Hematopoiesis integrates cytokine signaling, metabolism, and epigenetic modifications to regulate blood cell generation. These processes are linked, as metabolites provide essential substrates for epigenetic marks. In this study, we demonstrate that ATP citrate lyase (Acly), which metabolizes citrate to generate cytosolic acetyl-CoA and is of clinical interest, can regulate chromatin accessibility to limit myeloid differentiation. Acly was tested for a role in murine hematopoiesis by small-molecule inhibition or genetic deletion in lineage-depleted, c-Kit–enriched hematopoietic stem and progenitor cells from Mus musculus. Treatments increased the abundance of cell populations that expressed the myeloid integrin CD11b and other markers of myeloid differentiation. When single-cell RNA sequencing was performed, we found that Acly inhibitor–treated hematopoietic stem and progenitor cells exhibited greater gene expression signatures for macrophages and enrichment of these populations. Similarly, the single-cell assay for transposase-accessible chromatin sequencing showed increased chromatin accessibility at genes associated with myeloid differentiation, including CD11b, CD11c, and IRF8. Mechanistically, Acly deficiency altered chromatin accessibility and expression of multiple C/EBP family transcription factors known to regulate myeloid differentiation and cell metabolism, with increased Cebpe and decreased Cebpa and Cebpb. This effect of Acly deficiency was accompanied by altered mitochondrial metabolism with decreased mitochondrial polarization but increased mitochondrial content and production of reactive oxygen species. The bias to myeloid differentiation appeared due to insufficient generation of acetyl-CoA, as exogenous acetate to support alternate compensatory pathways to produce acetyl-CoA reversed this phenotype. Acly inhibition thus can promote myelopoiesis through deprivation of acetyl-CoA and altered histone acetylome to regulate C/EBP transcription factor family activity for myeloid differentiation.
Cell differentiation allows long-term maintenance of populous short-lived cell types derived from more durable and rare stem cells. Stem cells respond to a combination of cytokine innervation, metabolic activation, and epigenetic changes to generate various cell types as needed (1–3). Hematopoietic stem cells reside within hypoxic, perivascular bone marrow (BM) niches surrounded by mesenchymal stromal cells, nonmyelinating Schwann cells, megakaryocytes, macrophages, CXCL12-abundant reticulocytes, and other cells (4). These cells and local nutrients provide the mixture of microenvironmental metabolites and cytokines that promote hematopoiesis (5). Step-by-step relationships between hematopoietic progenitors demarcated by surrogate markers have been painstakingly established over time. However, hematopoiesis is a fluid landscape with a gradient of differentiation regulated by intersecting systems of cellular metabolism and epigenetic homeostasis that remain incompletely understood. Metabolic hubs including the tricarboxylic acid (TCA) cycle are closely connected to epigenetic regulation of gene expression and supply essential substrates such as S-adenosyl methionine, acetyl-CoA, and α-ketoglutarate, which facilitate methylation, acetylation, and demethylation, respectively (6). Understanding how metabolic and epigenetic stimuli influence hematopoiesis will help build a more comprehensive paradigm of cellular development and stimulate new therapeutic approaches against disease.
Although cytokines are the primary drivers of hematopoiesis, metabolic activation and epigenetic intervention also influence these pathways to guide hematopoietic cell fate decisions (6–8). It has been recognized that metabolic activation itself can also decrease stem cell potential and induce differentiation. Acetyl-CoA provides the substrate for lipid synthesis and protein acetylation reactions and derives from numerous bioenergetic pathways including glycolysis, fatty acid oxidation, amino acid deamination, TCA cycle flux, and acetate. Histone acetyltransferases (HATs) catalyze the addition of acetyl groups to histone tails and are sensitive to changes in the concentration of their primary substrate, acetyl-CoA. When the intracellular concentration of acetyl-CoA is at its nadir, the enzyme velocities of HATs decrease to alter the steady-state maintenance of histone acetylation and chromatin accessibility (6). Aberrant acetylation in hematopoiesis by HATs such as GCN5, CBP/p300, MOZ, and TIF2 has been implicated in transforming healthy hematopoietic stem cells and precursors into hematologic malignancies (9).
ATP citrate lyase (Acly) catalyzes the reaction of ATP, citrate, and CoA into ADP, inorganic phosphate, acetyl-CoA, and oxaloacetate. Acly reactions are key contributors to the cytosolic nuclear pools of acetyl-CoA and are the primary path for acetyl-CoA derived from mitochondrial citrate. Acly represents one of several sources of cellular acetyl-CoA that contribute to histone acetylation (10, 11). The relative roles of these pathways, however, are poorly understood. Acly expression may play an important role in myeloid differentiation and myeloid cell function, as increased PU.1 was associated with reduced Acly expression and regulation of the cell cycle in a myeloid cell line (12). The impact of Acly is not limited to myeloid cells, as Acly has been noted as a regulator of CD8+ T cell function, early CD8+ T cell differentiation, and osteoclast differentiation (13–15).
