The aberrant transcriptional program of myeloid malignancies with poor prognosis
Gerritsen, Myléne
DOI:
10.33612/diss.113503008
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Publication date:
2020
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Citation for published version (APA):
Gerritsen, M. (2020). The aberrant transcriptional program of myeloid malignancies with poor prognosis:
the effects of RUNX1 and TP53 mutations in AML. Rijksuniversiteit Groningen.
https://doi.org/10.33612/diss.113503008
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and block granulocytic differentiation in
human in vitro models and primary AMLs
M. Gerritsen, G. Yi, E Tijchon, J Kuster, J.J. Schuringa, J.H.A. Martens and E. Vellenga
This research was originally published in Blood Advances. © the American Society of Hematology.
ABSTRACT
To unravel molecular mechanisms by which RUNX1 mutations contribute to leukemic transformation, we introduced the RUNX1-S291fs300X mutation in human CD34+ stem/
progenitor cells and in human induced pluripotent stem cells (iPSC). In both models, RUNX1mut
overexpression strongly impaired myeloid commitment. Instead, self-renewal was enhanced as shown by increased long term culture-initiating cell frequencies and enhanced Colony-Forming-Cell replating capacity. Long-term suspension cultures with RUNX1mut-transduced cordblood (CB)
CD34+ cells continued for over 100 days during which the cells displayed an immature GMP-like
CD34+/CD123+/CD45RA+ phenotype. The CD34+/CD38- Hematopoietic Stem Cell (HSC) population
most likely acted as cell of origin since HSCs provided the best long-term proliferative potential upon overexpression of RUNX1mut. CEBPA expression was reduced in RUNX1mut cells, and
re-expression of CEBPA partly restored differentiation. RNA-seq analysis on CB/iPSC systems and on primary patient samples confirmed that RUNX1 mutations induce a myeloid differentiation block, and that a common set of RUNX1mut-upregulated target genes was strongly enriched for GO terms associated with nucleosome assembly and chromatin structure. Interestingly, in comparison to AML1-ETO binding in AMLs we found significantly distinct genomic distribution and differential expression for RUNX1mut of genes such as TCF4, MEIS1 and HMGA2 that may
potentially contribute to the underlying difference in clinical outcomes between RUNX1mut and AML1-ETO patients. In conclusion, RUNX1mut appears to induce a specific transcriptional
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INTRODUCTION
Aberrations in the functionality of transcription factor Runt-related transcription factor 1 (RUNX1) are linked to myeloid malignancies. Mutations in the RUNX1 gene (RUNX1mut) have been identified in myelodysplasia (MDS)1,2,3,4, in de novo and secondary acute myeloid leukemia (AML)5,
and are generally associated with unfavourable clinical outcomes6. Familial RUNX1 mutations
are associated with familial thrombocytopenia and patients have a predisposition to AML development7,8. Besides the point mutations in the RUNX1 gene, a number of fusion partners
of RUNX1 and their deregulating functions have been described in detail, including AML1-ETO9,
while mutations in RUNX1 have hardly been studied.
RUNX1 is essential during multiple stages in the hematopoietic development, including the formation of definitive HSCs5,10, B-cell maturation11, megakaryocyte maturation and granulocytic
differentiation12. RUNX1 mutations, both heterozygous and homozygous, are located along the
full length of the protein. This usually results in single amino-acid substitutions in the DNA-binding domain or in C-terminal truncation mutations lacking all or part of the activation domain 13, 14, 15, 16.
Loss-of-function studies in mice have demonstrated that RUNX1-deficiency is associated with an expansion of the common myeloid progenitors (CMP) and granulocyte-macrophage progenitor (GMP) pool, which could be rescued by inactivation of Hmga217. In addition, RUNX1 loss leads to
an increased susceptibility to AML development in collaboration with MLL-ENL and N-Ras, which is probably because RUNX1 has a tumour-suppressor function18,19. However, recent work has
illustrated the importance of RUNX1wt expression in AML1-ETO and MLL-AF9 cells, suggesting a pro-survival function in leukemogenesis20.
Here, we studied the in vitro growth, the RUNX1 binding pattern and expression profile induced by the C-terminal truncating RUNX1-S291fs300X mutant in cord blood (CB) CD34+ cells and induced
pluripotent stem cells (iPSC). We then compared these findings to those from primary RUNX1mut AMLs. The results indicate that a single RUNX1 mutation leads to increased self-renewal, enhanced long-term-culture initiating-cell (LTC-IC) frequency and long-term maintenance of CD34+ cells. ChIP-seq and RNA-seq experiments, including primary AML samples, indicate that
RUNX1 mutations – independent of the model – induce a unique transcriptional program. By comparing the effects of RUNX1mut with the programs triggered by AML1-ETO, we found that RUNX1mut targets several other genes that possibly affect the clinical outcomes of these two types of leukemia.
MATERIALS & METHODS
Human cells
Umbilical cord blood (CB) was derived after healthy full-term pregnancies from the obstetrics departments of the Martini Hospital and University Medical Centre in Groningen, the Netherlands. Normal bone marrow cells were derived from healthy volunteers or extracted from healthy hipbone following hip-replacement surgery after informed consent. Peripheral blood stem cells (PBSCs) were extracted after stem cells mobilization from patients after informed consent. AML blasts from peripheral blood or bone marrow from de novo AML patients were studied after informed consent was obtained in accordance with the Declaration of Helsinki. The protocol was approved by the Ethical Review Board of the University Medical Center Groningen, Groningen, The Netherlands. To obtain relative pure cell populations and the largest fraction of leukemic cells cell sorting (FACS) based on expression of cell surface markers CD33 or CD34 was performed. An overview of all mutations can be found in Supplementary table 2 and more information about these AMLs is described in Yi et al. (submitted). The megakaryoblast-derived iPSCs were obtained from Sanquin Amsterdam, the Netherlands21. Full culture methods and differentiation of
RUNX1mut iPSCs towards the granulocytic lineage are described in the Supplementary Methods.
Flow cytometry
Surface marker analysis was performed on the LSR-II (BD Biosciences). A list of antibodies can be found in the Supplementary Methods.
Immunoblotting
Whole-cell extracts were prepared by boiling an appropriate number of cells in Laemmli sample buffer for 5 minutes with subsequent separation on a 10% sodium dodecyl sulphate (SDS)– acrylamide gels. Proteins were transferred to PVDF (Millipore) using wet-transfer. RUNX1 was detected with α-RUNX1 antibody (abcam 23980) 1:2000 and -Actin (abcam 16039).
CEBPA rescue experiments
Week 8-10 CB RUNX1mut cells were transduced with retroviral CEBPA-ER vector (described
previously22). Transduced cells were sorted and plated in growth medium with and without 100
nM 4-hydroxytamoxifen. After 3 days, expression of cell surface markers and RNA expression was determined.
RNA extraction and cDNA synthesis
RNA was extracted from 1106 iPSC control and RUNX1mut cells using the RNeasy mini kit (Qiagen)
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minutes in fragmentation buffer (200 mM Tris-acetate, 500 mM Potassium Acetate, 150 mM Magnesium Acetate (pH 8.2)). First strand cDNA synthesis was performed using superscript III (Life Technologies), followed by synthesis of the second cDNA strand.
Chromatin Immunoprecipitation
Cells were crosslinked with 1% formaldehyde for 10 minutes at room temperature at a concentration of 15106 cells/ml. The fixed cells were sonicated for 5 minutes (30 sec on; 30 sec off) using the
Diagenode Bioruptor. Sonicated chromatin was centrifuged for 10 minutes at maximum speed and then incubated with 5 µg RUNX1 antibody (Abcam ab23980) that recognizes both RUNX1wt
and RUNX1mut proteins, or 2 µg flag M2 antibody that recognizes the overexpressed RUNX1mut
protein (Sigma F3165). Beads were washed with four different wash buffers and chromatin was eluted from the beads. DNA-proteins were de-crosslinked and samples were purified using the Qiaquick MinElute PCR purification kit. Library preparation, Illumina high-throughput sequencing and ChIP-seq and RNA-seq data analysis is described in detail in the supplementary materials.
