Mutant p53 promotes clonal hematopoiesis by generating a chronic inflammatory microenvironment

Sisi Chen; Sergio Barajas; Sasidhar Vemula; Yuxia Yang; Ed Simpson; Hongyu Gao; Rudong Li; Farzaneh Behzadnia; Sarah C. Nabinger; David A. Schmitz; Hongxia Chen; Wenjie Cai; Shiyu Xiao; Ruyue Luo; Mohammed Abdullahel Amin; Maegan L. Capitano; James P. Ropa; Aidan Fahey; Shuyi Zhou; Tiffany M. Mays; Magdalena Sotelo; Hao Pan; Sophie K. Hu; Sophia Veranga; Moiez Ali; Maria Shumilina; Reuben Kapur; Kehan Ren; Yuzhi Jia; Huiping Liu; Irum Khan; Yasmin Abaza; Jessica K. Altman; Elizabeth A. Eklund; Lucy A. Godley; Christine R. Zhang; Peng Ji; Seth L. Masters; Ben A. Croker; H. Scott Boswell; George E. Sandusky; Zhonghua Gao; Lindsey D. Mayo; Sharon A. Savage; Stephanie Halene; Yali Dou; Leonidas C. Platanias; Madina Sukhanova; Yunlong Liu; Omar Abdel-Wahab; Yan Liu

doi:10.1172/JCI184285

Mutant p53 promotes clonal hematopoiesis by generating a chronic inflammatory microenvironment

Sisi Chen,^1,2 Sergio Barajas,^3,4 Sasidhar Vemula,⁵ Yuxia Yang,^5,6 Ed Simpson,⁷ Hongyu Gao,⁷ Rudong Li,⁷ Farzaneh Behzadnia,⁷ Sarah C. Nabinger,⁵ David A. Schmitz,⁵ Hongxia Chen,^3,8 Wenjie Cai,^3,4 Shiyu Xiao,³ Ruyue Luo,³ Mohammed Abdullahel Amin,³ Maegan L. Capitano,⁹ James P. Ropa,⁹ Aidan Fahey,⁵ Shuyi Zhou,¹ Tiffany M. Mays,³ Magdalena Sotelo,³ Hao Pan,³ Sophie K. Hu,³ Sophia Veranga,³ Moiez Ali,³ Maria Shumilina,³ Reuben Kapur,⁵ Kehan Ren,¹⁰ Yuzhi Jia,¹¹ Huiping Liu,^11,12 Irum Khan,^3,12 Yasmin Abaza,^3,12 Jessica K. Altman,^3,12 Elizabeth A. Eklund,^3,12,13 Lucy A. Godley,^3,12 Christine R. Zhang,^10,12 Peng Ji,^10,12 Seth L. Masters,¹⁴ Ben A. Croker,¹⁵ H. Scott Boswell,¹⁶ George E. Sandusky,¹⁷ Zhonghua Gao,¹⁸ Lindsey D. Mayo,⁵ Sharon A. Savage,¹⁹ Stephanie Halene,²⁰ Yali Dou,²¹ Leonidas C. Platanias,^3,12,13 Madina Sukhanova,^10,12 Yunlong Liu,⁷ Omar Abdel-Wahab,² and Yan Liu^3,12

Authorship note: SC, SB, S Vemula, and YY contributed equally to this work.

Published December 30, 2025 - More info

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Abstract

Older individuals with somatic TP53 mutations manifest clonal hematopoiesis (CH) and are at high risk of developing myeloid neoplasms. However, the underlying mechanisms are not fully understood. Here, we show that inflammatory stress confers a competitive advantage to p53 mutant hematopoietic stem and progenitor cells (HSPCs) by activating the NLRP1 inflammasome and increasing the secretion of pro-inflammatory cytokines such as IL-1β, inhibiting WT HSPC fitness in a paracrine fashion. During aging, mutant p53 dysregulates pre-mRNA splicing in HSPCs, leading to enhanced NF-κB activation and increased secretion of IL-1β and IL-6, thereby generating a chronic inflammatory bone marrow microenvironment. Furthermore, blocking IL-1β with IL-1β neutralizing antibody or inhibiting IL-1β secretion using gasdermin D inhibitor decreases the fitness of p53 mutant HSPCs. Thus, our findings uncover an important role for mutant p53 in regulating inflammatory signaling in CH and suggest that curbing inflammation may prevent the progression of TP53-mutant CH to myeloid neoplasms.

Introduction

Human aging is associated with increased occurrence of clonal hematopoiesis (CH) both with normal blood counts (CH of indeterminate potential [CHIP]) and with clonal cytopenias of unknown significance (CCUS), where mutated hematopoietic stem and progenitor cells (HSPCs) undergo clonal expansion in older individuals without evidence of frank neoplasia (1–4). CH is associated with increased risks of developing myelodysplastic syndromes (MDSs) and acute myeloid leukemia (AML) (5–7). However, the underlying mechanisms are not fully understood.

Chronic inflammation has been implicated in cancer initiation, progression, and metastasis (8). Chronic and low-grade inflammation develops during aging (9, 10), which may contribute to the pathogenesis of CH and MDS (11, 12). Indeed, some older individuals with CH display an aberrant systematic inflammatory milieu compared with older individuals without CH (13, 14). Inflammatory stress has been implicated in the expansion of Tet2, Dnmt3a, and Asxl1 mutated HSPCs in animal models of CH (15–17). However, whether chronic inflammation promotes the progression of CH to myeloid neoplasms is largely unexplored.

The tumor suppressor gene TP53 is frequently mutated in human cancer (18, 19). Acquired somatic gain-of-function mutant p53 proteins, such as p53^R248W and p53^Y220C, were identified in patients with CH (5–7). We and others discovered that genotoxic stresses, including radiation and chemotherapy, promote TP53-mutant HSPC expansion (20–23). Mechanistically, mutant p53 drives CH through modulating epigenetic pathways (24). Somatic TP53 mutations in CH are associated with an increased incidence of MDS and AML (5–7). We found that mutant p53 promotes leukemia development through enhancing leukemia-initiating cell self-renewal (25). TP53 mutations are present in 10%–15% of primary and in 20%–30% of secondary MDS cases (26, 27). CH and, in particular, MDS are frequently caused by mutations in key factors of the spliceosome, including SF3B1, SRSF2, U2AF1, and ZRSR2 (28, 29). Altered splicing has been implicated in pathogenesis of hematological malignancies (28–30). Notably, both older human and mouse HSPCs display dysregulated pre-mRNA splicing (31–33). However, whether mutant p53 alters pre-mRNA splicing in HSPCs and drives MDS development is not known.

In this study, we report that mutant p53 increases chromatin accessibility to the NLRP1 inflammasome gene and dysregulates pre-mRNA splicing in key regulators of NF-κB in HSPCs, thereby generating a chronic inflammatory microenvironment to promote CH.

Results

Inflammatory stress confers a competitive advantage to p53 mutant HSPCs. Both viral and bacterial infections activate inflammatory response and induce chronic inflammation (8). The double-stranded RNA mimetic polyinosinic-polycytidylic acid (pI:pC) mimics the viral genome of many RNA viruses, and prolonged pI:pC treatment induces chronic inflammation in mice (34). To determine whether viral infection expands p53 mutant HSPCs in vivo, we performed competitive bone marrow (BM) transplantation assays. Eight weeks following transplantation, recipient mice were treated with PBS or pI:pC twice a week for 4 weeks (Figure 1A). While pI:pC treatment had no effect on the engraftment of WT BM cells, it significantly increased the engraftment of p53 mutant cells in peripheral blood (PB) of the recipient mice compared with those treated with PBS (Figure 1B). Given that pI:pC induces DNA damage in hematopoietic cells (34) and the canonical function of p53 is a mediator of DNA damage response (18, 19), we examined γH2AX in PBS- or pI:pC-treated Lin⁻ BM cells. p53 mutant HSPCs displayed more γH2AX foci compared with WT HSPCs at steady state. However, pI:pC treatment increased the number of γH2AX foci in WT HPSCs but decreased H2AX phosphorylation in mutant cells (Supplemental Figure 1, A and B; supplemental material available online with this article; https://doi.org/10.1172/JCI184285DS1), suggesting that pI:pC stimulation enhances DNA damage repair in mutant HSPCs.

Figure 1

Inflammatory stress confers a competitive advantage to p53 mutant HSPCs. (A) Schematic illustration of experimental design where recipient mice underwent competitive transplantation and were treated with either PBS or pI:pC (25 μg/kg, i.p.). MNC, mononuclear cell; TBI, total body irradiation. (B) Percentage of donor-derived cells (CD45.2⁺) in PB of recipient mice at 16 weeks following PBS or pI:pC treatment; n = 5 mice per group. (C) Schematic illustration of experimental design where recipient mice underwent competitive transplantation and were treated with PBS or LPS (1 mg/kg, i.p.). (D) Percentage of donor-derived cells in PB of recipient mice following PBS or LPS treatment; n = 6–8 mice per group. (E) Schematic illustration of experimental design where BM from primary mice treated with PBS or LPS (1 mg/kg, i.p.) were used for competitive transplantation. (F) Percentage of donor-derived cells in PB of recipient mice; n = 10–15 mice per group. (G) The frequency of donor-derived cells in PB of secondary recipient mice; n = 9 mice per group. (H) The frequency of donor-derived LSKs in the BM of secondary recipient mice; n = 9 mice per group. (I) The number of donor-derived LSKs in the BM of secondary recipient mice; n = 9 mice per group. (J) p53^+/+ and p53^R248W/+ macrophages were stimulated with LPS (100 ng/mL), and the levels of secreted cytokines and chemokines were assayed using a mouse 31-Plex Cytokine/Chemokine array; n = 4 biological replicates. (K) p53^+/+ and p53^R248W/+ mice were treated with LPS (1 mg/kg, i.p.), and PB serum was collected on days 1 and 3 after LPS treatment. The levels of cytokines and chemokines were assayed by Cytokine/Chemokine array; n = 4 mice per group. The comparison among multiple groups was evaluated by 1-way ANOVA (B and G–I) or 2-way ANOVA (D and F). The comparison between 2 groups was evaluated by 2-tailed t test (J and K). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

LPS is a ligand for TLR4, and LPS treatment activates inflammatory response in hematopoietic cells, promoting HSPC proliferation while reducing competitive fitness (35). To determine the impact of LPS on p53 mutant HSPCs, we performed competitive BM transplantation assays to determine whether LPS treatment expands p53 mutant HSPCs in vivo. Four weeks following transplantation, recipient mice were treated with LPS or PBS every other day for 4 weeks (Figure 1C). We found that LPS treatment significantly increased the engraftment of p53^R248W/+ cells in PB of recipient mice compared with those treated with PBS (Figure 1D).

