Pathophysiological significance of impaired KAT7-dependent histone H3K14 acetylation during zinc deficiency

Abstract

Zinc is an indispensable micronutrient for optimal physiological functions, and zinc deficiency has been implicated in the pathogenesis of various human diseases. One potential mechanism underlying such pathogenic effects is the alteration of gene expression caused by zinc deficiency; however, the details of this process remain largely unexplored. Here, we show that during zinc deficiency, the histone acetyltransferase KAT7 loses its enzymatic activity, leading to the attenuated acetylation of histone H3 at Lys14 (H3K14ac) at enhancer regions. Physiologically, the decrease in H3K14ac leads to the upregulation of the expression of ZIP10, a plasma membrane-localized zinc transporter, thereby facilitating the import of extracellular zinc to maintain cellular zinc homeostasis. Moreover, prolonged zinc deficiency in mice induced by a zinc-deficient diet or high-fat diet, accompanied by decreased H3K14ac levels in the liver, upregulated the expression of genes associated with intracellular lipid droplet formation, leading to the accumulation of lipids within liver tissue. Our findings demonstrate that cells respond to zinc deficiency by converting it into an epigenetic signal that drives physiological or pathophysiological biological processes.

This study identifies KAT7 as a key factor converting zinc deficiency into reduced H3K14ac. This signal drives both physiological zinc homeostasis and, upon prolonged deficiency, pathophysiological liver fat accumulation.

Introduction

Zinc is an essential micronutrient that plays a pivotal role in maintaining the structural integrity and enzymatic activities of numerous proteins. Computational analysis revealed that as much as 10% of the human genome encodes a zinc-binding proteins¹, suggesting the importance of maintaining zinc homeostasis in the human body. Nevertheless, zinc deficiency occurs, arising from various factors, such as inadequate zinc intake, impaired zinc absorption, and aging². The clinical manifestations of zinc deficiency in humans were first reported in the 1960s^3,4, and since then, numerous studies have shown that zinc deficiency is strongly associated with the development of various diseases. The classical symptoms of zinc deficiency include skin diseases, mental disorders, immune dysfunction, and growth retardation⁵. Recently, emerging evidence has highlighted correlations between reduced zinc levels and diseases such as cancer^6–12, inflammatory bowel disease¹³, and chronic kidney disease¹⁴. However, the role of zinc in the pathogenesis of these conditions and the underlying molecular mechanisms remain to be fully elucidated.

To overcome zinc-deficient conditions, cells need to import zinc from extracellular sources. The Zrt-/Irt-like protein (ZIP) family is a group of zinc ion transporters¹⁵. Among the 14 human ZIP family proteins, 10 ZIPs (ZIP1-6, 8, 10, 12, and 14) are thought to be localized on the plasma membrane and to be involved in the import of extracellular zinc. Upon zinc deficiency, the abundance of ZIPs on the plasma membrane increases to facilitate zinc uptake as a stress response to maintain cellular zinc homeostasis. One related mechanism involves the upregulation of gene expression through transcriptional control. For example, our previous research demonstrated that zinc deficiency activates the transcription factor ATF6 via the endoplasmic reticulum (ER) stress response, thereby promoting the transcription of ZIP14¹⁶. In addition, it has been reported that ZIP10 expression increases during zinc deficiency through transcriptional control mediated by the transcription factor MTF1¹⁷. However, the detailed molecular mechanisms remain largely unclear.

Histone posttranslational modifications (PTMs) are among the key regulators that govern transcriptional control¹⁸. Each histone PTM is balanced via the regulation of a myriad of enzymes referred to as “writers” and “erasers”; writers catalyse the formation of specific types of PTMs, and erasers remove the PTMs. Interestingly, multiple writers and erasers possess zinc-binding domains¹⁹, suggesting the critical role of zinc in regulating histone PTMs. Nevertheless, little is known about the dynamics of these writers and erasers during zinc deficiency.

In this study, we observed a drastic decrease in histone H3K14ac during zinc deficiency. Mechanistically, this decrease was driven by the loss of activity of the zinc-dependent writer KAT7, which catalyses H3K14ac formation. In cultured cells, a decrease in H3K14ac led to the transcriptional upregulation of ZIP10, thereby importing zinc from extracellular sources to maintain cellular zinc homeostasis. At the organismal level in mice, chronic zinc deficiency in the liver induced the upregulation of the expression of genes involved in lipid droplet synthesis pathways and promoted lipid accumulation in liver tissue via a reduction in KAT7 activity and H3K14ac. Additionally, we showed that a high-fat diet led to a decrease in hepatic zinc and H3K14ac levels, suggesting that these reductions may more universally contribute to hepatic lipid accumulation. Our data demonstrate that cells respond to zinc deficiency stress by converting it into an epigenetic signal to drive cellular responses.

Results

Global decrease in histone H3K14ac during zinc deficiency

We previously reported that the activation of the transcription factor ATF6 elicits the upregulation of ZIP14 expression in HepG2 cells during zinc deficiency¹⁶. To investigate the impact of ATF6 on global expression changes in genes other than ZIP14, we examined the presence of ATF6-binding motifs on the promoters of genes whose expression was upregulated under zinc-deficient conditions. The Gene Expression Omnibus (GEO) repository contains four datasets reporting gene expression profiles in various types of cells during zinc deficiency, including a dataset from our laboratory^16,20–22. We found that, across all the datasets, fewer than 2% of the genes presented an ATF6-binding motif²³ (ER stress-responsive element, ERSE) within their promoter regions (Fig. S1a). These findings indicate that mechanisms other than the ATF6 axis contribute to the alterations in gene expression provoked by zinc deficiency.

In this study, we focused on the role of histone PTMs, which are critical factors responsible for gene expression changes whose association with zinc deficiency remains unexplored. First, we systematically analysed whether zinc deficiency stress affects the major histone PTMs implicated in gene expression in HEK293A cells (promoter marks: H3K4me3, H4K5ac, and H4K12ac; enhancer marks: H3K4me, H3K14ac, and H3K27ac; gene body marks: H3K36me3 and H3K79me3; and heterochromatin marks: H3K9me3 and H3K27me3). We found that treatment with N,N,N’,N’-tetrakis(2-pyridylmethyl)ethylenediamine (TPEN), a cell-permeable zinc chelator, selectively attenuated the signal for H3K14ac; however, no other marks were affected (Fig. 1a). This reduction in H3K14ac was observed across diverse cellular contexts (Fig. S1b, c); this reduction was also evident when cells were cultured in zinc-deficient medium (Fig. 1b), where the reduction was reversed by the addition of zinc and remained unaltered by the addition of other metals (Fig. 1c, d). Consistently, treatment with zinc-saturated TPEN failed to reduce H3K14ac levels (Fig. S1d), highlighting that zinc availability is indispensable for maintaining the H3K14ac level. Furthermore, the H3K14ac signal was restored in a time-dependent manner by the resupplementation of zinc under conditions of zinc deficiency (Fig. 1e). Collectively, these results suggest that zinc reversibly regulates H3K14ac.

Fig. 1. Global decrease in histone H3K14ac during zinc deficiency.

a Immunoblot analysis of HEK293A cells treated with TPEN for the indicated times. b Immunoblot analysis of HEK293A cells cultured in the indicated medium. c Immunoblot analysis of HEK293A cells cultured in the indicated medium supplemented with each metal at 20 µM for 48 h. d Immunoblot analysis of HEK293A cells cultured in zinc-deficient medium supplemented with the indicated concentration of zinc for 48 h. The graph shows the mean ± SEM of the H3K14ac level normalized to H3. The H3K14ac level in zinc-containing normal medium was normalized to 1; x-axis, logarithmic scale. Statistical annotations below the ZD curve represent comparisons of each ZD treatment group to the 0 µM ZD reference group. Statistical annotations above the ZN curve represent comparisons between the ZD group and the ZN control group at the indicated concentrations. e Immunoblot analysis of HEK293A cells cultured in zinc-deficient medium for 36 h and additional cultures supplemented with 20 µM zinc for the indicated times. The graph shows the mean ± SEM of the H3K14ac level normalized to H3. The H3K14ac level in the culture medium was normalized to 1. Data at 0.5 h were omitted from the graph to avoid complexity. The data omitted from the graph were included in the statistical analysis. Statistical annotations below the ZD curve represent comparisons of each time point to the 0 h ZD reference group. Statistical annotations above the ZN curve represent comparisons between the ZD group and the ZN control group at the corresponding time point. The immunoblot images are representative of three independent experiments (a–e). The graphs are presented as means ± SEM (n = 3 biological replicates). Statistical analysis was performed using two-sided Student’s t-test followed by Bonferroni post hoc correction (d, e). n.s. not significant, ZN zinc-normal medium, ZD zinc-deficient medium. Source data are provided with this paper.

HDACs deacetylate H3K14ac during zinc deficiency

We next investigated the molecular mechanisms underlying the reduction in H3K14ac during zinc deficiency. In general, the histone acetylation status is balanced by histone deacetylases (HDACs) and histone acetyltransferases (HATs). We hypothesized that the activation of HDACs or the inactivation of HATs results in a decrease in H3K14ac during zinc deficiency.

First, we investigated the involvement of HDACs. HDACs are classified into two distinct categories: the classic HDAC family (HDAC1-11) and the sirtuin family of NAD-dependent HDACs (SIRT1-7). To identify which HDACs are responsible for the deacetylation of H3K14ac during zinc deficiency, we treated cells with specific inhibitors of each HDAC family. Treatment with nicotinamide (NAM), a paninhibitor of the sirtuin family, failed to inhibit the reduction in H3K14ac (Fig. 2a). In contrast, treatment with trichostatin A (TSA), a paninhibitor of the HDAC family, completely prevented this decrease (Fig. 2b), suggesting that the classic HDAC family deacetylates H3K14ac during zinc deficiency.