Statistical analysis
A paired or unpaired two-sided Student’s t-test was used to calculate statistical differences. A p-value of <0.05 was considered statistically significant. Error bars represent standard deviation: *
p < 0.05; ** p < 0.01; *** p < 0.001.
RESULTS
RUNX1
mutcells display impaired erythroid differentiation but increased myeloid
replating capacity and expansion.
Cordblood (CB) CD34+ cells were transduced with the RUNX1 mutant S291fs300X (RUNX1mut,
Figure 1A) and overexpression was confirmed by RT-q-PCR (Figure S1A). The progenitor frequency was determined by colony-forming cell (CFC) assay. A decline in BFU-E formation was observed without a difference in CFU-G or CFU-M colony formation (Figure 1B). Besides a reduction in BFU-E numbers, the colony morphology reflected an immature appearance (data not shown), which was linked to a reduced expression of GATA1, an important regulator of erythroid differentiation (Figure S1B). In addition, a strong reduction in expansion was observed when CB CD34+ RUNX1mut cells were cultured under erythroid-permissive conditions (Figure S1C). In
contrast, the CFC replating capacity of myeloid progenitors was enhanced upon overexpression of RUNX1mut (Figure 1B). This was especially the case in the more primitive CD14-/CD15- cell
Figure 1. RUNX1mut increases replating capacity. A. Schematic overview of RUNX1wt and RUNX1 S291fs300X mutant. B. CFC and replating potential of RUNX1mut vs control. At start, 500 cells were plated per dish in duplicates. Between day 10-14 the number of colonies were scored and all colonies were harvested. 25,000 cells per dish were plated for each replating.
C. Replating potential of control vs RUNX1mut in the CD14- / CD15- fraction and (D) replating potential of control vs RUNX1mut in the CD14+ / CD15+ fraction. At start, 500 cells were plated per dish in duplicates. Between day 10-14 the number of colonies were scored, colonies were harvested and stained for CD14 and CD15. 25000 CD14+ or CD15+ cells and 25000 CD14-/CD15 -cells were FACS sorted in a new CFC. After 10-14 days this was repeated for another replating round.
To determine whether RUNX1mut transduced cells could also be propagated for a longer period
in the context of a bone marrow microenvironment, CB CD34+ RUNX1mut cells were cultured on
MS5 bone marrow stroma (Figure 2A, S2A). No significant growth advantage was observed for the RUNX1mut cells compared to control CB CD34+ cells. However, RUNX1mut transduced cells were
maintained under the stroma, without significant expansion, for up to 10-12 weeks. FACS analysis of these stroma-adherent RUNX1mut cells after 10 weeks of culture revealed that a high proportion
of the cells, in contrast to the control cells (Figure S2B), had a persistent immature phenotype, as shown by expression of CD34+/CD38- and high expression of CD123 and CD45RA (Figure 2B).
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Figure 2. RUNX1mut transduced CB CD34+ cells have increased growth potential, LTC-IC-frequency and CD34+ maintenance. A. Relative expansion in MS5 co-culture of control vs RUNX1mut. Growth curve of MS5 co-culture (5 weeks). After 5 weeks the MS5 was replated (see Figure S2A for continued growth). B. Marker expression analysis of adherent fraction of RUNX1mut cells in MS5 co-culture after 10 weeks. Expansion of suspension fraction was very limited and did not allow for surface marker expression after 10 weeks. C. Relative expansion in liquid culture over a course of several weeks in control vs RUNX1mut (n=3). D. Marker expression analysis of RUNX1mut cells in liquid culture after 10 weeks. E. CD34+ expression of 3 CB control vs RUNX1mut cells in time. Each line represents a different culture. F. LTC-IC frequency of control vs RUNX1mut cells.
In addition, long-term myeloid suspension cultures were initiated to study the growth potential of CB CD34+ RUNX1mut cells over a longer period without a bone marrow microenvironment
(Figure 2C). Control transduced cells fully differentiated to macrophages after 8 weeks of culture, whereas the RUNX1mut CB cell culture had a persistent population of more immature monoblastic
cells that expanded for over 3 months (Figure S2C). Further characterization of RUNX1mut cells
by FACS analysis at week 10 identified a mixture of CD34+ and CD34- cells (Figure 2D). CD34
-cells were CD14+/CD15- (Figure S2D) whereas the CD34+ cell population consisted of a CD38+
and a large CD38- sub-fraction suggesting a HSC phenotype. The CD34+CD38+ fraction strongly
expressed both CD123 and CD45RA, suggesting an accumulation at the GMP stage (Figure 2D). Besides these phenotypic markers, at week 10 RUNX1mut cells also showed increased expression
of several markers that have been described on leukemic stem cells such as CD116, IL1RAP and CD135 (Figure S2E)23. We observed an initial loss of the CD34+, which re-appeared after 6-7 weeks
CD38- in our MS5 co-culture, we hypothesized that RUNX1mut had induced distinct phenotypes in
the immature stem cell fraction (CD34+CD38-) relative to the mature (CD34+/CD38+) progenitor
cell fraction. To test this hypothesis, CB cells were transduced with RUNX1mut and sorted for
CD34+CD38- and CD34+CD38+ cells, and then expanded in suspension cultures over time (Figure
S2F). The immature cell population with blast cell morphology originated mostly in the CD34+/
CD38- fraction whereas the CD34+/CD38+ fraction had a higher amount of fully differentiated
macrophages (Figure S2G), suggesting that the HSC population acts as the cell of origin. This is in line with the findings of an increased stem cell-frequency of CB CD34+ RUNX1mut cells after 4
weeks of stromal co-cultures (Figure 2F). To exclude the possibility that the long-term expansion was limited to foetal-CB-derived CD34+ cells, adult bone marrow (n=2, not shown) and adult
peripheral blood (n=2) CD34+ cells were transduced with the RUNX1mut construct (Figure S2H).
In all the experiments, long-term expansion for 10 weeks was obtained and provided a similar phenotype. These findings indicate that expression of the RUNX1mut impairs myeloid and erythroid
maturation, results in a differentiation block at the GMP stage and enhances CFC replating and stem-cell frequencies.
The RUNX1
mutphenotype is characterized by low CEBPA expression and can be
partially rescued by re-expression of CEBPA.
We hypothesized that the differentiation block could be abolished by culturing RUNX1mut cells
in granulocytic differentiation medium containing G-CSF. Interestingly, G-CSF did not induce granulocytic differentiation, in contrast to control cells (Figure S3A). FACS analyses showed that the RUNX1mut cells lost the expression of the G-CSF receptor (CD114) over time (Figure 3A). We
therefore analysed the expression levels of several known RUNX1 target genes important for granulocytic differentiation, such as CEBPA and SPI1. This analysis showed a two-fold decline in
CEBPA expression, a key regulator of granulocytic differentiation (Figure 3B). With the publicly
available TCGA dataset we identified 16 AMLs with a RUNX1 mutation, and confirmed that a reduced expression of CEBPA is a common feature of RUNX1mut-expressing cells in comparison to RUNX1wt (Supplementary Table 1, Figure 3C)24. To determine whether CEBPA downregulation
was the main cause of the observed GMP differentiation block, we overexpressed a CEBPA-ER fusion construct in week 10 RUNX1mut CD34+ cells. Stimulation with 4OH-tamoxifen resulted in a
decline in expansion, increased expression of CD15 and CD14 and decreased expression of CD34 (Figure 3D). Morphological studies revealed a rapid decline in monoblastic-like cells after 3 days of culture (Figure 3E), but no granulocytic cells were observed. RT-q-PCR and FACS revealed only a limited increase of CD114 mRNA levels and cell surface expression (Figure S3B, S3C), while
CTSG and MPO, 2 important genes involved in granulocytic differentiation were not altered,
suggesting that CEBPA is an important downstream target of RUNX1mut, but also other targets
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Figure 3. CEBPA expression is decreased in RUNX1mut cells. A. CD114 expression of CB control vs RUNX1mut cells in time in cells cultured with G-CSF. B. RT-q-PCR analysis of CEBPA and SPI1 expression in control vs RUNX1mut after ~2 weeks of expansion in liquid culture medium. C. Relative CEBPA expression in RUNX1wt vs RUNX1mut AMLs in the TCGA dataset. AML1-ETO or inv(16) AMLs are excluded from this analysis. D. Immature marker (CD34) and immature markers (CD14 and CD15) expression before and after 3 days of overexpression of CEBPA-ER induced by Tamoxifen. E. Percentage of mature and immature appearing cells in RUNX1mut and RUNX1mut + CEBPA cells after 3 days of overexpression of CEBPA. 200 cells were score a rated mature or immature based on their morphology.