To determine the impact of LPS on HSPC function in vivo, we treated p53^+/+ and p53^R248W/+ mice with LPS or PBS every other day for a month. We then isolated live BM cells from LPS- or PBS-treated donor mice and performed competitive transplantation assays (Figure 1E). Consistent with a previous report (35), LPS-treated WT BM cells showed decreased engraftment in PB and BM in both primary and secondary transplantation assays, whereas LPS-treated p53 mutant BM cells continued to show increased engraftment (Figure 1, F and G, and Supplemental Figure 1, C and D). Importantly, LPS treatment increased both the frequency and absolute number of p53 mutant LSKs in the BM of secondary recipient mice compared with PBS treatment (Figure 1, H and I).

To understand how LPS-induced inflammatory stress promotes p53 mutant HSPC expansion, we examined the impact of mutant p53 on cytokine secretion from BM-derived macrophages (36). We treated p53^+/+ and p53^R248W/+ macrophages with 100 ng/mL LPS for 12 h to induce pro-inflammatory activation. Supernatants were then collected and subjected to cytokine array analysis. Notably, p53 mutant macrophages showed increased secretion of multiple pro-inflammatory cytokines, including IL-1β, IL-1α, IL-12, KC, and CXCL-9, compared with WT macrophages following LPS stimulation (Figure 1J and Supplemental Figure 1E). Next, we treated p53^+/+ and p53^R248W/+ mice with 1 dose of LPS and examined cytokine and chemokine in PB serum of mice on day 1 (24 h) and day 3 (72 h) after LPS treatment. We observed increased levels of IL-1β in PB serum of p53^R248W/+ mice compared with that of the p53^+/+ mice on both days 1 and 3 after LPS treatment (Figure 1K). Thus, we demonstrated that mutant p53 enhances LPS-induced inflammatory response.

p53 mutant HSPCs show enhanced NLRP1 activation. To investigate how mutant p53 regulates inflammatory response in HSPCs, we performed ATAC-seq (assay for transposase-accessible chromatin with sequencing) assays in LSK cells to identify differential regions of chromatin accessibility (37). There were approximately 600 peaks significantly upregulated in p53^R248W/+ LSKs compared with p53^+/+ LSKs (Figure 2A). Furthermore, unbiased DAVID (Database for Annotation, Visualization, and Integrated Discovery) pathway analysis of ATAC peaks significantly upregulated in p53^R248W/+ LSKs revealed enrichment in hematopoietic stem cell (HSC) maintenance and self-renewal pathways (Figure 2B) (38). On the contrary, the p53 pathway, canonical Wnt signaling, and negative regulation of immune system process were enriched in p53^+/+ LSKs (Supplemental Figure 2A). To characterize chromatin regions with differential accessibility in p53^+/+ and p53^R248W/+ LSKs, we performed a motif search of the gained openly accessible peaks in p53^R248W/+ LSKs (39). Motif analysis identified a strong enrichment for transcription factor binding sites, including CTCF, ETS1, MEIS1, and FOXO1, in p53^R248W/+ LSKs (Figure 2C), some of which are known regulators of HSPCs.

Figure 2

p53 mutant HSPCs show increased NLRP1 activation. (A) LSK cells were subjected to ATAC-seq assays to identify differential regions of chromatin accessibility. Heatmap shows differential active regions in p53^+/+ and p53^R248W/+ LSKs. (B) Enriched biological processes of genes associated with significantly upregulated ATAC-seq peaks in p53^R248W/+ LSKs analyzed by DAVID Functional Annotation Tool. (C) Enrichment of transcription factor (TF) motifs in peaks that gain open accessibility in p53^R248W/+ LSKs compared with p53^+/+ LSKs determined by HOMER de novo Known Motif Analysis. (D) Nlrp1a and Nlrp1b acquired significant open chromatin peaks in p53^R248W/+ LSKs. (E) Nlrp1a and Nlrp1b were significantly upregulated in p53^R248W/+ LSKs compared with p53^+/+ LSKs; n = 3 biological replicates. (F) The level of Nlrp1b was increased in p53^R248W/+ Lin⁻ cells. (G) The levels of Nlrp1b and Nlrp3 in BM-derived macrophages following LPS stimulation were determined by immunoblotting. (H) GFP or p53^R248W was introduced into Nlrp1^+/+ and Nlrp1^−/− Lin⁻ BM cells using retrovirus. Transduced cells (GFP⁺) were differentiated into macrophages. Macrophages were stimulated with LPS (100 ng/mL) for 12 hours, and the levels of secreted cytokines and chemokines were assayed using a Cytokine/Chemokine array; n = 4 biological replicates. The comparison among multiple groups was evaluated with 1-way ANOVA (E and H). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

Mutant p53 increases chromatin accessibility to chemokine gene Cxcl9 (Supplemental Figure 2B). Indeed, Cxcl9 is upregulated in p53 mutant HSPCs (Supplemental Figure 2C). Notably, mutant p53 increases the chromatin accessibility to inflammasome genes, including Nlrp1a and Nlrp1b (Figure 2D), and we confirmed the upregulation of both Nlrp1a and Nlrp1b in p53^R248W/+ LSKs compared with p53^+/+ LSKs (Figure 2E). While NLRP3 inflammasome is activated in HSPCs from low- to high-risk MDS (40), mutant p53 affects neither the chromatin accessibility nor the expression of Nlrp3 in LSKs (Figure 2, D and E). Inflammasomes are multiprotein complexes that activate caspase-1 and increase the release of pro-inflammatory cytokines such as IL-1β, leading to caspase-1–dependent death, known as pyroptosis (41). NLRP1 was one of the first inflammasome-forming pattern recognition receptors to be identified (42, 43). To examine inflammasome activation in HSPCs, we treated p53^+/+ and p53^R248W/+ Lin⁻ cells with LPS and performed immunoblotting analysis to examine the levels of Nlrp1b and Nlrp3. We observed increased levels of NLRP3 and active NLRP1B (NLRP1B-UPA-CARD) in mutant HSPCs at baseline and following LPS stimulation, whereas LPS stimulation increased NLRP3 levels in both p53^+/+ and p53^R248W/+ HSPCs (Figure 2F). In addition, we found increased levels of NLRP1B-UPA-CARD in mutant macrophages compared with WT cells (Figure 2G). Consistent with our findings in HSPCs, LPS stimulation increased NLRP3 levels in both p53^+/+ and p53^R248W/+ macrophages (Figure 2G). LPS stimulation also increased levels of cleaved caspase-1 in mutant macrophages compared with WT cells, whereas LPS increased levels of mature gasdermin D (GSDMD-NT) and cleaved caspase-3 in both p53^+/+ and p53^R248W/+ HSPCs (Supplemental Figure 2D). NLRP1 is known to sense and be activated by dsRNA (42). To determine the involvement of the cGAS/STING pathway in NLRP1 activation in mutant cells, we treated p53^+/+ and p53^R248W/+ Lin⁻ cells with pI:pC and examined levels of cGas and Sting. pI:pC stimulation increased the levels of cGas and Sting but decreased levels of p-Sting in both p53^+/+ and p53^R248W/+ cells (Supplemental Figure 2E).

To determine the contribution of NLRP1 to the clonal advantage of p53 mutant HSPCs, we crossed Nlrp1^−/− mice with p53^R248W/+ mice. As Nlrp1 and Trp53 are close to each other on mouse chromosome 11, we failed to generate Nlrp1^−/− p53^R248W/+ mice (43). We then ectopically expressed GFP or p53^R248W in WT and Nlrp1^−/− Lin⁻ BM cells using retroviruses. Transduced cells (GFP⁺) were differentiated into macrophages and stimulated with LPS for 12 h. Macrophage supernatants were then subjected to multiplex cytokine analysis. Loss of Nlrp1 significantly decreased the secretion of multiple pro-inflammatory cytokines, including IL-1β, IL-1α, IL-12, and KC, from p53 mutant macrophages (Figure 2H). Thus, we demonstrated that p53 mutant HSPCs show enhanced NLRP1 activation and that NLRP1 is a key mediator of mutant p53-driven inflammation in hematopoietic cells.

Resistance to inflammatory stress underlies enhanced fitness seen in p53 mutant HSPCs. To further understand how inflammatory stress confers a competitive advantage to p53 mutant HSPCs, we performed in vitro cell competition assays (Figure 3A). WT competitor cells (CD45.1⁺) show decreased competitiveness after co-culture with p53 mutant cells (CD45.2⁺) for 7 days (Figure 3B). Importantly, WT CD45.1⁺ cells co-cultured with p53 mutant cells show decreased replating potential compared with WT CD45.1⁺ cells co-cultured with WT CD45.2⁺ cells (Figure 3C). However, the number of apoptotic WT CD45.1⁺ cells were comparable between WT and mutant groups (Figure 3D). To understand how p53 mutant cells inhibit the fitness of WT competitor cells in vitro, we measured the levels of cytokines in the condition media utilizing cytokine arrays and observed increased levels of IL-1β, IL-6, and CXCL-9 in mutant cell cultures compared with that of the WT cells (Figure 3E). Notably, WT CD45.1⁺ cells co-cultured with mutant cells showed increased pyroptosis (Figure 3F and Supplemental Figure 3A). Importantly, the enhanced pyroptosis of WT CD45.1⁺ cells seen in the mutant group appeared to be IL-1β dependent, as adding IL-1β to the culture further increased pyroptosis, whereas adding IL-1β–neutralizing antibody brought the level of pyroptosis back to normal (Figure 3F).