Fig. 2. HDACs deacetylate H3K14ac during zinc deficiency.

a Immunoblot analysis of HEK293A cells treated with 10 µM TPEN or zinc-saturated TPEN for two h after pretreatment with NAM for 1 h. b Immunoblot analysis of HEK293A cells treated with 10 µM TPEN or zinc-saturated TPEN for 2 h after pretreatment with TSA for 1 h. c Immunoblot analysis of HEK293A cells. The cells were treated with the indicated siRNA and cultured for 48 h. The cells were treated with 10 µM TPEN for the indicated times and subjected to immunoblot analysis. d Cell lysate–based HDAC activity assay. Cell lysates treated with 10 µM TPEN for the indicated times were prepared. The lysates were incubated with purified histones with or without 2 nM TSA for the indicated times. The graph shows the mean ± SEM of the H3K14ac level normalized to H3. The H3K14ac level in the absence of cell lysate was normalized to 1. The results of statistical analyses comparing the reference group (no lysate) to other groups are shown on the bars. The results of statistical analyses comparing data from the same incubation times are also shown. e In vitro HDAC assay. HDAC-FLAG immunoprecipitated from HEK293A cells treated with 10 µM TPEN or zinc-saturated TPEN for two h was subjected to an in vitro HDAC assay. f In vitro HDAC assay. HDAC1-FLAG or HDAC6-FLAG immunoprecipitated from HEK293A cells treated with or without 10 µM TPEN for the indicated h was subjected to an in vitro HDAC assay. The graph shows the mean ± SEM of the H3K14ac level normalized to H3. The H3K14ac level in the absence of immunoprecipitates was normalized to 1. The results of statistical analyses comparing data among different TPEN treatment conditions at the same incubation time point are shown. The immunoblot images are representative of three independent experiments (a–f). The graphs are presented as means ± SEM (n = 3 biological replicates). Statistical analysis was performed using two-sided Student’s t-test followed by Bonferroni post hoc correction (d, e). n.s. not significant. Source data are provided with this paper.

Next, we attempted to identify the specific member of the HDAC family responsible for the deacetylation of H3K14ac during zinc deficiency. However, the knockdown of individual members of the HDAC family did not prevent the TPEN-induced deacetylation of H3K14ac (Figs. 2c, S2a). In addition, SAHA, a paninhibitor of the HDAC family, but not other subtype-specific HDAC inhibitors (TMP195: HDAC4/5/7/9; tubastatin: HDAC6; UF010: HDAC1/2/3/6/8/10; MS275: HDAC1/2/3/8) effectively inhibited the TPEN-induced deacetylation of H3K14ac (Fig. S2b, c). These data suggest that several HDAC family members cooperatively contribute to the deacetylation of H3K14ac during zinc deficiency.

To examine the possibility that zinc-deficient stress enhances the deacetylation activity of members of the HDAC family towards H3K14ac, we measured HDAC activity via a cell lysate–based HDAC activity assay using purified histones. The basal HDAC activity for H3K14ac in cell lysates did not differ in the presence or absence of TPEN, suggesting that zinc deficiency in cells enhances the deacetylation of H3K14ac through an independent mechanism that regulates HDAC activity (Fig. 2d). To further investigate whether zinc deficiency directly alters HDAC enzymatic activity, we next examined the activity of individual HDACs toward H3K14ac using an in vitro HDAC assay. While HDAC1 and HDAC6 exhibited measurable deacetylase activity, their enzymatic functions were not enhanced under zinc-deficient conditions (Fig. 2e), which was further supported by a more detailed time-course analysis (Fig. 2f). Moreover, simultaneous knockdown of HDAC1 and HDAC6 did not prevent the reduction of H3K14ac induced by zinc-deficient stress (Fig. S2d). These data suggest that certain members of the classic HDAC family are needed to deacetylate H3K14ac during zinc deficiency but are not direct regulatory targets of zinc deficiency.

Loss of KAT7 activity during zinc deficiency

Given that HDAC activity is not enhanced under zinc-deficient conditions, we hypothesized that zinc deficiency suppresses the activity of HATs towards H3K14. First, to identify the specific HAT responsible for H3K14ac in HEK293A cells, we performed an siRNA screen using siRNAs targeting 17 human HATs and found that the knockdown of K (lysine) acetyl transferase 7 (KAT7) led to a significant decrease in H3K14ac (Figs. 3a, S3a). This decrease was also observed when the cells were treated with WM-3835, a specific inhibitor of KAT7²⁴ (Fig. S3b), indicating the crucial role of KAT7 in the acetylation of H3K14. Thus, we focused on the regulatory mechanisms of the KAT7 HAT activity during zinc deficiency.

Fig. 3. Zinc coordination in the KAT7 MYST domain regulates HAT activity towards H3K14.

a Immunoblot analysis of HEK293A cells treated with the indicated siRNAs for 48 h. b FLAG-KAT7 immunoprecipitated from HEK293A cells transfected with the indicated plasmids and treated with or without 10 µM TPEN for 2 h was subjected to an in vitro HAT assay. c Upper panels, FLAG-KAT7 WT immunoprecipitated from HEK293A cells cultured in the indicated medium for 48 h was subjected to an in vitro HAT assay. Lower panels, immunoblot analysis of HEK293A cells transfected with FLAG-KAT7 WT and incubated with the indicated medium. ZN, zinc-normal medium; ZD, zinc-deficient medium; ZS, zinc-deficient medium supplemented with 20 µM ZnCl₂. d Left, schematic representation of the KAT7 domain structure and amino acid sequences in zinc finger domains. Right, 3D structures of the KAT7 MYST domain (Protein Data Bank, 6MAJ). Light blue, zinc finger domain. The amino acids coordinating zinc are highlighted. e FLAG-KAT7 WT or mutants immunoprecipitated from HEK293A cells were subjected to an in vitro HAT assay. f RecKAT7^HAT with or without mutation was subjected to an in vitro HAT assay. g ZnAF-2 assay. The zinc content in recKAT7^HAT was measured. h FLAG-KAT7 WT or C371A immunoprecipitated from HEK293A cells treated with or without 10 µM TPEN for two h was incubated with the indicated concentration of ZnCl₂ for 30 min. The samples were subjected to an in vitro HAT assay. The results of statistical analyses comparing the reference group (with no additional zinc) to other groups are shown on the plots. The results of statistical analyses comparing data collected following 10 pM zinc supplementation are shown. The immunoblot images are representative of three independent experiments (a–c, e, f, h). The graphs are presented as means ± SEM (n = 3 biological replicates). Statistical analysis was performed using two-sided Student’s t-test (g), two-sided Student’s t-test followed by Bonferroni post hoc correction (h). n.s. not significant. Source data are provided with this paper.

Previous reports have shown that KAT7 undergoes degradation in response to DNA damage and lipopolysaccharide stimulation via the ubiquitin‒proteasome pathway^25,26. However, our investigation indicated that the effect of TPEN on the protein level of KAT7 was limited (Fig. S3c). While treatment with the proteasome inhibitor MG132, but not the autophagy inhibitor bafilomycin A1, inhibited the TPEN-induced slight decrease in the protein amount of KAT7, both inhibitors had little effect on the decrease in H3K14ac. These results suggest that the degradation of KAT7 is not involved primarily in the reduction in H3K14ac during zinc deficiency.

Several reports have demonstrated that KAT7 localizes to the nucleus to acetylate histones^27,28. We investigated the possibility that the subcellular localization of KAT7 changes during zinc deficiency. However, immunofluorescence analysis of endogenous KAT7 revealed that KAT7 localized to the nucleus after TPEN treatment (Fig. S3d), suggesting that a change in subcellular localization is not involved in the regulation of KAT7 during zinc deficiency.

Interactome analyses demonstrated that KAT7 forms a multisubunit complex that includes BRPF1/2/3, JADE1/2/3, ING4/5, and MEAF6^29–35. Given that these interactors have been reported to directly bind to KAT7 to increase its HAT activity, we hypothesized that zinc deficiency leads to the dissociation of KAT7 from the complex. The systematic knockdown of each interactor revealed that BRPF2 and MEAF6 play critical roles in the acetylation of H3K14 in HEK293A cells (Fig. S3e). However, treatment with TPEN did not dissociate KAT7 from each component (Fig. S3f). These findings suggest that the disruption of the KAT7 complex is not a cause of the reduction in H3K14ac during zinc deficiency.

Since working hypotheses based on previous reports were not applicable in this case, we next investigated whether zinc deficiency directly affects KAT7 activity by employing an in vitro HAT assay. The treatment of cells with TPEN strongly inhibited the HAT activity of immunoprecipitated KAT7 to a similar level as that of the HAT-dead EQ mutant²⁴ (Fig. 3b). In addition, the KAT7 HAT activity in cells cultured in low-zinc-medium was attenuated in a time-dependent manner, which was followed by the deacetylation of H3K14ac (Fig. 3c). Consistently, no reduction in KAT7 activity was observed upon zinc-saturated TPEN treatment (Fig. S3g). These results suggest that KAT7 loses its enzymatic activity under zinc deficiency.

Zinc coordination in the KAT7 MYST domain regulates HAT activity towards H3K14

KAT7 possesses two zinc finger domains: one is located within the MYST-type HAT domain (MYST domain), and the other is in the N-terminal region (Fig. 3d). We speculated that the loss of zinc coordination during zinc deficiency affects KAT7 HAT activity. To test this possibility, we examined whether the zinc finger domains in KAT7 are needed for KAT7 HAT activity. Depleting KAT7 using the CRISPR‒Cas9 system resulted in decreased H3K14ac levels in HEK293A cells (Fig. S3h). Adding back KAT7 wild-type (WT) or KAT7 mutants with mutations in the N-terminal zinc finger (C185A, C190A, H203A, and C209A) into the KAT7 knockout HEK293A cell line restored the H3K14ac level (Fig. S3h). In contrast, the reintroduction of KAT7 mutants with mutations in the zinc finger within the MYST domain (C368A, C371A, H384A, and C388A) failed to do so. In support of that finding, immunoprecipitated KAT7 mutants harbouring a zinc finger mutation in the MYST domain showed no HAT activity for H3K14 in vitro, whereas KAT7 mutants with a zinc finger mutation in the N-terminal region presented HAT activity similar to that of WT KAT7 (Fig. 3e). These results suggest that the zinc finger within the MYST domain and zinc-binding amino acid residues are needed for KAT7 HAT activity.