RUNX1
mutexpression in differentiating iPSC model results in a differentiation
block of the myeloid lineage by direct regulation of RUNX1
muttarget genes
As second method to model the effect of the RUNX1mut without interference from additional
mutations, we used induced pluripotent stem cells (iPSCs). In these cells, expression of the RUNX1mut was induced during hematopoietic development in vitro (Figure 4A). The TET on/off
promotor enables effective RUNX1mut expression due to the addition of doxycycline (Figure S4A
and S4B). Expression of the RUNX1mut was induced directly after mesoderm differentiation (day
0) or at various stages during hematopoietic development (day 6, 10, 14). Phenotyping the cells over time revealed a retained expression of CD34 upon RUNX1mut expression (Figure 4B and S4C)
and a decreased number of differentiated (CD15+) cells in RUNX1mut-expressing cells as compared
to non-induced cells (Figure 4C). Induction of RUNX1mut at early time points (day 0) resulted in a
short delaying effect on HSC formation, but otherwise no abnormal differentiation was observed. In combination with the monoblastic appearance of these cells (Figure S4D), this suggests a
block in differentiation, similar to that observed for the CB data. These data were corroborated by RNA-seq analysis of control versus dox induced iPSCs, which revealed, similar as for the CB model, reduced levels of CEBPA in the RUNX1mut expressing cells (Figure 4D).
Figure 4. iPSC differentiation is impaired when expressing RUNX1mut. A. Time schedule of iPSC differentiation protocol and marker expression analysis. At the beginning of the haematological differentiation (blue arrow) doxycycline is added, inducing expression of RUNX1mut. Cell surface marker expression on CD34 / CD45 was performed on different time points (red arrows). B. Example of marker analysis and a representable figure of CD34+/CD45+ and CD34-/CD45+ expression on different time points during differentiation. C. Marker analysis and a representable figure of CD15 and CD16 expression in control vs
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Identification of a RUNX1
mutspecific gene program
To determine the relevance of the RUNX1 target genes in primary AML cells, we then compared transcriptional differences initiated by RUNX1mut by RNA-seq in CB-RUNX1mut cells, iPSC-RUNX1mut
cells, normal bone marrow CD34+ (GSE63569)25, monocytes, macrophages and granulocytes,
as well as to patient AML cells harbouring RUNX1mut (Figure S5A). PCA analysis based on
expression revealed clustering of specific cell types and CD34+ cells: most of the undifferentiated
cell populations, including the iPSC and CB models, clustered on one side of the PCA and the differentiated cells on the opposite side (Figure 5A). To map the compositional difference of
RUNX1mut expressing cells systematically, we used the CIBERSORT deconvolution method26. This
showed that a large proportion of the cell population was not only carrying a CD34+ progenitor
signature, but also a substantial monocytic signature, while a granulocytic signature was nearly absent (Figure 5B). We also found that genes normally expressed in granulocytes showed decreased expression levels in RUNX1mut-expressing models and primary AMLs, whereas the
expression levels of monocytic-related genes were comparable (Figure 5C and Figures S5B, S5C). This indicates that RUNX1mut cells maintain a progenitor phenotype featured by monocytic
characteristics, but only granulocytic genes were dramatically repressed by aberrant RUNX1 protein, suggesting that RUNX1mut predominantly affects granulocytic differentiation.
Using k-means clustering, we visualized differentially expressed genes between RUNX1mut
expressing cells and normal CD34+ HSPCs (Figure 5D). This revealed 4 clusters, of which cluster
4 represents 556 genes that are downregulated in all RUNX1mut-expressing groups compared
to normal CD34+ cells. This cluster comprises enrichment for genes involved in inflammation
and immune response, suggesting an effect on myeloid cell maturation (Figure 5E). Cluster 3 represents a set of 542 genes commonly upregulated in all RUNX1mut expressing cells, but not
in CD34+ cells. This RUNX1mut-specific gene cluster is enriched for genes related to chromatin
organization, including several small nucleolar RNAs (snoRNAs), thus suggesting a role in RNA processing.
This transcriptome-wide exploration indicates that in vitro RUNX1mut models can recapitulate
the altered transcriptional landscape induced by RUNX1mut in patient AML cells. Moreover, it
identified the RUNX1mut specific program in the primary AMLs, suggesting that the additional transcriptional alterations in these AML cells (for example in cluster 1 and 2) might be due to other genetic lesions.
Figure 5. The RUNX1mut gene program. A. PCA plot using RNA-seq results of the indicated cell types. Hollow circle indicates RNA-seq data from published primary AMLs carrying RUNX1mut (https://leucegene.ca/research-development/). Normal cells are indicated within the blue circles. B. Based on RNA-seq predicted average fraction of CD34+ cells, granulocytes, monocytes and macrophages in RUNX1mut expressing cells using CIBERSORT. C. Differential expression of monocytic and granulocytic signature genes in normal CD34 cells, RUNX1mut expressing CB cells, iPSC models and primary AML cells. D. Heatmap of gene expression by k-means clustering in RUNX1mut expressing CB, iPSC and primary AML cells vs control CD34+ cells resulting in 4 main clusters. E. Biological process enrichment for each of the 4 clusters identified in (D).
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Global RUNX1
mutbinding in CB and iPSC models binding in primary AMLs with
RUNX1 mutations
To define binding sites of RUNX1 and compare these within our CB and iPSC models and primary AMLs, we performed ChIP and ChIP-seq analysis using an antibody recognizing both RUNX1wt
and RUNX1mut (Figure 6A, S6A). This revealed binding of RUNX1 in RUNX1mut-transduced CB
cells (CB-RUNX1mut) and in RUNX1mut-expressing iPSC (iPSC-RUNX1mut) at similar sites as those in
primary RUNX1mut AMLs (Figure 6A). We found similar RUNX1 occupancy profiles in control iPSCs,
although with lower density (Figure 6A-B). We identified 895 high-confidence RUNX1 binding sites shared between CB-RUNX1mut, iPSC-RUNX1mut and primary AMLs with a RUNX1 mutation
(Figure S6B). Functional analysis of the genes associated with overlapping RUNX1 binding sites revealed enrichment for processes associated with transcription and translation (Figure S6C). As RUNX1 binding has been associated with regulating the local epigenetic environment, we investigated the epigenetic state of the common RUNX1 binding sites by examining epigenetic data of RUNX1mut AMLs (Yi et al., submitted). In line with the observation that most of the common
RUNX1mut binding sites are in promoters or near genes (Figure S6D, S6E), our analysis revealed
enrichment for active chromatin marks such as DNaseI accessibility, H3K4me3 and H3K27ac (Figure S6F) and association of RUNX1 binding with genes that are actively transcribed (Figure S6G). This is in accordance with the reduced levels of repressive marks such as H3K27me3.