Figure 3

Resistance to inflammatory stress underlies enhanced fitness seen in p53 mutant HSPCs. (A) Schematic of in vitro cell competition assays using BM cells from p53^+/+ and p53^R248W/+ mice (CD45.2⁺) and WT competitor cells (CD45.1⁺). Three groups of BM cells, CD45.1, CD45.1/WT, and CD45.1/p53 mutant, were cultured in serum-free medium supplemented with SCF, TPO, FLT3 ligand, IL-3, and GM-CSF. 7 days later, live CD45.1⁺ cells from each group were sorted and used for proliferation, serial replating, apoptosis, and pyroptosis assays. In addition, cytokines and chemokines in the conditioned media were assayed by Cytokine/Chemokine array. (B) Proliferation of live CD45.1⁺ cells from 3 cell culture groups shown in A; n = 3 independent experiments performed in duplicate. (C) Serial replating assays of live CD45.1⁺ cells purified from 3 cell culture groups shown in A; n = 3 independent experiments performed in duplicate. (D) CD45.1⁺ cells purified from 3 cell culture groups were assayed for apoptosis; n = 3 independent experiments. (E) Increased levels of IL-1β, IL-6, and CXCL-9 were observed in the conditioned media from the CD45.1/p53 mutant group compared with the CD45.1/WT cell group; n = 3 independent experiments performed in duplicate. (F) Pyroptosis assays of CD45.1⁺ cells purified from 3 cell culture groups in the presence of PBS, IL-1β, or IL-1β–neutralizing antibody; n = 3 independent experiments. (G) Serial replating assays of p53^+/+ and p53^R248W/+ BM cells in the presence or absence of IL-1β; n = 3 independent experiments performed in duplicate. (H and I) MYC targets (H) and inflammatory response (I) gene signatures were significantly upregulated in IL-1β–treated p53^R248W/+ LSKs compared with IL-1β–treated p53^+/+ LSKs; n = 3 biological replicates. The comparison among multiple groups was evaluated with 1-way ANOVA (B, D, and E) or 2-way ANOVA (C, F, and G). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

IL-1β is known to exert suppressive effects on WT HSPC activity (44, 45). We found that IL-1β treatment decreased the replating potential of WT BM cells, but not that of p53 mutant cells (Figure 3G). To determine the mechanism by which p53 mutant HSPCs are resistant to IL-1β treatment, we performed RNA-seq in p53^+/+ andp53^R248W/+ LSKs treated with IL-1β (10 ng/mL) or PBS in vitro for 90 minutes (Supplemental Figure 3B). GSEA revealed that HSPC differentiation genes are significantly upregulated in p53 mutant LSKs compared with WT LSKs following IL-1β stimulation (Supplemental Figure 3C). Notably, p53 mutant LSKs treated with IL-1β are enriched with MYC target genes (Figure 3H and Supplemental Figure 3D). IL-1β treatment also increased the expression of genes of the inflammatory response pathway, including IFN-α, IL-6, and IFN-γ (Figure 3I and Supplemental Figure 3E).

The NLRP1/IL-1β/GSDMD axis is critical for p53 mutant HSPC fitness. To determine the role of Nlrp1 in p53 mutant HSPCs, we performed competitive transplantation assays using Nlrp1^+/+ and Nlrp1^−/− Lin⁻ cells expressing GFP or mutant p53. We found that loss of Nlrp1 significantly decreased the repopulating potential of HSPCs expressing mutant p53 in vivo (Figure 4, A and B). To determine the role of Il-1b in p53 mutant HSPCs, we generated Il-1b^−/− p53^R248W/+ mice (46). While Il-1b deficiency did not affect colony formation of WT HSPCs, loss of Il-1b significantly decreased the colony formation of p53 mutant HSPCs (Figure 4C). IL-1β has been shown to be a growth factor for AML cells (47), and we found that IL-1β–neutralizing antibody treatment (48, 49) decreased the replating potential of p53 mutant BM cells compared with p53 mutant cells treated with IgG (Figure 4D).

Figure 4

The NLRP1/IL-1β/GSDMD signaling axis is important for p53 mutant HSPC fitness. (A) Competitive transplantation experiments using Nlrp1^+/+ and Nlrp1^−/− Lin⁻ BM cells expressing GFP or p53^R248W. The frequency of donor-derived cells (CD45.2⁺) in PB of recipient mice was examined every 5 weeks for 15 weeks; n = 7–10 mice per group. (B) Loss of Nlrp1 decreases the engraftment of p53 mutant HSPCs in the BM of recipient mice 15 weeks after transplantation; n = 7–10 mice per group. (C) Loss of Il-1b decreases the colony formation of p53 mutant HSPCs; n = 3 biological replicates. (D) Serial replating assays of p53^+/+ and p53^R248W/+ BM cells in the presence of IgG or IL-1β–neutralizing antibody; n = 3 independent experiments performed in duplicate. (E) p53^+/+ and p53^R248W/+ BM cells were treated with DMSO or inflammasome activator VbP for 24 h and assayed for pyroptosis; n = 3 independent experiments. (F) VbP treatment enhances GSDMD maturation in BM cells. (G) GSDMD inhibitor dimethyl fumarate (DMF) treatment decreases the colony formation of p53 mutant BM cells; n = 3 independent experiments. The comparison among multiple groups was evaluated with 1-way ANOVA (B, C, E, and G) or 2-way ANOVA (A and D). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

Given that NLRP1 activation increases IL-1β secretion, inducing pyroptosis of WT HSPCs (43), we treated p53^+/+ and p53^R248W/+ BM cells with Val-boroPro (VbP), a small molecule activator of inflammasomes (50), and performed pyroptosis analysis. VbP treatment significantly increased pyroptosis of WT BM cells compared with p53 mutant BM cells (Figure 4E). Cytosolic sensing of pathogens assembles inflammasomes, which activate inflammatory caspases to IL-1β and GSDMD. Cleaved GSDMD forms membrane pores, leading to cytokine release and pyroptosis (51, 52). p53 mutant BM cells show increased GSDMD maturation, which is further enhanced by VbP treatment (Figure 4F). While GSDMD inhibitor dimethyl fumarate treatment had no effect on WT BM cells (53), it significantly decreased the colony formation of p53 mutant BM cells (Figure 4G). Collectively, our data suggest that the NLRP1/IL-1β/GSDMD signaling axis is important for the enhanced fitness of p53 mutant hematopoietic cells.

Heterozygous p53 mutant mice developed MDS with age. Most homozygous p53^−/− and p53^R248W/R248W mice develop spontaneous tumors, including lymphoma, thymoma, and sarcoma, and die within 3 to 6 months after birth (54). We aged p53^+/+ and heterozygous p53^R248W/+ mice for more than 1 year and monitored their survival and tumor development. p53^R248W/+ mice showed reduced survival compared with p53^+/+ mice and died within 15–18 months after birth (Figure 5A). Approximately 60% of p53^R248W/+ mice developed MDS based upon pathological analysis of BM and PB smears, while the remainder of p53^R248W/+ mice developed lymphoma and sarcoma (Figure 5B). Aged p53^R248W/+ mice showed hypercellular BM compared with age-matched p53^+/+ mice (Supplemental Figure 4A). Specifically, aged p53^R248W/+ mice showed morphological dysplasia on myeloid precursors, erythroid precursors, and megakaryocytes in the BM (Figure 5, C–I). We also observed morphological dysplasia on p53^R248W/+ PB smears, including Pseudo-Pelger-Huët, hypersegmented neutrophils, and nucleated red cells (Supplemental Figure 4B). Significant leukopenia, thrombocytopenia, and anemia were seen in aged p53^R248W/+ mice with MDS compared with age-matched p53^+/+ mice (Figure 5J). p53^R248W/+ mice also show splenomegaly (Supplemental Figure 4, C and D). Moreover, the spleens of aged p53^R248W/+ mice have increased frequency of HSPCs (Supplemental Figure 4E). Collectively, the p53^R248W/+ mice developed MDS phenotypes, recapitulating some clinical situations seen in MDS patients with TP53 mutations.

Figure 5

Heterozygous p53 mutant mice developed MDS with age. (A) Kaplan-Meier survival curve of p53^+/+ and p53^R248W/+ mice; n = 12–15 mice per group. (B) p53^R248W/+ mice developed various diseases during aging. Disease types and frequencies are shown. (C–I) H&E-stained BM sections of p53^R248W/+ mice developed MDS. Representative images show binucleated myeloid precursor (C and D), karyorrhexis in erythroid precursors (E), erythroid precursors with nuclear budding (F), binucleated erythroid precursor (G), and a giant megakaryocyte (H) compared with a megakaryocyte from age-matched p53^+/+ mice (I). Scale bars: 10 μm. (J) WBC, platelet, and RBC count as well as hemoglobin levels in PB of p53^R248W/+ developed MDS and age-matched p53^+/+ mice; n = 6 mice per group. (K) Kaplan-Meier survival curve of recipient mice repopulated with BM cells from aged p53^+/+ or p53^R248W/+ mice that developed MDS; n = 6 mice per group. (L) WBC and RBC counts in PB of recipient mice following BM transplantation; n = 5 mice per group. The survival curve was determined by the log-rank (Mantel-Cox) test (A and K). The comparison between 2 groups was evaluated by 2-tailed t test (J and L). Statistical significance: **P < 0.01, ***P < 0.001.

MDS is a clonal disease from mutant MDS stem cells (55, 56). To determine whether mutant p53 driven MDS is transplantable, we injected BM cells from aged p53^R248W/+ mice with MDS or age-matched p53^+/+ mice into lethally irradiated recipient mice. All recipient mice repopulated with p53^R248W/+ cells died within 4 months after transplantation, whereas no mice transplanted with aged p53^+/+ BM cells died during this period (Figure 5K). Recipient mice repopulated with p53^R248W/+ cells showed decreased WBC and RBC counts compared with recipients of WT BM cells (Figure 5L). Importantly, transplantation of BM cells from p53^R248W/+ mice with MDS resulted in lethal MDS and BM failure in all transplant recipients within 4 months (Supplemental Figure 4F), suggesting that the MDS phenotype induced by mutant p53 is transplantable.

Mutant p53 generates a chronic inflammatory microenvironment during aging. To decipher the mechanism by which mutant p53 drives MDS development, we first measured the levels of cytokines and chemokines in PB serum of young (2–4 months old) p53^+/+ and p53^R248W/+ mice utilizing cytokine arrays. Cytokine/chemokine levels in young p53^+/+ and p53^R248W/+ mice were comparable (Figure 6A). Given that chronic and low-grade inflammation develops during aging (9, 10), we then tested whether p53 mutant mice show enhanced systematic inflammation in an age-dependent manner. Notably, we observed increased levels of several pro-inflammatory cytokines, including IL-1β, L-17, and M-CSF, in PB serum of old (15–20 months old) p53 mutant mice compared with that of the age-matched p53^+/+ mice (Figure 6A). Given that dysregulated inflammatory signaling within the BM microenvironment has been implicated in MDS development (12), we examined the levels of cytokines and chemokines in BM fluid of old p53^+/+ and p53^R248W/+ mice and observed increased levels of IL-1β, IL-6, and GM-CSF in the BM of old p53 mutant mice compared with that of age-matched WT mice (Figure 6B).

Figure 6

Mutant p53 generates a chronic inflammatory microenvironment during aging. (A) Cytokines and chemokines in PB serum of p53^+/+ and p53^R248W/+ (young and old) mice were assayed using a mouse 31-Plex Cytokine/Chemokine array; n = 9 mice per group. (B) The levels of cytokines and chemokines in BM fluid of old p53^+/+ and p53^R248W/+ mice; n = 5 mice per group. (C) Patients with MDS or AML with TP53 mutations have increased levels of pro-inflammatory cytokines in their BM compared with the control group (control group, n = 12; TP53-mutated MDS or AML, n = 28). The comparison among multiple groups was evaluated by 2-way ANOVA (A). The comparison between 2 groups was evaluated by 2-tailed t test (B and C). Statistical significance: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001.

