive controls, and β2-microglobulin, Fpn1, HPRT and 18S rRNA are used as negative controls. See also Figure S6

Gene Relative Expression
IgG		 Relative Expression
TTP		 Fold Enrichment
Over IgG Control
Transferrin Receptor
   TfR1-1 281.81 426.31 1.51
   TfR1-2 528.61 672.54 1.27
   TfR1-3 268.16 381.56 1.42
   TfR1-4 266.13 369.66 1.45
Positive Controls
   VEGF-1 1931.18 3678.42 1.90
   VEGF-2 2048.95 3982.25 1.94
   Pitx2-1 2082.60 1960.18 0.94
   Pitx2-2 690.53 984.72 1.43
Negative Controls
   β2MG 22080.25 19099.78 0.87
   Fpn1 836.35 828.85 0.99
   HPRT 39322.53 34227.51 0.87
   18S 52.92 55.49 1.04
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mTOR regulates iron homeostasis through TTP

Finally, we asked whether TTP mediates the changes in TfR1 expression observed with mTOR modulation. TTP KO MEFs were incubated with rapamycin, and the expression of TfR1 was assessed. In the TTP WT MEFs, rapamycin led to the expected ~50% decrease in TfR1 levels; however, in TTP KO MEFs the reduction in TfR1 expression was significantly blunted (Figure 5E). Furthermore, while the expression of ARE-containing iron-requiring proteins was reduced by rapamycin in WT MEFs, this response was greatly attenuated in TTP KO MEFs (Figure 5F). Finally, TfR1 mRNA stability was reduced in WT MEFs incubated with rapamycin, but not in TTP KO MEFs (Figure 5G). These results establish TTP as the downstream target of mTOR, whose activity is at least partially responsible for the mTOR-dependent regulation of TfR1.

While we have shown that TTP regulates TfR1 mRNA stability, the significance of TfR1 downregulation in the setting of iron deficiency and in respect to the IRP1/2 pathway is unclear. We hypothesized that TTP acts as a “brake” on the IRP1/2 system to prevent the unnecessary over-activation of iron uptake as the cell is reducing its iron use through TTP-dependent downregulation of iron-containing proteins. If correct, this hypothesis predicts exaggerated induction of TfR1 by iron deficiency in the absence of TTP, while higher levels of TTP would blunt the TfR1 upregulation by iron chelation. Consistently, we found that the degree of TfR1 upregulation was significantly greater in DFO-treated TTP KO MEFs compared to the chelated TTP WT MEFs (Figure 5H), and a combination of DFO and rapamycin, which had an additive effect on TTP upregulation (Figure S4F), led to attenuation of TfR1 induction compared to DFO alone (Figure S5C).

To assess the functional importance of TTP in regulation of cellular iron, we measured the survival of TTP KO MEFs in iron-deficient conditions. We hypothesized that TTP is protective in low iron states, thus its deletion will result in increased cell death upon iron chelation. Consistently, we observed a significant drop in cell viability of TTP KO MEFs treated with 250µM DFO and 250µM 2,2-BPD for 40 hrs (Figure 5I). In addition, TTP KO MEFs were exquisitely sensitive to serum deprivation (Figure S5D), which is the main source of iron in the growth medium (Kakuta et al., 1997), but not to glutamine deprivation (Figure S5D), indicative of tolerance of TTP KO MEFs to general cellar stressors. Supplementation of TTP KO MEFs with transferrin-bound iron has significantly, although not completely, reversed the effects of serum deprivation on cell viability, but had no effect on viability of TTP KO MEFs grown in serum-replete medium (Figure S5E). These findings suggest that TTP KO MEFs have reduced cell viability when deprived of iron compared to TTP WT MEFs.

To determine whether cell death in iron-deficient TTP KO MEFs is mediated by mTOR pathway, we pre-treated TTP KO MEFs with rapamycin for 6 hours, followed by co-incubation with DFO and 2,2-BPD. Surprisingly, rapamycin has significantly attenuated cell death due to iron chelation (Figure 5J). Analysis of cell cycle progression has revealed significantly higher baseline division rates in TTP KO MEFs compared to TTP WT MEFs (Figure S5F), and a defect in cell cycle arrest in response to DFO (Figure S5G,H). On the other hand, pre-treatment of TTP KO MEFs with rapamycin prior to chelation with DFO has equally suppressed cell cycle progression in TTP KO and TTP WT MEFs (Figure S5G,H), suggesting that rapamycin-dependent preservation of viability of TTP KO MEFs is due to the suppression of cell division in TTP KO MEFs.