To determine whether this state is different in RUNX1wt AML cells, we then compared the epigenetic
state at RUNX1 binding sites common in RUNX1wt and RUNX1mut AMLs and the transcriptional
state of their target genes (Figure S6H). This showed no global differences between RUNX1wt
and RUNX1mut AMLs in the presence of active or repressive marks at the common RUNX1 binding
sites (Figure S6I), suggesting the absence of a common epigenetic deregulating mechanism by RUNX1mut. Interestingly, target gene expression still varied significantly between RUNX1wt and
RUNX1mut AMLs, with approximately half of the target genes up- and half of the target genes
downregulated in RUNX1mut AML cells (Figure 6C, middle). These alterations in gene activity at
the local level did correlate with epigenetic alterations, as exemplified for NUCB2 (up in RUNX1wt)
and TAL1 (up in RUNX1mut) (Figure 6C, left and right).
For a more general view, we zoomed into the genes that have differential expression upon RUNX1mut presence (Figure 5D, clusters 3 and 4), and found that the levels of active chromatin
marks (mainly H3K27ac and H3K4me3) were higher in genes that are upregulated compared to the genes that are downregulated (Figure S6J). Levels of H3K27me3 were low overall and were not altered. For example, for CTSG we found decreased expression coinciding with less H3K27ac (Figure 6D), while this was not observed for genes that are not differentially expressed (Figure S6K). Expression of CEBPA was also decreased, but not significantly, possibly due to a limited number of studied AMLs. However, we did see a decrease in H3K27ac (Figure 6E), supporting our cell biological data.
Figure 6. RUNX1 binding in model cells and primary AMLs. A. Heatmap of RUNX1mut binding sites in RUNX1mut transduced CB cells, control and RUNX1mut induced iPSCs and primary AMLs carrying a RUNX1mut. B. Screenshot of RUNX1 binding in CB-RUNX1mut and control cells, iPSC-RUNX1mut cells with and without dox, and primary AMLs expressing RUNX1mut. C. Differential expression of genes that are bound by RUNX1 in both RUNX1wt and RUNX1mut AMLs. NUCB2 (left) is higher expressed in
RUNX1wt AMLs whereas TAL1 (right) is expressed more in RUNX1mut AMLs. Differential levels of H3K27ac are associated with the levels of expression. D. Expression levels and levels of DNaseI, H3K27ac, and H3K27me3 at the CTSG locus in RUNX1wt vs
RUNX1mut AMLs. E. Expression levels and levels of DNaseI, H3K27ac, and H3K27me3 at the CEBPA locus in RUNX1wt vs RUNX1mut AMLs.
These results suggest that RUNX1mut does not have a global effect on the epigenetic state at
its binding sites but has specific effects on genes important for granulocytic and monocytic differentiation.
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AML1-ETO peaks in our primary AMLs and found partial overlap for RUNX1mut and AML1-ETO
(Figure S7A). Subsequently, we included recently published data on AML1-EVI1 binding28. When
comparing peak distributions from these datasets, we found that RUNX1mut peaks are mostly
located in promoters whereas AML1-ETO and AML1-EVI1 are located intergenic, suggesting distinct occupancy and regulatory roles due to the three types of mutations (Figure 7A). Also, analysis of the genes in the direct vicinity (+5kb, -1kb; Figure 7B) identified only a small number of direct AML1-ETO and AML1-EVI1 target genes. In this study we showed that in our iPSC model RUNX1mut, similar as shown for AML1-ETO (Tijchon et al.in preparation, Mandoli et al.27,
Figure S7B), is involved in repressing granulocytic differentiation. However, when comparing specific gene expression changes, we observed that key granulocytic genes like MPO and CTSG are more deregulated in the RUNX1mut expressing cells (Figure 7C, S7C). Upregulated genes in RUNX1mut iPSCs – but not in AML1-ETO iPSCs – include MEIS1, TCF4, ERG and HMGA2, genes
previously described as being upregulated in leukemogenesis (Figure S7C, Supplementary table 3,). Interestingly, comparing RUNX1 binding differences in RUNX1mut and AML1-ETO primary
AMLs, showed decreased binding of RUNX1 at many of these genes, including TCF4, HMGA2 and
MN1, in AML1-ETO AMLs (Figure 7D). In addition, we found that RUNX1mut AMLs have decreased
expression of granulocytic genes like CEBPE and ELANE (Figure 7E, Supplementary table 4). Upregulated genes in RUNX1mut AMLs include genes important for signal transduction and
hematopoiesis including again TCF4, MN1 and MEIS1 (Figure 7E and 7F).
Finally, we were interested if we could rescue the overexpression of these genes by CEBPA re-expression as we had conducted in our CB models. These results show that MEIS1, HMGA2 and
CD34 were downregulated upon overexpression of CEBPA ( Figure S7D), suggesting that CEBPA
mediated differentiation might results in a decreased stem cell phenotype.
In conclusion, our results suggest that RUNX1mut occupies and regulates different loci compared to
AML1-ETO and triggers different gene expression patterns at a number of critical genes relevant for granulocytic differentiation.
Figure 7. RUNX1mut vs. AML1-ETO AMLs. A. Genomic distribution of RUNX1mut, AML1-ETO and AML1-EVI1 binding sites. All peaks are assigned to promoters (2 kb away from TSS), exons, introns and intergenic regions based on the physical distance. The priority rule of category assignment: promoter exon intron intergenic region. B. Gene overlap of RUNX1mut, AML1-ETO and AML1-EVI1 binding sites selected by +5kb, -1kb distance. C. Differential expression in RUNX1mut and AML1-ETO iPSC. Several significant genes regulated by RUNX1 or AML1-ETO are highlighted. D. Differential binding of RUNX1 in RUNX1mut and AML1-ETO primary AML samples. E. Differential expression between four RUNX1mut AMLs and three AML1-ETO AMLs. F. Biological processes enriched for genes upregulated in RUNX1mut AMLs compared to AML1-ETO AMLs.
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DISCUSSION
To unravel the mechanisms by which RUNX1 mutations contribute to leukemic transformation we investigated the cell biological and molecular consequences of a RUNX1 mutation as single genetic hit in healthy CB-derived CD34+ cells and in human iPSCs. The resulting data were compared to
that from RUNX1-mutated and wt-primary AML patient cells, and from patient samples expressing the AML1-ETO fusion protein, which display rather different molecular and clinical characteristics. This comparison showed that the RUNX1 truncation mutant S291fs300X induces a differentiation block when introduced into CB CD34+ cells or iPSCs. This results in the emergence of GMP-like
cells, which also express IL1-RAP, with increased self-renewal potential in vitro.
The differentiation block at the GMP stage is linked to a reduced expression of CEBPA in the myeloid lineage, which has been described as a main deregulated gene in AML1-ETO29. In our
study, re-expression of CEBPA partially rescued the differentiation block, which is also in line with findings in AML1-ETO AMLs30. RNA-seq analysis showed transcriptional differences regarding the
presence of the RUNX1mut in human in vitro models and primary AMLs when compared to normal
bone marrow CD34+ cells, thereby identifying several clusters. PCA analysis of our RNA-seq data
revealed an enrichment of stem cell genes and a loss of genes for granulocytic differentiation, reflecting the differentiation block induced by the RUNX1mut. This suggests that RUNX1mut
dictates a distinct transcriptional program in favour of transformation.
We analysed model systems that have only one RUNX1 mutation, thus excluding all secondary effects caused by additional mutations, and identified genes directly deregulated by RUNX1mut.
Our approach therefore has little overlap with a recent study that compared RUNX1mut AMLs with RUNX1wt AMLs 31. This is in line with the findings that RUNX1mut requires additional mutations
frequently found together with RUNX1mut, including ASXL132 or elevated BMI1 expression33, that
epigenetically influence the expression of another set of target genes.
By using ChIP-seq, we linked RUNX1mut binding to differences in histone modifications, revealing
that RUNX1mut did not completely reshape the localization of epigenetic marks. The analysis showed that RUNX1mut can both induce and repress target genes, and that this is associated with
the amount of activating histone modifications, mainly H3K27ac, suggesting a more gene-specific regulation of transcription. This differs from the activity of AML1-ETO, which has been shown to bind mainly to introns and intergenic regions that are enriched in accessible chromatin, marked with p300, but are generally low in acetylation34. Also, upon direct gene binding AML1-ETO leads
to gene repression and thus has a dominant negative effect relative to native AML1 by recruiting HDACs, NCoR and mSin3A35–38.
