Evidence for AKT-independent regulation of FOXO1 and FOXO3 in haematopoietic stem and progenitor cells

<p>Transcription factors FOXOs (1, 3, 4) are essential for the maintenance of haematopoietic stem cells. FOXOs are evolutionary conserved substrates of the AKT serine threonine protein kinase that are also phosphorylated by several kinases other than AKT. Specifically, phosphorylation by AKT is known to result in the cytosolic localization of FOXO and subsequent inhibition of FOXO transcriptional activity. In addition to phosphorylation, FOXOs are regulated by a number of other post-translational modifications including acetylation, methylation, redox modulation, and ubiquitination that altogether determine these factors' output. Cumulating evidence raises the possibility that in stem cells, including in haematopoietic stem cells, AKT may not be the dominant regulator of FOXO. To address this question in more detail, we examined gene expression, subcellular localization, and response to AKT inhibition of FOXO1 and FOXO3, the main FOXO expressed in HSPCs (haematopoietic stem and progenitor cells). Here we show that while FOXO1 and FOXO3 transcripts are expressed at similar levels, endogenous FOXO3 protein is mostly nuclear compared to the cytoplasmic localization of FOXO1 in HSPCs. Furthermore, inhibition of AKT does not enhance nuclear localization of FOXO1 nor FOXO3. Nonetheless AKT inhibition in the context of loss of NAD-dependent SIRT1 deacetylase modulates FOXO3 localization in HSPCs. Together, these data suggest that FOXO3 is more active than FOXO1 in primitive haematopoietic stem and multipotent progenitor cells. In addition, they indicate that upstream regulators other than AKT, such as SIRT1, maintain nuclear FOXO localization and activity in HSPCs.</p

Bacteria integrate stimuli from the environment and decide whether to make biofilms or to move using the c-di-GMP network.

<p>A: Bow-tie architecture of c-di-GMP signaling network: c-di-GMP is synthesized by diguanylate cyclase (DGC) proteins with GGDEF domains such as WspR, DipA, and SadC, and degraded by phosphodiesterases (PDE) proteins with EAL or HD-GYP domains such as BifA, and SadR. The DGCs and PDEs could sense stimuli—such as chemoattractants which could be a signal for motility, or mechanical contact with surfaces which could be a signal for biofilm formation—and change intracellular c-di-GMP levels in response; c-di-GMP effectors—such as c-di-GMP binding proteins and riboswitch RNAs—then sense c-di-GMP levels and control phenotype outputs such as biofilm formation, motility, virulence and cell division. B: At low levels of c-di-GMP the bacteria express flagella genes and go into motile mode. C: At high levels of c-di-GMP the bacteria repress flagella genes, express biofilm genes and go into biofilm mode.</p

Specialist strains produced by strong selection in laboratory evolution have large-effect alleles in c-di-GMP network.

<p>A: Bulk c-di-GMP levels measured for evolved mutants, collected from bacterial colonies. B: Diagram of drip flow biofilm reactor used in biofilm selection. C: Biofilm levels quantified by the crystal violet assay. D: Production of extracellular polymers required for biofilm formation, measured using the Congo-red assay. E: Expression of the gene <i>fliC</i> required for flagella synthesis, measured as GFP expressed by the reporter fusion P_<i>fliC</i>-GFP. The data of three evolved mutants <i>fleN*dipA*</i>, <i>fleN*dipA**</i> and <i>fleN*wspF*</i> in B-E are statistically different from ancestral strain <i>fleN*</i> (P<0.05). F: Phylogenetic representation of the mutants evolved in laboratory experiments showing the tradeoff between biofilm and swarming.</p

Experimental tests reveal new mutations that regain hyperswarming to a biofilm specialist.

