Runx1 and Runx3 drive progenitor to T-lineagetranscriptome conversion in mouse T cell commitmentvia dynamic genomic site switchingBoyoung Shina,1

, Hiroyuki Hosokawaa,b,1, Maile Romero-Wolfa,2, Wen Zhoua, Kaori Masuharab,

Victoria R. Tobinc, Ditsa Levanond

, Yoram Gronerd, and Ellen V. Rothenberga,3

aDivision of Biology and Biological Engineering, California Institute of Technology, Pasadena, CA 91125; bDepartment of Immunology, Tokai UniversitySchool of Medicine, Isehara, Kanagawa 259-1193, Japan; cSchool of Veterinary Medicine, University of California, Davis, CA 95616; and dDepartment ofMolecular Genetics, Weizmann Institute of Science, Rehovot 76001, Israel

Edited by Nancy A. Speck, Raymond and Ruth Perelman School of Medicine at the University of Pennsylvania, Philadelphia, PA, and approved December 14,2020 (received for review September 22, 2020)

Runt domain-related (Runx) transcription factors are essential forearly T cell development in mice from uncommitted to committedstages. Single and double Runx knockouts via Cas9 show that tar-get genes responding to Runx activity are not solely controlled bythe dominant factor, Runx1. Instead, Runx1 and Runx3 are coex-pressed in single cells; bind to highly overlapping genomic sites;and have redundant, collaborative functions regulating genes piv-otal for T cell development. Despite stable combined expressionlevels across pro-T cell development, Runx1 and Runx3 preferen-tially activate and repress genes that change expression dynamicallyduring lineage commitment, mostly activating T-lineage genes andrepressing multipotent progenitor genes. Furthermore, most Runxtarget genes are sensitive to Runx perturbation only at one stageand often respond to Runx more for expression transitions than formaintenance. Contributing to this highly stage-dependent gene reg-ulation function, Runx1 and Runx3 extensively shift their binding sitesduring commitment. Functionally distinct Runx occupancy sites asso-ciated with stage-specific activation or repression are also distin-guished by different patterns of partner factor cobinding. Finally,Runx occupancies change coordinately at numerous clustered sitesaround positively or negatively regulated targets during commitment.This multisite binding behavior may contribute to a developmental“ratchet” mechanism making commitment irreversible.

Runx transcription factors | early T lymphocyte development |transcriptional regulation | DNA binding site choice | functional genomics

Runt domain-related (Runx) family transcription factors reg-ulate transcriptional and epigenetic programs essential for

multiple developmental processes (1–6). T cell development isparticularly dependent on Runx family factor activity from itsearliest pro-T cell stages (7, 8). Both Runx-dependent activationand Runx-dependent repression are important (9–12). The rec-ognition motif for Runx factors is also frequently enriched in cis-regulatory regions of genes used in early T cell development (11,13–17), although the targets most sensitive to Runx activity arestill not fully defined.As Runx1 is the predominant Runx factor expressed in early

T-lineage development (17, 18), it could be assumed to mediatethese activities. Runx1 is critical for hematopoietic developmentfrom the earliest embryonic stem cell stage onward (19, 20), andamong hematopoietic cells tested, intrathymic T cell precursorsexpress the highest levels of Runx1 (17, 18). Runx1 plays a long-established role in opening the T cell receptor (TCR) coding locifor recombination (21–24). Runx1 is also a functional regulatorycollaborator both of transcription factors that are active after pro-T cell lineage commitment (11) and of those acting before (refs.14 and 15; reviewed in ref. 25). However, although pro-T cellstages are all nearly eliminated when core binding factor β(CBFβ), the common partner of all Runx factors, is down-regulated (8), the acute disruption of Runx1 caused only mild

transcriptome effects in pro-T cells (14). Furthermore, impacts ofRunx1 deletion in vivo are mainly seen in postcommitment pro-Tcell stages (10, 26), later than the effect of deleting Cbfb. There-fore, we have tested whether other Runx family members com-pete, complement, or collaborate with Runx1 in shaping the generegulatory programs involved in pro-T cell specification.Three Runx paralogs Runx1, Runx2, and Runx3 (4, 27) have

diverged to distinct, often reciprocal tissue expression patterns,which restrict their functional redundancy (28–31). DifferentRunx paralogs can mediate nonredundant functions (in NaturalKiller cell responses), maintain mutual exclusion (in B cells), orcompensate for one another (in leukemia) (32–34). The rela-tionship and division of labor between different Runx proteinsthus need to be defined in a context-dependent manner.Early T cell lineage development actually provides more than

one regulatory context in which Runx factors can work (25).Driven by Notch signaling, the early precursor cells in the thy-mus, CD4−CD8− double-negative (DN) pro-T cells, progress in

Significance

T-lineage specification requires waves of gene network changesto terminate multipotency and establish T identity. Circumstantialevidence has suggested roles for Runt domain-related (Runx)factors in these regulatory events; Runx1 is strongly expressed,but Runx1 knockouts have previously shown little impact duringstages around T-lineage commitment. We show that Runx1 andRunx3 function together to drive T-lineage commitment, bindingthe same sites. Double knockouts block both activation ofT-lineage genes and repression of progenitor genes. Contrastingwith the stable activity of Runx proteins throughout these stages,Runx factors preferentially regulate target genes that sharplychange expression during T commitment. We show that this re-flects pronounced, global redistributions of Runx binding choicesacross the genome before and after T-lineage commitment.

Author contributions: B.S., H.H., and E.V.R. designed research; B.S., H.H., M.R.-W., W.Z.,and K.M. performed research; D.L. and Y.G. contributed new reagents/analytic tools; B.S.,H.H., M.R.-W., W.Z., V.R.T., and E.V.R. analyzed data; and B.S., H.H., and E.V.R. wrotethe paper with contributions from D.L. and Y.G

Competing interest statement: E.V.R. is a member of the Scientific Advisory Board ofCentury Therapeutics, LLC.

This article is a PNAS Direct Submission.

Published under the PNAS license.1B.S. and H.H. contributed equally to this work.2Present address: Department of Stem Cell Biology, University of Southern California, LosAngeles, CA 90033.

3To whom correspondence may be addressed. Email: evroth@its.caltech.edu.

This article contains supporting information online at https://www.pnas.org/lookup/suppl/doi:10.1073/pnas.2019655118/-/DCSupplemental.

Published January 21, 2021.

PNAS 2021 Vol. 118 No. 4 e2019655118 https://doi.org/10.1073/pnas.2019655118 | 1 of 12

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stages from DN1 or “early thymic progenitor” (ETP) to DN2a,DN2b, and DN3 (Fig. 1A) and after acquiring a form of TCR, toDN4 and beyond (35–38). In ETP and DN2a stages, pro-T cellsare still developmentally multipotent, express the transcriptionfactor PU.1, and epigenetically resemble prethymic multipotentprogenitors (15, 17, 39–42). These uncommitted stages are re-ferred to as “Phase1.” Commitment to the T cell fate occurs intransition from DN2a to DN2b stage and coincides with epige-netic changes defining entry into “Phase2” (25). This depends, atleast in part, on up-regulation of the transcription factor Bcl11b(41, 43, 44), which mediates changes in chromatin states andtranscriptional signatures (11, 42). In Phase2, committed DN2band DN3 cells establish expression of T cell identity genes andbegin TCR assembly (Fig. 1A). Importantly, signature tran-scription factors for Phase1 and Phase2, PU.1 and Bcl11b, re-spectively, both use Runx1 as a cofactor (11, 14).This study focused on three central questions. First, is Runx1

the sole member of the Runx family that contributes to earlyT cell development, or is its activity reinforced or modulated byactions of other Runx family members? Second, does the Runxfamily regulate a constant core of target genes throughout earlyT cell development, or does its mode of activity shift fromPhase1 to Phase2? As the three Runx paralogs are expressed indifferent developmental patterns in early pro-T cells, the answersto these two questions may be linked. Finally, how does genomicbinding of Runx factors explain their actions?Our results show that at each pro-T cell stage, Runx1 and

Runx3 are largely concordant in their binding and effects on target

genes, but they bind to substantially different sites before and aftercommitment. Their target genes are highly stage specific in theirfunctional responses and are highly developmentally dynamic innormal expression. Thus, despite apparently constant activitylevels, Runx factors work via developmentally changing genomicsites and preferentially drive developmental state transitions.