Transcriptional differences between RUNX1mut and AML1-ETO in our iPSC models and primary
AML cells include HMGA217, TCF439 and MEIS140. Previous studies have shown that the expression
of these genes is increased in AMLs, which makes them potentially important targets of RUNX1mut,
but not for AML1-ETO in leukemogenesis.
In summary, the results of cellular assays, transcriptomic and epigenomic profiling of human CB and iPSC models expressing RUNX1 mutations demonstrate that RUNX1 binding and RNA expression recapitulate the effects of RUNX1-mutated primary AMLs. This indicates a change in the RUNX1-dependent cellular programming that is independent of additional mutations. This programming differs between RUNX1mut and AML1-ETO-positive AMLs. This could potentially
explain the different clinical outcomes of AML patients harbouring an RUNX1 mutation (poor risk) and those harbouring an AML1-ETO (good risk) translocation.
Acknowledgements
2
REFERENCES
1. Harada, Y. & Harada, H. Molecular pathways mediating MDS/AML with focus on AML1/RUNX1 point mutations. J. Cell. Physiol. 220, 16–20 (2009).
2. Matsushita, H. et al. Development of human and murine myelodysplastic syndromes RUNX1 / AML1 mutant collaborates with BMI1 overexpression in the development of human and murine myelodysplastic syndromes. Blood 121, 3434–3446 (2013).
3. Haferlach, T. et al. Landscape of genetic lesions in 944 patients with myelodysplastic syndromes. Leukemia 28, 241–247 (2013). 4. Galili, N., Nilsson, B., Garcia-manero, G., Neuberg, D. & Ebert, B. L. Clinical Effect of Point Mutations in Myelodysplastic Syndromes. N Engl J
Med. 364:2496-2506 (2011).
5. Ichikawa, M. et al. A role for RUNX1 in hematopoiesis and myeloid leukemia. Int. J. Hematol. 97, 726–734 (2013).
6. Haferlach, T. et al. The new provisional WHO entity ‘ RUNX1 mutated AML ’ shows speci fi c genetics but no prognostic in fl uence of dysplasia. Leukemia 30, 2074–2125 (2016).
7. Song, W. et al. Haploinsufficiency of CBFA2 causes familial thrombocytopenia with propensity to develop acute myelogenous leukaemia.
Nature Genetics 23, 166-75 (1999).
8. Liew, E. & Owen, C. Familial myelodysplastic syndromes: A review of the literature. Haematologica 96, 1536–1542 (2011).
9. Gaidzik, V. I. et al. RUNX1 mutations in acute myeloid leukemia: Results from a comprehensive genetic and clinical analysis from the AML study group. J. Clin. Oncol. 29, 1364–1372 (2011).
10. Tober, J., Yzaguirre, A. D., Piwarzyk, E. & Speck, N. a. Distinct temporal requirements for Runx1 in hematopoietic progenitors and stem cells.
Development 140, 3765–76 (2013).
11. Dowdy, C. R. et al. A germline point mutation in Runx1 uncouples its role in definitive hematopoiesis from differentiation. Exp. Hematol. 41, 980–991 (2014).
12. Guo, H., Ma, O., Speck, N. A. & Friedman, A. D. Runx1 deletion or dominant inhibition reduces Cebpa transactivation via conserved promoter and distal enhancer sites to favor monopoiesis over granulopoiesis. Blood 119, 4408–4418 (2012).
13. Osato, M. et al. Biallelic and heterozygous point mutations in the runt domain of the AML1/PEBP2alphaB gene associated with myeloblastic leukemias. Blood 93, 1817–1824 (1999).
14. Osato, M. Point mutations in the RUNX1/AML1 gene: another actor in RUNX leukemia. Oncogene 23, 4284–4296 (2004).
15. Preudhomme, C. et al. High incidence of biallelic point mutations in the Runt domain of the AML1/PEBP2 alpha B gene in Mo acute myeloid leukemia and in myeloid malignancies with acquired trisomy 21. Blood 96, 2862–2869 (2000).
16. Tang, J. et al. AML1 / RUNX1 mutations in 470 adult patients with de novo acute myeloid leukemia : prognostic implication and interaction with other gene alterations. Blood 114, 5352–5361 (2009).
17. Lam, K. et al. Hmga2 is a direct target gene of RUNX1 and regulates expansion of myeloid progenitors in mice. Blood 124, 2203–2212 (2014).
18. Nishimoto, N. et al. Loss of AML1/Runx1 accelerates the development of MLL-ENL leukemia through down-regulation of p19ARF. Blood
118, 2541–2550 (2011).
19. Motoda, L. et al. Runx1 Protects Hematopoietic Stem/Progenitor Cells from Oncogenic Insult. Stem Cells 25, 2976–2986 (2007). 20. Goyama, S. & Schibler, J. Transcription factor RUNX1 promotes survival of acute myeloid leukemia cells. J. Clin. Invest. 123, 3876–3888
(2013).
21. Hansen, M. et al. Generation and characterization of human iPSC line MML-6838-Cl2 from mobilized peripheral blood derived megakaryoblasts. Stem Cell Res. 18, 26–28 (2017).
22. Wierenga, A. T. J., Schepers, H., Moore, M. A. S., Vellenga, E. & Schuringa, J. J. STAT5-induced self-renewal and impaired myelopoiesis of human hematopoietic stem / progenitor cells involves down-modulation of C / EBPa. Blood 107, 4326–4333 (2006).
23. Bonardi, F. et al. A Proteomics and Transcriptomics Approach to Identify Leukemic Stem Cell ( LSC ) Markers . Mol Cell Proteomics 12, 626–637 (2013).
24. Network, T. C. G. A. Genomic and Epigenomic Landscapes of Adult De Novo Acute Myeloid Leukemia. N. Engl. J. Med. 368, 2059–74 (2013). 25. Dolatshad, H. et al. Disruption of SF3B1 results in deregulated expression and splicing of key genes and pathways in myelodysplastic
syndrome hematopoietic stem and progenitor cells. Leukemia 29, 1092–1103 (2015).
26. Newman, A. M. et al. Robust enumeration of cell subsets from tissue expression profiles. Nat. Methods 12, 453–457 (2015).
27. Mandoli, A. et al. The Hematopoietic Transcription Factors RUNX1 and ERG Prevent AML1-ETO Oncogene Overexpression and Onset of the Apoptosis Program in t(8;21) AMLs. Cell Rep. 17, 2087–2100 (2016).
28. Loke, J., Assi, S. A., Rosaria, M., Cockerill, P. N. & Reprogram, R. D. RUNX1-ETO and RUNX1-EVI1 Differentially Reprogram the Chromatin Landscape in t(8;21) and t(3;21) AML. Cell Reports 19, 1654–1668 (2017).
29. Pabst, T. et al. AML1 – ETO downregulates the granulocytic differentiation factor C / EBP α in t ( 8 ; 21 ) myeloid leukemia. Nat. Med. 7, (2001). 30. Loke, J. et al. C/EBP a overrides epigenetic reprogramming by oncogenic transcription factors in acute myeloid leukemia SET. Blood
Advances 2, 271–284 (2018).
31. Simon, L. et al. Chemogenomic landscape of RUNX1-mutated AML reveals importance of RUNX1 allele dosage in genetics and glucocorticoid sensitivity. Clin. Cancer Res. 23, 6969–6981 (2017).
32. Nagase, R. et al. Expression of mutant Asxl1 perturbs hematopoiesis and promotes susceptibility to leukemic transformation. J. Exp. Med.
33. Matsushita, H. et al. development of human and murine myelodysplastic syndromes RUNX1 / AML1 mutant collaborates with BMI1 overexpression in the development of human and murine myelodysplastic syndromes. Blood 121, 3434–3446 (2013).