In summary, our results suggest that mTOR regulates cellular iron homeostasis and TfR1 expression through TTP and its inhibition may function as a negative feedback loop to fine-tune the iron deficiency response and maintain cell viability in low-iron states.

mTOR and TTP regulate iron homeostasis in vivo

So far our data suggest that mTOR and TTP regulate expression of TfR1 and iron homeostasis in MEFs and H9c2 cells. To determine if these pathways play a role in regulation of iron homeostasis in vivo, we treated c57 black WT mice with 5 daily intraperitoneal (IP) injections of rapamycin to inhibit mTOR, and also studied iron regulation in triple TTP- and TNFα receptors 1 and 2 knockout (TTP/TNFR1/2 KO) mice. While mice with the deletion of TTP alone display profound inflammation primarily due to stabilization of TNFα (Taylor et al., 1996), TTP/TNFR1/2 triple-KO mice show no inflammatory phenotype for up to one year of age (Carballo and Blackshear, 2001) and thus represent a suitable model to study iron regulation.

Treatment of WT mice with IP rapamycin successfully inhibited mTOR signaling, as evidenced by the absence of pS6 expression (Figure S6A). Similar to the in vitro findings, nonheme and heme iron levels were increased (Figure 6A,B), while the expression of TfR1 and Fpn1 was reduced (Figure 6C,D), in the hearts of rapamycin-treated mice. TTP expression was also induced by rapamycin in the mouse hearts (Figure 6D), suggesting that this protein may potentially mediate TfR1 expression in vivo. Surprisingly, assessment of rapamycin effects on hepatic iron homeostasis revealed reduced non-heme iron levels (Figure S6B) with a decrease in TfR1, but preservation of Fpn1 expression (Figure S6C,D). This suggests that rapamycin primarily reduces the uptake of iron into the liver, leading to an export-dependent iron loss. Finally, rapamycin had no effect on the expression of systemic iron-regulatory hormone hepcidin and two of its upstream regulators, bone morphogenic protein 6 (BMP6) and hemojuvelin (HJV) (Figure S6E). Thus, rapamycin appears to primarily affect iron regulation on a cellular level both in vitro and in vivo.

Figure 6. In vivo regulation of iron homeostasis by mTOR and TTP.

Figure 6

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Non-heme (A) and heme (B) iron levels in the hearts of c57 black WT mice treated with 5 daily IP injections of rapamycin, 10mg/kg (n=6), or equal volume of DMSO vehicle. (C) TfR1 and Fpn1 mRNA levels in the hearts of rapamycin-treated mice (n=6). (D) Western blot analysis of rapamycin-treated mouse hearts. Densitometry analysis is presented below the Western blots (n=3). (E). Western blot analysis of TfR1 protein in the hearts of TTP/TNFR1/2 KO (designated as TTP KO) and matched littermate WT control mice (designated as TTP WT). Densitometry analysis is presented on the right to the Western blot (n=3). (F) Non-heme iron levels in the hearts of TTP KO and WT mice (n=4). (G) Western blot analysis of ferritin light and heavy chains (FtL + FtH) and Fpn1 proteins in the hearts of TTP KO and WT mice. Densitometry analysis is presented below the Western blots (n=4). (H) Model of cellular iron regulation by mTOR and TTP. Data are presented as mean ± SEM. * p<0.05. See also Figure S6.