In this study, we sought to test the role of Acly on differentiation of primary hematopoietic stem and progenitor cells (HSPCs). Inhibiting Acly with the small-molecule inhibitor SB204990 or genetically deleting Acly in lineage-depleted and c-Kit–enriched HSPCs led to increased myeloid differentiation in cytokine-replete methylcellulose media. A single-cell assay for transposase accessible chromatin sequencing (scATAC-seq) of HSPCs treated with SB204990 showed altered gene expression and chromatin accessibility with increased myeloid cell abundance consistent with a role for Acly in the epigenetic regulation of HSPCs. Mechanistically, multiple members of the C/EBP transcription factor family were affected and may contribute to increased myelopoiesis. Acly inhibition resulted in increased Cebpe and reduced Cebpa and Cebpb gene expression by quantitative RT-PCR (qRT-PCR) and increased chromatin accessibility at C/EBP transcription factor binding sites by scATAC-seq. Mitochondrial metabolism was also altered, and acetate restored normal myeloid differentiation to suggest that Acly inhibition resulted in limiting levels of acetyl-CoA. Acly thus contributes as one pathway to acetylation in hematopoiesis and can regulate myeloid differentiation through metabolic regulation of epigenetic marks and myeloid gene expression.
Materials and Methods
Wild-type C57BL/6 mice (strain 000664) were obtained from The Jackson Laboratory. Bones from UBC-Cre ERT2;Acly f/f mice were provided by K. Wellen. Animals were maintained under specific pathogen-free conditions and handled per the Association for Assessment and Accreditation of Laboratory Animal Care guidelines. The Institutional Animal Care and Use Committee at Vanderbilt University approved the experiments. Six- to 16-wk-old female mice were used in all experiments.
BM from Acly f/f Vav1-cre+/− and Acly f/f mice was injected into lethally irradiated (2 × 5 Gy separated by 3 h), BL/6 CD45.1 (The Jackson Laboratory strain 002014) recipients and were monitored for 4 wk posttransplant. Each mouse received 1 × 107 whole BM cells via retro-orbital injection after being placed on sulfamethoxazole/trimethoprim-treated water. Groups of five mice per condition (Acly f/f Vav1-cre+/− or Acly f/f) were taken down at 7, 14, 21, and 28 d posttransplant.
Methylcellulose culturing of lineage-negative c-Kit HSPCs
BM was taken from C57BL/6 mouse bones, strained through a 70-μm MACS SmartStrainer (Miltenyi Biotec, catalog no. 130-110-916), and lysed for RBCs using ACK (ammonium-chloride-potassium) lysis buffer (Thermo Fisher Scientific, catalog no. A1049201). Cells were counted and depleted of cells expressing CD5, CD45R (B220), CD11b, Gr-1 (Ly-6G/C), 7-4, and Ter-119 using a MACS lineage cell depletion kit (Miltenyi Biotec, catalog no. 130-090-858) according to the manufacturer’s protocol. Cells were then counted again and enriched for cells expressing c-Kit/CD117 using MACS CD117 MicroBeads (Miltenyi Biotec, catalog no. 130-091-224). Unless otherwise stated, cells were counted and resuspended at a concentration of 1 × 104/ml in PBS. Cells were added to tubes of MethoCult GF M3434 (STEMCELL Technologies) cytokine-replete methylcellulose media to dilute the original cell concentration (1:10) for a final cell concentration of 1 × 103 cells/ml. One microliter of this mixture was then added to a 35-mm culture dish (STEMCELL Technologies, catalog no. 27150) and cultured for 2 wk in incubators with 5% CO2 at 37°C. Cells were then harvested and resuspended in 2% FBS (VWR, catalog no. 97068-085)/PBS for flow cytometry. When applicable, unless otherwise stated, 30 μM Acly inhibitor (Aclyi), 4 μM (Z)-4-hydroxytamoxifen, ≥98% Z isomer (Sigma-Aldrich, catalog no. H7904-5MG), 5 mM acetate (sodium acetate trihydrate [BioXtra], ≥99.0%; Sigma-Aldrich, catalog no. S7670-250G), 1.25–5 mM N-acetyl-l-cysteine (NAC; ≥99% [TLC], powder; Sigma-Aldrich, catalog no. A7250-10G), 1–4 mM dimethyl 2-oxoglutarate (96%; Sigma-Aldrich, catalog no. 349631-5G), and/or 10 μM Acyl-CoA synthetase short-chain family member 2 (Acss2) inhibitor (Selleck Chemicals, catalog no. S8588) were added and vortexed well into M3434 methylcellulose media before cells were added.
Staining of cells for flow cytometry
Cells were Fc blocked in 1:50 TruStain FcX PLUS (BioLegend, catalog no. 156604) in 2% FBS/PBS for 10 min, followed by a stain of at least 30 min of 1:200 CD11b-FITC (Thermo Fisher Scientific, catalog no. 11-0112-82), 1:400 CD117-PE (Thermo Fisher Scientific, catalog no. 12-1171-82), 1:200 Sca-1-PE/Cy7 (BioLegend, catalog no. 122514), 1:1000 Ghost Dye Red 780 fixable viability dye (Cell Signaling Technology, catalog no. 18452), and/or 1:200 lineage-FITC (BioLegend catalog no. 133301). Cells were then washed twice with 2% FBS/PBS and run on the flow cytometer. For compensation, UltraComp eBeads compensation beads (Thermo Fisher Scientific, catalog no. 01-2222-42) were stained at the same concentration for the same amount of time as the cells to be analyzed, washed twice with 2% FBS/PBS, and run on the flow cytometer. For viability dye compensation, an aliquot of cells was stained only with 1:1000 Ghost Dye Red 780 fixable viability dye (Cell Signaling Technology, catalog no. 18452) for the same duration as the cells to be analyzed, washed twice with 2% FBS/PBS, and run on the flow cytometer.