To further understand the role of chronic inflammation in pathogenesis of TP53-mutated myeloid neoplasms, we interrogated cytokine levels in the BM of MDS and AML patients with TP53 mutations using cytokine arrays. The comparative control group included BMs of lymphoma patients without involvement in disease. Consistent with our observations in p53 mutant mice (Figure 6, A and B), we found increased levels of IL-1β, IL-6, and TNF-α in the BM of MDS or AML patients with TP53 mutations compared with the control group (Figure 6C). Thus, we demonstrated that mutant p53 generates a chronic inflammatory BM microenvironment during aging, which may contribute to pathogenesis of MDS.

Murine HSPCs expressing mutant p53 display aberrant pre-mRNA splicing. To understand how mutant p53 modulates inflammatory response during aging, we performed RNA-seq studies to compare gene expression in HSPCs isolated from middle-aged (12–14 months old; pre-MDS) p53^+/+ and p53^R248W/+ mice. Spliceosome genes were significantly downregulated in middle-aged p53^R248W/+ HSPCs compared with age-matched p53^+/+ HSPCs (Supplemental Figure 4G). We confirmed that the expression of splicing factors, such as Prpf3 and Prpf4, was significantly downregulated in middle-aged (pre-MDS) p53^R248W/+ HSPCs (Figure 7A). As Prpf3 and Prpf4 play critical roles in spliceosome assembly (57, 58), we speculated that downregulation of Prpf3 and Prpf4 in p53 mutant HSPCs may alter pre-mRNA splicing. We conducted alternative splicing analysis utilizing the rMATS algorithm to identify differences in exon inclusion ratio between WT and mutant HSPCs (59, 60). We observed differential splicing of all classes of alternative splicing events, including Cassette Exons, Competing 5′ and 3′ splice sites, Mutually Exclusive Exons, and Retained Introns in p53^R248W/+ HSPCs (Figure 7B).

Figure 7

Murine HSPCs expressing mutant p53 display aberrant pre-mRNA splicing. (A) Splicing factors Prpf3 and Prpf4 were significantly downregulated in middle-aged p53 mutant HSPCs; n = 3 biological replicates. (B) Table of splicing events altered in middle-aged p53 mutant versus WT HSPCs. (C) Sashimi plots across splice junctions surrounding differentially spliced cassette exons in Usp15. (D and E) RT-PCR analysis revealed a significant change in pre-mRNA splicing of Usp15 in p53 mutant HSPCs; n = 3 biological replicates. (F) p53 mutant BM cells show enhanced NF-κB activation. (G) p53 mutant macrophages show enhanced NF-κB activation following LPS stimulation. (H) Loss of Il-1b did not alter p65 phosphorylation in p53 mutant BM cells. The comparison between 2 groups was evaluated by 2-tailed t test (A and E). Statistical significance: *P < 0.05, **P < 0.01.

NF-κB is a key mediator of IL-1β/IL-1R and LPS/TLR-4 signaling in immune cells (61). The NF-κB family consists of several proteins, including RelA (commonly known as p65), RelB, c-Rel, p50, and p52, operating as heterodimers or homodimers. Upon IL-1β or LPS stimulation, p65 is phosphorylated and translocated into the nucleus to activate NF-κB target genes (62). Notably, p53 mutant HSPCs show dysregulated pre-mRNA splicing in key regulators of NF-κB such as Usp15 (Figure 7C). p53 mutant HSPCs mainly express the short isoform of Usp15 (Figure 7, D and E). Usp15 is a known activator of NF-κB (63), and we found that p53 mutant BM cells have high levels of p65 phosphorylation in steady state (Figure 7F). We also observed that p53 mutant macrophages displayed elevated levels of p65 phosphorylation in steady state compared with WT cells, which was further enhanced by LPS stimulation (Figure 7G). To determine whether the baseline increase of NF-κB in p53^R248W/+ BM cells is due to elevated IL-1β systemically or is a cell intrinsic mechanism, we examined p65 phosphorylation in p53^R248W/+Il-1b^−/− BM cells and found that loss of Il-1b did not alter the level of p65 phosphorylation in mutant cells (Figure 7H).

Human HSPCs and primary MDS cells with TP53 mutations display aberrant pre-mRNA splicing. To determine the role of mutant p53 in human HSPCs, we introduced GFP or p53^R248W into human CD34⁺ cells using retroviruses (Figure 8A) (64). The transduction efficiency was approximately 20%–40% (Supplemental Figure 5A). Mutant p53 maintained CD34 expression in human HSPCs (Supplemental Figure 5, B and C). We sorted transduced CD34⁺ cells (GFP⁺) and performed CFU assays. The ectopic expression of p53^R248W enhanced the colony formation of human CD34⁺ cells compared with control viruses (MIGR1) transduced with CD34⁺ cells (Figure 8, B and C). To determine the response of human HSPCs to irradiation, we irradiated human CD34⁺ cells in vitro and performed apoptosis and CFU assays. Human CD34⁺ cells expressing mutant p53 show decreased apoptosis compared with human cells expressing WT p53 (Figure 8D). Mutant p53 also inhibited erythroid differentiation of human HSPCs (Supplemental Figure 5, D and E). To assess the long-term reconstituting ability of human CD34⁺ cells expressing mutant p53, we transplanted human CD34⁺ cells expressing mutant p53 or empty vector (MIGR1) into sublethally irradiated (2.5 Gy) NSGS mice (65) and examined the engraftment of human cells (hCD45⁺) in the PB and BM of recipient mice. Ectopic expression of mutant p53 increased the engraftment of human hematopoietic cells in PB, BM, and spleen of NSGS mice at 16 weeks after transplantation (Figure 8, E–G, and Supplemental Figure 6, A–E). Furthermore, NSGS mice repopulated with p53 mutant HSPCs showed splenomegaly (Supplemental Figure 6, F and G). THP1 is a human monocyte cell line that can differentiate into macrophages following PMA treatment. We introduced MIGR1 or p53^R248W into THP1 cells and differentiated them into macrophages. We treated macrophages with LPS for 4 h and examined the levels of full-length NLRP1 and active NLRP1 (NLRP1-NT) in these cells. Consistent with our findings in murine macrophages (Figure 2G and Supplemental Figure 2D), THP1 macrophages expressing mutant p53 had increased levels of full-length NLRP1 at baseline and LPS treatment increased the level of NLRP1-NT in mutant cells (Supplemental Figure 7A). These findings suggest that the impact of mutant p53 on NLRP1 activation in hematopoietic cells is conserved across species.

Figure 8

Human HSPCs and primary human MDS cells with TP53 mutations display dysregulated pre-mRNA splicing. (A) Schematic of in vitro and in vivo experiments using human HSPCs expressing mutant p53. CFC, colony-forming cell. (B and C) Mutant p53 enhances the colony formation of human HSPCs; n = 3 biological replicates. (D) Human HSPCs expressing mutant p53 show decreased apoptosis following irradiation; n = 3 biological replicates. (E) Ectopic mutant p53 expression increases the engraftment of human HSPCs in PB of NSGS mice; n = 9 mice per group. (F) Ectopic mutant p53 expression increases the engraftment of human HSPCs in the BM of NSGS mice at 16 weeks; n = 9 mice per group. (G) Mutant p53 expands human HSPCs in the BM of NSGS mice at 16 weeks; n = 9 mice per group. (H) Human HSC and multipotent progenitor cell gene signatures are significantly enriched in human CD34⁺ cells expressing mutant p53. (I) Table of splicing events altered in human CD34⁺ cells expressing mutant p53 compared with cells WT for p53; n = 3 biological replicates. (J) Sashimi plots across splice junctions surrounding differentially spliced cassette exons in IKBKE. (K and L) RT-PCR analysis revealed significant changes in pre-mRNA splicing of IKBKE in human HSPCs expressing mutant p53; n = 4 biological replicates. (M) IKBKE-L, but not IKBKE-S, enhances p65 phosphorylation in MDSL cells. (N) Table of splicing events altered in human MDS cells with TP53 mutations compared with the control group (control, n = 2; TP53 mutated MDS, n = 3). The comparison between 2 groups was evaluated by 2-tailed t test (B–D, F, G, and L). The comparison among multiple groups was evaluated by 2-way ANOVA (E). Statistical significance: *P < 0.05, **P < 0.01.

To determine the mechanism by which mutant p53 enhances the repopulating potential of human HSPCs, we performed RNA-seq studies in human CD34⁺ cells purified from the BM of NSGS mice at 16 weeks following transplantation. GSEA revealed that HSC and multipotent progenitor gene signatures are positively enriched in human HSPCs expressing mutant p53 compared with those of the WT cells (Figure 8H and Supplemental Figure 7B). We then conducted alternative splicing analysis to identify differences in exon inclusion ratio between human CD34⁺ cells expressing GFP and mutant p53 (59, 60). Consistent with our findings in murine HSPCs (Figure 7B), human HSPCs expressing mutant p53 displayed aberrant pre-mRNA splicing events compared with WT HSPCs (Figure 8I). Notably, human HSPCs expressing mutant p53 showed dysregulated pre-mRNA splicing in IKBKE (inhibitor of nuclear factor kappa B kinase subunit epsilon) (Figure 8J), a key regulator of NF-κB (66, 67). p53 mutant HSPCs mainly expressed the long isoform of IKBKE (IKBKE-L) (Figure 8, K and L). To determine the impact of IKBKE isoforms on NF-κB activation, we introduced HA-tagged IKBKE-L or IKBKE-S into human MDSL cells using retroviruses and examined NF-κB activation in transduced cells. We found that IKBKE-L, but not IKBKE-S, enhanced p65 phosphorylation in MDSL cells (Figure 8M).

To determine whether mutant p53 alters pre-mRNA splicing in primary human MDS cells, we performed RNA-seq in primary MDS cells with TP53 mutations and a control group with WT TP53. All human samples were free of known splicing factor mutations. Splicing analysis revealed that primary human MDS cells with TP53 mutations display aberrant pre-mRNA splicing compared with control hematopoietic cells with WT TP53 (Figure 8N).

Discussion

TP53 mutations are present in individuals with CHIP and CCUS (1–5). TP53 is also frequently mutated in myeloid neoplasms (26, 27). Some individuals with TP53-mutant CHIP and CCUS develop hematological malignancies later in their life (3–7). However, how inflammatory stress inhibits WT HSPC fitness and confers a competitive advantage to p53 mutant HSPCs is largely unexplored. We found that inflammatory stress enhances the fitness of p53 mutant HSPCs in transplantation assays. Notably, p53 mutant HSPCs manifest a hyperactive NLRP1/IL-1β signaling pathway, promoting cell proliferation and survival. While WT HSPCs are sensitive to inflammatory stress induced by LPS stimulation, p53 mutant HSPCs are resistant to LPS treatment. Thus, the growth advantage seen in p53 mutant HSPCs is likely due to elevated NLRP1/IL-1β signaling. Our data also suggest that p53 mutant HSPCs benefit from inflammatory signals as the colony formation capability of mutant HSPCs depends on IL-1β.