ResultsRunx Motifs Are Highly Enriched in the Open Chromatin Regions ofThymic Precursor Cells before and after Commitment. The sequencemotif recognized by Runx proteins has been found enriched atDNA sites occupied by E2A, PU.1, GATA3, or Bcl11b inT-lineage precursor cells (11, 13–16, 45). To measure the prev-alence of Runx motifs in an unbiased way across all genomicregions likely to be active in early pro-T cells, we screened for themost enriched transcription factor binding motifs in the openchromatin sites reported for thymic ETP, DN2a, DN2b, andDN3 cells (17) (SI Appendix, Fig. S1A and Dataset S1). Globally,there is a sharp transition in genomic activity and conformationfrom Phase1 (ETP and DN2a) to Phase2 (DN2b and DN3), al-though many sites maintain accessibility through both stages(Phase1&2 in common) (Fig. 1A) (17, 42). Motifs for Spi1(PU.1) and Tcf7 (high mobility group [HMG] box) family factorsshowed strong reciprocal changes in enrichment betweenPhase1-specific and Phase2-specific sites (Fig. 1B). However,consensus binding motifs for Runx factors were consistentlyamong the top three at open sites at all these stages (Fig. 1B and

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Fig. 1. Enrichment of Runx motif sequences at accessible chromatin (ATAC) peaks from pro-T cells and Runx1 and Runx3 coexpression in individual thymicprogenitor cells. (A) Schematic of T cell development and ATAC peak groups. P1: Phase 1; P2: Phase 2. (B) Motif density histograms show frequency of motifoccurrences around ATAC peaks of indicated sites. (C) Gene expression kinetics of Spi1, Bcl11b, Tcf7, Runx1, and Runx3 during T cell development (http://www.immgen.org/) are shown. (D and E) ETP-DN2 (cKithigh CD44+) and DN3 (cKitlow CD44− CD25+) cells were isolated from 4-wk-old animals. Expression ofRunx1 and Runx3 transcripts was measured by seqFISH in ETP (Kit copies ≥ 5, Il2ra copies ≤ 3, n = 890), DN2 (Kit copies ≥ 5, Il2ra copies >3, n = 1,984), andseparate DN3 (n = 1,587) cells. Transcript counts of Runx1 and Runx3 are enumerated and shown with median (D). Gene–gene Pearson distance heat maps ofRunx1 and Runx3 coexpression in thymic progenitor cells are displayed (E). LT-, ST-HSC: long-term, short-term hematopoietic stem cell; DP: double positive; SP:single positive.

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SI Appendix, Fig. S1A), supporting roles for Runx motif bindingfactors in pro-T cells before and after commitment.

Runx1 and Runx3 Are Coexpressed in Individual Thymic T ProgenitorCells. All Runx paralogs use a Runt homology domain to bind tothe same motif (1, 4). Whereas in most cell types, distinct Runxparalogs are expressed in different patterns, all three Runxparalogs are expressed in bone marrow hematopoietic precursorsbefore they enter the thymus, in data from http://www.immgen.org/ (Fig. 1C and SI Appendix, Fig. S1B) (17). Similar levels ofRunx1, Runx2, and Runx3 RNAs all appear to be expressed inthe ETP population, which also expresses Tcf7 and high Spi1(encoding PU.1) but little if any Bcl11b (Fig. 1C). Phase1 pro-Tcells also express Runx3 transcripts from the distal promoter (SIAppendix, Fig. S1C) (17), which may be preferentially translatedinto protein (46). After T-lineage commitment, as Spi1 falls andBcl11b rises, levels of Runx1 increase, whereas Runx3 expressiondeclines and Runx2 expression disappears completely (17, 18, 47)(Fig. 1C and SI Appendix, Fig. S1B) (http://www.immgen.org/).Runx1;Runx3 double knockouts (dKOs) lose all T cell devel-

opment in vivo (48), but Runx1 and Runx3 have contrasting rolesand expression in later thymocytes (46). Although both appearedhighly expressed in ETP and DN2a populations, we askedwhether Runx1 and Runx3 expression might be mutually exclu-sive in single cells. For maximally sensitive detection of Runxtranscripts, we used sequential fluorescent in situ hybridization(seqFISH) datasets (39) for efficient detection of low-copynumber RNA transcripts, important for genes encoding tran-scription factors (49–51). In fact, single-molecule transcriptcounts in single cells (Dataset S2) agreed with bulk RNA se-quencing (RNA-seq) for Runx1 and Runx3 expression profilesoverall (Fig. 1D) and confirmed that both Runx1 and Runx3transcripts are expressed simultaneously in the same individualpro-T cells (Fig. 1E). As expected, the ratio of Runx3 to Runx1transcript counts among individual cells in the population de-creased with developmental stage (Fig. 1E), with Runx3 tran-script counts slightly higher in most ETP and Runx1 transcriptcounts higher in most DN3 cells. Even so, >85% of ETPsand >93% of DN2 cells expressed three or more copies of both(57% of ETPs and >74% of DN2 cells had five or more copies ofboth) (Dataset S2). Runx1 and Runx3 proteins were also clearlycoexpressed in thymic pro-T cells, especially in DN2 stages (SIAppendix, Fig. S1D). Thus, despite changing developmentalpatterns, Runx1 and Runx3 are substantially coexpressed innearly all individual early intrathymic T cell precursors.

Runx1 and Runx3 Bind Shared Genomic Loci in Pro-T Cells but HaveStage-Specific Patterns of Occupancy. Despite coexpression, Runxfactors in pro-T cells might have paralog-specific functions ifthey interacted with distinct genomic regions. We comparedRunx1 and Runx3 occupancies directly by chromatin immuneprecipitation and deep sequencing (ChIP-seq) for Runx1 andRunx3 in primary Phase1 (cKithigh CD25low) and Phase2 (cKitlow