<p>A: The <i>wspF</i>* biofilm specialist which has a repeat-insert in the <i>wspF</i> gene, initially cannot swarm but regains swarming by losing the repeat-insert when in swarming selection. B: An engineered <i>fleN</i>*Δ<i>wspF</i> strain also regains swarming despite lacking the <i>wspF</i> gene entirely. Survival analysis reveals that this mutant takes significantly longer than the <i>wspF</i>* to start swarming, but does so eventually. C: A spontaneous mutant in <i>wspA</i> regained swarming in the <i>fleN</i>*Δ<i>wspF</i> background. D-F: Diagram explaining how Wsp mutations enable switching between extremes of biofilm and swarming. When WspA senses an attachment signal, it transduces the signal to other Wsp proteins that phosphorylate protein WspR, which then produces c-di-GMP and the cells form biofilm (D). When WspF gains the insertion mutation, it fails to demethylate. WspR therefore is hyper-phosphorylated even in the absence of an attachment signal. (E). A Δ<i>wspF</i> mutant phenocopies <i>wspF*</i>. However, a spontaneous mutation in <i>wspA</i> enables cell to swarm. This mutation impairs biofilm formation even when the cells are placed under biofilm forming condition (F). G: Compilation of mutations identified from the mass swarming selection experiment started with the <i>fleN</i>*Δ<i>wspF</i> strain that revealed 43 new Wsp-disabling mutations.</p

Bacteria integrate stimuli from the environment and decide whether to make biofilms or to move using the c-di-GMP network.

<p>A: Bow-tie architecture of c-di-GMP signaling network: c-di-GMP is synthesized by diguanylate cyclase (DGC) proteins with GGDEF domains such as WspR, DipA, and SadC, and degraded by phosphodiesterases (PDE) proteins with EAL or HD-GYP domains such as BifA, and SadR. The DGCs and PDEs could sense stimuli—such as chemoattractants which could be a signal for motility, or mechanical contact with surfaces which could be a signal for biofilm formation—and change intracellular c-di-GMP levels in response; c-di-GMP effectors—such as c-di-GMP binding proteins and riboswitch RNAs—then sense c-di-GMP levels and control phenotype outputs such as biofilm formation, motility, virulence and cell division. B: At low levels of c-di-GMP the bacteria express flagella genes and go into motile mode. C: At high levels of c-di-GMP the bacteria repress flagella genes, express biofilm genes and go into biofilm mode.</p

Phenotypic diversity in 28 <i>P</i>. <i>aeruginosa</i> isolates from acutely infected cancer patients at MSKCC explained by many small-effect alleles in c-di-GMP network.

<p>A: Bulk c-di-GMP levels collected from bacterial colonies, including for the laboratory strain PA14. B: Biofilm levels measured in microtiter plates using the crystal-violet assay. C: Motility measured as swarm area after 16 h of incubation. D: Phylogenetic tree reconstructed from 88,347 genetic variants identified in core genes, including PA14 and two other laboratory strains PAO1 and PA7. Numbers shown represent the number of open-reading frames (ORFs) identified with c-di-GMP related motifs: GGDEF domain for synthesizing c-di-GMP, EAL for degrading c-di-GMP, and effector for sensing c-di-GMP. Some ORFs encode both GGDEF and EAL domains. E: Explaining diversity in c-di-GMP, biofilm and swarming required many alleles of small-effect in c-di-GMP genes identified within the 28 genomes. Model selection using LASSO revealed that a model that explains 85% of the phenotypic deviance requires including at least 21 genetic variants in c-di-GMP related genes. E’ shows a detail of LASSO model selection, which increases the tuning parameter λ and selects variants to include in the model. F: Each of the 21 genetic variants by itself explains 27% or less of the phenotypic variance, even in the best model selected by LASSO. The analysis supports that the phenotypic diversity observed among clinical isolates is due to small-effect alleles.</p

Promoter methylation of <i>miR-124-1</i> and expression of <i>miR-124</i> in primary samples.

<p>(A) Methylation of <i>miR-124-1</i> in primary samples. (B) M-/U-MSP analysis of <i>miR-124-1</i> promoter methylation status and (C) Stem-loop qRT-PCR analysis of the mature <i>miR-124</i> expression in 25 primary NHL samples with matched DNA and RNA. ΔC_t, C_t<i>miR-124</i> -C_t<i>RNU48</i>.</p

Methylation of <i>miR-124-1</i>.