We next assessed the effects of TTP deletion on iron regulation in TTP/TNFR1/2 KO and WT littermate control mice. Consistent with our in vitro findings, TfR1 protein levels were significantly induced in TTP-deficient mouse hearts (Figure 6E), which corresponded to an increase in cardiac non-heme iron content (Figure 6F) and elevated levels of iron storage protein ferritin (light and heavy chains) (Figure 6G). However, there was a reduction in the levels of Fpn1 protein with TTP deletion (Figure 6G). Analysis of iron homeostasis in TTP-deficient livers revealed no significant change in non-heme iron content (Figure S6F), TfR1or ferritin light chain protein (Figure S6G), while, similar to the heart, Fpn1 protein expression was suppressed. To determine if the suppression of Fpn1 protein was due to increased expression and subsequent degradation by hepcidin, we measured the levels of this peptide in TTP-deficient mouse livers. However, consistent with the unaltered hepatic iron levels, the protein content of hepcidin was similar in TTP/TNFR1/2 KO and WT mice (Figure S6H), suggesting a different mechanism for Fpn1 downregulation in the heart and liver. Taken together, our data show that both mTOR and TTP regulate TfR1 and iron homeostasis in vitro and in vivo, thus providing an iron-regulatory pathway that likely complements cellular functions of IRP1/2.

DISCUSSION

Iron is essential for the catalysis of many reactions and there appears to be an intimate link between iron and energy metabolism. Short-term iron deficiency is known to promote insulin sensitivity, while iron overload may lead to glucose intolerance (Dongiovanni et al., 2008; Liu et al., 2009). Iron accumulation is also linked to non-alcoholic fatty liver disease, and may cause alteration in the function of adipocytes and their ability to oxidize glucose (Dongiovanni et al., 2011). On a cellular level, circumstantial evidence points towards a cross-talk between energy sensing and iron regulatory pathways, as exemplified by a study by Galvez et. al. that identifies mTOR as a positive regulator of transferrin-bound iron uptake (Galvez et al., 2007), while iron deficiency was shown to inhibit mTOR signaling (Ndong et al., 2009; Ohyashiki et al., 2009). However, the direct and systematic studies of mTOR and iron regulation are lacking. Here, we describe a pathway that links mTOR, a major metabolic hub of the cell, to regulation of cellular iron uptake and flux via the modulation of TfR1 expression. The mechanism for the observed changes is at least partially mediated by the TZF protein TTP, which is induced by rapamycin, interacts with TfR1 mRNA and leads to its degradation (Figure 6H). Moreover, we provide the first evidence that TTP functions in mammals are similar to those of yeast Cth1p/Cth2p in regulating cellular iron through a reduction in the levels of iron-containing proteins and preservation of cell viability in iron deficiency.

While it is possible that in vitro changes in cellular iron content with mTOR modulation can be solely explained by differential rates of cellular division and “iron-dilution” effect, the increased iron levels in rapamycin-treated terminally-differentiated mouse hearts argue against this possibility. Moreover, the repressive effects of rapamycin on genes involved in cellular iron regulation are specific to this pathway and are not due to global transcriptional inhibition, as we found the levels of iron-independent genes either unaltered or even increased with rapamycin treatment. The magnitude of change in iron levels with mTOR modulation is modest, suggesting that mTOR may fine-tune cellular iron utilization to match the metabolic requirements of a cell. Alternatively, the modest change in iron with mTOR inhibition is in agreement with our hypothesis that TTP is involved in iron conservation response. While IRP1/2 activation enhances cellular iron uptake and reduces export leading to iron accumulation, TTP primarily affects the distribution of iron molecules already present in the cell without a large induction of iron import or storage. The exact role of TTP in regulation of iron homeostasis under various environmental conditions remains to be determined; however, our data highlight the importance of this pathway in low-iron states, as evidenced by the severe loss of cell viability in iron deficiency with TTP knockout.

