A PML-PPARδ pathway for fatty acid oxidation regulates haematopoietic stem cell maintenance

Ppard plays a central role in HSC maintenance

The contribution of lipid catabolism to HSC function has not yet been defined. PPARs have been established as critical regulators of the transcriptional program underlying FAO, however. We therefore investigated whether PPAR and FAO could be relevant to HSC biology.

We first analysed the expression levels of the PPAR family in the HSC compartment, and found that Pparδ was the predominant receptor expressed there (Supplementary Figs. S1a, b). We next assessed whether PPARδ function played a role in HSC maintenance. To this end, we conditionally deleted Ppard in Kit^posSca^posLin^neg (KSL) cells ex vivo through retroviral Cre transduction and sorting of GFP-positive infected cells, in order to evaluate the characteristics of Ppard-deficient long-term HSCs (CD34^negKSL; Fig. 1a). We initially investigated Ppard-deleted HSC function in vivo in the context of bone marrow transplantation (BMT). Upon transplantation, Ppard loss did not affect homing of HSCs (Fig. 1b and Supplementary Fig. S1c), but profoundly impacted long-term repopulating capacity (Fig. 1c, d and Supplementary Fig. S1d). Critically, the defective repopulation capacity of Ppard-deleted HSCs was further evidenced in the second and third rounds of serial BMT (Fig. 1e, f). It is worth noting that no obvious differences in the expression of previously described genes involved in the maintenance of the HSC compartment were observed in Ppard-deleted HSCs (Supplementary Fig. S1e).

Consistent with these data, deletion of Ppard also led to a dramatic decrease in the function of HSCs in vitro, with reduced cobblestone area-forming cells (CAFC), a surrogate indicator of HSC maintenance (Supplementary Fig. S1f)²³ and colony forming ability in long-term culture (defined as long-term culture initiating cell assay, LTC-IC assay; Supplementary Fig. S1g–i), together with a profound reduction in ATP levels (Supplementary Fig. S1j). However, Cre expression itself did not have a significant effect on the proliferation and differentiation of stem/progenitor cells (Supplementary Fig. S1k).

It has been shown that several HSC-regulatory factors modulate the stem cells’ exit from quiescence; therefore, we evaluated the relevance of Ppard for the maintenance of a quiescent state. Interestingly, Ppard-deletion in the HSC resulted in exit from quiescence (Fig. 1g and Supplementary Fig. S1l).

PPARδ activators enhance HSC function

We then tested whether pharmacological PPARδ activation would lead to increased maintenance and function in vivo and in vitro. First, we treated 2-month-old mice with GW501516 (GW) daily for 28 days (Supplementary Fig. S2a), and observed that GW administration did not alter the proliferation, differentiation, or overall number of progenitor cells at the steady state, even as it increased long-term culture initiating cell (LTC-IC) capacity (Supplementary Fig. S2b–d and data not shown).

Next, we employed two different pharmacological activators of PPARδ at low doses in vitro (L165,041; L165 and GW)^24–26. As predicted, incubation with PPARδ activators increased the number of cobblestone areas (Supplementary Fig. S2e), in line with the increased expression of PPARδ targets (Supplementary Fig. S2f) and ATP levels (Supplementary Fig. S2g). Critically, we found that culturing HSCs in the presence of PPARδ agonists increased their ability to generate colonies of differentiated haematopoietic cells in LTC-IC assays (Supplementary Fig. S2h, i).

To provide a definitive proof of the potential benefit of PPARδ activators in the HSC compartment, we then tested the effect of GW in BMT. To this end, we transplanted 1.5x10³ KSL cells in lethally irradiated mice, and we subjected them to treatment with vehicle or GW daily during the period of HSC homing, lodgement and engraftment to the BM niche^27–29 (Fig. 2a). Six weeks after transplantation, when a major donor contribution to haematopoiesis is observed (data not shown), we harvested the BM and analysed the number of remaining HSCs. Treatment with GW significantly increased the number of long-term HSCs in the BM (Fig. 2b and Supplementary Fig. S2j). Notably, similar results were obtained with shorter treatments of 2-weeks with GW (Supplementary Fig. S2k, l). Importantly, GW treatment significantly increased long-term BM and multi-lineage haematopoietic reconstitution capacity after a second BMT of KSL cells or MNCs (Fig. 2c–e).