Cells were incubated with CM-H2DCFDA (chloromethyl derivative of 2′,7′-dichlorodihydrofluorescein diacetate; general oxidative stress indicator) (Thermo Fisher Scientific, catalog no. C6827), tetramethylrhodamine ethyl ester (TMRE) reagent (mitochondrial membrane potential) (Thermo Fisher Scientific, catalog no. T-669), MitoTracker Green (mitochondrial mass) (Thermo Fisher Scientific, catalog no. M7514), or MitoSOX (superoxides) (Thermo Fisher Scientific catalog no. M36008). Cells were incubated for 30 min at 37°C, covered from light, washed twice with 2% FBS/PBS, and run on the flow cytometer.
Analysis of flow cytometry data
Data were analyzed using FlowJo 10.5.3. The percentage CD11b+ gate indicates live (gated on Ghost Dye Red 780 fixable viability dye), singlet (gated on forward scatter area and forward scatter height) cells positive for CD11b.
Sample preparation for single-cell RNA sequencing
C57BL/6 lineage-depleted BM cells were seeded at 1 × 103 cells per 35-mm dish in STEMCELL Technologies’ Methocult GF M3434 methylcellulose media and cultured for 2 wk with either 30 μM SB204990 (Tocris Bioscience, catalog no. 4962) or an equivalent volume of DMSO. Cells were harvested and resuspended in PBS alongside freshly isolated lineage-depleted BM cells from a C57BL/6 mouse. Cells were then stained for Ghost Dye Red 780 fixable viability dye and flow sorted for viable cells. Cells were then submitted to Vanderbilt University’s VANTAGE core, where they were prepared for single-cell 5′ RNA sequencing using the 10x Genomics chromium system. Libraries were prepared using PN 1000014, 1000020, 1000080, and 120262 according to the manufacturer’s protocol. The libraries were sequenced using the NovaSeq 6000 with 150-bp paired-end reads. RTA (version 2.4.11; Illumina) was used for base calling, and analysis was completed as described below.
Analysis of single-cell RNA sequencing data
Cellranger software (v3.0.2, https://github.com/10XGenomics/cellranger) was used with default parameters for library demultiplexing, aligning reads, fastq file generation, and unique molecular identifier (UMI) quantification to create the gene expression matrix. Gene expression matrices containing total numbers of UMIs per cell per gene were filtered to optimize data quality. To enrich for live cells, cells were retained with at least 200 genes detected and <5% mitochondrially derived reads out of total UMIs. All detected genes were retained for the following analysis. Principal component analysis and uniform manifold approximation and projection (UMAP) and clustering were applied to the filtered matrix using Seurat version 4.0.3 with default parameters, except the top 20 dimensions of principal component analysis were used for UMAP dimensional reduction (16, 17). Cell type assessment was performed using SingleR (v1.8.1) (18) inferring the most transcriptionally similar bulk transcriptome from sorted cell populations from the Immunological Genome Project (ImmGen) database (19) executed in tandem with expert gating. Gating validation was performed using Seurat to overlay gene expression onto cell type clusters and create heatmaps of cell type–associated genes. Data were visualized using Seurat-specific tools or Rathmell laboratory scripts using ggplot2. Datasets have been deposited in Gene Expression Omnibus and are publicly available under the accession number GSE217080 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE217080).
Regulatory network analysis was performed using the Docker image aertslab/pySCENIC (v0.9.18) using the default parameters. The coexpression modules were based on cisTarget databases (mm10: mm10__refseq-r80__10kb_up_and_down_tss.mc9nr.feather). Then, the scored regulon of cells was imported and integrated into Seurat object with other analysis and visualized under R (v3.6.1).
Sample preparation for scATAC-seq
C57BL/6 lineage-depleted BM cells were seeded at 1 × 103 cells per 35-mm dish in STEMCELL Technologies’ Methocult GF M3434 methylcellulose media and cultured for 2 wk with either 30 μM SB204990 (Tocris Bioscience, catalog no. 4962) or an equivalent volume of DMSO. Cells were harvested and resuspended in PBS alongside freshly isolated lineage-depleted BM cells from a C57BL/6 mouse. Nuclei were then isolated and submitted to Vanderbilt University’s VANTAGE core, where they were prepared for scATAC-seq following the manufacturer’s protocol using PN 1000111, 1000086, and 1000084.
Analysis of scATAC-seq data
Files were imported into R using ArchR (filterTSS = 8, filterFrags = 2819), with the filterFrags value set based on the distribution of transcription start site (TSS) versus the log transform of unique fragments generated by ArchR (20). The ArchR project was created, doublets were filtered, LSI dimensional reduction was performed, clusters were plotted, and UMAP dimensional reduction was embedded using default settings. Gene scores were also calculated using ArchR, representing chromatin accessibility within 100 kb on either side of a gene, using default values. Cell type labels were assigned to scATAC-seq using ArchR’s addGeneIntegrationMatrix function. Cells from scATAC-seq were directly aligned with cells from scRNA-seq by comparing the scATAC-seq gene score matrix with the scRNA-seq gene expression matrix. Chromatin accessibility tracks were plotted using default settings.