Next, we analysed the properties of recombinant proteins purified from E. coli. We attempted to obtain full-length KAT7 but were unsuccessful. Thus, we purified the recombinant KAT7 HAT domain (recKAT7^HAT), which includes the MYST domain. Consistent with the results above, recKAT7^HAT WT, but not the recKAT7^HAT EQ mutant or recKAT7^HAT C371A mutant, exhibited HAT activity towards H3K14 (Fig. 3f). Notably, a ZnAF-2 assay, a method to quantify the zinc content coordinated within recombinant proteins, demonstrated that recKAT7^HAT C371A coordinates with substantially less zinc than recKAT7^HAT WT does (Fig. 3g), suggesting the critical role of proper zinc coordination in the MYST domain for KAT7 HAT activity.

To further assess the role of zinc in KAT7, we manipulated the zinc metalation status of KAT7 in vitro and examined the effect on HAT activity. First, we attempted to demetallate zinc from KAT7 in vitro. However, even when recKAT7^HAT WT was treated with TPEN in vitro, recKAT7^HAT WT still coordinated with zinc and exhibited HAT activity (Fig. S3i, j). Consistent with these findings, in vitro TPEN treatment of KAT7 immunoprecipitated from HEK293A cells did not affect HAT activity (Fig. S3k). These results suggest that KAT7 coordinates zinc much more strongly than TPEN does and that the elimination of zinc from KAT7 in vitro was unsuccessful.

Given that in vitro TPEN treatment did not change the KAT7 activity, we considered that improper zinc coordination during the production or folding of KAT7 may lead to a decrease in its intracellular activity. To investigate this possibility, we treated cells with the proteasome inhibitor MG132 and the protein synthesis inhibitor cycloheximide (CHX) to block protein degradation and synthesis, respectively, and then subjected the cells to zinc depletion. Although the differences were not statistically significant, both H3K14ac levels and KAT7 activity showed a slight increase at 1 h after TPEN treatment when cells were co-treated with MG132 and CHX compared with the control, but by 2 h both parameters had declined to control levels despite the inhibitor treatment (Fig. S3l). These observations suggest that improper zinc coordination during KAT7 production or folding contributes partly to the observed activity loss, yet the predominant contribution likely comes from zinc dissociation in mature KAT7.

Next, we examined the effect of in vitro zinc repletion. As expected, in vitro zinc addition to KAT7 WT immunoprecipitated from HEK293A cells treated with TPEN, but not KAT7 C371A, restored the HAT activity in a dose-dependent manner (Fig. 3h). Collectively, these results suggest that KAT7 loses zinc from the MYST-type zinc finger domain during zinc deficiency, which leads to decreased HAT activity towards H3K14.

Loss of H3K14ac on the enhancer induces ZIP10 expression to maintain cellular zinc homeostasis

Next, we investigated the physiological role of transcriptional changes through the drastic reduction in H3K14ac during zinc deficiency. We and others have previously shown that H3K14ac regulates gene expression at enhancers^36–38 and at promoters^39,40. ChIP–seq analysis revealed that H3K14ac signals were markedly decreased at enhancer and promoter regions in cells cultured under zinc-deficient conditions (Figs. 4a, S4a). The zinc deficiency–sensitive loci showed substantial overlap with those exhibiting H3K14ac loss following KAT7 knockdown, indicating that zinc deficiency attenuates KAT7 activity (Fig. S4b). Notably, a greater reduction of H3K14ac at enhancer regions was associated with larger gene expression changes, involving both up- and down-regulated genes (Fig. 4b). In contrast, for genes with H3K14ac loss at promoters or at both promoters and enhancers, stronger H3K14ac reduction correlated with transcriptional down-regulation (Fig. S4c). These correlations highlight the functional importance of KAT7-mediated H3K14 acetylation in regulating gene expression.

Fig. 4. Loss of H3K14ac on the enhancer induces ZIP10 expression to maintain cellular zinc homeostasis.

a H3K14ac ChIP-seq reads around enhancers. b Correlation between H3K14ac signal and gene expression changes during zinc deficiency. Central line: median; boxes: 25th–75th percentiles (IQR); whiskers: 1.5 × IQR. c Heatmaps showing gene expression changes during zinc deficiency and H3K14ac signal at enhancers. d, e qPCR of HeLa cells cultured in the indicated medium for 48 h (d) or 36 h (e) (n = 3 biological replicates). f Membrane protein biotinylation assay of HeLa cells cultured in the indicated medium for 48 h. The immunoblot images are representative of three independent experiments. g Left: H3K14ac ChIP-seq data. Right: ChIP‒qPCR of HeLa cells cultured in the indicated medium for 48 h (n = 3 biological replicates). h, i Left: ZnDA-3H/HTL-TMR ratio images. HeLa cells were cultured in the indicated medium for 12 h and treated with 0.5 µM TSA (h), or transfected with the indicated siRNA for 24 h and cultured in ZD for 48 h (i). Changes in zinc concentration before and 10 min after addition of ZnCl₂ are shown. Scale bars, 10 µm. Middle: Cytoplasmic labile zinc influx. Triangles, individual cells; circles, experiment means; rectangles, overall mean. Right: Cytoplasmic labile zinc concentration. Triangles, individual cells; circles, experiment means; diamonds, overall mean. j Immunoblot of HeLa cells transfected with the indicated siRNAs and cultured in the indicated medium for 48 h. ZD→Zinc samples were further incubated with 10 µM ZnCl₂ for 4 h. k HeLa cells transfected with the indicated siRNAs were cultured in the indicated medium for 48 h, treated with or without 10 µM ZnCl₂ for 9 h, followed by EdU assay (n = 3 biological replicates). White lines indicate nuclear boundaries. l Schematic of the model reported in this study. TSA was added every 12 h (e–i). The graph presents the mean ± SEM. Statistical analysis was performed using two-sided Mann–Whitney U test (b, j), two-sided Student’s t-test (d), Student’s t-test followed by Bonferroni post hoc correction (g), one-way ANOVA followed by Dunnett’s test (e, h, i). n.s. not significant. ZN, zinc-normal medium, ZD zinc-deficient medium, ZS zinc-deficient medium supplemented with 10 µM ZnCl₂.

Next, we speculated that genes responsible for maintaining cellular zinc homeostasis are regulated by H3K14ac. The Molecular Signatures Database (MSigDB) lists genes involved in zinc homeostasis (GO:0006829, zinc ion transport)^41,42. We systematically integrated our gene expression profile data with H3K14ac ChIP-seq data under conditions of zinc deficiency and KAT7 knockdown. This analysis revealed that multiple genes were transcriptionally altered, with decreased H3K14ac signals at their enhancer regions under zinc-deficient conditions and upon KAT7 knockdown (Fig. 4c), suggesting that these genes may be regulated by a reduction in H3K14ac. Among these genes, we became interested in plasma membrane-localized ZIPs, which play a key role in importing zinc from extracellular sources. Quantitative PCR (qPCR) analysis results confirmed our previous microarray findings (Fig. 4d). In particular, the notable upregulation of ZIP10 expression led us to focus on the physiological role of upregulated ZIP10 expression during zinc deficiency.

In HeLa cells, zinc supplementation suppressed the decrease in H3K14ac during zinc deficiency, which was strongly correlated with KAT7 HAT activity (Fig. S4d, e). Moreover, treatment with TSA also suppressed the decrease in H3K14ac but did not restore KAT7 HAT activity. Under these experimental conditions, we examined whether the zinc-dependent regulation of H3K14ac regulates ZIP10 expression. Zinc supplementation or TSA treatment counteracted the upregulation of ZIP10 expression during zinc deficiency (Fig. 4e). Using a membrane protein biotinylation assay, we confirmed that zinc deficiency led to the expression of ZIP10 at the plasma membrane, an effect that was suppressed by treatment with zinc or TSA (Fig. 4f). In addition, ChIP-seq and ChIP‒qPCR analyses revealed that the H3K14ac signal in the enhancer region of ZIP10, which was predicted by EnhancerAtlas 2.0⁴³, significantly decreased during zinc deficiency, whereas the signal in the promoter region did not decrease (Figs. 4g, S4f). This decrease in the enhancer regions was completely suppressed by treatment with zinc or TSA. Moreover, this decrease in the enhancer region of H3K14ac and increase in ZIP10 mRNA were dependent on KAT7, as confirmed after KAT7 knockdown (Fig. S4g-i). These results suggest that ZIP10 expression is regulated by KAT7-catalysed H3K14ac at enhancer regions.

To investigate whether the upregulation of ZIP10 expression contributes to maintaining zinc homeostasis, we assessed cellular zinc levels indirectly by measuring MT2A mRNA, the expression of which is well known to change depending on the concentration of cellular zinc^44,45. Consistent with previous reports, zinc supplementation drastically upregulated MT2A expression in cells cultured in low-zinc medium, an effect that was suppressed by the knockdown of ZIP10 (Fig. S4j). Notably, the knockdown of ZIP10 but not other ZIPs significantly attenuated the upregulation of MT2A expression (Fig. S4k), suggesting the critical role of ZIP10 in the import of zinc during zinc deficiency. To directly assess zinc uptake through ZIP10, we employed ZnDA-3H, a cell-permeable fluorescent Zn²⁺ probe that can be localized to specific subcellular compartments, via HaloTag technology⁴⁶. ZnDA-3H was localized in the cytosol when Halo-NES was expressed, and zinc influx was measured under these experimental conditions. The import of zinc (Δ[Zn²⁺] = 80.8 ± 3.0 pM) was significantly greater in the cells cultured in the low-zinc medium than in the cells cultured in the normal medium (42.4 ± 4.0 pM) (Fig. 4h). In contrast, TSA treatment suppressed zinc uptake (44.5 ± 8.4 pM), suggesting a critical role for H3K14ac in zinc influx. Notably, zinc uptake was significantly lower after ZIP10 knockdown (siRNA#1, 18.7 ± 5.3 pM; siRNA#2, 16.2 ± 4.3 pM) than after control knockdown (91.9 ± 6.2 pM) (Fig. 4i). Furthermore, zinc replenishment after deficiency led to recovery of H3K14ac levels in the control knockdown condition, whereas no such recovery was observed under ZIP10 knockdown (Fig. 4j). Collectively, these data suggest that the zinc influx induced by the addition of zinc to zinc-deficient cells is driven primarily by the H3K14 deacetylation-dependent increase in ZIP10 expression.