In order to demonstrate that PPARδ agonists did exert their beneficial activity through the selective cell autonomous activation of PPARδ in the HSCs, we treated Ppard-Wild type (WT) or Knockout (KO) HSCs with the PPARδ agonist GW. As predicted, Ppard-KO HSCs were refractory to the effect of GW in both CAFC and LTC-IC assays (Supplementary Fig. S3a, b), and the repopulation capacity of KSL cells and MNCs upon BM transplant (Fig. 2f and Supplementary Fig. S3c). Furthermore, in vitro treatment with GW in the absence of stromal cells still increased function of WT stem cells (Supplementary Fig. S4).

Taken together, these data reveal PPARδ as a critical druggable modulator of HSC maintenance and function in a haematopoietic stem cell-autonomous manner.

Inhibition of mitochondrial FAO impairs maintenance of the HSC

PPAR transcription factors are central regulators of nutrient sensing, metabolic reprogramming and differentiation¹¹. In particular, PPARδ, together with PPARα, plays a critical role in the sensing of fatty acids and the activation of the FAO transcriptional program^13,30. We therefore evaluated the requirement of active FAO for HSC maintenance.

We first measured FAO in both undifferentiated and differentiated haematopoietic cells. KSL cells exhibited detectable and measurable FAO, which was determined by incubating the cells in the presence or absence of maximal doses of Etomoxir (a pharmacological inhibitor of mitochondrial beta-oxidation of long-chain fatty acids, which does not affect the oxidation of short-chain FA nor peroxisomal FAO, 100μM) in the assay (Fig. 3a)^31,32. However, FAO analysis with the same cell number of lineage positive cells did not render a signal that was significantly over the counts obtained upon incubation with Etomoxir (Fig. 3a). Notably, FAO analysis with a greater number of differentiated cells (up to 2.5 fold more Lineage positive cells than KSL cells) yielded similar results (data not shown). This data is consistent with the hypothesis that PPARδ signalling is downregulated during the course of haematopoietic differentiation (Supplementary Fig. S5a).

To further evaluate whether mitochondrial FAO plays a role in stem cell biology, we first evaluated in vivo the impact of Etomoxir on the ability to repopulate the BM. Notably, we did not observe any toxic effect of Etomoxir in our experimental settings, in line with previous reports³³. Four-week treatment with Etomoxir after BMT in lethally irradiated mice led to a significant reduction in long-term HSCs in BM (Fig. 3b). Even a shorter treatment (of 2 weeks) with Etomoxir after BMT resulted in a profound alteration of long-term CD34^negFlt3⁻KSL cells in the BM and donor-derived cells in peripheral blood 24 weeks after transplantation (Fig. 3c,d and Supplementary Fig. S5b). Moreover, we observed reduced long-term BM and haematopoietic reconstitution capacity in secondary BMT with two different protocols of serial BMT (Fig. 3e–h). Importantly, both in vitro and in vivo, the doses of Etomoxir employed in these experiments initially increased the number of total HSCs, but did not affect cell survival, nor did they display toxicity in the HSC compartment (Supplementary Fig. S5c–e).

We also analysed the impact of pharmacological mitochondrial FAO inhibition in vitro. Incubation with Etomoxir led to a reduction in the number of cobblestone areas (Supplementary Fig. 5f). We then tested the impact of Etomoxir treatment on an LTC-IC assay, and found that Etomoxir triggered an opposite response to that observed with PPARδ activators, but similar to that observed in Ppard-deficient cells, i.e., an initial expansion (coherent with increased exit from quiescence) followed by exhaustion of the compartment (Supplementary Fig. S5g–i). In accord with previous data, Etomoxir also led to a reduction in levels of FAO and ATP (Supplementary Fig. S5j, k). Importantly, we confirmed the cell-autonomous effect of Etomoxir on sorted haematopoietic stem cells in vitro (Supplementary Fig. S4).