Other R packages used for data analysis and visualization include dplyr, patchwork, formattable, ggplot2, gridExtra, sctransform, GenomicRanges, reticulate, loomR, scater, pheatmap, celldex, scRNAseq, scuttle, stringr, plyr, scales, devtools, Rcpp, presto, msigdbr, fgsea, SummarizedExperiment, and chromVARmotifs (Refs. 21–43 and G. Korotkevich, V. Sukhov, N. Budin, B. Shpak, M.N. Artyomov, and A. Sergushichev, manuscript posted on bioRxiv, DOI: 10.1101/060012).
Extracellular flux analysis
Cells were resuspended in Seahorse XF media (Agilent Technologies, catalog no. 103576-100) supplemented with 2 mM glutamine (Agilent Technologies, catalog no. 103579-100), 1 mM pyruvate (Agilent Technologies, catalog no. 103578-100), and 10 mM glucose (Agilent Technologies, catalog no. 103577-100) at 4 × 106/ml, seeding 2 × 105 cells per well in the Seahorse assay plate (Agilent Technologies, catalog no. 102416-100). The rest of the protocol was conducted according to the manufacturer’s XF Cell Mito Stress Test kit (Agilent Technologies, catalog no. 103015-100) user’s guide. Seahorse XF calibrant solution (Agilent Technologies, catalog no. 100840-000) was used to calibrate the extracellular flux analyzer. Values were normalized to bright-field cell counts taken per well by a BioTek Cytation 5 cell imaging multi-mode reader.
RNA was isolated using Qiagen’s RNeasy mini kit (Qiagen, catalog no. 74104) according to the manufacturer’s protocol. cDNA was then synthesized using the Bio-Rad iScript cDNA synthesis kit (Bio-Rad, catalog no. 1708890) according to the manufacturer’s protocol. Finally, qRT-PCR was performed using the Bio-Rad SsoAdvanced Universal SYBR Green supermix (Bio-Rad, catalog no. 1725271) according to the manufacturer’s protocol. The amplification program consisted of 95°C for 30 s followed by 40 cycles of the set of 95°C for 15 s, annealing time for 30 s, and, finally, 95°C for 5 s. Each Cebp isoform’s reaction was run alongside a control β-actin reaction with different annealing time values for each Cebp isoform, depending on the specific primer characteristics.
qRT-PCR primers used included the following: β-actin (forward, 5′-AAGTGTGACGTTGACATCCGTAA-3′; reverse, 5′-TGCCTGGGTACATGGTGGTA-3′) (44), Cebpa (forward, 5′-AATGGCAGTGTGCACGTCTA-3′; reverse, 5′-CCCCAGCCGTTAGTGAAGAG-3′) (45), Cebpb (forward, 5′-TTGATGCAATCCGGATCAAACG-3′; reverse, 5′-CAGTTACACGTGTGTTGCGTC-3′) (46), Cebpg (forward, 5′-TTCGTAACCGTCGCTCCTCC-3′; reverse, 5′-TCAGAGCAATGTGATCCGAGG-3′), Cebpd (forward, 5′-GAACCCGCGGCCTTCTAC-3′; reverse, 5′-GAAGAGTTCGTCGTGGCACA-3′) (47), Cebpe (forward, 5′-CCCTTCTAGGTCCCCAGAGT-3′; reverse, 5′-TCATTTGGTCCCGACCTTCC-3′), Cebpz (forward, 5′-AGCCAGATCCCAGTGGATGA-3′; reverse, 5′-GTGGGAAGCAGTTGTCGTCT-3′).
Quantification and statistical analysis
Statistical analyses were performed with GraphPad Prism software version 8.1.0. For data involving two groups, analysis was performed using a Student t test.
Inhibition of Acly promotes myelopoiesis
To test the role of Acly in hematopoiesis, HSPCs were fist isolated by lineage depletion and c-Kit enrichment. We depleted cells expressing a surrogate marker for myeloid differentiation CD11b from BM (Fig. 1A) (48–50). HSPCs were then cultured in cytokine-replete methylcellulose media (MC-cultured) for 2 wk in the presence of vehicle or Aclyi SB204990 at 30 μM, a concentration previously used to provide a near-complete inhibition of glucose-derived lipid synthesis (51, 52). Cells were stained for lineage markers anti-CD3, anti-Gr-1, anti-CD11b, anti-CD45R, and anti-Ter-119 as well as Sca-1 and c-Kit that indicate the canonical LSK compartment that is enriched in stem cells (52, 53). Aclyi resulted in a significant decrease in lineage-negative, Sca-1+, c-Kit+ (LSK+) cells (Fig. 1B). We then asked which lineages Aclyi increased by assessing CD11b expression in MC-cultured HSPCs. Despite a modestly decreased viability, live cell numbers were unchanged and Aclyi significantly increased the frequency and number of myeloid CD11b+ cells to indicate shift toward myelopoiesis in culture (Fig. 1C). This effect was observed as early as day 4 of cell culture with high doses of SB204990 (Supplemental Fig. 1).
