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. 2021 Jan-Mar;17(1):26–32. doi: 10.4183/aeb.2021.26

Regulation of Activating Transcription Factor 4 (ATF4) Expression by Fat Mass and Obesity-Associated (FTO) in Mouse Hepatocyte Cells

TM Mizuno 1,*, PS Lew 1
PMCID: PMC8417484  PMID: 34539907

Abstract

Context.

Abnormally increased hepatic glucose production contributes to hyperglycemia in diabetes. Interventions that suppress hepatic gluconeogenesis should be beneficial in improving glycemic control in patients with diabetes.

Objectives.

It has been suggested that hepatic FTO is involved in glycemic control by regulating gluconeogenesis. Both FTO and activating transcription factor 4 (ATF4) positively regulate the expression of gluconeogenic genes in the liver, suggesting the possibility that ATF4 mediates the stimulatory effect of FTO on hepatic gluconeogenesis. The present study aimed to determine the effect of altered expression or activity of FTO on Atf4 and gluconeogenic gene expression in hepatocyte cells.

Methods.

Mouse hepatocyte AML12 cells were treated with the FTO inhibitor rhein or transfected with an FTO-expressing plasmid. Levels of gluconeogenic glucose-6-phosphatase (G6pc) and Atf4 mRNA and protein were measured.

Results.

Rhein treatment significantly reduced G6pc mRNA levels as well as Atf4 mRNA and protein levels. Conversely, enhanced FTO expression caused an increase in G6pc and Atf4 mRNA levels.

Conclusions.

These findings support the hypothesis that hepatic FTO participates in the regulation of hepatic gluconeogenic gene and ATF4 expression. Reducing the activity of the hepatic FTO-ATF4 pathway may be beneficial in reducing hepatic glucose production and ameliorating hyperglycemia in diabetes.

Keywords: liver, gluconeogenesis, demethylase, transcription factor, type 2 diabetes

Introduction

Hepatic metabolism is critical for the maintenance of whole body glucose homeostasis. Gluconeogenesis is the major metabolic process that causes an increase in hepatic glucose production. Abnormally increased hepatic gluconeogenesis contributes to an impaired glycemic control in diabetes. Thus, understanding the mechanism underlying both normal and abnormally increased hepatic glucose production may suggest new and better anti-diabetes drug candidates.

Common genetic variants of the fat mass and obesity-associated (FTO) gene are associated with obesity and type 2 diabetes (1,2). The association between FTO variants and risk of type 2 diabetes involves obesity-dependent and –independent components (3,4). FTO is ubiquitously expressed and previous studies have suggested that FTO participates in the regulation of metabolism in a tissue specific manner (5). Levels of FTO (the rodent homologue of human FTO) mRNA and protein are altered by nutrient and hormonal signals in mouse liver and hepatocyte cells (6, 7). Enhanced FTO expression leads to increased levels of gluconeogenic genes, while FTO knockdown causes an opposite effect (7-9). Moreover, liver-specific FTO overexpression results in an increased blood glucose level and impaired glucose tolerance in mice (8). Conversely, FTO inhibitor treatment or liver-specific FTO deficiency reduced blood glucose level and improved glucose tolerance in mice (9). These findings suggest that hepatic FTO participates in the regulation of whole body glucose metabolism through alterations in hepatic gluconeogenic gene expression by responding to nutrient and hormonal signals.

Activating transcription factor 4 (ATF4) is a transcription factor that belongs to the cAMP-responsive element–binding (CREB) protein family and plays a pivotal role in the regulation of glucose homeostasis. Atf4 (the rodent homologue of human ATF4) deficiency causes a reduction in gluconeogenic gene expression in the liver and improves glucose homeostasis in mice (10-13). A recent study showed that levels of ATF4 protein are elevated in the liver of transgenic mice with enhanced liver specific FTO expression (14). These findings led to the hypothesis that ATF4 mediates the stimulatory effect of FTO on hepatic gluconeogenesis by up-regulating gluconeogenic gene expression. Conversely, it is hypothesized that reduced FTO activity in liver causes a reduction of gluconeogenic gene expression by suppressing ATF4 expression. To address this hypothesis, we investigated the effect of altered FTO expression and activity on Atf4 mRNA and gluconeogenic gene expression in mouse hepatocyte cell line.