CD25+) pro-T cells. Populations were generated using theOP9-Delta-like 1 (Dll1; a Notch ligand) coculture in vitro dif-ferentiation system (52) (Fig. 2A). This mimics closely the earlypro-T cell stages in vivo before TCR gene assembly; a directtranscriptomic comparison is also shown (SI Appendix, Fig. S4D).Total Runx peak counts in Phase1 and Phase2 (reproduciblepeaks, as defined in Materials and Methods) were broadly con-sistent with the Runx gene expression patterns (Phase1: 21,069Runx1; 29,541 Runx3; Phase2: 32,796 Runx1; 27,172 Runx3).In each phase, Runx1 and Runx3 occupied most of the same

sites (Fig. 2 B–E) and usually bound to those sites with similarintensities (Fig. 2B, subgroups denoted as “R1∼R3”). However,despite similar overall levels of Runx binding at both phases, thepatterns of occupancy for both factors shifted substantially be-tween Phase1 and Phase2 (Fig. 2B). The statistics were different

for promoter and nonpromoter regions, in accord with theglobally stronger correlation reported between nonpromoter(enhancer) activity and developmentally dynamic gene expres-sion (17). Most of the promoter regions (22% of Runx-interacting sites) were occupied similarly in both Phase1 andPhase2, in common by Runx1 and Runx3 (Fig. 2C and SI Ap-pendix, Fig. S2 A and B). In contrast, nonpromoter sites com-prised multiple groups with distinctive stage-specific Runx bindingpatterns, and we focused on these (Fig. 2 B,D, and E). Three majorgroups were distinguished: a Phase1-specific group (8,810 peakstotal, 27% of nonpromoter sites), a Phase2-specific group(10,781 peaks total, 34% of nonpromoter sites), and a separateStable group common to both Phase1 and Phase2 (10,724 peakstotal, 34% of nonpromoter sites). The Phase1-specific peakswere also highly enriched for PU.1 (Spi1) motifs as well as Runxmotifs, whereas at the Phase2-specific peaks, Runx, E26 trans-formation-specific (ETS), Tcf7 (HMG), and basic helix-loop-helix (bHLH) motifs were all overrepresented (SI Appendix, Fig.S2 C and D). Thus, Runx1 and Runx3 behaved similarly inDNA binding activity, and they collectively occupied similarnumbers of sites before and after T cell lineage commitmentbut markedly shifted their binding site choices across commit-ment in a way that was not explained by availability of Runxfactors alone.

Functional Differences between Subsets of Runx1 and Runx3 Sites.Despite sharing occupancy of 10,000 to 16,000 peaks, Runx1 orRunx3 showed preferential occupancy at some genomic regions(Fig. 2 B and E, “R1 > R3” and “R1 < R3”). In Phase1 whenRunx3 expression is higher, ∼47% of all Phase1 peaks (11,573peaks of 24,622) showed stronger binding of Runx3 than ofRunx1, and very few bound Runx1 preferentially. In Phase2,∼28% of the Phase2 peaks (7,758 of 27,281) bound more Runx1than Runx3. Interestingly, these select, paralog binding-biasedgenomic regions displayed slightly different motif enrichments(Dataset S1). Runx3-preferring sites always gave the Runx motifas the most frequently discovered consensus sequence, both inPhase1 and Phase2, whereas Runx1-preferring sites showedsimilar or higher enrichments for motifs of other partners suchas bHLH or ETS factors (SI Appendix, Fig. S2 E and F) (cf ref.14). Hence, despite the preponderance of common targets forboth Runx factors in both stages, Runx1-specific DNA inter-actions may be more influenced by its binding partners thanthose of Runx3.

Runx1 and Runx3 Are Necessary and Functionally Redundant for Pro-TCell Development in Phase1. Even when binding to the same sites,Runx1 and Runx3 might exert complementary, antagonistic, ormutually reinforcing impacts. These could be distinguished byeffects of acute, stage-specific Cas9-mediated single- or double-gene deletion. To test this, therefore, bone marrow progenitorcells from B6.ROSA26-Cas9 knock-in mice with a Bcl2 transgene(B6.Cas9;Bcl2) were cocultured with OP9-Dll1 cells to initiateT cell development, and guide RNAs (sgRNAs) against Runx1and/or Runx3 or control sgRNAs were retrovirally delivered toinduce deletion at specific times of development (Fig. 3). TheBcl2 transgene was included to enhance viable recovery of cellswith regulatory perturbations without altering development (53),and a Bcl11bmCh allele (54), also included, provided an mCherryfluorescent marker for Bcl11b up-regulation, which coincideswith T commitment in late DN2a stage (41). We induced acutedeletion before commitment (Phase1 deletion) or after com-mitment (Phase2 deletion) and then analyzed the cells 3 d later(Fig. 3A). The introduction of sgRunx1 and/or sgRunx3 causedspecific deletion at targeted sites within 3 d, as shown by point-focused deletions in RNA transcripts and loss of protein de-tected by intracellular immunostaining in the pro-T cells andsupported by western blotting in acute Runx knockout (KO) cell

Shin et al. PNAS | 3 of 12Runx1 and Runx3 drive progenitor to T-lineage transcriptome conversion in mouse T cellcommitment via dynamic genomic site switching

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lines (SI Appendix, Fig. S3 A–C). The developmental status ofRunx-disrupted pro-T cells was scored with markers cKit andCD25, which distinguish pro-T stages as shown in Fig. 1A,and using Bcl11b-mCherry expression to mark commitment.Phase1 pro-T cells with single disruptions (single knockout

[sKO]) of Runx1 or Runx3 were minimally affected. They stillprogressed to DN2 stage and activated levels of Bcl11b expres-sion comparable with those of sgControl transduced cells(Fig. 3 B–D), despite slightly lower overall cell recoveries (SIAppendix, Fig. S3D). In contrast, deletion of both Runx1 andRunx3 simultaneously (dKO) severely impaired progression ofPhase1 cells, yielding a significantly lower percentage of DN2cells (Fig. 3 B and D, Left) and failure to induce Bcl11b ex-pression (Fig. 3 C and D, Right). The recovered cell numberswere also more profoundly reduced in dKO than in control orsingle deletion groups, indicating that both Runx1 and Runx3normally contribute to proliferation or survival (SI Appendix, Fig.S3D). Thus, Runx1 does not act alone in Phase1 pro-T cell de-velopment; it shows substantial redundancy and compensationwith Runx3.

Runx1 Supports Optimal Expression of Bcl11b in Phase2. The knockouteffect was not superficially as obvious in Phase2 as in Phase1. Whenwe delivered sgRNAs to B6.Cas9;Bcl2 cells after progenitor cellshad become committed to the T cell fate (Fig. 3A), most cellsappeared to reach DN3 stage by the time of analysis (Fig. 3E). Runxgene knockouts now caused no significant impacts on the CD25 andcKit expression phenotypes at day 3 postinfection, and the fre-quency of Bcl11b+ CD25+ (presumptively committed) cells wasnot altered either (Fig. 3 E–G). Also, unlike Phase1, the recoveryof these Phase2 cells was almost insensitive to Runx disruption.Only Runx1, Runx3 double deletion yielded slightly lower numbersof cells than the other groups, and this was not statistically sig-nificant (SI Appendix, Fig. S3E). In agreement with the high per-centage of Bcl11b-mCherry+ cells, the Phase2-deleted cellsremained functionally committed to the T cell lineage, as shownby their response on transfer to conditions lacking Notch ligand(SI Appendix, Fig. S3F). However, the expression level of Bcl11b(detected by mCherry reporter) per cell was modestly but con-sistently reduced in both the Runx1 sKO and the Runx1, Runx3dKO samples (Fig. 3H) (P value < 0.05), in agreement with aprevious report (41). More substantial effects of Runx disruption

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Fig. 2. Runx1 and Runx3 interact with shared genomic sites that shift at different developmental stages. (A) Progenitor cells from normal bone marrow (BM)were cultured with OP9-Dll1 stromal cells for 5 d (Phase1) or for 10 d (Phase2); then, pro-T cells in Phase1 (cKithigh CD25low) or Phase2 (cKitlow CD25+) werefluorescence-activated cell-sorted for chromatin immunoprecipitation with sequencing (ChIP-seq) as described in Materials and Methods. Experiment sche-matic and gating strategy are shown. (B) Tag count distributions for Runx1 and Runx3 in Phase1 and Phase2 stages are represented by peak-centered heatmap. P1: Phase1; P2: Phase2; R1: Runx1; R3, Runx3. (C–E) Area-proportional Venn diagrams illustrate the peak overlaps of Runx1 and Runx3 ChIP-seq peaks inPhase1 and Phase2, with numbers of peaks in each category. Scatterplots compare log2 normalized tag counts per 10 million tags for peaks in indicatedcategories with Pearson correlation r. (C) Promoter region peaks (TSS: transcription start site); (D) nonpromoter region peak comparison for Runx1 and Runx3;and (E) nonpromoter region comparison for Phase1 and Phase2. Data are based on ChIP-seq peaks scored as reproducible in two replicate samples (B–E).