Genetic deletion of Acly promotes CD11b with and without myelopoiesis while suppressing mast-like cells
To corroborate that small-molecule inhibition of MC-cultured HSPCs resulted in increased myeloid differentiation, we tested whether acute genetic deletion of Acly led to similar shifts toward myelopoiesis (Fig. 2). Acly was genetically deleted using a UBC-Cre-ERT2+/− Acly f/f model with tamoxifen-inducible Acly deletion by treating isolated HSPCs with 4-hydroxytamoxifen. This approach to delete Acly resulted in a significant increase in the percentage of CD11b+ cells that mirrored the percentage of CD11b+ myeloid cell increases induced by Acly small-molecule inhibition. Small-molecule inhibition of Acly in combination with Acly deletion further increased the proportion of myeloid CD11b+ cells in MC-cultured HSPCs. This additive effect may be due to incomplete deletion of Acly or non-Acly effects of SB204990. Interestingly, although inhibition of Acly did not result in decreased overall cell numbers, treatment with 4-hydroxytamoxifen and genetic deletion of Acly reduced total live cell numbers, but with an increased frequency of CD11b+ cells. Both Aclyi and genetic deletion of Acly resulted in a significant reduction in the numbers and percentage of c-Kit (CD117)–expressing cells (Fig. 2). These CD117+ cells most likely represent stem and progenitor or mast cells, which often comprise HSPC-descended cells in methylcellulose cultures (54, 55).
To determine the role of Acly in vivo, we assessed demand-induced myelopoiesis in Acly-deficient BM chimeras using hematopoietic-specific, Acly f/f Vav1-cre+/−, Acly-deficient BM chimeras. Acly f/f Vav1-cre+/− or Acly f/f BM was transplanted into lethally irradiated C57BL/6 CD45.1 hosts and recipient BM was characterized by flow cytometry at 14 and 21 d posttransplant. Acly deficiency in Acly f/f Vav1-cre+/− BM recipients resulted in a modest initial increased CD16/32 (FcγRIII) expression, consistent with Acly suppression of myelopoiesis, although B and NK cells may have also contributed to increased CD16/32 expression (Supplemental Fig. 2). One week later, myelopoiesis had increased in both control and Acly-deficient cultures and this difference was no longer apparent. The effect of genetic Acly deficiency was modest and transient, demonstrating that Acly-independent pathways can compensate for Acly loss to generate cytosolic acetyl-CoA in vivo. Acly may thus act in concert with other pathways, such as Acss2-mediated conversion of acetate to acetyl-CoA, to influence hematopoietic differentiation under stressful or non–steady-state conditions following stem cell transplant with the effect of Acly compensated over time.
Acly inhibition promotes macrophage and neutrophil differentiation
To better understand how Aclyi affected HSPC populations and gene expression, scRNA-seq and scATAC-seq were performed. Freshly isolated and lineage-depleted C57BL/6 BM and MC-cultured lineage-depleted BM cells were treated with either Aclyi or vehicle prior to scRNA-seq (Fig. 3A–C). Cell cluster identities were assigned using the software package SingleR, which compares scRNA-seq transcriptomes to bulk RNA-seq transcriptomes of expert-gated, sorted immune populations from the ImmGen database (18, 19). Although use of SingleR assigns clusters of differentiating cells from our cultures to defined and often mature cell populations that may not be fully equivalent, it nevertheless provides an unbiased approach to infer cell lineage and identity. These clusters consisted of B cells, basophils, eosinophils, macrophages, mast cells, monocytes, neutrophils, HSPCs, and “other,” with other comprising a minority of cells that had no viable comparisons to the ImmGen database using SingleR (Fig. 3B, 3C). Most cells in the vehicle-treated HSPCs were categorized as mast cells, and treatment with Aclyi increased the proportion of macrophages while suppressing mast cell differentiation (Fig. 3C). These MC-cultured HSPCs share gene expression similarities with canonical myeloid gene expression profiles, and expert gating of scRNA-seq data revealed that expression of cell type–associated genes correlated with SingleR-identified cell type clusters (Fig. 3D, 3E) (56). Although the HSPC-descended cell types did not fully match canonical immune cell transcriptomes, we refer to each based on SingleR categorization. Transcription factor binding-motif analysis of differentially expressed genes using SCENIC identified a variety of transcriptional drivers of myeloid differentiation in the presence of Aclyi that corroborated the SingleR-assigned cell clusters. These included the macrophage-associated transcription factor-coding genes Spi1, as well as Nfe2l2 (NRF2) and binding partners Maf and Mafb (57–59) (Supplemental Table I).
Chromatin accessibility was measured by scATAC-seq and closely matched the scRNA-seq results. Cells from the same sample subjected to scRNA-seq were split and assessed by scATAC-seq, with the freshly isolated and lineage-depleted murine BM clustering separately from the MC-cultured, lineage-depleted BM (Fig. 4A). The software package ArchR was used to map the gene expression signatures of the SingleR-identified populations from the scRNA-seq onto the scATAC-seq data (20). SingleR identified populations that correlated with expected populations based on scRNA-seq. HSPCs were enriched in the fresh lineage-depleted, undifferentiated sample, whereas mature myeloid populations such as basophils, eosinophils, mast cells, neutrophils, and macrophages were concentrated in the MC-cultured, lineage-depleted conditions (Fig. 4B).