Materials and Methods

Cell culture

The immortalized mouse hepatocyte cell line AML12 (ATCC®, CRL-2254™) was maintained in DMEM/Ham’s F-12 (1:1) medium supplemented with insulin (5 ng/mL), dexamethasone (40 ng/mL), sodium selenite (5 ng/mL), transferrin (5 µg/mL), fetal bovine serum (10%) and antibiotics at 37ºC in humidified 5% CO2 atmosphere.

To determine the effect of FTO inhibition on the expression of gluconeogenic genes and Atf4 mRNA and protein, subcloned cells were grown to 60-70% confluency and treated with the FTO inhibitor rhein (4,5-dihydroxyanthraquinone-2-carboxylic acid, Cayman Chemical, Ann Arbor, MI) at various concentrations (10 fM, 100 pM, 10 nM, 1 µM, 20 µM or 50 µM) for 24 h. Rhein is one of the major components of Rheum palmatum L and has the ability to inhibit demethylase activity of FTO (15,16). Cells were harvested at the end of the 24-h treatment. To determine the time course, cells were treated with rhein at 1 µM and harvested 0, 2, 4, 6, 12 and 24 h later. Rhein was initially dissolved in dimethyl sulfoxide (DMSO) and further diluted in 0.2-µm filter sterilized water to prepare stock solutions. Working solutions of rhein were freshly prepared by further diluting the stock solution in maintenance media immediately before use. DMSO solution (0.1%) was used as control vehicle.

To determine the effect of increased FTO level on the expression of gluconeogenic genes and Atf4 mRNA, cells were subcultured and grown overnight in the regular DMEM/F-12 (1:1) to 60-70% confluency and transfected with an FTO-expression vector (pAdTrack-CMV-FTO, 1 µg) using TurboFect (Thermo Fisher Scientific, Ottawa, ON). Mock-transfected cells were used as control. Cells were harvested 24 h after transfection.

RNA analysis

Total RNA was extracted from cells using TRIzol reagent (T9424, Sigma-Aldrich, St Louis, MO) and converted to cDNA using iScript™ Reverse Transcription Supermix (170-8840, Bio-Rad Laboratories, Hercules, CA). The mRNA expression levels were measured by real-time PCR using the ABI 7500 Fast thermal cycler (Applied Biosystems, Foster City, CA) as described previously (17). All primer pairs were designed using Primer Express software (Version 3.0, Applied Biosystems). Primer sequences used in this study were as follows: FTO (Accession number: NM_011936) forward: 5’-GACATCGAGACACCAGGATTAACA-3’ (exon 4),

FTO reverse: 5’-GTGAGCCAGCCAAAACACAGT-3’ (exon 5), G6pc (Accession number: NM_008061) forward: 5’-TCTTTCCCATCTGGTTCCATCT-3’ (exon 1), G6pc reverse: 5’-AATACGGGCGTTGTCCAAAC-3’ (exon 2), phosphoenolpyruvate carboxykinase 1 (Pck1, Accession number: NM_011044) forward: 5’-GCTGGCCCCGGGAGTCACC-3’ (exon 7),

Pck1 reverse:5’-TGCCGAAGTTGTAGCCGAAGAAGG-3’ (exon 10), Atf4 (Accession number: X61507) forward: 5’-CAGCGTTGGCCTTTGCA-3’ (exon 1), Atf4 reverse: 5’-CGCCAACACTTCGCTGTTC-3’ (exon 2), cyclophilin (Accession number: X52803) forward: 5’-AAGCATACAGGTCCTGGCATCT-3’ (exon 4), and cyclophilin reverse: 5’-TGCCATCCAGCCATTCAGT-3’ (exon 4/5). Levels of mRNA were normalized to cyclophilin mRNA levels, and are expressed as means (% of the control group) ± standard error of the mean (S.E.M.). All experiments were performed in triplicates and the coefficient of variation was less than 5% for each triplicate.

Assessment of cytotoxicity

Cytotoxicity of rhein was evaluated by the amount of lactate dehydrogenase (LDH) released from the cytoplasm. In brief, AML12 cells (1.7 x 105 cells in 100 µL medium) were plated in a 96-well culture plate and grown overnight under normal culture condition. Cells were treated with 0.1% DMSO as a vehicle control or rhein (10 nM or 1 µM) for 24 h. The culture plate was centrifuged at 514 x g for 4 min and 50 µL cell-free medium (culture supernatant) was used for LDH assay. Medium LDH levels were measured with the Pierce™ LDH Cytotoxicity Assay Kit (cat no. 88953, Thermo Scientific, Waltham, MA) and absorbance was measured at 490 nm with the background correction at 655 nm using a microplate reader (iMark Absorbance Microplate Reader, Bio-Rad Laboratories). Cytotoxicity(%) was calculated as follows, percent cytotoxicity(%) = [(Experimental OD490 – Spontaneous OD490)/(Maximum OD490 – Spontaneous OD490)] x 100.