Next, we examined the biological role of upregulated ZIP10 expression during zinc deficiency. A previous study showed that zinc deficiency induces cell cycle arrest and that subsequent zinc repletion facilitates cell cycle re-entry⁴⁷. By employing an EdU assay, we confirmed that zinc deficiency indeed arrests the cell cycle and that subsequent zinc supplementation induces cell cycle re-entry (Fig. 4k, siCtrl). Notably, ZIP10 knockdown inhibited the process of cell cycle re-entry (Fig. 4k, siZIP10#1 and siZIP10#2). These findings reinforce the importance of upregulated ZIP10 expression during zinc deficiency in cellular biological processes (Fig. 4l).

Lipid accumulation in the liver during zinc deficiency

Next, we investigated the biological significance of the zinc deficiency-induced reduction in H3K14ac in vivo. Given the central role of the liver in zinc metabolism, we examined the expression of ZIP10 in the livers of mice fed a zinc-deficient diet. Consistent with the findings in HeLa cells, ZIP10 expression was significantly upregulated (Fig. S5a), suggesting that zinc uptake through the increased expression of ZIP10 may also contribute to maintaining zinc homeostasis in vivo. Moreover, if zinc deficiency persists due to a prolonged unbalanced diet, the limited availability of zinc for cellular uptake may disrupt cellular zinc homeostasis, potentially leading to abnormalities in biological functions. Thus, we investigated the possibility that prolonged zinc deficiency contributes to disease development by inducing sustained changes in gene expression associated with reduced H3K14ac levels in vivo.

We utilized the gene expression database Expression Atlas⁴⁸ to investigate diseases potentially linked to a reduction in zinc levels. Focusing on the metallothionein 2 (Mt2) gene, whose gene expression changes depending on the cellular zinc level, we examined the gene expression profile datasets of the mice with reduced Mt2 expression. There were 174 datasets in which Mt2 expression was significantly decreased. Among them, we focused on the “control vs. high-fat diet” data for the liver, which reported the most significant decrease in Mt2 expression (Fig. 5a). On the basis of these publicly available data, a high-fat diet in mice may lead to a significant reduction in hepatic zinc levels. This finding was confirmed by inductively coupled plasma‒mass spectrometry (ICP‒MS) analysis, which revealed a marked reduction in hepatic zinc concentrations in mice fed a high-fat diet (Fig. 5b). To explore the molecular mechanisms underlying this reduction, we examined the expression of zinc transporters. Specifically, we analyzed ZIP10, ZIP14—which primarily mediates zinc uptake in the liver—and ZNT1, which is responsible for zinc efflux. However, none of these transporters showed significant changes in expression (Fig. S5b). Thus, while the molecular mechanism regarding the cause of zinc reduction remained unresolved, we shifted our focus to the pathological consequences of hepatic zinc reduction. Since a high-fat diet induces hepatic lipid accumulation and promotes the development of fatty liver, we investigated the possibility that the reduction in liver zinc levels is involved in hepatic steatosis.

Fig. 5. Lipid accumulation in the liver during zinc deficiency.

a Log₂ fold change values of Mt2 expression under various conditions in mouse tissues. Gene expression data were obtained from the Expression Atlas database. b ICP‒MS analysis of liver samples from 8-week-old mice (male, n = 5; female, n = 3) fed the indicated diets for 2 weeks. c Lipid staining analysis of liver samples from 9-week-old mice (male, n = 3; female, n = 3) fed the indicated diet for the indicated periods. A total of 652 (HFD) and 623 (HFD + ZD) cells in males and 425 (HFD) and 459 (HFD + ZD) cells in females were evaluated. Scale bar, 30 µm, and 10 µm (inset). d Lipid staining analysis of liver samples from 8-week-old mice (male, n = 8 images from five mice; female, ND, n = 5; HFD, n = 7; HFD+zinc, n = 9 images from three mice) fed the indicated diets for two weeks. ZnCl₂ (1 mg/mL) was supplemented with drinking water. A total of 750 (ND), 543 (HFD), and 589 (HFD + ZD) cells were evaluated in males, and 470 (ND), 760 (HFD), and 975 (HFD + ZD) cells were evaluated in females. Scale bar, 30 µm, and 10 µm (inset). e ICP‒MS analysis of liver samples from 8-week-old mice (male, ND, n = 8; ZD, n = 8; female, ND, n = 6; ZD, n = 7) fed the indicated diets for 2 weeks. f Lipid staining analysis of liver samples from 8-week-old mice (male, n = 4; female, n = 4 images from three mice) fed the indicated diets for two weeks. A total of 216 (ND) and 234 (ZD) cells were evaluated in males, and 441 (ND) and 381 (ZD) cells were evaluated in females. Scale bar, 30 µm, and 10 µm (inset). g Triglyceride quantification assay of liver samples from 8-week-old mice (male, n = 4; female, n = 3) fed the indicated diets for two weeks. The graph presents the mean ± SEM. Statistical analysis was performed using two-sided Student’s t-test (b, c, e, f), one-way ANOVA followed by Dunnett’s test (d, g). n.s. not significant. ND normal diet, HFD high-fat diet, ZD low-zinc diet LD lipid droplet. Source data are provided with this paper.

First, to examine the effect of zinc deficiency on hepatic steatosis, mice fed a normal or zinc-deficient diet were fed a high-fat diet. Comparative analysis revealed a pronounced exacerbation of hepatic lipid accumulation in mice fed the zinc-deficient diet, as demonstrated by oil red O staining and BODIPY staining (Figs. 5c, S5c). Conversely, dietary zinc supplementation in mice fed a high-fat diet attenuated hepatic lipid accumulation (Figs. 5d, S5d). These findings implicate zinc deficiency as a contributing factor to the development of fatty liver.

To elucidate the mechanisms underlying the contribution of zinc deficiency to hepatic steatosis, we investigated the hepatic characteristics of a zinc-deficient mouse model. Mice fed a zinc-deficient diet presented a reduction in hepatic zinc levels (Fig. 5e). This reduction was accompanied by notable hepatic lipid accumulation (Figs. 5f, S5e) and elevated triglyceride levels to a similar extent as those observed with a high-fat diet (Fig. 5g). Collectively, these results suggest that diminished hepatic zinc levels facilitate lipid accumulation in the liver, thereby contributing to the development of hepatic steatosis.

Loss of H3K14ac promotes lipid accumulation in liver tissue via gene expression upregulation

Next, we investigated the mechanism underlying hepatic steatosis induced by zinc deficiency. Mice fed a zinc-deficient diet presented reduced levels of H3K14ac (Fig. 6a, b). Similarly, mice fed a high-fat diet presented reduced H3K14ac levels, whereas zinc supplementation counteracted this reduction (Figs. 6c, S6a). Notably, KAT7 immunoprecipitated from mouse livers fed a zinc-deficient diet presented significantly decreased HAT activity (Fig. 6d). Moreover, the intraperitoneal treatment of mice with the KAT7 inhibitor WM-3835 resulted in lipid accumulation in the liver (Fig. 6e). These results suggest that a reduction in KAT7 activity and the subsequent decrease in H3K14ac levels in the liver may induce hepatic steatosis. To assess whether zinc deficiency in the liver promotes hepatic steatosis in a cell-autonomous manner, we conducted experiments using mouse primary hepatocytes. TPEN treatment of these cells resulted in a decrease in H3K14ac levels and an increase in lipid droplets, effects that were suppressed by TSA treatment (Fig. 6f, g). Furthermore, treatment with WM-3835 or the knockdown of KAT7 led to increased lipid droplet formation (Fig. S6b-e). Taken together, these results suggest that zinc deficiency contributes at least in part to hepatic steatosis through decreased KAT7 activity and a reduction in H3K14ac.

Fig. 6. Loss of H3K14ac promotes lipid accumulation in liver tissue via gene upregulation.

a Immunoblot of liver samples from 8-week-old male mice, with representative data and H3K14ac quantification from six mice. b Immunofluorescence of liver samples from 8-week-old male mice. A total of 210 (ND) and 312 (ZD) cells were evaluated. Scale bar, 30 µm. c Immunoblot of liver samples from 8-week-old male mice (ND and ZD, n = 8; HFD+zinc, n = 7). d KAT7 immunoprecipitated from liver samples of 8-week-old male mice (n = 3) was subjected to an in vitro HAT assay. e Lipid staining of liver samples from 8-week-old male mice (DMSO, n = 7; WM-3835, n = 9 images from three mice) treated with DMSO or 10 mg/kg WM-3835 for two weeks via daily intraperitoneal injections. Scale bar, 50 µm (oil red O staining), 20 µm (BODIPY staining), and 10 µm (inset). A total of 511 (DMSO) and 422 (WM-3835) cells were evaluated. Immunoblot (f) or lipid staining (g) of mouse primary hepatocytes treated with 3 µM TPEN or zinc-saturated TPEN for 10 h after pretreatment with 0.5 µM TSA for one hour. The immunoblot images are representative of three independent experiments. A total of 36 (DMSO), 57 (TPEN), 46 (Zn + TPEN), and 51 (TPEN + TSA) cells were evaluated. Scale bar, 30 µm, 10 µm (inset). h Volcano plot of RNA-seq. i Average plot and heatmap of H3K14ac ChIP-seq reads around enhancers. j Heatmaps showing gene expression changes during zinc deficiency and the H3K14ac signal in enhancer regions of the male mouse liver. k Read density of H3K14ac around the genes involved in lipid droplet synthesis in liver samples of male mice. l Schematic of the model reported in this study. m Meta-analysis of the zinc concentration in the livers of patients with steatosis-related diseases. The mice were fed for two weeks with the indicated diet (a–e, h–k). The graph presents the mean ± SEM. Statistical analysis was performed using two-sided Student’s t-test (a, b, d, e), Student’s t-test followed by Bonferroni post hoc correction (c, g). n.s., not significant. ND normal diet, ZD low-zinc diet, LD lipid droplet. Source data are provided with this paper.