Gene scores, as defined by ArchR to measure chromatin accessibility within 100,000 bp of genes of interest, were used to uncover populations with distinct regions of chromatin accessibility (Fig. 4C) (20). When overlaid onto the UMAP plot, expression of cell type–associated genes including Cd34, Mmp8, and Itgam (CD11b) revealed close association with canonical cell-type gene expression patterns (Fig. 4D, 4E). As in the scRNA-seq dataset, the proportion of mast cells in the vehicle was significantly reduced whereas macrophages increased in the presence of Aclyi. Furthermore, one of the most differentially expressed genes between vehicle and Aclyi and between fresh lineage-depleted and Aclyi was Gpnmb, with an average 4.65-fold increase in expression with Aclyi compared with fresh lineage-depleted and a 3.34-fold increase in expression with Aclyi compared with vehicle, respectively. Gpnmb, or glycoprotein nonmetastatic B, is a soluble glycosylated transmembrane protein highly expressed in macrophages that negatively regulates inflammation (60, 61). There was also evidence that Aclyi promoted neutrophil differentiation, as the top differentially expressed gene between fresh lineage-depleted BM and Aclyi was Ngp, a gene coding for neutrophilic granule protein that accumulates in the cytoplasmic granules of neutrophilic precursors. Furthermore, Ngp is known to be regulated by C/EBPε and PU.1, which have transcription factor motifs enriched in the Aclyi scATAC-seq condition (62). Our single-cell data demonstrate a promyelopoietic effect of Aclyi that promotes macrophage and neutrophil differentiation by altering chromatin accessibility and gene expression in MC-cultured HSPCs.
Acly inhibition drives Cebpe and suppresses Cebpa and Cebpb expression
After Aclyi was found to promote macrophage differentiation at the expense of mast cell development, scATAC-seq of Aclyi-induced myeloid differentiation was analyzed to identify potential sequence motifs and transcriptional regulatory families that may respond to Aclyi. Through ArchR-based analysis of scATAC-seq data, transcription factor motifs were identified that were enriched in fresh lineage-depleted BM, Aclyi-treated, and vehicle-treated MC-cultured lineage-depleted murine BM. ArchR searched for transcription factor binding motifs enriched in the open chromatin of analyzed samples. ArchR identified several transcription factors, including Nfe2l2 and the Cebpa, Cebpb, Cebpg, Cebpd, Cebpe, and Cebpz members of the C/EBP family of transcription factors, as enriched in the Aclyi condition (Fig. 5A). These transcription factors are considered master regulators of hematopoiesis and cellular differentiation (63). C/EBP family transcription factors are also known regulators of the top differentially expressed gene between fresh lineage-depleted BM and Aclyi by scRNA-seq, Ngp (64) (Supplemental Table II). Although all family members share a high binding affinity for the promoter CCAAT box sequence, each member has unique structural elements that render its function unique, and all play a role to balance hematopoiesis (65).
To identify which C/EBP family member was most changed in expression in response to Aclyi, we performed qRT-PCR for Cebpa, Cebpb, Cebpg, Cebpd, Cebpe, and Cebpz on RNA isolated from MC-cultured HSPCs treated with Aclyi or vehicle. Aclyi resulted in significant reductions in Cebpa and Cebpb with a simultaneous significant increase in Cebpe (Fig. 5B). Furthermore, Cebpe’s chromatin region is most open in neutrophils and HSPCs retained in methylcellulose culture when the gene score is overlaid on the UMAP plot (Fig. 5C). Although the proportion of neutrophils is consistent between vehicle and Aclyi, the substantial increase in Ngp expression and enrichment of Cebpe binding motif accessibility with Aclyi suggest that Aclyi could promote neutrophil differentiation (Fig. 5A, Supplemental Table II). Additionally, neutrophils and HSPCs from Aclyi had greater chromatin accessibility at its Cebpe locus compared with vehicle and fresh lineage-depleted BM cells, suggesting Aclyi may open chromatin at Cebpe and poise HSPCs for neutrophilic differentiation (Fig. 5D). These data support a potential role for C/EBP transcription factors to drive macrophage differentiation in response to Acly, but that Cebpe is not essential for this process upon Acly inhibition.
Acly inhibition impacts mitochondrial health and superoxide production
Acly catalyzes the conversion of mitochondrially derived citrate to acetyl-CoA, and inhibition of this pathway may affect mitochondrial metabolism. C/EBP family transcription factors are also known regulators of cellular metabolism, differentiation, and immune function (66, 67). We thus tested whether Aclyi altered metabolic and mitochondrial characteristics of MC-cultured HSPCs by measuring extracellular flux. The oxygen consumption rate and extracellular acidification rate of MC-cultured HSPCs were not significantly altered with Aclyi (Fig. 6A). While the respective rates of oxidative phosphorylation and glycolysis did not change, the mitochondrial potential sensitive dye TMRE mean fluorescence intensity significantly decreased with Aclyi, showing that mitochondrial membrane potential and electron transport chain efficiency were decreased (Fig. 6B). General oxidative stress as measured by 2′,7′-dichlorodihydrofluorescein diacetate mean fluorescence intensity was not also significantly changed with Aclyi (Fig. 6C), but mitochondrial superoxide and mass increased with Aclyi, suggesting a worsening of mitochondrial quality and health (68) (Figs. 3E, 6D).