The fact that TTP reduces TfR1 mRNA levels and stability is surprising. We show that TTP is strongly induced by low iron levels in two different cell lines and mouse hearts, and also with mechanistically distinct chelators. Thus, it appears counterintuitive that TTP signals to degrade TfR1 message while IRP1/2 concurrently stabilize this mRNA. A similar situation was described in yeast cells where Aft1p (the yeast homolog of IRP1/2) activates the transcription of FIT1 and FIT2 genes involved in iron uptake, whereas Cth2p promotes their degradation in response to iron deficiency (Puig et al., 2005). We hypothesize that a parallel activation of the two pathways serves to fine-tune TfR1 levels based on the severity of iron deficiency and the degree of a reduction in cellular iron utilization by TTP. As levels of iron-containing proteins decrease and more iron becomes available, the need for additional import of iron may go down, corresponding to a reduction in TfR1 levels. In other words, TTP may act as a “brake” on the IRP1/2 response to better match the rate of iron import with the actual needs of the cell and spare exogenous iron for use by other tissues. The relative contribution of TTP and IRP1/2 pathways to iron deficiency response remains to be determined; however, a significant sequence overlap between AREs and IREs, the binding sites for TTP and IRP1/2, respectively, suggests a possibility of steric hindrance effects. Moreover, the rapid activation of IRP1/2 and slower transcriptional induction of TTP by iron deficiency may provide an additional degree of regulation, with IRP1/2 system serving as a rapid response aimed at correcting iron deficiency by enhancing iron uptake. If unsuccessful, the TTP-mediated iron-conservation program will subsequently get induced, allowing cells to survive in chronic iron deficiency.

In addition to TTP, our data suggest the existence of another, yet unidentified, pathway mediating the mTOR-dependent changes in iron homeostasis. First, we observed a significant attenuation, but not a complete reversal of rapamycin effects in TTP KO MEFs. Second, rapamycin administration caused a very rapid and progressive reduction in TfR1 mRNA levels, reaching significance at four hours of treatment (n=18). Since the induction of TTP by rapamycin represents a later-stage response, a more acute mechanism, such as phosphorylation event or microRNA processing, may explain these findings. Moreover, it is presently unclear how TTP expression is regulated by mTOR. We found that induction of TTP by rapamycin is unaltered in MEFs with defective HIF or IRP1/2 signaling, suggesting that these pathways are not involved in mTOR-dependent regulation of TTP. It is also unlikely that rapamycin regulates TTP levels indirectly through an increase in cellular iron levels, since rapamycin and iron chelation with DFO have an additive, rather than an opposing, effect on TTP induction. Thus, other pathways downstream of mTOR may play a role in the regulation of TTP and cellular iron homeostasis. Moreover, our data do not exclude a possibility of independent roles for the mTOR and TTP pathways in regulation of TfR1 and iron homeostasis. Extensive characterization of molecular networks, environmental signals and downstream effectors of these pathways will shed light on their biological role in iron regulation.

In summary, we identified mTOR and TTP as new players in cellular iron homeostasis that regulate mRNA stability of TfR1 and iron-containing genes. Moreover, we described the existence of a Cth1p/Cth2p-like pathway in mammalian cells which may represent an important iron-deficiency response mechanism that functions in parallel with a well-established IRP1/2 regulatory system.

EXPREIMENTAL PROCEDURES

Cell culture and reagents

MEFs were grown in complete DMEM medium (Cellgro, VA) supplemented with 10% FBS (Invitrogen, CA) and 1% penicillin-streptomycin (P/S). For glutamine-deprivation experiments Cellgro DMEM without L-glutamine (Cat. # 15-013-CV) was used. Each of the genetically modified MEF lines was compared to the matched WT MEFs obtained from a littermate control mouse. TSC2 KO-V and TSC2 KO-TSC2 MEFs were described previously (Zhang et al., 2003). TTP KO (line 66) and TTP WT (line 67) MEFs were derived from littermate E14.5 embryos as described (Taylor et al., 1996). ARNT KO MEFs were a generous gift of Dr. Celeste Simon (Maltepe et al., 1997). IRP1 KO and IRP2 KO MEFs were kindly provided by Dr. Tracey A. Rouault. H9c2 cardiac myoblasts were purchased from ATCC and kept in complete DMEM medium (ATCC, VA) with 10% FBS and 1% P/S. H9c2 and MEF cells were treated with 20nM rapamycin (LC Labs, MA) for 24 or 48 hours, 150µM DFO or 200µM 2,2-BPD for 16–24 hours unless specifically noted, and 50µg/ml of FAC for 16 hrs (Sigma-Aldrich, USA).

siRNA treatment

H9c2 cells were transfected with siRNA using DharmaFect (Thermo Scientific, CO) reagent according to the manufacturer’s protocol. Rat TSC2 siGENOME siRNA (Thermo Scientific, CO) and rat TTP FlexiTube siRNA (Quiagen, CA) were used.