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in Phase2 became evident from transcriptome analysis, which isdiscussed next.

Runx1 and Runx3 Exert Parallel Functions in Gene Regulation inPhase1 and Phase2. RNA-seq analysis at 3 d post-sgRNA intro-duction (Dataset S3) showed that sKO of either Runx factor alonecaused little change in transcriptomes of Phase1 pro-T cells(Fig. 4 A and B). In contrast, when Runx1 and Runx3 were dis-rupted together, 342 genes were differentially expressed (differ-entially expressed genes [DEGs]; adjusted P value < 0.05, log2 foldchange either direction >0.5) (Fig. 4B and Dataset S3). Moregenes in the dKO were derepressed (213 genes up-regulated) thandown-regulated (129), implying that Runx factors normally havesubstantial direct or indirect roles in these cells in repression(“inhibition”) as well as activation (“dependency”). Similar resultswere seen using a more stringent threshold, |log2 fold change| > 1(210 inferred to be Runx inhibited, 118 dependent). The synergyobserved implies that previously observed Runx1 effects in Phase1cells only appeared weak due to paralog redundancy, primarilywith Runx3, in these stages.

In Phase2 after T commitment, delivery of sgRunx1 alone wassufficient to change gene expression substantially (Fig. 4C andDataset S3). This transcriptome impact was much stronger thanwas obvious from the CD25, cKit, and Bcl11b marker analysis (cfFig. 3 E–G). Because the knockouts had little impact on cellrecovery at this stage, the effects seen could not be artifacts ofselective survival. The Runx1 sKO gave >300 differentially reg-ulated genes (DEGs), of which 186 were down-regulated (Runxdependent) and 114 were up-regulated (Runx inhibited) in thesKO. In a sharp contrast, Runx3 deletion alone at this stage didnot cause any changes in messenger RNA (mRNA) levels.However, Runx1, Runx3 dKO cells again showed the greatestnumber of DEGs. Runx-inhibited genes increased to 224 sig-nificant DEGs, while Runx-dependent DEGs increased to 244(Fig. 4C) (with more stringent threshold, 168 inhibited and 167dependent).Runx1 and Runx3 worked in the same direction on individual

genes in Phase2 cells, as shown by comparing the log2 foldchange values of individual DEGs obtained from Runx1 sKO andfrom Runx1, Runx3 dKO cells. Effects in Runx1 sKO and Runx1,

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Fig. 3. Runx factors are essential for thymic T cell development at Phase1 and Phase2. (A) Experiment schematic for Phase1 and Phase2 perturbation isdisplayed. (B–D) For Phase1 perturbation, bone marrow (BM) progenitor cells from Cas9;Bcl2 mice or Cas9;Bcl2 mice with Bcl11b-mCherry reporter werecultured with OP9-Dll1 cells for 2 d before transduction with sgRNAs (Phase1). (B and C) Expression levels of cKit, CD25, and Bcl11b of pro-T cells as analyzedby flow cytometry 3 d after Phase1 perturbation, gated on 7AAD− Lin− CD45+ infection marker (CFP&NGFR)+ population. Frequencies of DN2 (cKithigh CD25+)population and CD25+ Bcl11b+ population are shown (D). (E–H) Expression levels of cKit, CD25, and Bcl11b are shown 3 d after Phase2 perturbation, gated on7AAD− Lin− CD45+ infection marker (CFP&NGFR)+ population. (E and F) Representative flow cytometry plots. (G) Frequencies of DN3 (cKitlow CD25+) cells andCD25+ Bcl11b+ populations. (H) Histogram of geometric mean fluorescence intensity (gMFI) of Bcl11b expression following Phase2 perturbation, comparingrelative gMFI with control group. n = 7 for cKit, CD25 analysis (B and E ), and n = 3 to 4 for Bcl11b analysis (C and F ). Graphs show the average ± SD. (D,G, and H) One-way ANOVA. *P value < 0.05; **P value < 0.01.

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Runx3 dKO Phase2 cells were positively correlated for bothRunx-dependent and -inhibited targets (Pearson r = 0.60 fordependent genes; r = 0.82 for inhibited genes). Comparing dKOwith sKO effects on the same genes, dKO effects were moreintense for both Runx-dependent (slope = 1.332) and Runx-inhibited DEGs (slope = 1.524), whereas best-fit line slopes fornon-DEGs were <1, ruling out nonspecific effects (Fig. 4D).Thus, even in Phase2, the dKO augmented changes in Runx-sensitive (dependent or inhibited) gene expression, in the samedirections, relative to effects of the sKO.These data, with Fig. 3, demonstrate that Runx1 and Runx3

function similarly when they are coexpressed in early T cell de-velopment. They show strong functional redundancy in Phase1,and in Phase2, where Runx1 becomes the major contributor,even lowly expressed Runx3 plays a milder but still parallel rolein transcriptional regulation. Thus, the sum of Runx1 and Runx3activity appears most important.

Runx Factors Drive Activation of the T-Lineage Program and SuppressAlternative Lineage Signatures. If Runx1 and Runx3 act equiva-lently, it was notable that the genes affected by Runx1, Runx3dKO were a select minority of the T cell precursor transcriptomethat was most developmentally dynamic. Empirical cumulativedistribution frequency (ECDF) plots and histograms showed thatabout 60% of total Runx-responsive DEGs normally underwentmore than twofold changes in expression levels between Phase1and Phase2, much higher than the fraction of expressed Runxnon-DEGs undergoing such changes (about 15%; P value < 9e-30) (Fig. 5A). Furthermore, Runx-dependent and Runx-inhibitedgenes had reciprocally biased developmental profiles: 80% ofRunx-dependent genes (defined by dKO at either phase) weremore highly expressed normally in Phase2, whereas 70% of

Runx-inhibited genes (defined by dKO at either phase) hadhigher gene expression normally in Phase1 (P value = 3.1e-15)(Fig. 5B). Thus, despite near-constant total Runx factor DNAbinding (Fig. 2 C and E), Runx activities were most importantto cause gene expression changes during the Phase1–Phase2transition.Runx-dependent DEGs were enriched for T cell “identity”

genes associated with DN2 and DN3 Phase2 cells in Gene SetEnrichment Analysis (GSEA). For example, Bcl11b, Cd3d, Cd3e,Cd3g, and Lck, key T cell program genes highly activated duringT-lineage commitment (Fig. 5C and SI Appendix, Fig. S4A), weredown-regulated by loss of Runx whether deleted in Phase1(Phase1 sensitive) or in Phase2 (Phase2 sensitive). In contrast,Runx-inhibited DEGs were enriched for stemness signaturegenes (ETPs and hematopoietic stem cells; e.g., Meis1, Tal1, andKlf1 [Phase1 sensitive]) and for genes associated with alternativelineages, including Csf1, Ly9, Cd86, and Id2 (Phase1 sensitive),and Spi1, Id1, Id2, Id3, and Cd81 (Phase2 sensitive) (Fig. 5C andSI Appendix, Fig. S4B). In agreement with GSEA, the ImmGendatabase (17) showed that the Runx-dependent genes wereexpressed more highly in early T-lineage populations (SI Ap-pendix, Fig. S4C, red labels) than in myeloid cells, whereas theRunx-inhibited genes had the reverse pattern (SI Appendix, Fig.S4C). Therefore, Runx1 and Runx3 are essential to establish theT cell program and inhibit myeloid and stem cell gene expressionin this process.Runx-gene dKO at different phases distorted the normal de-

velopmental trajectory, as shown by a supervised principalcomponent analysis (PCA) of control and Runx KO pro-T celltranscriptomes (Fig. 5D). We used 65 highly informative devel-opmental regulatory genes from a reference single-cell RNA-seqanalysis of normal in vivo thymocytes (39) to define a fixed frame