Acss2 inhibition, reactive oxygen species, and α-ketoglutarate do not affect myelopoiesis
Acly catalyzes the reaction between cytosolic citrate derived primarily from the TCA cycle into acetyl-CoA in an ATP-dependent manner. Acss2 can also generate cytosolic acetyl-CoA from cytosolic acetate and may complement Acly to replenish cellular acetyl-CoA from acetate in an ATP-dependent reaction (69) (Fig. 7A). When MC-cultured HSPCs were treated with Acss2i, the proportion of CD11b+ cells did not increase as with Aclyi. Two metabolic signaling mechanisms that may be influenced by Aclyi and the failure to convert citrate to acetyl-CoA are reactive oxygen species or accumulation of citrate and conversion to α-ketoglutarate, which regulates histone and DNA demethylation reactions and has been shown to impact hematopoietic differentiation. MC-cultured HSPCs were supplemented with NAC, a precursor to the reactive oxygen species/reactive nitrogen species scavenger molecule glutathione. No significant effect of NAC was observed on myeloid differentiation as measured by the frequency of cells expressing CD11b (Fig. 7B). Similarly, cultures supplemented with dimethyl α-ketoglutarate to directly increase α-ketoglutarate had no significant difference in the frequency of cells expressing CD11b+ cells in MC-cultured HSPCs versus vehicle (Fig. 7C).
In addition, combining Acss2i and Aclyi resulted in a trend toward increased frequency and number of myeloid CD11b+ cells relative to Aclyi alone (Fig. 7D). Although Acss2i alone did not impact myelopoiesis in our MC-cultured HSPCs, we next tested whether Acss2 could rescue the Aclyi-induced myelopoiesis in the presence of exogenous acetate. MC-cultured HSPCs were then treated with Aclyi with or without exogenous acetate. Culture media supplemented with 5 mM exogenous acetate reversed the increase in the frequency and number of myeloid CD11b+ cells even in the presence of Aclyi to levels comparable to vehicle. These data suggest that although Acss2 is not essential, it can replenish lost acetyl-CoA and compensate for Acly deficiency when cells are provided excess acetate (Fig. 7E).
Hematopoiesis balances extracellular stimuli including cytokines and microenvironmental nutrients to affect intracellular changes to metabolism and gene expression that regulate stemness and differentiation of blood cells. In this study, we investigated the role of citrate metabolism and Acly in hematopoietic differentiation of HSPCs cultured in cytokine-replete methylcellulose. Small-molecule inhibition and LoxP-cre–mediated genetic deletion of Acly similarly demonstrated that Acly deficiency promotes increased myeloid differentiation. We characterized the methylcellulose-based differentiation system of HSPCs by flow cytometry for specific cell markers and by scRNA-seq and scATAC-seq for transcriptional and chromatin markers. Acly deficiency was found in each approach to promote the differentiation of cells that transcriptomically resemble macrophages when assessed using SingleR. In contrast, LSK+ cells were decreased in number and frequency by Aclyi when cell surface markers were directly measured by flow cytometry yet increased by Aclyi when assessed transcriptionally by scRNA-seq or scATAC-seq and SingleR. Taken together, these data show that Acly deficiency can promote myeloid differentiation although effects on HSPCs remain uncertain.
Cytosolic acetyl-CoA can be produced through several mechanisms. Most notably, Acly and Acss2 produce acetyl-CoA from citrate or acetate, respectively. Whereas Aclyi led increased myelopoiesis in vitro, the effect of Acly deficiency was modest and transient in vivo following BM transplant. These data suggest that Acss2 may have compensated for Acly deficiency in vivo. Consistent with Acss2-mediated compensation for Acly deficiency, supplementation with the Acss2 substrate acetate could reverse the effects of Aclyi in vitro. Because Aclyi is under study as a potential therapeutic in cancer and inflammatory diseases, it may be important to consider potential compensation, although this may also reduce on-target toxicity of such drugs.
Acly regulated several genes that may promote or influence myelopoiesis. Inhibition of Acly in MC-cultured HSPCs resulted in increased expression of Cebpe and decreased expression of Cebpa and Cebpb with concomitant increases in chromatin accessibility for C/EBP family transcription factor binding sites. Aclyi also increased activity of the transcription factor Nfe2l2, enriched in the SingleR-identified macrophage population in the scRNA-seq data and in the Aclyi condition in the scATAC-seq data. The increased Nfe2l2 activity suggests a role for redox regulation in Aclyi-promoted myelopoiesis (70–72). It will be important in future studies to further examine genes affected by Aclyi to determine which change directly in response to decreased cytosolic acetyl-CoA and which change as a secondary consequence of cell differentiation.