Western blot analysis

Cells were lysed in protein lysis buffer (50 mM HEPES, 150 mM NaCl, 10% glycerol and 1% Triton X-100) supplemented with EDTA-free proteinase inhibitor cocktail (Roche Diagnostics, Indianapolis, IN). Total protein concentrations were determined by Bradford assay using serial diluted bovine serum albumin as standard. Proteins (12 µg) were separated on 10% stain-free gels (Bio-Rad Laboratories) at 250V and transferred to pre-wet low fluorescent polyvinylidene difluoride membranes (Bio-Rad Laboratories) at 100V for 1.5 h. The blots were rinsed with Tris-buffered saline (TBS) and images of stain-free blots were acquired using the ChemiDoc MP imager (Bio-Rad Laboratories) to check protein transfer efficiency and quality. After blocking with 5% non-fat milk in 1x TBS with 0.1% Tween 20 (TBST) for 1 h at room temperature, membranes were probed with primary antibodies for ATF4 (sc-390063, Santa Cruz Biotechnology, Dallas, TX, 1:1,000, mouse monoclonal) or pan-actin (no. 4968, Cell Signaling Technology, Danvers, MA, 1:1,000, rabbit polyclonal) at 4ºC overnight. After three washes in TBST, membranes were incubated with horseradish peroxidase-conjugated secondary antibody (anti mouse or rabbit) at 1:10,000 dilution in blocking buffer for 1 h at room temperature. Protein bands were visualized using an enhanced chemiluminescence and imaged using the ChemiDoc MP imager. The intensity of the bands was quantified by densitometry using the ImageLab software ver. 6 (Bio-Rad Laboratories) and normalized to total protein from stain-free blot for each sample. Blots were stripped after incubation with ATF4 antibody using low pH glycine buffer and reprobed with pan-actin antibody.

Statistical analysis

Data are presented as means ± S.E.M. Statistical analyses were performed by one-way analysis of variance (ANOVA) followed by Dunnett’s test (vs. control group). When data were not normally distributed, data were analyzed by Kruskal–Wallis test followed by Wilcoxon test with a Bonferroni correction (vs. control group). Comparisons between two groups were done using Student’s t-test (for parametric data) or Wilcoxon test (for nonparametric data). In all cases, differences were taken to be significant if p-values were below 0.05. Outliers were identified by Discordance test and omitted from the analysis.

Results

Dose-dependent and time-dependent effect of rhein treatment on gluconeogenesis gene expression in AML12 cells

Rhein treatment at 100 pM, 1 µM, 20 µM and 50 µM for 24 h significantly reduced levels of G6pc mRNA by 67.2% (P < 0.05), 65.2% (P < 0.0001), 64.6% (P < 0.005) and 84.3% (P < 0.0001), respectively, compared to the control DMSO treatment (Dunnett’s test, Fig. 1A). Level of G6pc mRNA was not significantly altered by rhein treatment at a lower concentration (10 fM, P = 0.65). Similarly, rhein treatment at 10 fM (P = 1.00) and 100 pM (P = 0.70) did not cause significant changes in Pck1 mRNA level, while higher concentrations of rhein (1, 20 and 50 µM) significantly reduced Pck1 mRNA level by 67.6% (P < 0.0001), 46.5% (P < 0.01) and 79.4% (P < 0.001), respectively, compared to the control DMSO treatment (Dunnett’s test, Fig. 1B). Rhein treatment at 10 nM and 1 µM did not significantly increase cytotoxicity compared to the control DMSO treatment as assessed by LDH release into the culture medium in AML12 cells (F [2, 69] = 2.1875, P = 0.1199 by one-way ANOVA, Fig. 1C). Since rhein treatment at 1 µM produced a marked reduction in G6pc mRNA level without causing a significant cytotoxicity, this dose was used in a time course study. Rhein treatment (1 µM) caused significant changes in G6pc mRNA level in AML12 cells in a time-dependent manner (H(5) = 11.68, P < 0.05 by Kruskal–Wallis test) with significant reductions after 2-h (52.1% reduction, P < 0.05), 12-h (48.1% reduction, P < 0.01) and 24-h (59.5% reduction, P < 0.005) treatment (Fig. 1D). Rhein treatment for 4 and 6 h at the same concentration did not cause significant changes in G6pc mRNA level (4 h: P = 0.34, 6 h: P = 0.51, Student’s t-test, Fig. 1D).