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Fig. 4. Runx1 and Runx3 function in parallel in generegulation. (A) Experimental schematic for RNA-seqis displayed. RNA-seq measurements were performedon fluorescence-activated cell-sorted sgControl-,sgRunx1-, sgRunx3-, and sgRunx1 + sgRunx3-transduced pro-T cells on day 3 posttransduction(7AAD− Lin− CD45+ CFP+ NGFR+ cells) for Phase1(cKithigh population) and Phase2 (CD25+ population)perturbations. (B and C) Volcano plots show statis-tical significance (log10 P value) vs. log2 fold change(FC) of DEGs (average fragments per kilobase-millionreads ≥ 1, adjusted P value < 0.05, log2 FC > 0.5 ei-ther way) in Phase1 perturbation (B) and Phase2perturbation (C) groups. n = number of genes scor-ing as DEGs in each condition. (D) Dot plots showlog2 FC of Runx1 sKO vs. Runx1 + Runx3 dKO uponPhase2 perturbation. Trend lines show linear re-gression results with 95% CI for Phase2 perturbationDEGs (red) and Non-DEGs (gray). Best-fit values forslopes and Pearson correlation r are noted. Data arebased on two to three replicate samples of RNA-seq results.

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of principal component (PC) loadings (Fig. 5D, open symbols;also Materials and Methods and Dataset S4). As expected, RNA-seq results from control in vitro-generated pro-T cell populations(no sgRNA) mapped into this PC space close to their in vivocounterparts (SI Appendix, Fig. S4D), validating the similarity ofin vitro- and in vivo-generated pro-T cells. In this framework(Fig. 5D), transcriptomes of populations transduced in Phase1with sgControl, sgRunx1 alone, or sgRunx3 alone all clusteredtogether, between those of normal ETP and Bcl11b-negativeDN2a thymocytes. In contrast, transcriptomes from Phase1dKO populations (sgR1 + R3) were clearly shifted backwardtoward ETP, indicating a defect in progression. In Phase2 per-turbations, samples transduced with sgControl or sgRunx3mapped close to DN2b references as expected, but here, bothRunx1 sKO and Runx1, Runx3 dKO samples were retarded(Fig. 5D). Taken together (Dataset S4), these data confirm thatRunx regulates targets that promote T-lineage identity and de-velopmental progression before and after commitment.

Specific Phase-Dependent Regulation of Runx Factor Functional Targets.In principle, transcription factors stably expressed across a devel-opmental stage transition like Runx1 + 3 might be expected toregulate their target genes similarly in both stages. However, among312 Runx-dependent genes, only 61 (19%) were sensitive to Runxdisruption at both stages in common. The targets of Runx inhibitionshowed even less overlap, as only 27 of 410 (6.5%) Runx-inhibitedgenes were sensitive in both (Fig. 5E and SI Appendix, Fig. S4E).The rest responded in stage-specific ways.Runx-dependent and Runx-inhibited targets showed stage-

specific responses to Runx deletion, whether they were devel-opmentally increasing or decreasing in expression (Fig. 5E). Thevaried patterns argue against systematic detection biases, andRNA-seq tracks show the robustness of these effects (Fig. 5F andSI Appendix, Fig. S5). Many Runx-dependent targets showedstage-specific responses, including genes with key ongoingroles in T cell development (e.g., Tcf7, Thy1, Il2ra, and Cd5[Cluster10], Phase1 sensitive only), while Ahr, Hdac4, andHmgcs2 (Cluster12) were sensitive in Phase2 only (Fig. 5F andSI Appendix, Fig. S5 C and E; note scale changes betweenphases). Differentially stage-sensitive groups were especially obvi-ous among the repression targets, as many Runx-inhibited geneswere more up-regulated when Runx was removed in Phase1(Fig. 5F and SI Appendix, Fig. S5D) (Cluster5 and Cluster7; e.g.,Tal1, Sgk1, Meis1, Irf7, and Csf1), while others responded more inPhase2 (Cluster1 and Cluster9; Spi1, Pou2af1, Lyl1, CD34, Id1, andId3) (Fig. 5F and SI Appendix, Fig. S5F). The stage-specific targetsTcf7 (Runx dependent only in Phase1 [Cluster10]) and Spi1 (PU.1;Runx inhibited only in Phase2 [Cluster1]) (Fig. 5F) have particularlyimportant roles in T cell development. Thus, unexpectedly, a ma-jority of both Runx-dependent and Runx-inhibited targets respon-ded to Runx input selectively during specific developmental phases,often during expression changes, rather than for maintenance of agiven expression level.

Runx Binding Is Enriched at DEGs and Enriched at Phase of HighestDEG Expression. Given the near-constant levels of Runx1 +Runx3 activity across commitment, these results raised ques-tions. First, were most effects of Runx activity mediated directly,and were stage-specific changes in binding sites (Fig. 2) re-sponsible for developmental changes in action on different setsof target genes? Second, could the stage-specific positive andnegative impacts of Runx on particular target genes (Fig. 5E) belinked to Runx binding to different function-specific sites or toaltering Runx function at the same sites? For insight, we testedwhether particular clusters of genes showing different patterns ofdevelopmental expression and Runx responsiveness (Fig. 5 B andE) might be correlated with particular patterns of Runx factor

occupancies at sites linked to them, from Phase1 to Phase2(Dataset S5).We first determined whether each transcript in DEG and non-

DEG groups had any linked high-scoring promoter and non-promoter Runx peak(s) in its surrounding genomic regions(Materials and Methods). Then, we compared 1) the percentageof genes in each category linked to such Runx binding and 2) thez score showing positive or negative association, by comparingthe difference between random frequencies of Runx peaksamong all expressed genes and the observed frequencies in eachgroup (Fig. 6A). Non-DEGs were more likely than DEGs topossess Runx binding sites at promoters, indicating that Runxoccupancy near the promoter was a poor indicator of functionaltargets (SI Appendix, Fig. S6). In contrast, a higher percentage ofDEGs (70 to 90%) than of non-DEGs (∼50%) was linked to atleast one nonpromoter Runx peak (Fig. 6B). Both Runx-inhibited and Runx-dependent genes had more local Runxbinding at nonpromoter regions than random expressed genes (zscores +10.8 and +12.8), whereas expressed but Runx-insensitive non-DEGs showed less binding (z score −16.9)(Fig. 6B and Dataset S5). This was also evident in ECDF ofRunx occupancies per gene (Fig. 6C). Runx-dependent genesshowed higher numbers of linked Runx binding sites/gene thanthe Runx-inhibited genes (Fig. 6 B and C). This suggests thatboth activation and repression may often be direct and thathigher numbers of high-scoring nonpromoter peaks were moreassociated with positive transcriptional regulation.To test how Runx occupancy shifts (Fig. 2B) might be related