NLRP3 is activated in HSCs during aging, and aged WT HSCs show increased levels of pyroptosis (68). Tet2-deficient macrophages exhibited increased secretion of IL-1β that is mediated by NLRP3, and the clonal expansion of Tet2-deficient hematopoietic cells contributes to the exacerbated atherosclerosis in mice (69). Low- to high-risk MDS HSPCs, which usually have WT TP53, manifest an activated NLRP3 inflammasome (40). Notably, mutant p53 increases the chromatin accessibility to NLRP1, but not NLRP3, in HSPCs, leading to NLRP1 activation. Furthermore, loss of NLRP1 significantly decreased the levels of pro-inflammatory cytokines secreted from p53 mutant macrophages. Thus, our findings suggest that mutant p53 activates NLRP1 and NLRP1 is a key mediator of inflammatory response in p53 mutant HSPCs.

IL-1β is important for regulating the immune response in the acute setting; however, it can also promote disease development in the chronic setting (45, 51). We observed increased levels of IL-1β in p53 mutant macrophages following LPS stimulation. In addition, we found that increased pyroptosis seen in WT competitor cells co-cultured with p53 mutant BM cells depend on IL-1β. Thus, IL-1β is a key mediator of mutant p53–driven inflammatory response in hematopoietic cells. While IL-1β treatment inhibits WT HSPC function (45), we found that p53 mutant HSPCs are not sensitive to IL-1β exposure. To understand the underlying mechanisms, we performed RNA-seq studies and found that MYC target gene signatures are enriched in p53 mutant HSPCs following IL-1β treatment. Given that IL-1β treatment is known to decrease WT HSPC proliferation and reduce protein translation and synthesis (45), our findings suggest that altered response to inflammation may promote p53 mutant HSPC survival and provide these HSPCs with a competitive advantage over WT cells. Our GSEA also revealed that HSPC differentiation genes are significantly enriched in p53 mutant LSKs following IL-1β stimulation. Previous studies have shown that the increased expression of differentiation genes induced by Il-1β treatment is reversed by Cebpa deficiency, leading to selection for mutant cells (70). Thus, it appears that CEBPA and TP53 mutation can overcome Il-1b–mediated suppression of HSPC fitness through distinct but overlapping mechanisms.

A growing body of evidence, primarily from studies involving Tet2- and Dnmt3a-KO mice and stem cell transplantation models of CHIP (15, 16), suggests that dysregulated inflammation may contribute to mutant HSPC expansion. For example, we and others found that inflammatory stress promotes the clonal expansion of Tet2 mutant HSPCs (15, 36). TET2 appears to restrain inflammatory gene expression in macrophages, manifested by increased expression of LPS-induced genes in Tet2^−/− macrophages (36). Recently, IFN-γ signaling pathway has been implicated in promoting Dnmt3a-null HSPC expansion (16). Resistance to inflammation appears to underly enhanced fitness seen in Asxl1-deficient HSPCs (17). These studies suggest that the mechanisms by which inflammatory stress drives CHIP emergence and progression may be mutation specific.

We utilized a lethal dose of irradiation to generate chimeric mice for competition studies (24). However, irradiation is known to promote tissue damage and inflammation (11, 12). This inflammation could account for the selection for p53 mutant HSPC even in the absence of pI:pC or other additional inflammatory stimuli. To alleviate the impact of radiation on the host inflammatory state, we will utilize less inflammatory conditioning methods such as low-dose busulfan or nonmyeloablative whole BM transplants to establish mouse models of CHIP in the future.

There have been no studies on the functional impact of TP53 mutations on MDS pathogenesis (26, 27), so it is important to develop mouse models of myeloid neoplasms with TP53 mutations. We reported that mutant p53 drives MDS development in mice during aging and that mutant p53–driven MDS is transplantable. p53^R248W/+ mice developed diverse disease phenotypes after a long latency, suggesting that additional cooperating mutations may contribute to MDS development. It is possible that TP53 mutations may cooperate with DNMT3A, TET2, ASXL1, or other mutations to drive disease development (27). The majority of TP53-mutated MDS patients have multiple-hit TP53 mutations, and this state predicted risk of death and leukemic transformation (27). Thus, TP53 allelic state may be critical for prognosis and treatment response. Given that p53^R248W/+ mice are global knockin mice, and these mice also develop lymphoma and other tumors (24, 25, 54), conditional knockin mice of hotspot TP53 mutations will be needed to determine the impact of TP53 allelic state in MDS pathogenesis and develop preclinical models for testing MDS therapeutics.

During aging, chronic and low-grade inflammation develops (9, 10), and we observed increased levels of IL-1β and IL-6 in the BM of old p53 mutant mice and in patients with MDS or AML with TP53 mutations. Thus, mutant p53 generates a chronic inflammatory BM microenvironment during aging, which may contribute to pathogenesis of MDS. Li-Fraumeni syndrome (LFS) predisposes individuals to hereditary cancer linked to TP53 germline mutations (71). Many tumor types are found in LFS patients, including sarcomas, brain tumors, breast cancer, and hematological malignancies (71). It will be interesting to examine cytokine and chemokine levels in LFS patients and their healthy, mutation-negative relatives to better understand the role of TP53 mutations in cancer development in patients with LFS.

To determine the mechanism by which mutant p53 drives MDS development during aging, we performed RNA-seq studies and found that both murine and human HSPCs expressing mutant p53 exhibit aberrant pre-mRNA splicing. Notably, we found that primary human MDS cells with TP53 mutations show dysregulated pre-mRNA splicing, and the extent of changes in alternative splicing in primary human MDS cells with TP53 mutations is similar to what is seen in MDS patients with spliceosome mutations (72, 73), underscoring the important role that mutant p53 plays in regulating pre-mRNA splicing.

While chronic inflammation has been implicated in the clonal evolution of TP53-mutated therapy-related myeloproliferative neoplasms (74), how mutant p53 regulates inflammatory response in HSPCs remains elusive. We found that mutant p53 alters pre-mRNA splicing in key regulators of NF-κB, such as Usp15 and IKBKE, leading to enhanced NF-κB activation in hematopoietic cells, thereby driving MDS development. Altered RNA splicing by mutant p53 has been shown to activate oncogenic RAS signaling in pancreatic cancer (75); thus, our findings suggest that mutant p53 may dysregulate RNA splicing in a context-dependent manner to promote tumorigenesis.

Given that CHIP has become an emerging public health issue that affects 15%–20% of individuals age 70 or above (1–7), preventing the progression of CHIP to life-threatening diseases such as MDS and AML will have huge health benefits. We found that loss of NLRP1 decreases the engraftment of p53 mutant HSPCs in transplantation assays. As there is no NLRP1-specific inhibitor available (41, 42), the impact of pharmacological inhibition of NLRP1 on p53 mutant HSPCs awaits future investigation. We predict that p53 mutant HSPCs will be sensitive to NLRP1-specific inhibitor treatment both in vitro and in vivo. IL-1β has been shown to expand CEBPα-deficient hematopoietic progenitors in vivo (70). We found that IL-1β–neutralizing antibody treatment decreases the colony formation of p53 mutant HSPCs. Furthermore, p53 mutant HSPCs are sensitive to GSDMD inhibitor in vitro. IL-1β–neutralizing antibody (canakinumab) is in different phases of clinical trials (76) and several GSDMD inhibitors are FDA-approved drugs (77), so testing these drugs in individuals with TP53-mutant CHIP may be of clinical benefit.

Recently, TP53-mediated CH has been shown to confer increased risk for incident atherosclerotic disease (78). However, loss of p53 did not alter the expression of IL-1β and IL-6 in macrophages (78), suggesting that TP53 mutations may utilize distinct mechanisms to drive the development of different age-associated diseases.

Methods

Sex as a biological variable. Our study examined male and female animals, and similar findings are reported for both sexes.

Animals. p53^R248W/+ mice used in our studies have been described previously (20, 24, 25, 54). WT C57BL/6 (CD45.2⁺), Boy/J (CD45.1⁺), and F1 mice (CD45.2⁺CD45.1⁺) mice were obtained from an onsite core breeding colony. Nlrp1^−/− mice were generated in-house (43). Il-1b–KO (46) and NSGS (65) mice were obtained from The Jackson Laboratory.

Flow cytometry. Flow cytometry analysis of murine and human HSPCs in PB, BM, and spleen of recipient mice was performed as described previously (20, 24, 25, 64). Flow cytometry antibodies are listed in Supplemental Dataset 1.

Determination of the impact of inflammatory stress on HSPCs in vivo. To determine whether pI:pC treatment expands p53 mutant HSPCs in vivo, we performed competitive BM transplantation assays. To mimic CH, the ratio of donor cells/competitor cells was 1:10. We transplanted 1 × 10⁵ p53^+/+ and p53^R248W/+ BM cells (CD45.2⁺) together with 1 × 10⁶ competitor BM cells (CD45.1⁺) into lethally irradiated F1 mice (CD45.1⁺CD45.2⁺). Eight weeks following transplantation, recipient mice were treated with PBS or pI:pC (25 μg/kg, i.p.) twice a week for 4 weeks (34). The frequency of donor-derived cells (CD45.2⁺) in PB was assayed by flow cytometry analysis.

To determine whether LPS treatment expands p53 mutant HSPCs in vivo, we transplanted 1 × 10⁵ p53^+/+ and p53^R248W/+ BM cells (CD45.2⁺) together with 1 × 10⁶ competitor BM cells (CD45.1⁺) into lethally irradiated F1 mice (CD45.1⁺CD45.2⁺). Four weeks following transplantation, recipient mice were treated with PBS or LPS (1 mg/kg, i.p.) every other day for 4 weeks (35). The frequency of donor-derived cells (CD45.2⁺) in PB was assayed by flow cytometry analysis every 4 weeks for 16 weeks.

To determine the impact of LPS treatment on HSPC function, we treated p53^+/+ and p53^R248W/+ mice with PBS or LPS (1 mg/kg, i.p.) every other day for 4 weeks. We then transplanted 5 × 10⁵ BM cells from PBS or LPS-treated p53^+/+ and p53^R248W/+ mice (CD45.2⁺) plus 5 × 10⁵ competitor BM cells (CD45.1⁺) into lethally irradiated F1 mice (CD45.1⁺CD45.2⁺). The frequency of donor-derived cells (CD45.2⁺) in PB was assayed by flow cytometry analysis every 4 weeks for 16 weeks. At 16 weeks following primary transplantation, we transplanted 3 × 10⁶ BM cells isolated from primary recipient mice into lethally irradiated secondary recipient mice. The frequency of donor-derived cells (CD45.2⁺) in PB and BM was assayed by flow cytometry analysis.