Figure 1.

Figure 1.

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Does-dependent and time-dependent effect of rhein treatment on gluconeogenesis gene expression in AML12 cells. A and B: Cells were treated with rhein (10 fM, 100 pM, 1 µM, 20 µM or 50 µM) for 24 h. DMSO (0.1%)-treated cells were used as control. Levels of G6pc and Pck1 mRNA were measured by real-time PCR. Values in control group were set to 100%. C: Cells were treated with rhein (10 nM or 1 µM) or DMSO for 24 h. Cytotoxicity (%) was assessed by LDH release into the culture medium. D: Cells were treated with rhein (1 µM) or DMSO for 2-24 h. Levels of G6pc mRNA were measured by real-time PCR. Values in control group were set to 100% at each time point. Data are means ± S.E.M. (n = 7-12/group, 24/group and 10-17/group in A and B, C and D, respectively). *: P < 0.05, #: P < 0.01, †: P < 0.005, ¶: P < 0.0001 by one-way ANOVA followed by Dunnett’s test (vs. control group, A and B), Student’s t-test or Wilcoxon test (D).

Effect of rhein treatment on Atf4 mRNA and protein expression in AML12 cells

To further determine the effect of FTO inhibition on Atf4 gene and protein expression, AML12 cells were treated with rhein in a separate study. Rhein treatment at 100 pM, 10 nM and 1 µM significantly lowered levels of Atf4 mRNA as well as G6pc mRNA in AML12 cells compared to the control treatment (Fig. 2A). There was a significant positive correlation between Atf4 mRNA and G6pc mRNA levels (r = 0.3487, P < 0.05). Western blot analysis showed that ATF4 protein levels were significantly reduced by rhein treatment (20 µM, Fig. 2B and C). Since rhein treatment significantly altered levels of pan-actin (F[3, 8] = 5.2260, P < 0.05 by one-way ANOVA), total protein for each sample from the stain-free blot images was used to normalize the intensity of the ATF4 protein bands (Fig. 2B).

Figure 2.

Figure 2.

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Reduced Atf4 and G6pc mRNA and protein levels in rhein-treated AML12 cells. Cells were treated with rhein (100 pM, 10 nM, 1 µM, 20 µM or 50 µM) for 24 h. DMSO (0.1%)-treated cells were used as control. A: Levels of Atf4 and G6pc mRNA were measured by real-time PCR. Values in control group were set to 100%. Data are means ± S.E.M. (n = 7-14/group). *: P < 0.05, †: P < 0.005, §: P < 0.001, &: P < 0.0005, ¶: P < 0.0001 by one-way ANOVA followed by Dunnett’s test. B: Representative images of ATF4 Western blot and stain-free protein blot. C: Quantification data of ATF4 protein normalized to total protein in stain-free blot. Values in control group were set to 100%. Data are means ± S.E.M. (n = 8-9/group). Groups that do not share the same letter are significantly different (P < 0.05 by Kruskal–Wallis test followed by Wilcoxon test with a Bonferroni correction).

Effect of enhanced FTO expression on Atf4 mRNA expression in AML12 cells

To increase FTO level, AML12 cells were transfected with an FTO-expression vector for 24 h. FTO mRNA levels were significantly increased 24 h after transfection of the FTO-expressing vector compared to the control mock transfection (P < 0.0001 by Wilcoxon test, Fig. 3A). We have shown that this protocol produced a significant 143.9% increase of FTO protein level in AML12 cells in the earlier study (7). Levels of Atf4 and G6pc mRNA were significantly increased by 58.3% and 119.3%, respectively, in FTO-overexpressing cells compared to the control cells (P < 0.05, Student’s t-test, Fig. 3B and C).

Figure 3.

Figure 3.

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Increased Atf4 and G6pc mRNA levels in Fto overexpressing AML12 cells. Cells were transfected with an Fto-expressing plasmid (1 µg) for 24 h (+Fto). Mock transfected cells were used as control. Levels of Fto (A), Atf4 (B) and G6pc (C) mRNA were measured by real-time PCR. Values in control group were set to 100%. Data are means ± S.E.M. (n = 11-15/group). *: P < 0.05, ¶: P < 0.0001 by Student’s t-test or Wilcoxon test.