Next, we investigated whether zinc deficiency in the liver alters the expression of genes associated with lipid accumulation. RNA-seq analysis revealed that zinc deficiency in the liver induced a decrease in the expression of 1,193 genes and an increase in the expression of 1,781 genes (Fig. 6h). We then conducted ChIP-seq analysis for H3K14ac, the results of which revealed a global reduction in H3K14ac at enhancer regions during zinc deficiency (Fig. 6i). In line with the findings shown in Fig. 4, the RNA-seq analysis demonstrated an upregulation of ZIP14, a zinc transporter reported to be critical for maintaining hepatic zinc homeostasis (Fig. S6f). Moreover, the ChIP-seq analysis revealed a reduction of H3K14ac signals at the enhancer regions of both ZIP10 and ZIP14 (Fig. S6g). Together, these results suggest that the decrease in H3K14ac also contributes to hepatic zinc homeostasis through the regulation of zinc transporters under zinc-deficient conditions. In addition to the zinc transporters, the expression of several genes associated with lipid droplet synthesis pathways, such as triglyceride synthesis, intracellular lipid transport, and lipid droplet budding, was upregulated during zinc deficiency, along with a decrease in H3K14ac at enhancer regions (Fig. 6j, k). Additionally, the treatment of primary hepatocytes with WM-3835 increased the expression of key genes involved in lipid droplet formation (Fig. S6h). Collectively, these findings suggest that zinc deficiency drives lipid accumulation by upregulating the expression of genes involved in lipid droplet synthesis, a process mediated by reduced KAT7 activity and diminished H3K14ac levels (Fig. 6l).

Finally, we conducted a clinical literature review to examine the association between zinc and fatty liver diseases in humans. Several clinical studies have measured hepatic zinc concentrations in both healthy individuals and patients with diseases accompanied by hepatic steatosis. A meta-analysis of these studies revealed significantly lower hepatic zinc levels in the patient group than in the healthy control group (Fig. 6m). This finding suggests that zinc deficiency in the liver may serve as a potential risk factor for fatty liver diseases, including NAFLD.

Discussion

In this study, we elucidated the molecular mechanism by which zinc deficiency stress is converted to a decrease in H3K14ac. Moreover, we presented examples of genes that are controlled by this decrease in epigenetic signalling, including those related to zinc homeostasis and the accumulation of lipid droplets. We provided evidence that H3K14ac regulates gene expression via its enhancer regions; however, the mechanism remains unclear. H3K14ac acts as a binding site for numerous bromodomain proteins, referred to as “readers”⁴⁹. Our results suggest that transcriptional repressive readers may reside in the enhancer regions of ZIP10 and several genes implicated in lipid droplet synthesis in the basal state. Several groups, including our group, have reported that bromodomain-containing proteins such as ZMYND8 and BAZ2A recognize H3K14ac on enhancer regions to repress gene transcription^36–38, whereas at least 10 readers have been shown to recognize H3K14ac⁵⁰. The identification of the specific reader that recognizes H3K14ac on the enhancer is a future aim.

The results of our present study suggest that zinc deficiency may lead to a loss of zinc coordination in KAT7; however, how zinc is released from KAT7 remains unclear. Although TPEN is a chelator with a high affinity for zinc⁵¹, in vitro TPEN treatment did not affect the coordination of zinc or the HAT activity of KAT7 (Fig. S3i, j). This result suggests that zinc is tightly coordinated in KAT7 and that zinc is unlikely to be spontaneously released from KAT7 during zinc deficiency. One possible mechanism is that unidentified cellular molecule(s) may recognize the decreased concentration of zinc and facilitate zinc dissociation from KAT7 in the cell. Another possibility is that the activity of certain chaperones that help coordinate zinc in KAT7 may be reduced under zinc-deficient conditions. Recent reports have demonstrated that proper zinc transport mediated by a zinc chaperone regulates the activity of zinc-dependent enzymes^52,53. Moreover, in vitro zinc repletion of WT KAT7 immunoprecipitated from cells treated with TPEN did not fully restore KAT7 HAT activity (Fig. 3h). This raises the possibility that something irreversible other than the loss of zinc coordination, such as changes in the PTMs of KAT7, occurs in cells under zinc-deficient conditions. Additional studies are necessary to elucidate the precise regulatory mechanism of KAT7 HAT activity during zinc deficiency.

In this study, employing in vitro HDAC assays, we found that HDAC1 and HDAC6 are capable of deacetylating H3K14ac (Fig. 2e). However, for HDAC2, HDAC3, and HDAC11, we were unable to obtain sufficient amounts of protein, as their immunoprecipitation yielded markedly lower levels compared with other HDACs. Therefore, we speculate that these HDACs might also contribute to H3K14 deacetylation. Supporting this idea, the double knockdown of HDAC1 and HDAC6 did not suppress the TPEN-induced reduction of H3K14ac (Fig. S2d), suggesting that additional mechanisms beyond HDAC1 and HDAC6 are involved. Interestingly, although HDACs are known to be zinc-dependent enzymes, there are also reports indicating that zinc deficiency can enhance HDAC activity^54,55. Future studies investigating the activity of individual HDACs under zinc-deficient conditions will be crucial to elucidate the mechanisms underlying the reduction of H3K14ac during zinc deficiency.

Compared to the marked upregulation of ZIP10 expression under zinc deficiency (Fig. 4e), the increase observed upon KAT7 suppression was relatively modest (Fig. S4i). This suggests that mechanisms other than the reduction of H3K14ac contribute to the induction of ZIP10 under zinc-deficient conditions. It is possible that transcription factors contribute to this regulation. Indeed, transcription factors such as STAT3/5 have been reported to promote ZIP10 expression⁵⁶. However, zinc deficiency has been associated with conflicting reports regarding STAT3 activity regulation^57,58, and it remains unclear whether this pathway contributes to ZIP10 upregulation in the present context. Further investigation into the involvement of STAT3/5 and other transcription factors will therefore be required.

We showed that knockdown of ZIP10 suppressed the recovery of H3K14ac levels upon zinc replenishment (Fig. 4j). While zinc uptake is reduced under these conditions (Fig. 4i), it remains to be determined whether these effects result from insufficient ZIP10-mediated restoration of intracellular zinc. To address this question, it will be necessary to demonstrate that whether the zinc-transport activity of ZIP10 is required for these recoveries; however, given the limited reports on ZIP10, more detailed characterization of the protein is needed. ZIP-family transporters are thought to have conserved histidine residues essential for zinc transport⁵⁹. Although it is still unknown whether the corresponding residues in ZIP10 are similarly required for its transport function, analyses using ZIP10 mutants with impaired zinc-transport activity would enable direct evaluation of the necessity of this activity for the observed recoveries of the H3K14ac levels.

The regulation of ZIPs on the plasma membrane at the protein level contributes to the maintenance of zinc homeostasis. For example, ZIP4 is degraded via endocytosis under normal conditions, whereas endocytosis is suppressed during zinc deficiency, leading to an increase in the amount of ZIP4 on the plasma membrane and an increase in zinc influx^60,61; another example includes the phosphorylation-dependent regulation of ZIP7 activity⁶². However, little is known about the regulation of ZIP10 at the protein level, except that ZIP10 forms a heteromeric complex with ZIP6⁶³. The regulatory mechanisms of ZIP10 at the protein level need to be investigated.

We found that reduced H3K14ac in HeLa cells drives the upregulation of the zinc transporter ZIP10, thereby contributing to the maintenance of intracellular zinc homeostasis. Furthermore, in the mouse liver, zinc deficiency stress induced the expression of both ZIP10 and ZIP14 (Figs. S5a, S6f, g). Given that zinc serves as an essential cofactor for liver enzymes such as alcohol dehydrogenase and those involved in ammonia metabolism^64,65, its adequate regulation is indispensable for hepatic function. Furthermore, based on our findings that zinc deficiency induces hepatic lipid accumulation, ZIP10-mediated zinc uptake may influence the regulation of liver steatosis. Thus, the upregulation of ZIP10 and ZIP14 expression in response to decreased H3K14 acetylation under zinc-deficient conditions is likely to function cooperatively to maintain hepatic zinc homeostasis and thereby sustaining liver function and modulating hepatic lipid accumulation. However, whether ZIP10 contributes to maintaining liver function or the hepatic lipid accumulation during zinc deficiency in vivo remains unresolved, and further investigation is needed. As the whole-body Zip10 knockout model is lethal⁶⁶, generating a liver-specific Zip10 knockout mouse and assessing its effects on zinc uptake and hepatic lipid accumulation represent an important direction for future work.

We demonstrated that hepatic zinc levels are reduced in mice fed a HFD. The molecular mechanisms behind the reduction remain to be fully elucidated. One hypothesis that can be postulated is that HFD has the potential to compromise zinc absorption in the duodenum. Zinc is primarily absorbed from the intestinal lumen into enterocytes via the zinc importer ZIP4⁶⁷. It is subsequently exported to the circulation through ZNT1 for distribution to peripheral tissues⁶⁸. It is hypothesised that the modulation of the transcription or membrane trafficking of ZIP4 or ZNT1 by HFD could result in a reduction of zinc absorption and release into the bloodstream, which would lead to a decrease in hepatic zinc level. An alternative hypothesis is that the enhanced secretion of zinc-binding proteins, including specific apolipoproteins, induced by HFD contributes to the reduction in hepatic zinc content. The secretion of protein by hepatocytes is known to be subject to regulation by hormonal and peptide signals originating from the pancreas and adipose tissue⁶⁹. Such signals might be augmented under conditions of a HFD. Taken together, these observations suggest that a combination of altered zinc absorption, redistribution, and excretion pathways may underlie the reduction in hepatic zinc content induced by a HFD. Further investigation of these possibilities will advance a more comprehensive understanding of the mechanisms linking dietary fat intake to systemic zinc homeostasis.