Stem and progenitor cell assays. The CFU assays using murine or human HSPCs were performed as described previously (20, 24, 25, 64).

Generation of BM-derived macrophages. We cultured low-density BM mononuclear cells from p53^+/+ and p53^R248W/+ mice in IMDM supplemented with 20% FBS and 100 ng/mL GM-CSF (Pepro Tech) for 7 days (36). 5 × 10⁶ BM-derived macrophages were starved overnight in the absence of growth factors. Macrophages were treated with LPS (100 ng/mL) for 12 h (36). Supernatants were collected and subjected to cytokine array analysis (Eve Technologies, Mouse 31-Plex Cytokine/Chemokine Array). Cytokine array data are provided in Supplemental Dataset 2.

Generation of retroviruses and infection of murine HSPCs. Retroviruses were produced by transfection of Phoenix E cells with the MIGR1 control (MSCV-IRES-GFP) or MIGR1 full-length mutant p53 cDNA plasmid (MSCV-p53^R248W-IRES-GFP) according to standard protocols (20, 24, 25). Lineage-negative cells isolated from WT and Nlrp1^−/− mice were infected with high-titer retroviral suspensions in the presence of retronectin (Takara). Forty-eight hours after infection, the transduced cells (GFP⁺) were sorted by FACS.

In vitro cell competition assays. We cultured WT or mutant BM cells (CD45.2⁺) with competitor BM cells (CD45.1⁺) from B6.SJL mice. We generated 3 groups of BM cells in vitro: (a) 1 × 10⁶ WT CD45.1⁺ cells; (b) 5 × 10⁵ WT CD45.1⁺ + 5 × 10⁵ WT CD45.2⁺ cells; and (c) 5 × 10⁵ WT CD45.1⁺ + 5 × 10⁵ mutant CD45.2⁺ cells. We cultured 3 groups of cells in serum-free medium supplemented with hematopoietic cytokines, including FLT3 ligand, TPO, SCF, IL-3, and IL-6, for 7 days. We then sorted an equal number of live CD45.1⁺ cells from each group and performed proliferation, colony formation, apoptosis, and pyroptosis assays. In addition, we measured the levels of cytokines in conditioned medium utilizing cytokine arrays. Cytokine array data are provided in Supplemental Dataset 2.

RNA- and ATAC-seq assays. RNA-seq assays using murine HSPCs and data analysis were performed as described previously (24). We analyzed changes in chromatin accessibility in p53^+/+ and p53^R248W/+ LSKs by performing ATAC-seq (37). To ensure reproducibility, 2 biological replicates were performed, and 5 × 10⁴ LSK cells were pooled from age- and sex-matched littermates. Quality control of Fastq files was done by the FastQC tool. TopHat2 was used to align sequences on the mm9 genome. Reads from BAM files were filtered according to mapping quality value (mapq value, option -q 10). Then, peaks were detected on these BAM files by macs2. BigWig files were generated from bedgraph files (37). RNA- and ATAC-seq data are available in the Gene Expression Omnibus (GEO) under accession number GSE175370.

RNA splicing analysis. Alternative splicing events were identified by replicate Multivariate Analysis of Transcript Splicing (rMATS, version 4.1.0) (60). We adopted the Gencode annotation (GRCh38.p13, hg38) to define the splicing events (79). For each splicing event, the proportions of inclusion (I, the event occurred) and exclusion (S, otherwise) were quantified based on the junction reads supporting the respective isoforms in the RNA-seq alignment files. Changes in the inclusion level (L_I) across the control and mutant samples, as well as the statistical significance, were evaluated; this computational analysis was performed in murine HSPCs, human HSPCs, and primary human MDS cells. Differential alternative splicing events were selected under FDR < 0.05. In addition, differential splicing events that further satisfied (a) having ≥ 10 support junction reads in ≥ 60% of samples in the control and mutant group and (b) absolute fold change of L_I > 10% between control and mutant were highlighted.

See Supplemental Methods for additional information.

Statistics. Statistical analyses were performed with GraphPad Prism 9 software (GraphPad Software Inc). All data are presented as mean ± SEM. The sample size for each experiment and the replicate number of experiments are included in the figure legends. Statistical analyses were performed using 2-tailed t test where applicable for comparison between 2 groups, and a 1- or 2-way ANOVA test was used for experiments involving more than 2 groups. Representative Western blot and PCR results are shown from at least 3 biologically independent experiments. Representative flow cytometry profiles are shown from 3 biological replicates. Statistical significance was defined as P < 0.05, P < 0.01, P < 0.001, and P < 0.0001.

Study approval. MDS, AML, and control samples were collected at Northwestern University Memorial Hospital after informed consent. Protocols for sample handling and data analysis were approved by the Northwestern University Ethics Committee. All experiments involving primary human patient samples were performed in compliance with the Declaration of Helsinki. Furthermore, all animal experiments have received ethical approval from both Indiana and Northwestern University IACUCs. All mice were maintained in Indiana and Northwestern University animal facilities according to IACUC-approved protocols.

Data availability. Data are available in GEO (accession number GSE175370), in the Supporting Data Values file, or from the corresponding author upon request.

Author contributions

SC, SB, S Vemula, and YY: conceptualization, data curation, formal analysis, methodology, writing (original draft), writing (review), and editing. ES, HG, R Li, and FB: investigation, data curation, methodology, and formal analysis. SCN, PJ, and GES: investigation, data curation, and formal analysis. DAS, HC, WC, SX, R Luo, MAA, MLC, JPR, AF, SZ, TMM, M Sotelo, HP, SKH, S Veranga, MA, M Shumilina, KR, YJ, HL, and IK: investigation. RK, YA, JKA, EAE, LAG, CRZ, SLM, BAC, HSB, ZG, LDM, SAS, SH, YD, LCP, and SLM: resources. M Sukhanova, Yunlong Liu, and OAW: conceptualization, resources, formal analysis, supervision, investigation, visualization, writing (original draft), writing (review), and editing. Yan Liu: conceptualization, resources, formal analysis, investigation, visualization, supervision, funding acquisition, project administration, writing (original draft), writing (review), and editing.

Funding support

This work is the result of NIH funding, in whole or in part, and is subject to the NIH Public Access Policy. Through acceptance of this federal funding, the NIH has been given a right to make the work publicly available in PubMed Central.

National Institute of Diabetes and Digestive and Kidney Diseases Cooperative Center of Excellence in Hematology grant U54 DK106846 (to the Flow Cytometry Core and In Vivo Therapeutic Core Laboratories at the Indiana University School of Medicine).
NIH grants R01 CA298152, R01 HL150624, R56 DK119524, and R56 AG052501 (to Yan Liu).
Department of Defense awards W81XWH-18-1-0265 and DoD W81XWH-19-1-0575 (to Yan Liu).
The Leukemia & Lymphoma Society (LLS) Translational Research Program award 6581-20 (to Yan Liu).
St. Baldrick’s Foundation Scholar award (to Yan Liu).
LLS Career Development Special Fellow award (to SC).
American Society of Hematology Research Restart award (to SC).
Lady Tata Memorial International Awards for Research in Leukaemia (to SC).
NIH F31 award F31HL160120 (to SB).
NIH F32 award 1F32CA203049-01 (to SCN).

Acknowledgments

We thank Yang Xu at UCSD for providing the p53^R248W-knockin mice. We also acknowledge the Flow Cytometry Core, Pathology Core, NUSeq Core, Quantitative Data Science Core, Center for Comparative Medicine at Northwestern University Feinberg School of Medicine and Robert H. Lurie Comprehensive Cancer Center.

Address correspondence to: Yan Liu, Department of Medicine, Northwestern University Feinberg School of Medicine, Robert H. Lurie Comprehensive Cancer Center, 303 E. Superior Street, Chicago, Illinois 60611, USA. Phone: 312.503.1504; Email: yan.liu@northwestern.edu. Or to: Omar Abdel-Wahab, Human Oncology and Pathogenesis Program, Memorial Sloan Kettering Cancer Center, 1275 York Avenue, New York, New York 10065, USA. Email: abdelwao@mskcc.org. Or to: Yunlong Liu, Department of Medical Genetics, Indiana University School of Medicine, 1044 W. Walnut Street, Indianapolis, Indiana 46202, USA. Email: yunliu@iu.edu. Or to: Madina Sukhanova, Department of Pathology, Northwestern University Feinberg School of Medicine, 303 E. Superior Street, Chicago, Illinois 60611, USA. Email: madina.sukhanova@northwestern.edu.

Footnotes

Conflict of interest: OAW is a founder and scientific advisor of Codify Therapeutics; has served as a consultant for Foundation Medicine Inc., Merck, Prelude Therapeutics, Amphista Therapeutics, MagnetBio, and Janssen; and is on the Scientific Advisory Board of Envisagenics Inc., Harmonic Discovery Inc., and Pfizer Boulder. OAW has received prior research funding from H3B Biomedicine, Nurix Therapeutics, Minovia Therapeutics, and LOXO Oncology.

Reference information: J Clin Invest. 2026;136(3):e184285.https://doi.org/10.1172/JCI184285.