34. Saeed, S. et al. Chromatin accessibility, p300, and histone acetylation define PML-RARalpha and AML1-ETO binding sites in acute myeloid leukemia. Blood 120, 3058–3068 (2012).
35. Gelmetti, V. et al. Aberrant Recruitment of the Nuclear Receptor Corepressor-Histone Deacetylase Complex by the Acute Myeloid Leukemia Fusion Partner ETO. Mol. Cell. Biol. 18, 7185–7191 (1998).
36. Wang, J., Hoshino, T., Redner, R. L., Kajigaya, S. & Liu, J. M. ETO, fusion partner in t(8;21) acute myeloid leukemia, represses transcription by interaction with the human N-CoR/mSin3/HDAC1 complex. Proc. Natl. Acad. Sci. U. S. A. 95, 10860–5 (1998).
37. Lutterbach, B. et al. ETO, a target of t(8;21) in acute leukemia, interacts with the N-CoR and mSin3 corepressors. Mol. Cell. Biol. 18, 7176–84 (1998).
38. Hildebrand, D., Tiefenbach, J., Heinzel, T., Grez, M. & Maurer, A. B. Multiple Regions of ETO Cooperate in Transcriptional Repression. J. Biol.
Chem. 276, 9889–9895 (2001).
39. in ’t Hout, F. E. M., van der Reijden, B. A., Monteferrario, D., Jansen, J. H. & Huls, G. High expression of transcription factor 4 (TCF4) is an independent adverse prognostic factor in acute myeloid leukemia that could guide treatment decisions. Haematologica 99, 257-9 (2014). 40. Liu, J. et al. Meis1 is critical to the maintenance of human acute myeloid leukemia cells independent of MLL rearrangements. Ann. Hematol.
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SUPPLEMENTARY DATA
Supplementary Methods
Lentiviral transduction
CD34+ cells were isolated from these samples by MACS separation (Miltenyi). Cells were
maintained in HPGM (Lonza) and subsequently transduced with either control constructs or
RUNX1 S291fs300X mutant-containing(Flag-tagged) lentiviral constructs (pRRL-SFFV-iGFP). The
original constructs were kindly provided by T. Kitamura (Institute of Medical Science, University of Tokyo, Tokyo, Japan).
CFC, liquid cultures and MS5 stromal cultures
Colony forming cell (CFC) formation was tested by plating 1000 CD34+ cells in methylcellulose
(H4230, StemCell Technologies, Grenoble, France), supplemented with 20 ng/mL interleukin-3 (IL-3), 20 ng/mL IL-6, 20 ng/mL G-CSF, 20 ng/mL stem cell factor (SCF; c-kit ligand) and 6 U/mL erythropoietin (EPO). Colony forming unit-Granulocyte CFU-G, Colony forming unit-macrophage (CFU-M) and Burst-forming unit-erythroid (BFU-E) colonies were scored after 10-14 days of culture. Liquid cultures were performed by culturing cells in Iscove’s Modified Dulbecco’s Medium (IMDM; Lonza, Leusden, the Netherlands) supplemented with 20% heat-inactivated FCS (Sigma-Aldrich), 20ng/ml IL-3 and 20 ng/mL SCF for myeloid differentiation, 20ng/ml IL-3, 20 ng/mL SCF and 20 ng/mL G-CSF for granulocytic differentiation or 20 ng/mL SCF and 3 U/mL EPO for erythroid differentiation. MS5 stoma co-cultures were performed in α-modified Minimum Essential Media (αMEM; Lonza) supplemented with 12.5% heat-inactivated FCS (Sigma-Aldrich), heat-inactivated 12.5% horse serum (Sigma-Aldrich), 1% penicillin and streptomycin (PenStrep) 57.2 μM β– mercaptoethanol, and 1 μM hydrocortisone and with or without addition of cytokines (20ng/ml IL-3 and 20 ng/mL SCF). LTC-IC was performed by sorting transduced CB CD34+ cells in limiting
dilutions in a range between 1 and 243 cells per well on MS5 stromal cells in a 96-wells plates in Gartners medium supplemented with 20ng/ml IL-3. Methylcellulose was added to the wells after 4 weeks of culture. 2 weeks later, wells containing CFCs were scored as positive.
iPSC Culture
The iPSCs were cultured in E8 medium (Life technologies) supplemented with 1% pen/strep at 37°C. RUNX1S291fs300X mutant iPSCs were generated by knock-in of a AAVS1-RUNX1mut donor vector
and CRISPR-Cas91. Briefly, 2.5 million iPSCs were nucleofected with the AAVS1-RUNX1mut vector
and CRISPR-pCas9 using the P3 primary cell nucleofector kit (Lonza, Leusden, the Netherlands). Transfected iPSCs were plated on fibronectin-coated plates in E8 medium supplemented with 10 µM rock inhibitor for 24 hours. RUNX1mut iPSCs were selected 48 hours after transfection using
0.25 µg/ml puromycin. Positive clones were selected by PCR using primers that could discriminate between single and double integrations of RUNX1mut.
Hematopoietic (Granulocytic) differentiation
Construction of iPSCs has been previously described by Mandoli et al2. Briefly, iPSCs were seeded
as single cells on geltrex with a density of 18 colonies in a 10cm dish in E8 medium supplemented with 10 µM rock inhibitor. Cells were maintained for 14 days in E8 medium untill individual iPSC colonies reached a size of approximately 500μM. At day 0 of differentiation, the E8 medium was replaced by stemline II medium (Sigma Aldrich) supplemented with 1% pen/strep, 1:100 insulin-transferrin-selenium-ethanolamine (ITS-X) (Life technologies) and cytokines (20 ng/ml BMP4, 40 ng/ml VEGF and 50 ng/ml bFGF) (Miltenyi Biotec). After 6 days, the medium and cytokines were replaced by X-VIVO15 medium (Lonza) and a cytokine cocktail specific for granulocytic differentiation (50 ng/ml SCF, 50 ng/ml IL-3, 50 ng/ml G-CSF and 5 ng/ml TPO)3. The RUNX1mut was
induced by addition of 14 ng/ml doxycycline on day 0 of differentiation, and the expression was maintained by keeping the cells continuously in doxycycline. The differentiation medium was refreshed every 3-4 days or when the medium turned yellow.
FACS antibodies
For Flow cytometry we used the following antibodies: CD34-APC (clone 581, BD Biosciences); CD34-BV421 (clone 581, BD Biosciences); CD14-PE (clone HCD14, Biolegend); CD15 Pacific Blue (clone W6D3, Biolegend); CD123-Pe-cy7 (clone 6H6, Biolegend); CD45RA-BV421(clone 5H9, BD Biosciences); CD33-APC (clone WM53, Biolegend); CD11b-PE (clone M1/70, Biolegend); CD114-PE(clone LMM741, BD Biosciences) CD38-PE (clone HB7, Biolegend); IL1-RAP-PE (clone 89412, R&D Systems); CD97-PE (clone VIM3b, Biolegend); CD116-PE (clone 4H1, Biolegend); CD135-PE(clone A2F10, Biolegend).
Cytospins
For morphological analysis, 5104 iPS / CB cells were spinned for 10 min. at 800g on a glass slide
and air-dried for at least one hour at room temperature. Cells were fixed and stained for 5 minutes with May-Grünwald and 15 minutes with Giemsa staining. The slides were washed and mounted using permount.
Library preparation and Illumina high-throughput sequencing
Libraries were generated using the Kapa hyper prep kit. End repair and A-tailing was performed on the double strand DNA using end repair and A-tailing buffer. Subsequently, the adapters were ligated and a post-ligation clean-up was performed using Agencourt AMPure XP reagent. The libraries were amplified by PCR using the Kapa Hifi hotstart readymix and primer mix, 10 cycles. Samples were purified using the Qiaquick MinElute PCR purification kit and 300 bp fragments were selected from E-gel and the size was checked on the agilent bioanalyzer. Samples were sequenced on the Illumina HiSeq 2000.