Discussion

There is increasing evidence to support the role for FTO in the regulation of gluconeogenesis. However, the mechanism by which FTO regulates gluconeogenesis is not fully understood. It has been suggested that the regulation of gene expression by FTO is mediated by transcription factors such as forkhead box protein O1 (FOXO1), a major transcription factor controlling gluconeogenic gene expression (8,9,18). The transcription factor ATF4 belongs to the family of basic zipper-containing proteins and is a positive regulator of gluconeogenic gene expression. Levels of gluconeogenic genes and proteins were increased by enhanced Atf4 expression in primary mouse hepatocytes, while they were reduced by Atf4 knockdown (13). Complete absence of Atf4 resulted in improved glucose and pyruvate tolerance and reduced G6pc and Pck1 mRNA levels and fasting blood glucose in mice (10,19). Both hepatic Atf4 and extra-hepatic (i.e. osteoblastic) Atf4 contribute to these beneficial effects (11,13). Intriguingly, levels of ATF4 protein are elevated in the liver of transgenic mice in which FTO expression is selectively increased in the liver (14). In the present study, we demonstrated that FTO inhibition caused a reduction of Atf4 mRNA and protein levels in hepatocyte cells, while enhanced FTO expression led to an increase in Atf4 mRNA expression. These changes were paralleled by changes in gluconeogenic G6pc mRNA expression. These data support the hypothesis that hepatic ATF4 mediates the effect of FTO on glucose metabolism by modulating gluconeogenesis.

The mechanism by which ATF4 mediates FTO-induced gluconeogenic gene expression is currently unknown. Probable mechanisms involve the function of FTO as a nucleic acid demethylase. FTO is a Fe(II)- and 2-oxoglutarate-dependent dioxygenase that catalyzes demethylation of 3-methylthymine (m3T) in single-stranded DNA and 3-methyluracil (m3U) and 6-methyladenosine (m6A) in single-stranded RNA. To support the role of FTO in regulating Atf4 expression, there are 18 putative m6A sites with the consensus sequence RRACH in Atf4 mRNA (Genbank accession no: X61507). Consequently, FTO may promote demethylation at m6A sites on Atf4 mRNA, leading to an increased expression of Atf4 and its target genes such as gluconeogenic genes.

Secondly, a cooperative interaction between ATF4 and FOXO1 affects glucose metabolism. FOXO1 co-localizes with ATF4, physically interacts with ATF4, and promotes transcriptional activation of ATF4 (20). Although heterozygous null mutation in the FoxO1 gene alone or the Atf4 gene alone did not affect blood glucose level, double heterozygous resulted in reduced blood glucose level in mice (20). Thus, it is probable that FoxO1 and ATF4 regulates glucose metabolism in a synergistic manner by mediating FTO-induced gluconeogenesis.

Thirdly, ubiquitin-specific peptidase 14 (USP14) may be involved in the regulation of hepatic gluconeogenic gene expression by FTO and ATF4. USP14 is the deubiquitinating enzyme and stabilizes CREB to potentiate glucagon action, thereby promoting gluconeogenesis. Increased Usp14 expression in the liver led to increases in gluconeogenic gene expression and blood glucose in mice. Overexpression of hepatic Usp14 also caused impairments in glucose and pyruvate tolerance in mice, while hepatic Usp14 knockdown produced an opposite effect in high fat diet-induced obese mice (21). Enhanced Atf4 expression increases Usp14 promoter activity and levels of Usp14 mRNA and protein in mouse primary hepatocytes as well as mouse liver (21). Moreover, enhanced FTO expression promotes phosphorylation of CREB (18). Collectively, these findings suggest the possibility that USP14 mediates stimulatory effects of the FTO-ATF4 pathway on CREB-mediated gluconeogenic gene expression.

In conclusion, present findings suggest that hepatic FTO promotes gluconeogenesis by suppressing ATF4 expression. Abnormally increased activity of the FTO-ATF4 pathway may contribute to the increased hepatic glucose production, while the inhibition of this pathway (i.e. FTO inhibitor treatment) may be beneficial in suppressing hepatic gluconeogenesis and ameliorating hyperglycemia in diabetes. The present study opens a new avenue of research to determine whether hepatic FTO-ATF4s pathway is a viable target for diabetes treatment.

Conflict of interest

The authors declare that they have no conflict of interest.

Acknowledgment

This work was supported by a grant from University of Manitoba (University Research Grants Program).

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