to different functions, we classified nonpromoter binding sites bytheir occupancy with Runx factors in Phase1 only (Group 1),Phase2 only (Group 2), or both (Group 3) (cf Fig. 2B). We thenasked which site groups were preferentially enriched near Runx-activated and Runx-inhibited DEGs of distinct expression andsensitivity features (details and caveats are in SI Appendix, Fig.S7). In general, the Runx sites with stage-specific binding hadhigher enrichment near Runx-responsive target genes that wereexpressed most strongly in that stage. Phase2-specific (Group 2)peaks especially were highly enriched near targets expressedhighly in Phase2 (SI Appendix, Fig. S7 A and B). Peaks sharedbetween Phase1 and Phase2 (Group 3) were also more enrichedat Phase2-high DEGs. Runx binding was also slightly higher nearthe Runx-dependent targets with high expression in the matchingstage than near Runx-inhibited ones (SI Appendix, Fig. S7B),both for Phase2-specific Group 2 Runx peaks and for Phase1-specific Group 1 peaks.In contrast, the timing of DEG sensitivity to Runx was not

necessarily correlated with the time of highest Runx binding, andthis showed a much more complex pattern (SI Appendix, Fig.S7C), dissected further below. Of note, although sites gaining ormaintaining maximal Runx binding at Phase2 (Groups 2 and 3)showed the most biased linkage with genes that were Runx de-pendent, not Runx inhibited, this bias was surprisingly strongestamong genes that showed Runx dependence only in Phase1(Phase1-sensitive Dep vs. Inhib) (SI Appendix, Fig. S7 C, Middleand Bottom). These patterns suggest that a target gene’s maxi-mum sensitivity to Runx perturbation could occur during anRunx binding transition (e.g., during increase of local binding)rather than at the time of maximum Runx binding.

Association of Chromatin Accessibility with Stage-Specific RunxBinding. To explore how the developmental shifts in Runx bind-ing (Fig. 2B) could be associated with function, we asked whetherRunx factors require or induce chromatin opening, testing the re-lationship between chromatin accessibility (assay of transposase-accessible chromatin by sequencing [ATAC-seq]) and Runx occu-pancy (cf Fig. 2B) at nonpromoter Runx binding sites (SI Ap-pendix, Fig. S8). Not surprisingly, about 85% of the Group 3sites, which are commonly occupied by Runx factors both in

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Fig. 5. Runx1 and Runx3 function additively to direct pro-T cell program and suppress alternative lineage potentials. (A) RNA expression changes (log2 fold change [FC])between Phase1 and Phase2 in controls are shown, comparing Runx DEGs with non-DEGs in cumulative frequency (ECDF; Left) and histogram (Right) plots. Consistentlyexpressed genes are genes with |log2 FC| < 1 in either direction and average expression (as fragments per kilobase-million reads [FPKM]) ≥ 1. (B) ECDF plot comparingPhase2/Phase1 mRNA expression ratios (log2 FC) in Runx-dependent genes vs. Runx-inhibited genes. “Phase1 high” indicates genes with log2 FC of Phase2/Phase1 < 0.“Phase2 high” indicates genes with log2 FCs of Phase2/Phase1 > 0. (C) Top pathways identified by GSEA and normalized enrichment score (NES) are shown for Phase1-sensitive Runx-regulated genes (Left) and Phase2-sensitive genes (Right). Size of the dots indicates the number of genes in enriched gene set. Color of dot is NES. HSC:hematopoietic stem cell. (D) Effects of indicated perturbations on developmental progression, as shown by supervised PCA plot from RNA-seq data. Fixed PC loadings weredetermined based on single-cell RNA-seq data as described in Materials and Methods. (E) Heat map of RNA-seq displays expression levels of the DEGs either in Phase1perturbation and/or in Phase2 perturbation groups. Each cluster was determined by phase sensitivity (white bar, Phase1 sensitive only; gray bar, Phase2 sensitive only; blackbars, Phase1 and Phase2 sensitive), steady-state expression pattern, andmode of Runx factor effects (blue, Runx inhibited; red, Runx activated). (F) Representative genomebrowser tracks (http://genome.ucsc.edu) for DEGs sensitive in either Phase1 or Phase2 only. FPKM values are shown. Data are based on two to three replicate samples ofRNA-seq results (A–E) or are representative of two or three replicate samples (F).

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Phase1 and Phase2, were constantly open from ETP throughDN3. Group 1 sites also showed binding primarily associatedwith open chromatin, with about 46% of Group 1 sites losingaccessibility as Runx binding declined. In sharp contrast, themajority of Group 2 sites were either open (55% of Group 2peaks) or closed (25% of Group 2 peaks) at all times; less than12% opened specifically at the time of Runx binding. Thus,Phase2-specific Runx binding neither awaits nor causes chro-matin opening (SI Appendix, Fig. S8).

Distinct Stage-Specific Runx-Partner Factors Preferentially Associatewith Different Regulatory Effects. To distinguish between Runx-bound sites that mediate positive and negative gene regulation,we asked whether cobinding with prominent partner factorscould be important. As previously noted (11, 14, 15), most Runx1occupancy sites were co-occupied with PU.1 (53%) and/orBcl11b (66%), and Runx3 sites overlapped similarly with PU.1(43%) or Bcl11b (67%) (SI Appendix, Fig. S9A). We subdividedthe site groups based on ChIP-seq evidence for Runx cobindingwith PU.1 in Phase1 and/or with Bcl11b in Phase2 (Fig. 6D, SIAppendix, Fig. S9A, and Dataset S5), thus resolving seven sitegroups (Fig. 6D: 1a, 1b, 2a, 2b, 3a, 3b, and 3c). We then testedeach of these finer groups for preferential enrichment near DEGtypes defined in Fig. 5E (Fig. 6E and SI Appendix, Fig. S9B), firstusing the coarse DEG clusters defined in SI Appendix, Fig. S7B(SI Appendix, Fig. S9 C–E) and then using the finer DEG clusters(SI Appendix, Fig. S9 F–J). As simplified in Fig. 6E, genes inthese DEG clusters had separable responses (Fig. 6E, brokenlines) to Runx dKO at different stages (scissors), which weresuperimposed upon their different normal developmental pat-terns of expression (solid brown lines). Nearly all these Runx sitegroups were enriched in most DEG clusters relative to the non-DEG background (SI Appendix, Fig. S9 C–H, horizontal brokenlines). Some special functional relationships stood out withparticular DEG response types. This was shown when each sitegroup’s enrichment in linkage to genes of a particular DEG re-sponse type was compared with its enrichment among DEGsoverall (i.e., values expected if the frequency of that site groupwas randomized among all DEGs) (SI Appendix, Fig. S9 C–H,Rdm DEG bar; z scores for comparisons between DEGs areshown in color scale). These results showed that co-occupancy ofan Runx site with PU.1 or Bcl11b was associated with markedlydifferent linkages to specific DEG types (SI Appendix, Fig. S9 Cand D, Group 1a vs. 1b and Group 2a vs. 2b, respectively).Both cobinding partners, PU.1 and Bcl11b, were associated

with more frequent binding in DEGs as compared with non-DEGs (SI Appendix, Fig. S9 C–E; compare 1a vs. 1b, 2a vs. 2b,and 3a vs. 3b and 3c). However, the rare Phase1-high genespositively regulated by Runx were markedly enriched for Phase1-specific Runx sites without PU.1 cobinding (SI Appendix, Fig.S9C, Group 1a). Cluster C2 genes, Runx dependent in Phase1,were especially enriched for Group 1a (no PU.1) and not Group1b sites (with PU.1). In contrast, Phase1 Runx cobinding withPU.1 (Group 1b) was found more broadly with weaker, variedfunctional associations (SI Appendix, Fig. S9F).Cobinding with the Phase2 partner, Bcl11b, had markedly