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ChIP-Seq analyses
For each sample, all 42bp paired-end reads were aligned to the hg19 human genome using Burrows-Wheeler Aligner (BWA) program4 with default parameters. Only uniquely-mapped reads
were retained and then these reads were regarded as input for Picard MarkDuplicates (http:// broadinstitute.github.io/picard/), to remove potential PCR duplicates. Peak calling was performed using MACS25 with the narrow setting at a p-value cut-off of 1.00 10-6. All RUNX1mut alignment
files were normalized to Reads Per Kilobase per Million mapped reads (RPKM) coverage via deepTools6 with an extended fragment length of 200bp for visualization. BEDTools suite7 was used
to manipulate genomic features in BED format. To assess functional roles of RUNX1mut peaks, the GREAT tool (http://great.stanford.edu/public/html/) was conducted using default parameters. All ChIP-seq and RNA-seq data is available via GEO (GSE111821 and GSE111919).
To compare RUNX1 binding landscape with two types of aberrant RUNX1 rearrangement, AML1-ETO and AML1-EVI1, we retrieved their ChIP-seq data from GEO database (GSE23730 and GSE87283). All peak and bigwig files were generated using the same parameters as described previously. We used BEDTools intersect option to identify unique peak set for each type of mutation and displayed binding profiling with deepTools.
RNA-Seq analyses
RNA-Seq reads were mapped to the human genome hg19 using STAR aligner8 which could
enumerate gene-level read counts at the same time. The DESeq2 package9 was used to identify
differentially expressed genes by comparing two selected groups. Only those genes greater than 1.5-fold changed at Benjamini-Hochberg-corrected p-value 0.1 were considered significantly deregulated. Expression levels of RefSeq genes were quantified using Fragments Per Kilobase per Million aligned reads value (FPKM) values calculated by Cufflinks10. We used the CIBERSORT
method to systematically decompose the cellular population difference and cell-type signature genes for RUNX1mut-expressed cells in our two in vitro models and our primary AML samples. Dimensionality reduction and functional annotation
For visualization, the top 3,000 variable peaks or genes were first selected based on interquartile range (IQR) of normalized peak density or gene expression, and were then used to reduce dimensionality (principal component analysis and t-distributed stochastic neighbourhood embedding) of the dataset by the pca function and Rtsne package in R. To group genes showing similar expression patterns, we used the k-means clustering approach. Biological process ontology was assessed by the DAVID tool11 to gain insight into the biological functions for each
gene cluster. Only those functional terms with Benjamini-adjusted p-value 0.05 were considered significantly overrepresented.
Data retrieval
All CB materials, iPSCs (RUNX1mut and AML1-ETO) and Blueprint AML data are obtained using the
same platforms (HiSeq2000 and NextSeq500). AML1-EVI data from C. Bonifer has been performed on comparable platforms (HiSeq2500, NextSeq500). For all samples the same Illumina sample prep methods were used and yielded very comparable results12.
REFERENCES
1. Mali, P. et al. RNA-Guided Human Genome Engineering via Cas9. Science. 339, 823–826 (2013).
2. Mandoli, A. et al. The Hematopoietic Transcription Factors RUNX1 and ERG Prevent AML1-ETO Oncogene Overexpression and Onset of the Apoptosis Program in t(8;21) AMLs. Cell Rep. 17, 2087–2100 (2016).
3. Morishima, T. et al. Neutrophil differentiation from human-induced pluripotent stem cells. J. Cell. Physiol. 226, 1283–1291 (2010). 4. Li, H. & Durbin, R. Fast and accurate short read alignment with Burrows-wheeler Transform. Bioinformatics 25, 1754–1760 (2009). 5. Zhang, Y. et al. Model-based analysis of ChIP-Seq (MACS). Genome Biol. 9, (2008).
6. Ramírez, F. et al. deepTools2: a next generation web server for deep-sequencing data analysis. Nucleic Acids Res. 44, W160–W165 (2016). 7. Quinlan, A. R. & Hall, I. M. BEDTools: A flexible suite of utilities for comparing genomic features. Bioinformatics 26, 841–842 (2010). 8. Dobin, A. et al. STAR: Ultrafast universal RNA-seq aligner. Bioinformatics 29, 15–21 (2013).
9. Love, M. I., Huber, W. & Anders, S. Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2. Genome Biol. 15, 1–21 (2014).
10. Trapnell, C. et al. Transcript assembly and abundance estimation from RNA-Seq reveals thousands of new transcripts and switching among isoforms. Nat. Biotechnol. 28, 511–515 (2011).
11. Huang, D. W., Sherman, B. T. & Lempicki, R. A. Bioinformatics enrichment tools: Paths toward the comprehensive functional analysis of large gene lists. Nucleic Acids Res. 37, 1–13 (2009).
12. Aigrain, L. , Gu, Y., Quail, M.A. Quantitation of next generation sequencing library preparation protocol efficiencies using droplet digital PCR assays - a systematic comparison of DNA library preparation kits for Illumina sequencing. BMC Genomics 17, 458 (2016).
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Supplementary Table 1. Overview of RUNX1 mutations in the TCGA dataset.TCGA # RUNX1 mutation type of mutation Protein
TCGA-AB-2805 nonsense p.R201*
frame_shift_ins p.A142fs
TCGA-AB-2807 in_frame_del p.SG167in_frame_del
TCGA-AB-2821 frame_shift_ins p.D123fs TCGA-AB-2850 frame_shift_del p.G420fs TCGA-AB-2865 missense p.D198N missense p.S141L TCGA-AB-2899 missense p.R162G TCGA-AB-2912 missense p.R162K TCGA-AB-2927 nonsense p.R201* missense p.R162S TCGA-AB-2933 missense p.R162G TCGA-AB-2936 nonsense p.R320* TCGA-AB-2949 frame_shift_del p.S314fs TCGA-AB-2959 missense p.R201Q TCGA-AB-2970 missense p.P113L TCGA-AB-2978 nonsense p.R201*
TCGA-AB-2983 missense p.A149P
Supplementary Table 2. Baseline Characteristics of 40 Samples with Acute Myeloid Leukemia (AML).