The accumulation of intracellular lipid droplets observed in fatty liver progresses through multiple steps, including triacylglycerol synthesis at the endoplasmic reticulum membrane, lipid budding from the endoplasmic reticulum to the cytoplasm, and the maturation of lipid droplets in the cytoplasm. In this study, we showed that zinc deficiency upregulated the expression of genes involved in these steps, for example, Lpin1, Fabp1, and Fitm1. LPIN1 is an enzyme that catalyses the conversion of phosphatidic acid to diacylglycerol during triglyceride biosynthesis. Several reports have suggested that the inhibition of LPIN1 ameliorates hepatic steatosis via the suppression of triglyceride biosynthesis^70,71. FABP1 plays a critical role in the uptake and intracellular transport of fatty acids, and silencing Fabp1 has been shown to reduce lipid accumulation in NAFLD model mice⁷². FITM1 directly binds to triglycerides and induces lipid droplet formation⁷³. The overexpression of FITM1 has been shown to increase lipid droplet formation in cultured cells. On the basis of these previous reports, zinc deficiency may promote lipid synthesis at multiple stages by increasing the expression of these genes. However, the specific impact on each stage needs to be investigated further.

The findings of this study suggest that zinc deficiency contributes to hepatic lipid accumulation through a reduction in H3K14ac. On the other hand, lipid accumulation induced by the intraperitoneal injection of the KAT7 inhibitor was relatively mild compared with that caused by zinc deficiency. Previous research has shown that zinc deficiency can trigger oxidative stress and endoplasmic reticulum (ER) stress^16,74. Since oxidative and ER stress are known to promote lipid droplet synthesis^75,76, it is plausible that zinc deficiency further exacerbates hepatic lipid accumulation through these mechanisms. Additionally, studies have demonstrated that the knockout of GPR39, an extracellular zinc sensor, induces hepatic lipid accumulation in mice⁷⁷. This phenotype is consistent with the effects observed in zinc deficiency, which may inactivate GPR39. Together, these findings indicate that zinc deficiency may contribute to hepatic lipid accumulation through multiple interconnected pathways.

Although zinc deficiency has been implicated in numerous human diseases, the causal molecular mechanisms remain largely unexplored. This study provides evidence that zinc deficiency is associated with a loss of KAT7 activity, advancing our understanding of the mechanisms underlying various diseases caused by zinc deficiency. For example, zinc deficiency during pregnancy has devastating effects on newborns, such as growth impairment and low birth weight⁷⁸. From the viewpoint of KAT7, Kat7 knockout mice show decreased transcription of genes essential for embryonic development owing to a global reduction in H3K14ac, which leads to embryonic lethality⁷⁹. These findings suggest a link between zinc deficiency and the loss of KAT7 activity during embryonic development. Moreover, KAT7 plays key roles in immunity by regulating the functions and differentiation of T cells^80,81, which was observed in model rodents fed a zinc-deficient diet⁸². Thus, the biological phenomena observed during zinc deficiency are better understood from the perspective of the functional regulation of KAT7-catalysed H3K14ac.

Methods

A list of the primers, antibodies, and siRNAs used in this study is provided in Table S1-S7.

Plasmids

The cDNAs encoding human KAT7, BRPF1, BRPF2, BRPF3, JADE1, JADE2, JADE3, ING4, ING5, MEAF6, and HDACs were amplified by PCR using KOD One PCR Master Mix -Blue- (TOYOBO, KMM-201) and inserted into pcDNA3/GW with an HA or a FLAG tag (Invitrogen) or into pMAL-c6T (New England Biolabs). The mutated constructs were prepared by PCR-mediated site-directed mutagenesis. A single guide RNA (sgRNA) targeting KAT7 (5’-AGCCGCCGGCAATGCCGCGA-3’) was designed using CHOPCHOP. The sgRNA was inserted into pX459 (Addgene, 62988). The primer lists used for cloning and mutagenesis are provided in Tables S1 and S2.

Cell culture

HEK293A cells, U2OS cells, HCT116 cells, and Neuro2A cells were maintained in high-glucose Dulbecco’s modified Eagle’s medium (DMEM) (Sigma‒Aldrich, D5796; Wako, 044-29765). HeLa cells were maintained in low-glucose DMEM (Wako, 041-29775). Each type of culture medium was supplemented with 10% fetal bovine serum (FBS) (Gibco, 10270-106) and 100 units/ml penicillin G (Meiji Seika, 01028-85). All cell lines were grown in 5% CO₂ at 37 °C.

Preparation of zinc-deficient medium

FBS was incubated with Chelex-100 resin (Bio-Rad, 1422832) at a final concentration of 0.05 g/mL at 4 °C with rotation. After 12 h, the mixture was centrifuged at 500 × g for 5 min. The supernatant was passed through a Millex-GV 0.22 µm PVDF filter (Millipore, Cat#SLGVR33RS). Cell culture medium supplemented with Chelex-100 resin-treated FBS at a final concentration of 10% was used as zinc-deficient medium.

Preparation of Zn-saturated TPEN

Preparation of Zn-saturated TPEN was performed by mixing 10 µM TPEN with an equimolar concentration of ZnCl₂ (10 µM) and incubating the mixture overnight. The resulting solution was used as Zn-saturated TPEN for cell treatment.

Transfection

Plasmid transfection was performed using Polyethylenimine”MAX” (Polysciences, 24765) or Lipofectamine 2000 (Thermo Fisher Scientific, 11668019) according to the manufacturer’s instructions. siRNA-mediated knockdown was performed using Lipofectamine RNAiMAX (Thermo Fisher Scientific, 13778500) at a final siRNA concentration of 10 or 20 nM by reverse transfection, according to the manufacturer’s instructions. The list of siRNAs used in this study is provided in Tables S3 and S4.

Generation of knockout cell lines

To generate KAT7 knockout HEK293A cells or HeLa cells, cells were transfected with pX459 encoding a sgRNA targeting KAT7. Forty-eight hours after transfection using Lipofectamine 2000, the cells were selected by 1.0 µg/mL puromycin (Gibco, A11138-03) treatment for another 48 h. The selection medium was replaced with fresh standard medium, and the cells were grown for 3 days. Limiting dilution was performed to obtain single-cell clones. The knockout status of the clones was confirmed by immunoblotting.

Immunoblotting

Cells were lysed in a cell lysis buffer (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM EDTA, 1% Triton X-100, 5 µg/mL leupeptin, 1 mM phenylmethylsulfonyl fluoride (PMSF) and 2 nM TSA) for 10 min at 4 °C. The lysates were sonicated with 2 sets of 10 s pulses using an ultrasonic homogenizer (SMT, UH-50). The cell debris was cleared by centrifugation at 17,700 × g for 15 min. The supernatant was mixed with a sample buffer (80 mM Tris-HCl pH 8.8, 80 µg/mL bromophenol blue, 28.8% glycerol, and 4% SDS) supplemented with 10 mM DTT and incubated for 3 min at 98 °C. The samples were then resolved by SDS–PAGE and transferred onto Immobilon-P PVDF membranes (Millipore, IPVH00010). The SDS–PAGE gels used were either self-prepared polyacrylamide gels or NuPAGE® Bis-Tris Precast Gels (Thermo Fisher Scientific, WG1403BOX). Blocking was performed with 5% skim milk (Megmilk Snow Brand) in TBS-T (50 mM Tris-HCl pH 8.0, 150 mM NaCl and 0.05% Tween 20) for 30 min at room temperature. After incubation of the membranes with the indicated primary antibodies for 12–36 h at 4 °C and corresponding HRP-linked secondary antibodies for 1–2 h at room temperature, the chemiluminescent signals enhanced by ECL Select (Cytiva, RPN2235) were detected using a Fusion Solo 7S instrument (M&S Instruments). Quantification of the bands was performed using Fiji software⁸³. The list of antibodies used for immunoblotting is provided in Table S5.

Coimmunoprecipitation analysis

Cell lysates were incubated with anti-DYKDDDDK-tagged antibody beads (Wako, 016-22784) at 4 °C for 10 min. The beads were washed with washing buffer 1 (20 mM Tris-HCl pH 7.5, 500 mM NaCl, 5 mM EGTA and 1% Triton X-100) and washing buffer 2 (20 mM Tris-HCl pH 7.5, 150 mM NaCl and 5 mM EGTA). The washed beads were mixed with sample buffer (80 mM Tris-HCl pH 8.8, 80 µg/mL bromophenol blue, 28.8% glycerol, and 4% SDS) supplemented with 20 mM DTT for 5 min at room temperature and then incubated for 3 min at 98 °C. The samples were separated by SDS‒PAGE and immunoblotted with antibodies.

Immunofluorescence analysis

For confocal microscopy, cells were grown on glass coverslips (Matsunami Glass, C015001). For imaging with a high-content imaging system, cells were grown on a 96-well plate (Corning, 353072). The cells were fixed with 4% paraformaldehyde in PBS for 10 min and permeabilized with 0.2% Triton X-100 in PBS for 5 min. After blocking with 1% BSA (Iwai Chemicals, A001) in TBS-T for 1 h, the cells were incubated with the indicated primary antibodies overnight at 4 °C. After washing three times with PBS, the cells were incubated with fluorophore-conjugated secondary antibodies at room temperature for 2 h and then with Hoechst 33342 (Dojindo, 346-07951) for 20 min. After washing three times with PBS, the cells were mounted with Fluoromount (Diagnostic BioSystems, K024) for confocal microscopy. Images were acquired using an LSM880 with Airyscan (Zeiss) with a 63×/1.40 oil immersion objective or a CellInsight NXT HCS system (Thermo Scientific). The contrast and brightness of the images were adjusted using Fiji software. Quantification of the images was performed using Fiji software or CellProfiler⁸⁴.