Quantitiative RT-PCR

RNA was isolated with RNA STAT-60 (TEL-TEST, Inc, TX), reverse-transcribed with a Random Hexamer (Applied Biosystems, CA), and amplified on a 7500 Fast Real-Time PCR system with SYBR Green PCR Master Mix (Applied Biosystems, CA). Primers were designed using Primer3 (v. 0.4.0) software to target sequences spanning an exon-intron-exon boundary. mRNA levels were calculated by the comparative threshold cycle method and normalized to β-actin and/or HPRT gene.

Western blot

Fifteen-30 µg of protein were resolved on SDS-PAGE gels and transferred to nitrocellulose membranes (Invitrogen, CA) which were probed with antibodies against pS6 (Cell Signaling, MA), TfR1 (Invitrogen, CA), Slc40A1 (Fpn1, Novus Biologicals, CO), TTP (Cao et al., 2004) and Tubulin (Abcam, MA). HRP-conjugated donkey anti-rabbit and donkey anti-mouse were used as secondary antibodies (Santa Cruz, CA) and visualized by Pierce SuperSignal Chemiluminescent Substrates.

Plasmid transfections

Plasmids were transfected into MEFs using Lipofectamine reagent (Invitrogen, CA) in OptiMEM (Cellgro, VA) for four hours, followed by incubation in complete, antibiotic-free medium for another 20 hours.

Non-heme and heme iron assays

Non-heme iron was measured as described (Rebouche et al., 2004). Briefly, equal amounts of protein were mixed with protein precipitation solution (1:1 of 1N HCl and 10% trichloroacetic acid) and heated to 95°C for 1h to release iron. Precipitated protein was removed by centrifugation at 4°C at 16,000 × g for 10 minutes, and the supernatant was mixed with the equal volume of chromogen solution (0.5mM ferrozine, 1.5M sodium acetate, 0.1% (v/v) thioglycolic acid) and the absorbance was measured on Spectra Max Plus microplate reader at 562 nm. Heme was quantified as described (Ward et al., 1984). Briefly, equal amounts of protein were mixed with 2M oxalic acid, heated to 95°C for 30 minutes to release iron from heme and generate protoporphyrin IX. Samples were then centrifuged for 10 min at 1,000 × g at 4°C to remove debris, and the fluorescence of the supernatant was assessed at 405nm / 600nm on Spectra Max Gemini fluorescence microplate reader.

55Fe uptake studies

55Fe (Perkin-Elmer, MA) was conjugated to nitriloacetic acid (NTA, Sigma-Aldrich, USA) and dissolved to the final concentration of 100–300nM in complete, serum-containing medium 24 hours prior to the beginning of the experiments. MEFs and H9c2 were incubated for 1 hour or 24 hours in the 55Fe-containing complete medium, washed 3 times with ice-cold 500mM BPS in PBS to remove the membrane-associated 55Fe and lysed with 1% Triton-X100 in TBS. The radioactivity of each sample was determined on Beckman scintillation counter and normalized to the protein content of each sample.

mRNA stability assay

MEFs were incubated in the complete medium supplemented with 5µM actinomycin D (Sigma-Aldrich, USA) for 3–6 hours. No drop in cell viability was observed at the end of the treatment. RNA was collected and mRNA levels were analyzed by qRT-PCR as described above.

RNA-CoIP

The protocol was adapted from Emmons et. al (Emmons et al., 2008) with the following modifications. HEK-293 cells were resuspended in Buffer A (10mM Tris–HCl pH 7.6, 1mM KAc, 1.5mM MgAc, 2mM DTT, 10µl/ml Protease Arrest inhibitors), and lysed using Power Gen 500 homogenizer (Fisher Scientific, PA), followed by centrifugation at 12,000 × g for 10 min at 4°C to remove debris. Protein G Sepharose Fast Flow beads (Sigma-Aldrich, USA) were incubated with human TTP (kindly provided by Dr. William Rigby, Dartmouth University) or IgG antibody in the IP Buffer (10mM Tris-HCl pH 7.6, 1.5mM MgCl2, 100mM NaCl, 0.5% Triton X-100, 10µl/ml Protease Arrest inhibitors) at 4°C with continuous rotation for 4 hours. Beads were then washed 6 times with cold IP buffer and incubated with HEK-293 lysate at 4°C with continuous rotation for 2 hours. RNA was collected and equal amounts were amplified by qRT-PCR. Expression of each gene was normalized to that of 18S and data expressed as fold enrichment over IgG control.