Endogenous citrate derived from mitochondrial TCA cycle enzyme citrate synthase can catalyze the reaction of oxaloacetate and acetyl-CoA into citrate. Transported into the cytosol by citrate transporter protein and converted back into oxaloacetate and acetyl-CoA in the cytosol by Acly, citrate serves as a convenient acetyl-CoA shuttle from its mitochondrial origin to the cytosol and adjacent compartments (51, 73). After acetyl-CoA is derived from citrate in the cytosol, acetyl-CoA can then go on to serve as substrates in reactions including de novo lipid synthesis and histone acetylation (74). Intracellular acetyl-CoA when present in low concentrations limits the reaction rate of HATs, with the potential to alter histone acetylome homeostasis, chromatin accessibility, and transcription factor binding patterns (6, 75, 76). Altered acetylation and chromatin accessibility have also been shown to change stem cell behavior and fate (9). Acly itself has been recognized as a key regulator of hematopoiesis, disease outcome in hematologic malignancies, and mature myeloid cell effector ability to clear atherosclerotic plaques (77–79). Acetyl-CoA derived from citrate and other sources has wide-ranging intracellular activity, and the above results demonstrate an impact of Acly on chromatin accessibility patterns that result in altered gene expression and stem cell differentiation.
Cytokine-replete methylcellulose has been extensively used to assess in vitro hematopoietic stem cell differentiation and self-renewal (55, 80–83). The current classification of differentiation outcomes relies on light microscope–based colony morphology assessment, more recently aided by advances in imaging interpretation software (84). Potential lineages include but are not limited to granulocytic (CFU-G), erythrocytic (CFU-E), macrophage (CFU-M), and multipotent progenitor (CFU-GEMM) (85). Although flow cytometry has instead been used in the past to assess in vitro hematopoietic differentiation outcomes, this study, to our knowledge represents the first application of scRNA-seq and scATAC-seq to characterize populations derived from cytokine-replete methylcellulose culture (86). Several populations were identified in steady-state (vehicle) and perturbed (Aclyi) conditions using these methods, comprising cell types that transcriptionally resemble neutrophils, eosinophils, basophils, macrophages, monocytes, HSPCs, and mast cells. Our characterization represents a quantitative evaluation of the methylcellulose method that can be used to support traditional and computer-aided imaging techniques.
These data highlight a role for Acly and cytosolic acetyl-CoA in the epigenetic regulation of HSPC and myeloid differentiation. When Acly is inhibited and acetyl-CoA production CoA decreases, macrophage differentiation is favored, whereas mast cell differentiation occurs when Acly is active and cytosolic acetyl-CoA is not limiting. Two limitations to the study are that acetyl-CoA can be generated through multiple means that may compensate in context-specific manners and that Acly inhibitors may have off-target effects. The use of Acly genetic deletion, however, supports a direct and at least partially nonredundant role for Acly-derived acetyl-CoA in myelopoiesis. These data are like those of Rhee et al. (12), who showed that Acly played a role to regulate proliferation of myeloid cells in part through the transcription factor PU.1. It will be important in future studies of metabolic regulation of epigenetic marks, gene expression, and cell differentiation to further establish these associations and how these pathways may be modulated to promote myelopoiesis or differentiation of myeloid precursor cells.
We thank members of the Rathmell laboratory and co-authors for input.
This work was supported by the National Institutes of Health Grants R01DK105550 and R01CA217987 (to J.C.R.), R01DK116005 (to K.E.W.), F31HL152529 (to D.L.G.), F30CA239367 (to M.Z.M.), T32GM007347 (to A.S.), T32DK1011003 (to K.V.), K00CA234920 (to J.E.B.), and K23HL138291 (to P.B.F.), a Blavatnik Family Foundation predoctoral fellowship (to P.T.T.N.), and by a grant from the Vanderbilt–Incyte Alliance (to J.C.R., M.R.S., and P.B.F.).
The online version of this article contains supplemental material.
Abbreviations used in this article
ATP citrate lyase
Acyl-CoA synthetase short-chain family member 2
hematopoietic stem and progenitor cell
Immunological Genome Project
cultured in cytokine-replete methylcellulose media
single-cell assay for transposase-accessible chromatin sequencing
single-cell RNA sequencing
tetramethylrhodamine ethyl ester
uniform manifold approximation and projection
unique molecular identifier
M.R.S.: Membership on a board or advisory committee: Abbvie, Blackbird, Bristol Myers Squibb, CTI, Forma, Geron, Karyopharm, Novartis, Ryvu, Sierra Oncology, Taiho, Takeda, and TG Therapeutics; patents and royalties: Boehringer Ingelheim; research funding: ALX Oncology, Astex, Incyte, Takeda, and TG Therapeutics; equity ownership: Karyopharm and Ryvu. J.C.R.: Founder, scientific advisory board member, and stockholder of Sitryx Therapeutics; scientific advisory board member and stockholder of Caribou Biosciences; member of the scientific advisory board of Nirogy Therapeutics; has consulted for Merck, Pfizer, and Mitobridge within the past three years; and has received research support from Incyte, Calithera Biosciences, and Tempest Therapeutics. P.B.F.: Receives research funding from Incyte. The other authors have no financial conflicts of interest.