greater predictive value. Sites of Runx cobinding with Bcl11b(Group 2b, 3b, 3c) were strongly enriched for both overall DEGbinding and overall association with Phase2-specific, Runx-dependent DEGs as compared with those without Bcl11b (SIAppendix, Fig. S9 D and E, cf Groups 2a and 3a). Fine-grainedcluster comparison showed that sites with Bcl11b cobinding (SIAppendix, Fig. S9G, Group 2b) could be associated with Phase2timing of repressive as well as activating functions (i.e., both forPhase2-high Runx-dependent DEGs [C10, 12] and for the fewPhase2-high genes that were constrained by Runx inhibition inPhase2 [C9]). Of note, Bcl11b-linked Phase2 Runx binding didnot seem as important for silencing Phase1-high Runx repression

targets, as these sites were systematically underrepresented nearPhase1-high Runx-inhibited loci (SI Appendix, Fig. S9 D and E,Groups 2b and 3b; negative z scores). Their association fell tobackground levels near C5 DEGs, which were only Runx sensi-tive in Phase1 (SI Appendix, Fig. S9 G, Group 2b and H, Groups3b and 3c). Thus, “new” Runx binding sites in Phase2 areprobably not needed to maintain silence of genes originally re-pressed in Phase1. SI Appendix, Fig. S9 I and J summarizes allthese patterns, showing the representation of each peak groupwith each DEG cluster as its percentage of all peaks linked togenes in that cluster.We asked whether sites with preferences for Runx3 or Runx1

are linked to different functions (SI Appendix, Fig. S10 A–D).Most Runx-sensitive loci were linked to sites with both Runx1and Runx3 occupancy (Runx1∼Runx3), and overall results withthese and the Phase2-specific Runx1-preferring sites were asdescribed above (SI Appendix, Fig. S10 B and D). However,Phase1 peaks that preferred Runx3 over Runx1 distinctivelywere enriched near Runx-dependent genes with highest expres-sion in Phase1, not Runx-inhibited genes (SI Appendix, Fig.S10C). They thus resembled Phase1 occupancy sites withoutPU.1 (Group 1a: cf SI Appendix, Fig. S9 C and F). Thus, Runx3may have specific, limited positive roles in Phase1.Thus, different classes of Runx sites are associated with dif-

ferent target gene response types. Bcl11b cobinding potentiatesPhase2-specific Runx effects and initiation of Phase2-high ex-pression, whereas PU.1 cobinding may mask Phase1-specificRunx effects with its own generalized pioneering and regula-tory roles (15). In addition, the need for Runx to inhibit Spi1itself in Phase2 may create indirect pathways through whichPhase1-high genes that depend on PU.1-bound sites (C1) can beinhibited by Runx in Phase2.

Runx-Sensitive Target Genes Recruit Stage-Specific Runx Binding inClusters. The pattern of stage-specific Runx binding to functionaltarget loci showed one more contrast with the overall stability ofRunx activity. Occupancy patterns across the regulatory elementsof Runx DEGs (SI Appendix, Figs. S10E and S11) underwentcoordinated developmental changes in Runx binding duringcommitment, at multiple sites around target loci spanning largegenomic regions. Runx-dependent genes specifically expressed inPhase1 (e.g., Hhex [C2], Clnk [C4], and Itgax [C2]) displayeddramatic losses of Runx1 and Runx3 binding as clusters ofGroup1 Runx peaks disappeared after commitment (Fig. 6 F,Left and SI Appendix, Fig. S11A, orange and red arrowheads).Runx-inhibited genes with high expression in Phase1 also lostmultiple Phase1 occupancy peaks during transition to Phase2(e.g., Csf1 [C5], Meis1 [C5], and especially Plek [C6]) (SI Ap-pendix, Fig. S11 B and C), although statistical inference for thisgroup of DEG was weaker.The Runx-dependent loci highly expressed in Phase2 showed

the opposite pattern. The entire regulatory region of Bcl11b,spanning >1 Mb, was almost devoid of Runx binding in Phase1cells; then, >30 Group2 sites were occupied by Runx proteinsand Bcl11b, a few days later, in Phase2 (Fig. 6 F, Center, lightblue arrowheads). Thus, the early sensitivity of Bcl11b expressionto Runx dKO in Phase1 could reflect the need to assemble a newactivation structure for Bcl11b expression for transition toPhase2. Similar concerted Runx binding was found at Thy1(C10), the Cd3 cluster (C11), and Hdac4 (C12) (SI Appendix, Fig.S11D). Runx-inhibited genes normally expressed in Phase2 alsogained multisite binding in Phase2, as represented at Pou2af1and Id3 (Fig. 6F and SI Appendix, Fig. S11 C, Right). Suchconcerted binding of Group2 sites was striking, considering thatmost of these sites did not change in chromatin ATAC accessi-bility (SI Appendix, Fig. S8).Thus, taken together, Runx1 and Runx3 show phase-specific

DNA binding patterns around both positively and negatively

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regulated targets, which often appear as large groups of linkedsites that dynamically change occupancy together during T celldevelopment (SI Appendix, Fig. S10). These distinctive genomic

interactions in Phase1 and Phase2 are closely associated withstage-specific gene regulation, which is also correlated with pat-terns of co-occupancy with PU.1 or Bcl11b.

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Fig. 6. Phase-specific collaboration with PU.1 and Bcl11b is associated with Runx functional target genes. (A) Bar graph displays the percentage of the genesassociated with peaks, comparing observed values (Obs) and expected values (Exp). z scores (standardized residuals) across categories are shown by the colormap. (B) Cumulative frequency of the number of high score peaks found per gene in indicated group. Gray area indicates zero high-score peaks per gene.P values: Kolmogorov–Smirnov test. (C) Peak-centered heat map illustrates ChIP-seq tag count distributions for indicated transcription factors across15,051 high-quality genomic sites of Runx factor binding in Phase1 and Phase2 stages. High-score peaks indicate nonpromoter Runx binding sites with peakscores ≥ 30. (D) ChIP-seq and RNA-seq integrative analysis strategy is shown. As an example, observed numbers of genes in non-DEG, Runx-inhibited, andRunx-dependent categories with or without any Runx binding peaks are given, compared with a table of expected values if sites were distributed randomly.(E) Diagrams illustrate expression patterns of genes in different clusters from Fig. 5E and SI Appendix, Fig. S9B. C, cluster. (F) Representative genome browserprofiles (http://genome.ucsc.edu) are shown with highlighted stage-specific peaks. (Left) Phase1-high Runx-dependent gene (Hhex locus), (Center) Phase2-high Runx-dependent gene (Bcl11b locus), and (Right) Phase2-high Runx-inhibited gene (Pou2af1 locus). Data are based on ChIP-seq peaks scored as re-producible in two biological replicate ChIP-seq samples and two or three replicate samples of RNA-seq results.