In order to test whether PPARδ activation-mediated enhancement of HSC function depended on mitochondrial FAO, we treated HSCs with GW, Etomoxir, or both. Interestingly, we observed that inhibition of mitochondrial FAO partially or completely prevented the beneficial effect of GW in vitro and in vivo (Supplementary Fig. S6a–e). We also investigated whether the effect of Etomoxir depended on a functional PPARD. In line with our data, Etomoxir had a reduced effect on HSC maintenance accompanied with ATP reduction in vitro and in vivo in the absence of Ppard (Supplementary Fig. S6f–h).

Taken together, these data uncover a role for FAO downstream of PPARδ in the regulation of HSC maintenance.

PPARδ activation rescues the maintenance defect of Pml-KO HSCs

We have previously shown that deletion of Pml results in loss of HSC maintenance, and consequently exhaustion of the HSC compartment (Supplementary Fig. S7a–d and Ref. 16). Surprisingly, we found that Pml-deficient HSCs exhibit reduced expression of PPARD targets (Fig. 4a; similar results were obtained using 36B4 as a housekeeping endogenous control; data not shown). This observation, together with the notion that PML regulates the activity of several transcriptional factors and cofactors¹⁷, led us to hypothesize that regulation of PPAR signaling and FAO by PML in the HSC might be of relevance to its maintenance, and that, in turn, the defect in maintenance observed in Pml-KO HSCs could be rescued by pharmacologically enhancing the activity of PPARδ.

We ascertained the contribution of PML to the function of PPAR signaling and FAO in cells from haematopoietic origin. As stated above, Pml-deficient HSCs exhibited reduced expression of PPAR targets (Fig. 4a) and ATP levels (Supplementary Fig. S7e). Mechanistically, we found that PML expression resulted in decreased acetylation of PGC1A, a critical co-factor of PPARs, in line with the notion that PML controls key regulators of the acetylation machinery of PGC1A³⁴ (Supplementary Fig. S7f). Importantly, Pml-deficient HSC exhibited diminished FAO compared to Pml-WT cells (Fig. 4b), whereas no difference was observed in differentiated cells (Supplementary Fig. S7g), in agreement with the thesis that Pml expression is markedly reduced during the course of haematopoietic differentiation.¹⁶ Taken together these data suggest that PML regulates PPAR signaling and FAO in the HSC.

In order to evaluate the contribution of PPAR signaling to the defect observed in Pml-deficient cells, and since Pparδ is the main PPAR receptor expressed in this compartment (Fig. 1a and Supplementary Fig. S1a), we tested whether pharmacological activation of Pparδ would rescue the loss of maintenance observed in these cells. We first tested whether PPARδ activators would do so when administered systemically in vivo. As reported, transplantation of Pml-KO HSCs with WT competitor cells in lethally irradiated mice led to the expansion-exhaustion of the Pml-KO compartment (Fig. 4c–e and Supplementary Fig. S8)^16,35. Strikingly, treatment of mice with GW for 14 days resulted in the prevention of the expansion phase and the amelioration of exhaustion in the Pml-KO compartment (Supplementary Fig. S8a,b). Consistent with this data, GW led to a significant recovery in the number of Pml-KO HSCs in the BM of recipient mice (Fig. 4c–e and Supplementary Fig. S8c–f). A treatment as short as two weeks after first transplantation in Pml-KO cells significantly increased the number of HSCs in the BM after secondary BMT (Supplementary Fig. S8g–i; see Supplementary Fig. S8a for the experimental design). Importantly, 4-week GW treatment of mice upon transplantation with Pml-KO HSCs (see experimental design in Fig. 4c) rescued the number of resident HSCs in the BM 6 weeks after transplantation, and ameliorated Pml-KO HSC function in secondary transplantation (Fig. 4d,e).