Histone purification

HEK293A cells or HEK293A KAT7 KO cells were incubated in hypotonic lysis buffer for 30 min at 4 °C with rotation. After centrifugation at 10,000 × g for 10 min at 4 °C, the cell pellets were resuspended in 0.4 N H₂SO₄ and incubated for 30 min at 4 °C with rotation. After centrifugation at 15,100 × g for 10 min at 4 °C, the supernatant was transferred to a fresh 1.5 mL tube. Then, trichloroacetic acid (TCA) was added to the supernatant dropwise and incubated for 30 min on ice. After centrifugation at 15,100 × g for 10 min at 4 °C, the pelleted histones were washed twice with ice-cold acetone. The pellet histones were air-dried at room temperature and dissolved in TBS at a final concentration of 3 µg/µL.

Cell lysate–based HDAC activity assay

HEK293A cells treated with DMSO or 10 µM TPEN for 1 h or 2 h were lysed in a cell lysis buffer for in vitro HDAC assay (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM EDTA, 1% Triton X-100, 5 µg/mL leupeptin, and 1 mM PMSF) for 10 min at 4 °C. Purified histones were mixed with the cell lysates and incubated at 37 °C for 0.5, 1 or 2 h with DMSO or 10 nM TSA. Then, the samples were mixed with an equal volume of 2x sample buffer supplemented with 10 mM DTT and incubated for 3 min at 98 °C, followed by SDS‒PAGE and immunoblotting.

In vitro HDAC assay

HEK293A cells transfected with HDAC-FLAG were lysed in cell lysis buffer (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM EDTA, 1% Triton X-100, 5 µg/mL leupeptin, and 1 mM PMSF). For HDAC1, HDAC4, HDAC5, HDAC6, HDAC7, HDAC8, HDAC9, and HDAC10, cell lysates derived from 1.0 × 10^6 cells were used, whereas for HDAC2, HDAC3, and HDAC11, lysates from 1.0 × 10^8 cells were prepared. The cell lysates were incubated with anti-FLAG M2-agarose gel (Sigma‒Aldrich, A2220) at 4 °C for 5 min. The beads were washed twice with washing buffer 3 (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% Triton X-100) and once with TBS. Purified histones at a final concentration of 0.5 µM were mixed with the beads incubated for 5–15 min at 37 °C with frequent mixing (1200 rpm) using a Thermomixer C (Eppendorf). The samples were mixed with an equal volume of 2x sample buffer supplemented with 10 mM DTT and incubated for 3 min at 98 °C, followed by SDS‒PAGE and immunoblotting.

In vitro HAT assay

To measure the activity of KAT7 immunoprecipitated from cells, HEK293A cells transfected with FLAG-KAT7 WT or mutants were lysed in cell lysis buffer for an in vitro HAT assay (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 10 mM EDTA, 1% Triton X-100, 5 µg/mL leupeptin, and 1 mM PMSF). The cell lysates were incubated with anti-FLAG M2-agarose gel (Sigma‒Aldrich, A2220) at 4 °C for 5 min. The beads were washed twice with washing buffer 3 (20 mM Tris-HCl pH 7.5, 150 mM NaCl, 1% Triton X-100) and once with TBS. Purified histones at a final concentration of 0.5 µM were mixed with the beads and with 50 µM acetyl-CoA in TBS and incubated for 30 min at 30 °C with frequent mixing (1200 rpm) using a Thermomixer C (Eppendorf). The samples were mixed with an equal volume of 2x sample buffer supplemented with 10 mM DTT and incubated for 3 min at 98 °C, followed by SDS‒PAGE and immunoblotting. To measure the activity of the recombinant KAT7 HAT domain, 5 µM recombinant protein was used. To measure the activity of the KAT7 from mice liver, the organ lysates were incubated with KAT7 antibody at 4 °C for 16 h. Then, the lysates were incubated with Protein G SepharoseTM 4 Fast Flow (GE healthcare, 17061802) at 4 °C for 2 h.

Recombinant protein purification

The KAT7 HAT domain (a.a. 336-611) WT or C371A mutants encoded in pMAL-c6T were expressed in E. coli BL21 cells. Expression was induced by incubation in LB medium supplemented with 0.3 mM IPTG at 25 °C for 1.5 h. The cells were pelleted and resuspended in PBS supplemented with 1% Triton X-100 and 1 mM PMSF. After sonication with 4 sets of 30 s pulses using an ultrasonic homogenizer, the cell debris was cleared by centrifugation at 26,800 × g for 30 min. The supernatants were incubated with Ni Sepharose 6 Fast Flow (Cytiva, 17531806) at 4 °C for 8 h. The beads were then washed twice with PBS and twice with TBS and eluted with a 250 mM imidazole solution in TBS using Poly-Prep Chromatography Columns (Bio-Rad, 7311550). The eluates were loaded onto a Superose 6 Increase 10/300 GL column on an AKTA purifier (GE Healthcare). The purified His-MBP-KAT7 was incubated with TEV protease (NEB, P8118) at 30 °C for 4 h. The cleaved His-MBP fragment was removed by filtration through Ni Sepharose 6 Fast Flow resin. The purified KAT7 was dialyzed in TBS overnight at 4 °C. The purity and concentration of the final products were estimated by Coomassie blue staining of SDS–PAGE gels.

qPCR analysis

Total RNA was isolated from cells using Isogen (Nippongene, 319-90211) and reverse transcribed with ReverTra Ace qPCR RT Master Mix with gDNA Remover (TOYOBO, FSQ-301) according to the manufacturer’s instructions. Samples of the qPCR mixture were prepared using the KAPA SYBR Fast qPCR Kit (Kapa Biosystems, KK4602). qPCR was then performed on a LightCycler 96 (Roche) or QuantStudio 1 (Thermo Fisher Scientific) instrument. Data for each mRNA were normalized to that of RPS18. The list of primers used for qPCR is provided in Table S6.

ChIP‒qPCR

Chromatin samples for ChIP‒qPCR were prepared using a SimpleChIP Enzymatic Chromatin IP Kit (Cell Signaling Technology, 9003) according to the manufacturer’s instructions. The chromatin samples were incubated with anti-H3K14ac antibody or anti-histone H3 antibody (Cell Signaling Technology, 4499) at 4 °C for 8 h with rotation. Then, the samples were incubated with Dynabeads Protein G (VERITAS, DB10004) at 4 °C for 2 h with rotation. After washing three times with low-salt buffer (1% SDS, 1% Triton-X, 2 mM EDTA, 20 mM Tris-HCl pH 8.0 and 150 mM NaCl) and once with high-salt buffer (1% SDS, 1% Triton-X, 2 mM EDTA, 20 mM Tris-HCl pH 8.0 and 500 mM NaCl), the chromatin was eluted with elution buffer (1% SDS, 10 mM EDTA and 50 mM Tris-HCl pH 8.0) for 30 min at 65 °C with frequent mixing (1,200 rpm) using a Thermomixer C. The purified DNA was analysed as described in the “qPCR analysis” section. ChIP-seq data used for designing primers were downloaded from the ENCODE portal⁸⁵ with the following identifier: ENCFF002QJL. Data visualization was performed with Integrative Genomics Viewer (Broad Institute). The list of primers used for qPCR is provided in Table S7.

ZnAF2 assay

Recombinant proteins or ZnCl₂ solution for normalization was incubated with 10 µM ZnAF-2 (Goryo Chemical, SK2001-01) in TBS for 30 min at room temperature. The fluorescence signal was detected with a Varioskan microplate reader (Ex/Em = 492/515 nm, Thermo Scientific).

ERSE sequence analysis

ERSE sequences, CCAAT-(N)₉-CCAC[A/G], on the human genome GRCh/hg38 were retrieved from the GGGenome ultrafast sequence search browser. Among the sequences obtained (plus strand, 3764 sequences; minus strand, 3874 sequences), 1897 sequences were annotated as sequences on gene promoters by employing HOMER. The number of ERSE sequences on genes that were upregulated during zinc deficiency was counted programmatically. Gene expression profile data analysed by GEO2R were downloaded from the GEO repository (GSE49657, GSE99204, GSE108923, and GSE135873). Genes with a fold change in expression greater than 2 were considered upregulated.

EdU assay

HeLa cells were grown on a 96-well plate and analysed with a Click-iT EdU imaging kit (Invitrogen, C10340). The cells were incubated with 10 µM EdU for 1 h, fixed with 4% paraformaldehyde in PBS for 15 min at room temperature, and washed twice with PBS containing 3% BSA. The cells were permeabilized with 0.5% Triton X-100 in PBS for 20 min at room temperature and washed twice with PBS containing 3% BSA. Then, Click-iT reaction cocktail was added and allowed to react for 30 min at room temperature. After washing twice with PBS containing 3% BSA, the cells were incubated with Hoechst 33342 (1:2000) in PBS for 30 min at room temperature. Images were acquired with a CellInsight NXT automated microscope and analysed with CellProfiler.

Membrane protein biotinylation assay

Cells were incubated with 0.4 mg/mL biotin (EZ-Link Sulfo-NHS-SS-Biotin, Thermo Fisher Scientific, 21331) in PBS(+) (PBS containing 50 mM MgCl₂ and 100 mM CaCl₂) and washed twice with 150 mM glycine in PBS(+). Cells were lysed as described in the “Immunoblotting” section. The cell lysates were pulled down with streptavidin agarose resin (Thermo Fisher Scientific, 20353). The resin samples were washed twice with washing buffer 1 and once with washing buffer 2. The samples were subjected to immunoblotting.