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King KY, et al. Environmental influences on clonal hematopoiesis. Exp Hematol. 2020;83:66–73.
View this article via: CrossRef PubMed Google Scholar

Trowbridge JJ, et al. Innate immune pathways and inflammation in hematopoietic aging, clonal hematopoiesis, and MDS. J Exp Med. 2021;218(7):e20201544.
View this article via: CrossRef PubMed Google Scholar

Cook EK, et al. Comorbid and inflammatory characteristics of genetic subtypes of clonal hematopoiesis. Blood Adv. 2019;3(16):2482–2486.
View this article via: CrossRef PubMed Google Scholar

Bick AG, et al. Inherited causes of clonal haematopoiesis in 97,691 whole genomes. Nature. 2020;586(7831):763–768.
View this article via: CrossRef PubMed Google Scholar

Cai Z, et al. Inhibition of inflammatory signaling in Tet2 mutant preleukemic cells mitigates stress-induced abnormalities and clonal hematopoiesis. Cell Stem Cell. 2018;23(6):833–849.
View this article via: CrossRef PubMed Google Scholar

Hormaechea-Agulla D, et al. Chronic infection drives Dnmt3a-loss-of-function clonal hematopoiesis via IFNγ signaling. Cell Stem Cell. 2021;28(8):1428–1442.
View this article via: CrossRef PubMed Google Scholar

Avagyan S, et al. Resistance to inflammation underlies enhanced fitness in clonal hematopoiesis. Science. 2021;374(6568):768–772.
View this article via: CrossRef PubMed Google Scholar

Khoo KH, et al. Drugging the p53 pathway: understanding the route to clinical efficacy. Nat Rev Drug Discov. 2014;13(3):217–236.
View this article via: CrossRef PubMed Google Scholar

Brosh R, Rotter V. When mutants gain new powers: news from the mutant p53 field. Nat Rev Cancer. 2009;9(10):701–713.
View this article via: CrossRef PubMed Google Scholar

Chen S, et al. Genotoxic stresses promote clonal expansion of hematopoietic stem cells expressing mutant p53. Leukemia. 2018;32(3):850–854.
View this article via: CrossRef PubMed Google Scholar

Marusyk A, et al. Irradiation selects for p53-deficient hematopoietic progenitors. PLoS Biol. 2010;8(3):e1000324.
View this article via: CrossRef PubMed Google Scholar

Wong TN, et al. Role of TP53 mutations in the origin and evolution of therapy-related acute myeloid leukaemia. Nature. 2015;518(7540):552–555.
View this article via: CrossRef PubMed Google Scholar

Stoddart A, et al. Cytotoxic therapy-induced effects on both hematopoietic and marrow stromal cells promotes therapy-related myeloid neoplasms. Blood Cancer Discov. 2020;1(1):32–47.
View this article via: CrossRef PubMed Google Scholar

Chen S, et al. Mutant p53 drives clonal hematopoiesis through modulating epigenetic pathway. Nat Commun. 2019;10(1):5649.
View this article via: CrossRef PubMed Google Scholar

Nabinger SC, et al. Mutant p53 enhances leukemia-initiating cell self-renewal to promote leukemia development. Leukemia. 2019;33(6):1535–1539.
View this article via: CrossRef PubMed Google Scholar

Haase D, et al. TP53 mutation status divides myelodysplastic syndromes with complex karyotypes into distinct prognostic subgroups. Leukemia. 2019;33(7):1747–1758.
View this article via: CrossRef PubMed Google Scholar

Bernard E, et al. Implications of TP53 allelic state for genome stability, clinical presentation and outcomes in myelodysplastic syndromes. Nat Med. 2020;26(10):1549–1556.
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Yoshida K, et al. Frequent pathway mutations of splicing machinery in myelodysplasia. Nature. 2011;478(7367):64–69.
View this article via: CrossRef PubMed Google Scholar

Inoue D, et al. Spliceosomal gene mutations in myelodysplasia: molecular links to clonal abnormalities of hematopoiesis. Genes Dev. 2016;30(9):989–1001.
View this article via: CrossRef PubMed Google Scholar

Saltzman AL, et al. Regulation of alternative splicing by the core spliceosomal machinery. Genes Dev. 2011;25(4):373–384.
View this article via: CrossRef PubMed Google Scholar

Lee BP, et al. Changes in the expression of splicing factor transcripts and variations in alternative splicing are associated with lifespan in mice and humans. Aging Cell. 2016;15(5):903–913.
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Crews LA, et al. RNA splicing modulation selectively impairs leukemia stem cell maintenance in secondary human AML. Cell Stem Cell. 2016;19(5):599–612.
View this article via: CrossRef PubMed Google Scholar

Sun D, et al. Epigenomic profiling of young and aged HSCs reveals concerted changes during aging that reinforce self-renewal. Cell Stem Cell. 2014;14(5):673–688.
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Walter D, et al. Exit from dormancy provokes DNA-damage-induced attrition in haematopoietic stem cells. Nature. 2015;520(7548):549–552.
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Takizawa H, et al. Pathogen-induced TLR4-TRIF innate immune signaling in hematopoietic stem cells promotes proliferation but reduces competitive fitness. Cell Stem Cell. 2017;21(2):225–240.
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Cull AH, et al. Tet2 restrains inflammatory gene expression in macrophages. Exp Hematol. 2017;55:56–70.
View this article via: CrossRef PubMed Google Scholar

Buenrostro JD, et al. Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods. 2013;10(12):1213–1218.
View this article via: CrossRef PubMed Google Scholar

Dennis G Jr, et al. DAVID: database for annotation, visualization, and integrated discovery. Genome Biol. 2003;4(5):P3.
View this article via: CrossRef PubMed Google Scholar

Heinz S, et al. Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol Cell. 2010;38(4):576–589.
View this article via: CrossRef PubMed Google Scholar

Basiorka AA, et al. The NLRP3 inflammasome functions as a driver of the myelodysplastic syndrome phenotype. Blood. 2016;128(25):2960–2975.
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Guo H, et al. Inflammasomes: mechanism of action, role in disease, and therapeutics. Nat Med. 2015;21(7):677–687.
View this article via: CrossRef PubMed Google Scholar

Chavarría-Smith J, Vance RE. The NLRP1 inflammasomes. Immunol Rev. 2015;265(1):22–34.
View this article via: CrossRef PubMed Google Scholar

Masters SL, et al. NLRP1 inflammasome activation induces pyroptosis of hematopoietic progenitor cells. Immunity. 2012;37(6):1009–1023.
View this article via: CrossRef PubMed Google Scholar

De Mooij CEM, et al. Targeting the interleukin-1 pathway in patients with hematological disorders. Blood. 2017;129(24):3155–3164.
View this article via: CrossRef PubMed Google Scholar

Pietras EM, et al. Chronic interleukin-1 exposure drives haematopoietic stem cells towards precocious myeloid differentiation at the expense of self-renewal. Nat Cell Biol. 2016;18(6):607–618.
View this article via: CrossRef PubMed Google Scholar

Labow M, et al. Absence of IL-1 signaling and reduced inflammatory response in IL-1 type I receptor-deficient mice. J Immunol. 1997;159(5):2452–2461.
View this article via: CrossRef PubMed Google Scholar

Carey A, et al. Identification of interleukin-1 by functional screening as a key mediator of cellular expansion and disease progression in acute myeloid leukemia. Cell Rep. 2017;18(13):3204–3218.
View this article via: CrossRef PubMed Google Scholar

Cailleau C, et al. Treatment with neutralizing antibodies specific for IL-1beta prevents cyclophosphamide-induced diabetes in nonobese diabetic mice. Diabetes. 1997;46(6):937–940.
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Rondeau JM, et al. The molecular mode of action and species specificity of canakinumab, a human monoclonal antibody neutralizing IL-1β. MAbs. 2015;7(6):1151–1160.
View this article via: CrossRef PubMed Google Scholar

Johnson DC, et al. DPP8/DPP9 inhibitor-induced pyroptosis for treatment of acute myeloid leukemia. Nat Med. 2018;24(8):1151–1156.
View this article via: CrossRef PubMed Google Scholar

Shi J, et al. Cleavage of GSDMD by inflammatory caspases determines pyroptotic cell death. Nature. 2015;526(7575):660–665.
View this article via: CrossRef PubMed Google Scholar

He WT, et al. Gasdermin D is an executor of pyroptosis and required for interleukin-1β secretion. Cell Res. 2015;25(12):1285–1298.
View this article via: CrossRef PubMed Google Scholar

Li N, et al. Myoglobin promotes macrophage polarization to M1 type and pyroptosis via the RIG-I/Caspase1/GSDMD signaling pathway in CS-AKI. Cell Death Discov. 2022;8(1):90.
View this article via: CrossRef PubMed Google Scholar

Song H, et al. p53 gain-of-function cancer mutants induce genetic instability by inactivating ATM. Nat Cell Biol. 2007;9(5):573–580.
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Nimer SD. Myelodysplastic syndromes. Blood. 2008;111(10):4841–4851.
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Cazzola M. Myelodysplastic syndromes. N Engl J Med. 2020;383(14):1358–1374.
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Zhong Z, et al. Two novel mutations in PRPF3 causing autosomal dominant retinitis pigmentosa. Sci Rep. 2016;6:37840.
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Chen X, et al. PRPF4 mutations cause autosomal dominant retinitis pigmentosa. Hum Mol Genet. 2014;23(11):2926–2939.
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Katz Y, et al. Analysis and design of RNA sequencing experiments for identifying isoform regulation. Nat Methods. 2010;7(12):1009–1015.
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Shen S, et al. rMATS: robust and flexible detection of differential alternative splicing from replicate RNA-seq data. Proc Natl Acad Sci U S A. 2014;111(51):E5593–E5601.
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Taniguchi K, Karin M. NF-κB, inflammation, immunity and cancer: coming of age. Nat Rev Immunol. 2018;18(5):309–324.
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Luo H, et al. Mitochondrial stress-initiated aberrant activation of the NLRP3 inflammasome regulates the functional deterioration of hematopoietic stem cell aging. Cell Rep. 2019;26(4):945–954.
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Higa KC, et al. Chronic interleukin-1 exposure triggers selection for Cebpa-knockout multipotent hematopoietic progenitors. J Exp Med. 2021;218(6):e20200560.
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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

[15] Cai Z, et al. Inhibition of inflammatory signaling in Tet2 mutant preleukemic cells mitigates stress-induced abnormalities and clonal hematopoiesis. Cell Stem Cell. 2018;23(6):833–849.
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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

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View this article via: CrossRef PubMed Google Scholar

[20] Chen S, et al. Genotoxic stresses promote clonal expansion of hematopoietic stem cells expressing mutant p53. Leukemia. 2018;32(3):850–854.
View this article via: CrossRef PubMed Google Scholar

[21] Marusyk A, et al. Irradiation selects for p53-deficient hematopoietic progenitors. PLoS Biol. 2010;8(3):e1000324.
View this article via: CrossRef PubMed Google Scholar

[22] Wong TN, et al. Role of TP53 mutations in the origin and evolution of therapy-related acute myeloid leukaemia. Nature. 2015;518(7540):552–555.
View this article via: CrossRef PubMed Google Scholar

[23] Stoddart A, et al. Cytotoxic therapy-induced effects on both hematopoietic and marrow stromal cells promotes therapy-related myeloid neoplasms. Blood Cancer Discov. 2020;1(1):32–47.
View this article via: CrossRef PubMed Google Scholar

[24] Chen S, et al. Mutant p53 drives clonal hematopoiesis through modulating epigenetic pathway. Nat Commun. 2019;10(1):5649.
View this article via: CrossRef PubMed Google Scholar

[25] Nabinger SC, et al. Mutant p53 enhances leukemia-initiating cell self-renewal to promote leukemia development. Leukemia. 2019;33(6):1535–1539.
View this article via: CrossRef PubMed Google Scholar

[26] Haase D, et al. TP53 mutation status divides myelodysplastic syndromes with complex karyotypes into distinct prognostic subgroups. Leukemia. 2019;33(7):1747–1758.
View this article via: CrossRef PubMed Google Scholar

[27] Bernard E, et al. Implications of TP53 allelic state for genome stability, clinical presentation and outcomes in myelodysplastic syndromes. Nat Med. 2020;26(10):1549–1556.
View this article via: CrossRef PubMed Google Scholar