In vitro, PPARδ activators increased cobblestone area formation in Pml-KO HSCs (Fig. 4f and Supplementary Fig. S9a) to a greater extent than in WT HSCs (GW, 1.4 fold in WT versus 3.5 fold in KO and L165, 1.6 fold in WT versus 2.4 fold in KO), in line with the recovery in PPARδ signalling observed in conditions of GW treatment in Pml-KO HSCs (Supplementary Fig. S9b) and a recovery in ATP levels (Supplementary Fig. S9c). Strikingly, PPARδ activators normalized both the expansion and subsequent exhaustion phases observed in Pml-KO HSCs by LTC-IC assay in vitro (Fig. 4g and Supplementary Fig. S7a,b and S9d). These data collectively demonstrate that the defect observed in the Pml-KO HSCs arises, at least in part, from dysfunctional PPARδ signalling and FAO.

Pml-loss results in PPARδ-dependent loss of asymmetric division

We next asked what cellular outcomes might result from the beneficial effects of PML-PPARδ activation and FAO on HSC maintenance. Asymmetric division has recently been suggested as a regulator of cell fate decisions in the mammalian haematopoietic system, playing crucial roles in stem cell renewal⁷. Therefore we hypothesized that PML regulation of PPARδ signalling and FAO may be essential for asymmetric division.

We studied cell fate decision by utilizing a binary assay which takes advantage of two surface markers: Tie2 and CD48^2,36. In a purified CD150^posCD48^negCD41^negFlt3^negCD34^neg KSL population from wild type mice, we observed Tie2 positivity in up to 90% of cells (Fig. 5a). In single cell culture assays, after the first cell division, more than 40% of these cells asymmetrically divided to give rise to two distinct daughter cells (asymmetric division), Tie2^posCD48^neg (HSC) and Tie2^negCD48^pos (committed cell), while 20% provided two committed Tie2^negCD48^pos cells (symmetric commitment; Fig. 5b).

To assess the functional robustness of the assay, we investigated the functional asymmetry of daughter cells after the initial cell division. To this end, we employed a functional analysis of paired daughter cells in vitro by LTC-IC analysis (Fig. 5c). The two daughter cells after an initial cell division of CD150^posCD48^neg CD41^negFlt3^negCD34^neg KSL cells were separated, and their LTC-IC capacity was analyzed. Consistent with the immunophenotypical Tie2/CD48 characterization, the functional asymmetry, in which one daughter cell holds long-term repopulation capacity but another does not, was observed in more than 40% of divisions (Fig. 5c). Critically, we confirmed the functional asymmetry of daughter cells in vivo in BMT assays, where each daughter cell was transplanted into irradiated mice and its short-and long-term repopulating capacity were evalulated (Fig. 5d and Supplementary Fig. S10a and Supplementary Table 1, 2). Notably, the fact that symmetric divisions towards two long-term (LT)-HSCs are observed in vitro (through immunophenotyping), but not in in vivo functional assay (Supplementary Table 2), suggests that the production of LT-HSCs from a parental cell is a rather infrequent process, although there is potential for its occurrence.

After having validated the assay, we then investigated the effects of PPARδ activation and FAO on the cell fate decision of HSCs. Strikingly, Ppard-deletion resulted in a reduced asymmetric division rate and increased symmetric commitment, while no significant effect of Cre expression was observed in these experiments (Fig. 6a, b). Next, we corroborated the immunophenotypical asymmetric division features with functional analysis, and showed that Ppard-loss resulted in decreased function of paired daughter cells in vitro (Fig. 6c). We then tested whether Etomoxir would similarly affect asymmetric division, and we did indeed observe increased symmetric commitment and decreased asymmetric division in Etomoxir-treated HSCs (Fig. 6d). We validated this in vivo as well as in vitro in paired daughter cell assays (Fig. 6e, Supplementary Fig. S10b, and data not shown). In addition, and in line with our previous results, PPARδ activation with GW increased asymmetric division, an effect that was suppressed by pharmacological inhibition of mitochondrial FAO with Etomoxir (Fig. 6f). A higher dose of Etomoxir (20 μM) had a more profound effect than a lower dose of 10 μM, in line with a further reduction in FAO as well as FAO-derived ATP levels (Supplementary S10c, d), and in turn counteracted more effectively the action of GW. We also investigated the effect of another pharmacological FAO inhibitor, Perhexiline, on the division balance of HSCs. As predicted, Pehexiline treatment increased symmetric commitment, and importantly, counteracted the positive effect of GW treatment on asymmetric division (Supplementary Fig. S10e, f). These data provide a compelling explanation for the loss of maintenance observed in Etomoxir-treated HSCs, placing PPAR signaling and FAO at the regulatory core of asymmetric division and haematopoietic stem cell fate.