Zinc imaging by ZnDA-3H

Zinc measurement with ZnDA-3H was performed as described previously with minor optimization^46,86. HeLa cells were reverse-transfected with 10 µM siRNA. The cells were then transfected with Halo-NES and cultured for 24 hours. The cells were replated on 35 mmφ glass bottom dishes (Matsunami, D11130H) that were coated with 1% Cellmatrix Type I-C (Nitta Gelatin, KP-4100). After 24 hours, the cells were incubated with zinc-normal medium or zinc-deficient medium for an additional 48 hours. Cells were washed with Chelex-treated HEPES-buffered HBSS (30 mM HEPES-NaOH, pH 7.4, 5.36 mM KCl, 137 mM NaCl), and incubated with serum-free DMEM containing 250 nM ZnDA-3H and 5 nM HTL-TMR for 30 min. After labelling, the cells were incubated for an additional 30 minutes. After incubation, the cells were washed twice with Chelex-treated HEPES-buffered HBSS, and the medium was exchanged for Chelex-treated HBSS buffer. Multichannel time-lapse images were acquired with 4 averages per frame in 30-second intervals. ZnDA-3H or HTL-TMR was excited at 488 nm with an argon laser or at 561 nm with a DPSS laser and detected by a HyD detector (Leica). Images were acquired for 5 minutes, and then the cells were incubated with containing Chelex-treated HBSS 10 µM ZnCl₂ for an additional 10 min. To obtain additional images under zinc-deprived and zinc-saturated conditions, the cells were incubated with Chelex-treated HBSS containing 10 µM TPEN for 5 minutes, and then, the cells were incubated with Chelex-treated HBSS containing 400 µM ZnCl₂ and 2.0 µM zinc pyrithione for an additional 10 minutes. For presentation, the pseudocolour fluorescence ratio images of ZnDA-3H/HTL-TMR were generated using Fiji software. For quantification of the intensity, we used CellProfiler.

Meta-analysis

A systematic literature search was conducted in multiple electronic databases (PubMed, Web of Science, and Embase) using the predefined search terms “zinc” and “liver disease.” From the identified articles, the means of and variances in the liver zinc content in healthy subjects and patients with liver disease were extracted for analysis. All the statistical analyses were conducted using R (R Foundation for Statistical Computing, Vienna, Austria). For studies reporting only median and IQR data, the means and variances were estimated using the “estmeansd” package⁸⁷. A multilevel meta-analysis was performed to address possible correlations among several outcome statistics using the “meta” package⁸⁸.

Animal studies

All the experiments were performed following the experimental protocol approved by the animal ethics committee of the University of Tokyo. We complied with all relevant ethical regulations for animal use. Male and female C57BL/6 J mice were used. C57BL/6 J mice were purchased from Japan SLC Inc. The mice were maintained in a specific pathogen-free facility and housed under 25 °C, 40–60% humidity, 12/12 light/dark cycle. Age (8–10 weeks) and sex (both male and female) -matched mice were used for the experiments. Sample sizes determination was based on previous reports^89,90.

The mice were fed a ND (Zn: 6 mg/100 g, 13% calories from fat; CE-2, Clea Japan), ZD (Zn: 0 mg/100 g, 13% calories from fat; Clea Japan), HFD (Zn: 6 mg/100 g, 60% calories from fat; CE-2, Clea Japan), or HFD + ZD (Zn: 0 mg/100 g, 60% calories from fat; Clea Japan). Mouse livers were perfused and immersed O/N in 4% paraformaldehyde/PBS for fixation and then cleared with 30% sucrose solution. Fixed livers were embedded in CryoMount I (Muto PureChemicals), and 8 μm-thick sections were cut using a cryostat (Leica), followed by oil red O staining and BODIPY staining.

Primary hepatocyte isolation

Primary hepatocytes were isolated from male and female mice as previously described⁷⁵. Briefly, primary hepatocytes were isolated by two-step collagenase perfusion. After isolation, the cells were resuspended in low-glucose DMEM supplemented with 5% FBS and 100 units/mL penicillin G (Meiji Seika) and then seeded in a six-well plate precoated with rat collagen type I (Sigma, C3867). The culture medium was changed to maintenance medium after three hours. The maintenance medium contained William’s E medium (Gibco, 12551-032), 1% L-glutamine (Wako, 076-00521), and 1% penicillin G. After an overnight incubation, the hepatocytes were used for subsequent experiments.

Oil red O staining

Oil red O stock solution (oil red O (Wako, 154-02072) 0.03 g/10 mL isopropanol) was diluted 3:2 with Milli-Q water to make an Oil red O working solution. Frozen blocks were sectioned using a cryostat (Leica), and the sections were stained with Oil Red O working solution for 20 min. The sections were then washed three times for 5 min with Milli-Q water and transferred to haematoxylin solution (Sakura, 6187-2) for 1 min, followed by three washes with Milli-Q water. The sections were mounted with Fluoromount (Diagnostic BioSystems, K024) for confocal microscopy. Images were acquired using a DM2500B (Leica) with a 40× objective.

BODIPY staining

Mouse primary hepatic cells were seeded on glass coverslips, fixed with 4% paraformaldehyde in PBS for 30 min, and incubated with 0.5 μM BODIPY 493/503 (Cayman Chemical, 25892) for 20 min. The coverslips were mounted with Fluoromount (Diagnostic BioSystems, K024) for confocal microscopy. Frozen blocks were sectioned using a cryostat (Leica), and the sections were stained with 0.5 μM BODIPY 493/503 for 3 h. The sections were subsequently washed twice in 1x PBS and then mounted with Fluoromount (Diagnostic BioSystems, K024) for confocal microscopy. Images were acquired using a TCS SP8 confocal microscope with a 63× oil immersion objective. Lipid droplet counts were quantified using CellProfiler software.

Liver triglyceride quantification

Liver samples ranging from 40 to 60 mg were washed three times with 1x PBS. The samples were subsequently lysed with 10% NP-40. The livers were then homogenized using a homogenizer. The samples were boiled at 98 °C for 3 min with intermittent vortexing, cooled to room temperature, and then boiled again for complete solubilization of triglycerides. The samples were centrifuged at 10,000 × g for 10 min at 4 °C, after which the supernatant (including the lipid layer) was collected. The samples were diluted 1:20 in water for triglyceride measurements. Liver triglycerides were quantified using a triglyceride assay kit (Abcam, ab653363) in accordance with the manufacturer’s instructions.

ICP‒MS

The zinc concentration in liver tissues was measured via ICP‒MS (Agilent 7800, Agilent Technologies). Liver tissues were dried at 100 °C for 16 h. Thirty milligrams of dried liver was decomposed using concentrated nitric acid at 150 °C for 15 h. The decomposed product was dissolved in 0.08 M nitric acid. Zinc (m/z 66) was used to measure the zinc concentration, and indium (m/z 115) was used as an internal standard to correct the measurement results.

RNA-seq

Total RNA isolated by ISOGEN was used for library construction for RNA-Seq analysis. Library preparation was conducted using an NEBNext® Single Cell/Low Input RNA Library Prep Kit for Illumina® (NEB, E6420). Deep sequencing was performed on the Illumina NextSeq2000 platform to obtain 36-bp paired-end reads. Approximately 15 million sequences were obtained and mapped to the reference human genome (hg38) or mouse genome (mm10) via HISAT2 (v.2.2.1). Transcript read counts were determined using featureCounts (v.2.0.6). Differential gene expression analysis was performed using the R (v.4.3.2) package DESeq2 (v.1.42.1).

ChIP-seq

Chromatin samples were prepared using a SimpleChIP Enzymatic Chromatin IP Kit following the manufacturer’s instructions. For normalization, the samples were mixed with spike-in chromatin (Active Motif, 53083). The chromatin samples were then incubated with an anti-H3K14ac antibody (Abcam, ab52946) and a spike-in antibody (Active Motif, 61686) for 8 h at 4 °C with rotation. The samples were subsequently purified as described in the “ChIP‒qPCR” section. Library preparation was conducted using an NEBNext® Single Cell/Low Input RNA Library Prep Kit for Illumina® (NEB, E6420). Deep sequencing was performed on the Illumina NextSeq2000 platform to obtain 36-bp paired-end reads. Approximately 30 million sequences were obtained and mapped to the reference human genome (hg38) or mouse genome (mm10) via HISAT2 (v.2.2.1). The H3K14ac signals on the enhancers were calculated programmatically. BED files for enhancer regions were downloaded from the EnhancerAtlas2.0⁴³ and FANTOM5^91,92 databases. The H3K14ac signals on the enhancer regions were analysed via wigglescout, an R package (https://github.com/cnluzon/wigglescout/) on a supercomputer. The supercomputing resource was provided by the Human Genome Center (the Univ. of Tokyo) and the Institute of Statistical Mathematics.

Correlation analysis between H3K14ac ChIP-seq signal and gene expression

To investigate the relationship between transcriptional changes and histone modification alterations, microarray data were retrieved from the Gene Expression Omnibus (GEO) dataset GSE49657. This dataset comprises samples subjected to TPEN treatment or FBS starvation (FBS(−)), both of which represent zinc-deficient stimuli. Genes exhibiting a fold change > 1.5 under these zinc-deficient conditions compared to controls were defined as up-regulated and down-regulated. The change in H3K14ac ChIP-seq signal intensity was calculated for enhancer regions associated with these genes. Subsequently, genes were ranked based on the magnitude of H3K14ac signal reduction at their associated enhancers and classified into four quartiles (bins 1–4). Bin 1 comprises genes associated with enhancers showing the largest decrease in H3K14ac signal, whereas Bin 4 represents those with the smallest decrease.

Quantification and statistical analysis

All the statistical data were analysed via R (ver. 4.0.5) with RStudio (ver. 2022.12.0 + 353). P values < 0.05 were considered to indicate statistical significance. Detailed statistical information for each experiment, including the number of biological replicates (n), is shown in the figures and corresponding legends.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Author contributions

Conceptualization, T.F.; Methodology, T.F., T.K., T.M., S.M., Y.K., M.S., and H.N.; Investigation, T.F., S.T., L.M., M.O.; Formal analysis, T.F.; Visualization, T.F. and S.T.; Funding acquisition, T.F., S.T., M.S. and H.I.; Supervision, T.F. and H.I.; Writing – Original Draft, T.F. and S.T.; Writing—Review & Editing, S.T., L.M., M.O., T.K., T.M., S.M., I.N., and H.I.

Peer review

Peer review information

Nature Communications thanks the anonymous reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

Data availability

The RNA-seq and ChIP-seq data from this study have been deposited in the Gene Expression Omnibus (GEO) under accession numbers GSE288563, GSE288564, and GSE288565. Source data are provided with this paper.

Code availability

All numerical values used to generate the graphs, with the detailed statistical analysis procedures, have been deposited in the following GitHub repository: https://github.com/FujisawaGroup/zinc_h3k14ac.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41467-026-69476-z.