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DiscussionFunctional genomics and gene network predictions often assumethat the target genes of a transcription factor should be expressedin a pattern matching that of the transcription factor itself (55).Here, by dissecting the roles of Runx transcription factors in T celldevelopment, we find a strong counterexample to this prediction.The roles of Runx proteins in early T cell development have longbeen known to be important but were not clearly defined before.One problem was that the dominant factor, Runx1, was expressedvery broadly with little change across periods of strong tran-scriptome change; another was that Runx1 deletion in earlyT-lineage cells appeared to have only modest effects until laterstages. Here, RNA, protein, and functional data show that coex-pressed Runx1 and Runx3 work collaboratively and redundantly inindividual intrathymic pro-T cells before T-lineage commitment,providing a crucial input that activates or represses key genes todrive pro-T cell progression. Runx2, although not tested here,might also contribute during Phase1. Importantly, despite near-constant Runx activity levels, the target genes preferentially reg-ulated by Runx1 and Runx3 are stage specific and enriched forstrong expression changes across commitment. Runx action onthese targets reflects major redistributions of Runx binding acrossthe genome during commitment. A distinctive mode of Runx ac-tion is its stage-dependent, coordinate gain or loss of binding tomultiple neighboring sites in large genomic regions. Thus, Runxsite binding shifts, rather than changes in Runx expression, gen-erate momentous changes in T cell precursor gene expression.Paralogous factors Runx1 and Runx3 direct development and

function of various hematopoietic cells (3, 9, 12, 18, 46, 48, 56,57). If these factors were antagonists as in later thymocytes (46),decreasing Runx3:Runx1 ratios from Phase1 to Phase2 might bea switch between precommitment and postcommitment regula-tory patterns. Small differences between the RUNT1 andRUNT3 domains reportedly give Runx3 a stronger affinity forthe consensus Runx binding motif than Runx1 (27). Indeed,some of our results would support Runx3 acting more inde-pendently than Runx1 on Phase1 target sites. Conversely, finespecificity of binding by Runx1 may be more dependent onpartners. bHLH target motifs are enriched at Runx1-preferringsites, and partner Bcl11b may also interact preferentially withRunx1, even in an Innate Lymphoid Cell Type 2 contextexpressing more Runx3 than Runx1 (58). However, these smalldifferences between Runx1 and Runx3 were dwarfed in pro-Tcells by their coexpression, highly overlapping binding sites, andfunctional parallelism in activating T cell identity genes whilesuppressing alternative lineage genes.Although Runx1 and Runx3 together give pro-T cells nearly

constant levels of Runx activity across commitment, they preferen-tially regulate genes that change expression sharply between Phase1and Phase2. Furthermore, Runx target genes responded to Runxfactor perturbations with notably distinct stage sensitivities. Manygenes were Runx sensitive (for positive or negative regulation) onlyat one phase of their expression, often during a transition in theirexpression. This strongly recalled previous evidence for hit-and-runRunx activity at specific hematopoietic loci, where transient bindingof Runx1 to key target sites was sufficient to reorganize chromatinstructure and sustained presence of Runx1 was dispensable formaintaining chromatin accessibility (2). Thus, Runx1 + Runx3 madethe greatest difference for genes that were temporarily “destabilized”and poised for sharp developmental changes in expression. Thissuggests that Runx factors primarily regulate change rather thansteady-state expression levels in pro-T cells.A major contributor to this dynamism is the physical relocation

of Runx binding from an initial set of physiological genomic sitesto another set, often accompanied by different transcription factorpartners. We have previously shown that factor–factor interaction(“theft”) plays a role in this redistribution (14, 25), but not all

Phase1 sites are associated with the same “thieving” partner. BothPhase1 and Phase2 sites appear functional. Notably, also, some ofthe most highly enriched site–response associations suggest thattarget gene sensitivity to Runx perturbation is highest when Runxis making the site shift, more than when its configuration is stable.These results do not rule out indirect regulation, especially for

some classes of Runx-inhibited genes. Also, PU.1 or antagonismof E proteins could also cause indirect inhibition of some ap-parently Runx-dependent genes since loss of Runx factors inPhase2 unleashed Spi1 and ID protein expression (Id1, Id2, andId3). However, Runx binding overall and specific Runx siteclasses were clearly enriched with repressed as well as withactivated targets.Finally, the shifting of Runx factors between Phase1 and Phase2

showed striking group behavior. Multiple peaks of Runx bindingappeared or disappeared together across large genomic domains,such as Bcl11b distal enhancer regions, where >30 sites over >1 Mbdramatically appeared after T-lineage commitment. This suggeststhat Runx factors may recognize chromatin three-dimensionalstructures, although ATAC accessibility per se did not appear to bea major constraint. Notably, previous studies showed that Runx1regulates hematopoiesis by globally redistributing transcription factorsand remodeling chromatin structures itself (2, 59–61). Thus, as re-sponders or drivers, Runx factors are illuminating probes for theregulatory mechanisms that control remodeling of chromatin archi-tecture in the most transformative phases of cell-type development.

Materials and MethodsMouse strains, reagents, Cas9-mediated deletion, in vitro T-lineage differ-entiation, RNA-seq, ChIP-seq, and analytical methods used in this study wereessentially similar to those we have described previously (11, 14, 58, 62).Detailed methods and statistics are provided in SI Appendix. Newly gener-ated RNA-seq and ChIP-seq data for this study are deposited in the GeneExpression Omnibus (accession no. GSE154304). Mice were purchased fromthe Jackson Laboratory or generated by our laboratory at Caltech as de-scribed previously (54). Both male and female mice were used. All experi-ments shown were performed in at least two separate biological replicates(deep sequencing samples) or at least three independent experiments (allcell biological samples). Numbers of replicates, statistical tests, and P valuesare given in the figures.

Data Availability. ChIP-seq and RNA-seq data have been deposited in theGene Expression Omnibus (accession no. GSE154304).

ACKNOWLEDGMENTS. We thank Xun Wang (Caltech) and Joseph Lotem(Weizmann Institute) for helpful critiques of the manuscript; E. JanielleCuala and Suin Jo for preliminary experiments; Diana Perez, Jamie Tijerina,and Patrick Cannon of the Caltech Flow Cytometry and Cell Sorting Facilityfor cell sorting; Igor Antoshechkin and Vijaya Kumar of the Jacobs Geneticsand Genomics Center for sequencing; Henry Amrhein and Diane Trout forsequence curation and computer support; Ingrid Soto for animal care; MariaQuiloan for mouse breeding supervision; Rochelle Diamond for laboratorymanagement and sorting supervision; and the members of the group ofE.V.R. for sharing advice, reagents, and preliminary results. This work wassupported by Cancer Research Institute Irvington Postdoctoral FellowshipCRI.SHIN (to B.S.), the Japan Society for the Promotion of Science KAKENHIGrant JP19H03692 (to H.H.), The Mochida Memorial Foundation for Medicaland Pharmaceutical Research (H.H.), The Naito Foundation (H.H.), TheYasuda Medical Foundation (H.H.), the SENSHIN Medical Research Founda-tion (H.H.), the Takeda Science Foundation (H.H.), and US Public Health Ser-vice Grants R01AI135200 and R01HD076915 (to E.V.R.). V.R.T. was supportedby a Students Training in Advanced Research award from the University ofCalifornia, Davis. Flow cytometry, sequencing, and bioinformatics facilitysupport were from the Beckman Institute at Caltech. Support was also fromthe California Institute of RegenerativeMedicine Bridges to Stem Cell ResearchProgram (Pasadena City College and Caltech; M.R.-W.), the L. A. GarfinkleMemorial Laboratory Fund, the Al Sherman Foundation, and the Albert Bill-ings Ruddock Professorship (E.V.R.).

Shin et al. PNAS | 11 of 12Runx1 and Runx3 drive progenitor to T-lineage transcriptome conversion in mouse T cellcommitment via dynamic genomic site switching

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