<p>(A) Schematic diagram showing the distribution of CpG dinucleotides (solid vertical lines) over the precursor (solid black box) and mature <i>miR-124-1</i>. Sequence analysis of the M-MSP product from bisulfite-treated positive control DNA showed that the cytosine [C] residues of CpG dinucleotides were methylated and remained unchanged, whereas all the other C residues were unmethylated and were converted to thymidine [T], indicating complete bisulfite conversion and specificity of MSP. Grey bars indicated the amplification regions of the MSP, ChIP, and BGS primers. (B) U-MSP showed that the methylated positive control [P] was totally methylated, and all five normal controls (N1–N5) were unmethylated. In the M-MSP, the methylated control was positive (methylated) but all normal controls were negative (unmethylated). For the cell lines, SUP-T1, SUP-M2 (ALK+), SU-DHL-1 (ALK+), KARPAS-299 (ALK+), KMS-12-PE, LP-1, OPM-2, and WL-2 were completely methylated of <i>miR-124-1</i>. (C) Bisulfite genomic sequencing for the bisulfite-treated promoter region of <i>miR-124-1</i> of normal controls (N1–N5), lymphoma and myeloma cell lines of different methylation statuses (MM, UM, or UU), and the methylated positive control were depicted. Unmethylated (empty circle) and methylated (filled circle) CpG dinucleotides were shown by eight independent clones for each sample.</p

Effect of 5-Aza-2′-deoxycytidine (5-AzadC) treatment on lymphoma and myeloma cells.

<p>(A) M-/U-MSP analysis of <i>miR-124-1</i> promoter methylation status and stem-loop qRT-PCR analysis of the mature <i>miR-124</i> expression. 5-AzadC treatment resulted in progressive demethylation of <i>miR-124-1</i> promoter, and re-expression of the mature <i>miR-124</i> in cell lines harbouring homozygous <i>miR-124-1</i> methylation. (B) ChIP analysis for trimethyl H3K4, trimethyl H3K9, acetyl H3K9, trimethyl H3K27 in <i>miR-124-1</i> promoter. 5-AzadC treatment led to augmentation of euchromatin code of trimethyl H3K4. (C) Western blot analysis of CDK6 in response to 5-AzadC treatment. Bottom row showed densitometric quantization of the Western blot, indicating relative CDK6 expression under actin normalization.</p

A Systems Approach Identifies Essential FOXO3 Functions at Key Steps of Terminal Erythropoiesis

<div><p>Circulating red blood cells (RBCs) are essential for tissue oxygenation and homeostasis. Defective terminal erythropoiesis contributes to decreased generation of RBCs in many disorders. Specifically, ineffective nuclear expulsion (enucleation) during terminal maturation is an obstacle to therapeutic RBC production <i>in vitro</i>. To obtain mechanistic insights into terminal erythropoiesis we focused on FOXO3, a transcription factor implicated in erythroid disorders. Using an integrated computational and experimental systems biology approach, we show that FOXO3 is essential for the correct temporal gene expression during terminal erythropoiesis. We demonstrate that the FOXO3-dependent genetic network has critical physiological functions at key steps of terminal erythropoiesis including enucleation and mitochondrial clearance processes. FOXO3 loss deregulated transcription of genes implicated in cell polarity, nucleosome assembly and DNA packaging-related processes and compromised erythroid enucleation. Using high-resolution confocal microscopy and imaging flow cytometry we show that cell polarization is impaired leading to multilobulated <i>Foxo3</i>^<i>-/-</i> erythroblasts defective in nuclear expulsion. Ectopic FOXO3 expression rescued <i>Foxo3</i>^<i>-/-</i> erythroblast enucleation-related gene transcription, enucleation defects and terminal maturation. Remarkably, FOXO3 ectopic expression increased wild type erythroblast maturation and enucleation suggesting that enhancing FOXO3 activity may improve RBCs production. Altogether these studies uncover FOXO3 as a novel regulator of erythroblast enucleation and terminal maturation suggesting FOXO3 modulation might be therapeutic in disorders with defective erythroid maturation.</p></div