On this basis, we then tested whether Pml-loss would similarly result in increased symmetric commitment. As predicted, genetic or pharmacological (upon Arsenic trioxide treatment) depletion of Pml mimicked Ppard-loss and FAO inhibition, resulting in reduced asymmetric division and increased symmetric commitment (Fig. 6g and Supplementary Fig. S10g, h). Critically, the consequences of the aberrant division pattern alteration in HSCs were confirmed in vivo (Fig. 6h and Supplementary Fig. S10i–k). In agreement with our previous observations, the phenotype observed in Pml-KO HSCs was rescued upon treatment with the PPARδ agonist GW (Fig. 6i).

Finally, we evaluated whether the observed increase in symmetric commitment could be responsible for the loss of maintenance observed in vivo in Ppard and Pml-deleted HSCs. To this end, we analysed the fraction of CD48 positive cells within the CD34^negKSL fraction (Supplementary Fig. S11). In line with our hypothesis, Ppard and Pml deletion resulted in an initial increase of CD34^negKSL cells, with a clear enrichment in the fraction of cells that were positive for CD48. Conversely, GW treatment decreased the fraction of CD48 positive cells within the CD34^negKSL compartment of Pml-deficient cells, but had no effect on Ppard-deleted cells. As aforementioned, we have also found that loss of Pml or Ppard results in exit from quiescence without induction of apoptosis (Fig. 1g and Supplementary Fig. S1l, Fig. S5i, Fig. S7a,b, Fig. S10k, Fig. S12, and Ref ¹⁶). Therefore functional loss along the PML-PPARδ-FAO axis triggers loss in quiescence combined with excessive commitment, which results in the accumulation of committed HSCs (CD34^negCD48^posKSL) that fail to maintain haematopoiesis at later stages.

Mice

Generation of Pml-deficient mice has been described⁴⁷. C57BL/6 mice (B6-CD45.2) and C57BL/6 mice congenic for the CD45 locus (B6-CD45.1) were purchased from The Jackson Laboratory and crossed with 129Sv mice. F6 B6/Sv129 mice were used as recipients in transplantation assays as described¹⁶. GW501516 (5 mg per kg body weight per day; Enzo life sciences) and Etomoxir (25 mg per kg body weight, day ON day OFF; Sigma) were administered by oral gavage and i.p. injections, respectively. The animal protocol was reviewed and approved by the Beth Israel Deaconess Medical Center Institute Animal Care and Use Committee (IACUC). The Ppard floxed allele was generated as described previously⁴⁸. To delete Ppard in haematopoietic system, MSCV-Puro-IRES-GFP or Puro-IRES-GFP-Cre retroviral vectors were employed in purified KSL cells was utilized⁴⁹.

Reagents

GW501516 was dissolved for in vivo experiments as previously published³³, in Carboxymethylcellulose, and mice where treated with the GW solution or the Carboxymethylcellulose solution alone (vehicle). For in vitro experiments, GW and L165 were dissolved in DMSO, and DMSO was used in vehicle-treated cells. ATP levels in stem cell compartment was analysed by ATP lite luminescence ATP detection kit (Perkin Elmer, Inc., CA) according to manufacturer’s instruction.