Luciferase Assay

Luciferase assay of TfR1-3’UTR-Luc and various ARE-deletion constructs was performed using the Dual Glo Luciferase Assay system (Promega) according to the manufacturer’s protocol. 3’ UTR of TfR1 was cloned into the pMIR-report vector containing the gene for firefly luciferase (Ambion) and confirmed by sequencing. The luminescence was quantified on a Berthold Technologies Luminometer (Germany) and the Dual-Glo® Stop & Glo® reagent was added to quench firefly luminescence and provide substrate for renilla luminescence that was used to normalize for transfection efficiency.

Cell death studies

Cell death was assessed by labeling with PI (Sigma-Aldrich, USA) and/or Alexa Fluor® 350-conjugated Annexin V (Molecular Probes, NY), and analyzed by flow cytometry in a FacsCanto flow cytometer (BD Bioscience). For cell-cycle studies, cells were incubated with rapamycin for 6 hours and co-incubated with DFO and rapamycin for 10-hours prior to collection. Cells were then collected by trypsinization, washed, and fixed in 70% ethanol at −20°C for two hours, followed by incubation in the PI staining solution (50µg/mL PI, 0.2mg/ml RNAse A, 0.1% Triton X-100 in PBS) at 37°C for 20 minutes. Cell cycle was then analyzed by flow cytometry and FlowJo 7.6 software.

Mouse studies

Wild-type C57BL/6 females were purchased from Jackson labs and treated with 5 daily intraperitoneal injections of rapamycin at 10mg/kg or DMSO. Mice were then anesthetized with 250mg/kg dose of freshly-prepared Tribromoethanol (Avertin) and the harvested organs were flash-frozen in liquid nitrogen. The animal studies were conducted in accordance with Northwestern University animal care guidelines.

Yeast studies

Yeast cth1Δcth2Δ cells transformed with pRS416 vector alone or expressing CTH2, TTP or TTP-C124R under the control of CTH2 promoter region were grown in synthetic media containing 300µM iron (+ Fe) or 100µM of the Fe2+-specific chelator bathophenanthroline disulfonic acid (− Fe), and RNA was extracted and analyzed by RNA blotting as previously described (Puig et al., 2005). Actin (ACT1) mRNA levels were used as a loading control. Wild-type yeast cells in exponential phase of growth were treated with 1µg/mL of rapamycin for 16 hours, following which mRNA was collected and expression of Cth1/2 genes analyzed by qRT-PCR.

Statistical analysis

Data are expressed as mean ± SEM. Statistical significance was assessed with the unpaired Student t test; a P value of less than 0.05 was considered statistically significant.

Supplementary Material

01

HIGHLIGHTS.

  • mTOR regulates iron homeostasis by altering expression of iron transporters

  • TTP regulates iron homeostasis, similar to yeast Cth1p/2p

  • mTOR regulates TfR1 and iron homeostasis through TTP

  • TfR1 mRNA is a target of TTP

ACKNOWLEDGMENTS

We thank Dr. Seth A. Brooks for help with TTP RNA-CoIP experiments, Drs. Sarah Rice, Navdeep Chandel and Elizabeth Leibold for the critical review of the manuscript, and Drs. William F.C. Rigby, M. Celeste Simon and Tracey A. Rouault for their generous gifts of the human TTP antibody, ARNT KO MEFs, and IRP1 and 2 KO MEFs respectively. This work was supported by the Northwestern University Flow Cytometry Facility and a Cancer Center Support Grant (NCI CA060553). M.B. and H.A. designed the research. M.B., A.K. and S.P. performed the experiments and data analysis. P.J.B and S.P. provided tools and reagents. M.B. wrote the manuscript, which all authors commented on. H.A. supervised the project. M.B. is supported by the American Heart Association 10PRE4430021. H.A. is supported by NIH grants K02 HL107448 and R01 HL087149.

Footnotes

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The authors declare no conflicts of interest

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