ID Karyotype NPM1 FLT3 KIT NRAS JAK2 KRAS CALR CBL DNMT3A TET2 IDH1 IDH2 CEBPA RUNX1 WT1 TP53 ASXL1 EZH2 SRSF2 SF3B1
S00KPBH1 Complex S015UGH1 t(9;11) S00Q7NH1 Complex S00SB7H1 Complex S00Y21H1 Normal mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S013QWH1 Normal mt wt wt wt wt wt wt wt mt wt mt wt wt wt wt wt wt wt wt wt S013PYH1 Normal mt mt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S00D39H1 Normal mt wt wt wt wt wt wt wt mt mt wt wt mt wt mt wt wt wt wt wt S00D2BH1 Normal mt mt wt wt wt wt wt wt mt mt wt wt mt wt wt wt wt wt wt wt S005EJH1 Normal mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S008X6H1 Normal S015TIH1 t(9;11) S013M3H1 t(3;5) wt mt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S00Y6UH1 Complex mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S00XVLH1 Normal mt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S00D1DH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S00D0FH1 +8 wt mt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00CYPH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt mt wt wt wt wt wt S005FHH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S005GFH1 Normal S00XXHH1 Normal wt wt wt wt wt wt wt wt mt mt wt wt mt wt wt wt wt wt wt wt S00CWTH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00CXRH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00XUNH1 +11 wt wt wt wt wt wt wt wt wt wt wt mt wt mt wt wt wt wt mt wt S00XYFH1 Normal wt wt wt wt wt wt wt wt mt wt wt mt wt wt wt wt wt wt mt wt S00D47H1 Normal wt mt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt S00D63H1 Complex wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt mt wt S00FGCH1 Complex S00Y5WH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt mt mt wt wt S013SSH1 t(8;21) wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S013RUH1 inv16 wt wt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S013N1H1 inv16 wt wt mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00Y4YH1 t(8;21) wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S00Y05H1 Complex wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt mt S005CNH1 inv16 S00XWYH1 t(3;5) wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt
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Supplementary Table 2. Baseline Characteristics of 40 Samples with Acute Myeloid Leukemia (AML).ID Karyotype NPM1 FLT3 KIT NRAS JAK2 KRAS CALR CBL DNMT3A TET2 IDH1 IDH2 CEBPA RUNX1 WT1 TP53 ASXL1 EZH2 SRSF2 SF3B1
S00KPBH1 Complex S015UGH1 t(9;11) S00Q7NH1 Complex S00SB7H1 Complex S00Y21H1 Normal mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S013QWH1 Normal mt wt wt wt wt wt wt wt mt wt mt wt wt wt wt wt wt wt wt wt S013PYH1 Normal mt mt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S00D39H1 Normal mt wt wt wt wt wt wt wt mt mt wt wt mt wt mt wt wt wt wt wt S00D2BH1 Normal mt mt wt wt wt wt wt wt mt mt wt wt mt wt wt wt wt wt wt wt S005EJH1 Normal mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S008X6H1 Normal S015TIH1 t(9;11) S013M3H1 t(3;5) wt mt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt S00Y6UH1 Complex mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S00XVLH1 Normal mt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S00D1DH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S00D0FH1 +8 wt mt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00CYPH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt mt wt wt wt wt wt S005FHH1 Normal mt mt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt S005GFH1 Normal S00XXHH1 Normal wt wt wt wt wt wt wt wt mt mt wt wt mt wt wt wt wt wt wt wt S00CWTH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00CXRH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00XUNH1 +11 wt wt wt wt wt wt wt wt wt wt wt mt wt mt wt wt wt wt mt wt S00XYFH1 Normal wt wt wt wt wt wt wt wt mt wt wt mt wt wt wt wt wt wt mt wt S00D47H1 Normal wt mt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt S00D63H1 Complex wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt mt wt S00FGCH1 Complex S00Y5WH1 Normal wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt mt mt wt wt S013SSH1 t(8;21) wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S013RUH1 inv16 wt wt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S013N1H1 inv16 wt wt mt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt S00Y4YH1 t(8;21) wt wt wt mt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt S00Y05H1 Complex wt wt wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt mt S005CNH1 inv16 S00XWYH1 t(3;5) wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt wt wt wt wt S00D55H1 Normal wt wt wt wt wt wt wt wt wt wt wt wt wt mt wt wt wt wt wt wt S00Y13H1 +8 wt mt wt wt wt wt wt wt wt mt wt wt wt mt wt wt mt mt wt wt
Supplementary Table 3. Fold change list of genes deregulated in RUNX1mut iPSCs and AML1-ETO
iPSCs (Figure 7C) can be downloaded from https://ashpublications.org/bloodadvances.
Supplementary Table 4. Fold change list of genes in RUNX1mut AMLs vs AML1-ETO AMLs (Figure
7E) can be downloaded from https://ashpublications.org/bloodadvances.
Supplementary Figure 1. A. qPCR of RUNX1 overexpression in CB control vs RUNX1mut. B. GATA-1 expression levels in CB CD34+ transduced with control vector vs RUNX1mut. C. Expansion of control vs RUNX1mut expressing CB CD34+ cells under erythroid conditions.
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Supplementary Figure 2. A. Expansion of control vs RUNX1mut cells on MS5 after replating. B. May-Grünwald Giemsa stain of RUNX1mut cells after 10 weeks of liquid culture. C. Marker expression analysis (CD34 and CD38) of adherent fraction of control cells in MS5 co-culture after 10 weeks. D. Marker expression analysis of CD14, CD15, CD11b and CD33 in RUNX1mut cells from liquid culture after 10 weeks. E. Expression of leukemia stem cells markers CD116, CD135 and IL1RAP in the CD34+ population in PBSCs (red) and RUNX1mut cells (blue) after 10 weeks. Blank stains are depicted in black. F. Expansion of CB CD34+/CD38+ vs CD34+/CD38- sorted cells in control and RUNX1mut cells in liquid cultures. G. May-Grünwald Giemsa stains of CD34+/CD38+ vs CD34+/CD38- sorted cells in RUNX1mut cells in liquid cultures after 60 days. Approximately 100 cells were scored either mature or immature based on phenotype. H. Relative expansion of adult peripheral blood stem cells transduced with control of RUNX1mut for 10 weeks and May-Grünwald Giemsa stain transduced with control or RUNX1mut.
Supplementary Figure 3. A. May-Grünwald Giemsa stains of CB control vs RUNX1mut cells cultured in liquid cultures for ~6 weeks with addition of G-CSF to promote granulocytic differentiation. B. Analysis of CSF3R (CD114) marker expression of RUNX1mut before and after overexpression of CEBPA. C. RT-q-PCR analysis of CEBPA, CSF3R (CD114), CTSG and MPO before and after overexpression of CEBPA.
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Supplementary Figure 4. A. Fold induction of RUNX1mut after induction by doxycycline. B. RUNX1mut expression on WB after induction by doxycycline. C. Time schedule of iPSC differentiation protocol and marker expression analysis and representable figure of CD34+/CD45+ and CD34-/CD45+ expression at various time points during differentiation. At various time points (blue arrow) of the haematological differentiation, doxycycline was added to induce RUNX1mut expression. Cell surface marker expression on CD34/CD45 was performed at various time points (red arrows). D. May-Grünwald Giemsa stains of iPSC control or RUNX1mut expressing cells at day 17.
Supplementary Figure 5. A. Overview of mutations in RUNX1mut primary AMLs used in this study. AML1 and AML3 have 2
RUNX1 mutations. B. Expression of signature genes in the monocytic and granulocytic lineage over time in control iPSCs. C.
Expression of signature genes in the monocytic and granulocytic lineage over time in RUNX1mut iPSCs.
Supplementary Figure 6. A. ChIP-qPCR on two different loci bound by RUNX1 in K562 cells expressing either control vector.
ChIP was performed against either RUNX1 or Flag M2. B. Venn diagram of overlapping RUNX1mut binding sites in CB, iPSC and primary AML carrying a RUNX1 mutation. C. Biological processes enriched for genes commonly targeted by RUNX1mut. D. Genomic distribution of common RUNX1 binding sites. All peaks are assigned to promoters (2 kb away from TSS), exons,
introns and intergenic regions based on the physical distance. The priority rule of category assignment: promoter exon intron intergenic region. All expressed genes are grouped into five classes based on expression levels. E. Location of genes relative to the closest RUNX1mut peak. F. Levels of specific chromatin marks (DNaseI, H3K27ac, H3K4me3, H3K27me3 and H3K4me1) in RUNX1mut binding sites. G. Relative expression of genes containing a RUNX1 binding peaks in all peaks (black), distal genes(blue) and genes within 2kb of a RUNX1 peak (red). H. Genes bound by RUNX1 in RUNX1wt and RUNX1mut AMLs. I. Log2 (fold change) of different epigenetic markers (DNaseI, H3K27ac, H3K4me3, H3K27me3 and H3K4me1) at common RUNX1 binding sites in RUNX1wt vs RUNX1mut AMLs. J. Levels of DNaseI, H3K27ac, H3K4me3, H3k27me3 and H3K4me1 at genes that have been found deregulated in Figure 5D. Cluster 3 (upregulated expression =red), Cluster 4 (downregulated
Supplementary Figure 7.A. Heatmap of RUNX1mut (top) and AML1-ETO (bottom) peaks in RUNX1mut binding sites (green) and RUNX1mut (top) and AML1-ETO(bottom) peaks in AML1-ETO binding sites (purple). B. Fold induction of AML1-ETO after induction by doxycycline. C. qPCR validation of differential regulation of various target genes identified as differentially deregulated in RUNX1mut vs AML1-ETO AMLs. D. RT-q-PCR analysis of MEIS1, HMGA2 and CD34 before and after overexpression of CEBPA in CB cells.