Quantitative Real time PCR

Total RNA was extracted from cells using RNeasy kit (Qiagen) including DNAse digestion. cDNA was obtained with First-strand cDNA synthesis kit (USB). Taqman probes were obtained from Applied Biosystems and selected to avoid detection of genomic DNA. Amplifications were run in a 7900 Real-Time PCR System (Applied Biosystems). Each value was adjusted by using 36B4/Rplp0 or β-actin levels as reference.

Western blotting, immunoprecipitation and immunofluorescence

Cells were harvested and protein was extracted as previously described⁵⁰. Retroviral infection and acetylation experiments were performed as previously described^51,52. Briefly, 10⁶ K562 cells were transfected with Nucleofection reagent V (Lonza) using Amaxa nucleoporator, following manufacturers’ instructions (program T-16) and with the DNA ratio indicated above (3.5 μg of total DNA). After 24 hours, cells were lysed in 40 mM Tris (pH 7.6), 150 mM NaCl, 1 mM EDTA, 1 mM MgCl₂, 1% Triton X100, 1 mM sodium ortho-vanadate (Na3VO4), 1 mM NaF, 1 mM b-Glycerophosphate, 14 mM Nicotinamide, 400 nM trichostatin and protease inhibitor cocktail (Hoffmann–La Roche), cleared by centrifugation and subjected to immunoprecipitation with Flag-conjugated beads (Sigma) (in the same buffer with 0.1% Triton X100). After 2–3 hours, beads were washed, resuspended in laemmli loading buffer (Boston Bioproducts), boiled and the supernantant was subjected to SDS-PAGE. Proteins were detected using a rabbit polyclonal anti-PML antibody (Bethyl), anti-PGC1A antibody (H-300, Santa Cruz Biotechnology), anti-acetyl lysine (9441, Cell Signalling) and anti-β-actin monoclonal antibody (Sigma).

Fatty acid oxidation

Cells were incubated overnight in culture medium containing 100 μM palmitate (C16:0) and 1 mM carnitine. In the final 2 (MEFs) or 4 (Haematopoietic cells) hours of incubation, cells were pulsed for 4 hours with 1.7 μCi [9,10(n)-³H]palmitic acid (GE Healthcare) in the presence or absence of Etomoxir (100 μM, sigma), and the medium was collected to analyse the released ³H₂O, formed during cellular oxidation of [³H]palmitate^52,53. Briefly, medium was TCA precipitated, supernatants were neutralized with NaOH and loaded onto ion exchange columns packed with DOWEX 1X2-400 resin (Sigma). The radioactive product was eluted with water and quantitated by liquid scintillation counting. For haematopoietic cells, 70,000 cells (Fig. 3a and Fig. 4b) or 125,000 cells (Supplementary Fig. S7g) were assayed (as counted by sorting; the presence of an equal cell number at the time of the assay was validated by cell counting with haematocytometer for Fig. 4b). Etomoxir, a specific inhibitor of CPT1α, was used to specifically inhibit mitochondrial FAO. Alternatively, to determine FAO in small number of cells, oxygen consumption rate (OCR) immediately following the exposure of HSC to palmitate was used as an indicator of FAO in a XF96 Analyzer (Seahorse Bioscience)^54,55, where the appropriateness of the method is presented by the decrease in OCR by the incubation with increasing doses with the CPT1 inhibitor.

Flow cytometry

For flow cytometric analyses, we used monoclonal antibodies specific for the following: CD41 (eBioMWRag30), Flt-3 (Avas12a1), CD34 (RAM34), c-Kit (2B8), Sca-1 (E13-161.7), CD3e (145-2C11), CD4 (L3T4), CD8 (53-6.72), B220 (RA3-6B2), TER-119 (TER-119), Gr-1 (RB6-8C5), CD34 (RAM34), CD11b (M1/70), IgM (II/41), CD19 (eBio1D3) and NK-1.1 (PK136); all were from eBioscience. Anti-CD150 (TC15-12F12.2) and CD48 (HM48-1) antibodies were from BioLegend. We used a mixture of monoclonal antibodies against CD4, CD8, CD3e, B220, TER-119, CD11b, Gr-1, IgM, CD19 and NK-1.1 as a lineage marker (Lineage). We performed Pyronin Y staining as previously described².