[28] Yoshida K, et al. Frequent pathway mutations of splicing machinery in myelodysplasia. Nature. 2011;478(7367):64–69.
View this article via: CrossRef PubMed Google Scholar

[29] Inoue D, et al. Spliceosomal gene mutations in myelodysplasia: molecular links to clonal abnormalities of hematopoiesis. Genes Dev. 2016;30(9):989–1001.
View this article via: CrossRef PubMed Google Scholar

[30] Saltzman AL, et al. Regulation of alternative splicing by the core spliceosomal machinery. Genes Dev. 2011;25(4):373–384.
View this article via: CrossRef PubMed Google Scholar

[31] Lee BP, et al. Changes in the expression of splicing factor transcripts and variations in alternative splicing are associated with lifespan in mice and humans. Aging Cell. 2016;15(5):903–913.
View this article via: CrossRef PubMed Google Scholar

[32] Crews LA, et al. RNA splicing modulation selectively impairs leukemia stem cell maintenance in secondary human AML. Cell Stem Cell. 2016;19(5):599–612.
View this article via: CrossRef PubMed Google Scholar

[33] Sun D, et al. Epigenomic profiling of young and aged HSCs reveals concerted changes during aging that reinforce self-renewal. Cell Stem Cell. 2014;14(5):673–688.
View this article via: CrossRef PubMed Google Scholar

[34] Walter D, et al. Exit from dormancy provokes DNA-damage-induced attrition in haematopoietic stem cells. Nature. 2015;520(7548):549–552.
View this article via: CrossRef PubMed Google Scholar

[35] Takizawa H, et al. Pathogen-induced TLR4-TRIF innate immune signaling in hematopoietic stem cells promotes proliferation but reduces competitive fitness. Cell Stem Cell. 2017;21(2):225–240.
View this article via: CrossRef PubMed Google Scholar

[36] Cull AH, et al. Tet2 restrains inflammatory gene expression in macrophages. Exp Hematol. 2017;55:56–70.
View this article via: CrossRef PubMed Google Scholar

[37] Buenrostro JD, et al. Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods. 2013;10(12):1213–1218.
View this article via: CrossRef PubMed Google Scholar

[38] Dennis G Jr, et al. DAVID: database for annotation, visualization, and integrated discovery. Genome Biol. 2003;4(5):P3.
View this article via: CrossRef PubMed Google Scholar

[39] Heinz S, et al. Simple combinations of lineage-determining transcription factors prime cis-regulatory elements required for macrophage and B cell identities. Mol Cell. 2010;38(4):576–589.
View this article via: CrossRef PubMed Google Scholar

[40] Basiorka AA, et al. The NLRP3 inflammasome functions as a driver of the myelodysplastic syndrome phenotype. Blood. 2016;128(25):2960–2975.
View this article via: CrossRef PubMed Google Scholar

[41] Guo H, et al. Inflammasomes: mechanism of action, role in disease, and therapeutics. Nat Med. 2015;21(7):677–687.
View this article via: CrossRef PubMed Google Scholar

[42] Chavarría-Smith J, Vance RE. The NLRP1 inflammasomes. Immunol Rev. 2015;265(1):22–34.
View this article via: CrossRef PubMed Google Scholar

[43] Masters SL, et al. NLRP1 inflammasome activation induces pyroptosis of hematopoietic progenitor cells. Immunity. 2012;37(6):1009–1023.
View this article via: CrossRef PubMed Google Scholar

[44] De Mooij CEM, et al. Targeting the interleukin-1 pathway in patients with hematological disorders. Blood. 2017;129(24):3155–3164.
View this article via: CrossRef PubMed Google Scholar

[45] Pietras EM, et al. Chronic interleukin-1 exposure drives haematopoietic stem cells towards precocious myeloid differentiation at the expense of self-renewal. Nat Cell Biol. 2016;18(6):607–618.
View this article via: CrossRef PubMed Google Scholar

[46] Labow M, et al. Absence of IL-1 signaling and reduced inflammatory response in IL-1 type I receptor-deficient mice. J Immunol. 1997;159(5):2452–2461.
View this article via: CrossRef PubMed Google Scholar

[47] Carey A, et al. Identification of interleukin-1 by functional screening as a key mediator of cellular expansion and disease progression in acute myeloid leukemia. Cell Rep. 2017;18(13):3204–3218.
View this article via: CrossRef PubMed Google Scholar

[48] Cailleau C, et al. Treatment with neutralizing antibodies specific for IL-1beta prevents cyclophosphamide-induced diabetes in nonobese diabetic mice. Diabetes. 1997;46(6):937–940.
View this article via: CrossRef PubMed Google Scholar

[49] Rondeau JM, et al. The molecular mode of action and species specificity of canakinumab, a human monoclonal antibody neutralizing IL-1β. MAbs. 2015;7(6):1151–1160.
View this article via: CrossRef PubMed Google Scholar

[50] Johnson DC, et al. DPP8/DPP9 inhibitor-induced pyroptosis for treatment of acute myeloid leukemia. Nat Med. 2018;24(8):1151–1156.
View this article via: CrossRef PubMed Google Scholar

[51] Shi J, et al. Cleavage of GSDMD by inflammatory caspases determines pyroptotic cell death. Nature. 2015;526(7575):660–665.
View this article via: CrossRef PubMed Google Scholar

[52] He WT, et al. Gasdermin D is an executor of pyroptosis and required for interleukin-1β secretion. Cell Res. 2015;25(12):1285–1298.
View this article via: CrossRef PubMed Google Scholar

[53] Li N, et al. Myoglobin promotes macrophage polarization to M1 type and pyroptosis via the RIG-I/Caspase1/GSDMD signaling pathway in CS-AKI. Cell Death Discov. 2022;8(1):90.
View this article via: CrossRef PubMed Google Scholar

[54] Song H, et al. p53 gain-of-function cancer mutants induce genetic instability by inactivating ATM. Nat Cell Biol. 2007;9(5):573–580.
View this article via: CrossRef PubMed Google Scholar

[55] Nimer SD. Myelodysplastic syndromes. Blood. 2008;111(10):4841–4851.
View this article via: CrossRef PubMed Google Scholar

[56] Cazzola M. Myelodysplastic syndromes. N Engl J Med. 2020;383(14):1358–1374.
View this article via: CrossRef PubMed Google Scholar

[57] Zhong Z, et al. Two novel mutations in PRPF3 causing autosomal dominant retinitis pigmentosa. Sci Rep. 2016;6:37840.
View this article via: CrossRef PubMed Google Scholar

[58] Chen X, et al. PRPF4 mutations cause autosomal dominant retinitis pigmentosa. Hum Mol Genet. 2014;23(11):2926–2939.
View this article via: CrossRef PubMed Google Scholar

[59] Katz Y, et al. Analysis and design of RNA sequencing experiments for identifying isoform regulation. Nat Methods. 2010;7(12):1009–1015.
View this article via: CrossRef PubMed Google Scholar

[60] Shen S, et al. rMATS: robust and flexible detection of differential alternative splicing from replicate RNA-seq data. Proc Natl Acad Sci U S A. 2014;111(51):E5593–E5601.
View this article via: CrossRef PubMed Google Scholar

[61] Taniguchi K, Karin M. NF-κB, inflammation, immunity and cancer: coming of age. Nat Rev Immunol. 2018;18(5):309–324.
View this article via: CrossRef PubMed Google Scholar

[62] Karin M, et al. NF-kappaB in cancer: from innocent bystander to major culprit. Nat Rev Cancer. 2002;2(4):301–310.
View this article via: CrossRef PubMed Google Scholar

[63] Zhou Q, et al. USP15 potentiates NF-κB activation by differentially stabilizing TAB2 and TAB3. FEBS J. 2020;287(15):3165–3183.
View this article via: CrossRef PubMed Google Scholar

[64] Boitano AE, et al. Aryl hydrocarbon receptor antagonists promote the expansion of human hematopoietic stem cells. Science. 2010;329(5997):1345–1348.
View this article via: CrossRef PubMed Google Scholar

[65] Wunderlich M, et al. AML xenograft efficiency is significantly improved in NOD/SCID-IL2RG mice constitutively expressing human SCF, GM-CSF and IL-3. Leukemia. 2010;24(10):1785–1788.
View this article via: CrossRef PubMed Google Scholar

[66] Peters RT, et al. IKKepsilon is part of a novel PMA-inducible IkappaB kinase complex. Mol Cell. 2000;5(3):513–522.
View this article via: CrossRef PubMed Google Scholar

[67] Liu S, et al. The kinases IKBKE and TBK1 regulate MYC-dependent survival pathways through YB-1 in AML and are targets for therapy. Blood Adv. 2018;2(23):3428–3442.
View this article via: CrossRef PubMed Google Scholar

[68] Luo H, et al. Mitochondrial stress-initiated aberrant activation of the NLRP3 inflammasome regulates the functional deterioration of hematopoietic stem cell aging. Cell Rep. 2019;26(4):945–954.
View this article via: CrossRef PubMed Google Scholar

[69] Fuster JJ, et al. Clonal hematopoiesis associated with TET2 deficiency accelerates atherosclerosis development in mice. Science. 2017;355(6327):842–847.
View this article via: CrossRef PubMed Google Scholar

[70] Higa KC, et al. Chronic interleukin-1 exposure triggers selection for Cebpa-knockout multipotent hematopoietic progenitors. J Exp Med. 2021;218(6):e20200560.
View this article via: CrossRef PubMed Google Scholar

[71] de Andrade KC, et al. Cancer incidence, patterns, and genotype-phenotype associations in individuals with pathogenic or likely pathogenic germline TP53 variants: an observational cohort study. Lancet Oncol. 2021;22(12):1787–1798.
View this article via: CrossRef PubMed Google Scholar

[72] Papaemmanuil E, et al. Somatic SF3B1 mutation in myelodysplasia with ring sideroblasts. N Engl J Med. 2011;365(15):1384–1395.
View this article via: CrossRef PubMed Google Scholar

[73] Kim E, et al. SRSF2 mutations contribute to myelodysplasia by mutant-specific effects on exon recognition. Cancer Cell. 2015;27(5):617–630.
View this article via: CrossRef PubMed Google Scholar

[74] Rodriguez-Meira A, et al. Single-cell multi-omics identifies chronic inflammation as a driver of TP53-mutant leukemic evolution. Nat Genet. 2023;55(9):1531–1541.
View this article via: CrossRef PubMed Google Scholar

[75] Escobar-Hoyos LF, et al. Altered RNA splicing by mutant p53 activates oncogenic RAS signaling in pancreatic cancer. Cancer Cell. 2020;38(2):198–211.
View this article via: CrossRef PubMed Google Scholar

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Mutant p53 promotes clonal hematopoiesis by generating a chronic inflammatory microenvironment

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