Long-term cultures and colony-forming assays

For long-term cultures, we co-cultured 50 CD34^negKSL cells with OP9 cells in Minimum Essential Media, alpha modification (αMEM Sigma) containing 12.5% FCS (JRH Bioscience), 12.5% horse serum (Gibco BRL) and 1 nM dexamethasone. After 2, 4, 8 or 12 weeks of culture, we harvested cells and used them for haematopoietic colony forming assays as described^2,16. Frequencies of CAFC and LTC-IC were calculated by limiting-dilution method (maximum likelihood from the proportion of negative wells measured for each input dilution of cells tested)⁴⁷. In each experiment, 36 (CAFC) and 12 (LTC-IC) replicates of each dilution were performed, and a split-plot ANOVA analysis is utilized to investigate the difference between two conditions. For treatment experiments, 0.1 μM GW501516 (Enzo life sciences), 0.5 μM L165,041 (Sigma), 10 or 20 μM Etomoxir (Sigma), or 2 μM Perhexiline (Sigma) was added to cultures.

Competitive reconstitution assay

We sorted 1.5 x 10³ KSL cells from wild-type or Pml^−/− mice (CD45.2). We transplanted cells into lethally irradiated CD45.1 congenic mice in competition with 2 x 10⁵ bone marrow mononuclear cells from CD45.1 mice. We monitored reconstitution of donor (CD45.2) cells by staining blood cells with antibodies against CD45.2 and CD45.1. For treatment experiments, recipient mice were administered with GW501516 by oral gavage (everyday) and Etomoxir i.p. (every other day). For serial transplantation, donor-derived bone marrow mononuclear cells or KSL cells were collected from recipient mice 6, 12 or 24 weeks after BMT (first BMT) and transplanted into recipient mice with fresh competitor cells (second BMT). Calculation of competitive reconstitution unit (CRU) was conducted using L-Calc software (StemCell Technologies.

Asymmetric division and Paired daughter cell assays

Single cell deposition of CD150^posCD48^negCD41^negFlt3^negCD34^negKSL cells from 8-week old mice (total 160–320 cells/experiment) were directly performed onto retronectin-coated slides and cells were culture in StemSPAN (StemCell Technologies) supplemented with 50 ng/ml stem cell factor (SCF) and 50 ng/ml thrombopoietin (TPO). Cells were either maintained untreated or treated with vehicle, 0.1 μM GW501516 (Enzo life sciences) and/or 10 μM Etomoxir (Sigma) or 0.15 μM As₂O₃ (Sigma). Close to 50% of the cells (49.7±5.7 in 100 randomly selected cells) undergo initial cell division at day 3. Cells were stained with anti-mouse Tie2 antibody, anti-mouse CD48 antibody and DAPI at day 3. Student’s t test (as indicated in the figure) was utilized to determine statistical significance, and statistical significance in division pattern as well as the disturbance of division pattern were also confirmed by log-linear model and Chi-square test (data not shown). For paired daughter cell assay, the corresponding daughter cells derived from a single cell were separated followed by LTC-IC assay or BMT assay as previously described⁵⁶. Single daughter cell is transplanted into lethally irradiated recipient mice in a competitive assay. Long-term repopulation and short-term repopulation are achieved when more than 1% of haematological contribution in peripheral blood of recipient mice is observed, 16 weeks and 4 weeks after BMT respectively⁵⁷.

Homing analysis

Lethally irradiated Ly45.1 mice were transplanted with 2,000 CD34^negKSL cells from Ppard^F/F mice (Ly45.2) infected with Empty or Cre vector and 2x10⁵ competitor BMMNCs. Cells from the BM of recipient mice were analysed 24 hours post-transplantation to identify their derivation.

Statistical Analysis

Student’s t test was utilized to determine statistical significance unless otherwise specified. P values lower than 0.05 were considered statistically significant.