The X-linked gene DDX3X encodes an RNA helicase that is mutated at high frequencies in several types of human B-cell lymphoma. Females have two active DDX3X alleles and males carry a DDX3Y homolog on the Y chromosome. We show here that pan-hematopoietic, homozygous deletion of Ddx3x in female mice perturbs erythropoiesis, causing early developmental arrest. However, both hemizygous male and heterozygous female embryos develop normally, suggesting that one Ddx3x allele is sufficient for fetal hematopoietic development in females and that the Ddx3y allele can compensate for the loss of Ddx3x in males. In adult mice, DDX3X deficiency altered hematopoietic progenitors, early lymphoid development, marginal zone and germinal center B cells, and lymphomagenesis in a sex-dependent manner. Loss of both Ddx3x alleles abrogated MYC-driven lymphomagenesis in females, whereas Ddx3x deletion in males did not affect the formation of B-cell lymphoma in both mouse models. Moreover, tumors that appeared in male mice lacking DDX3X showed upregulated expression of DDX3Y, indicating a critical requirement for DDX3 activity for lymphomagenesis. These data reveal sex-specific roles of DDX3X in erythro- and lymphopoiesis as well as in MYC-driven lymphomagenesis.

Significance:

The sex-dependent effects of DDX3X deficiency in malignant transformation of B cells and the compensatory role of DDX3Y support inhibition of DDX3 as a treatment strategy for MYC-driven B-cell lymphoma.

Hematopoiesis is the process of blood cell formation that starts in the bone marrow (BM) where multipotent progenitor cells follow defined steps of differentiation during which, new cellular subsets are committed for the myeloid, lymphoid, or erythroid lineage. The common lymphoid progenitors (CLP) have the potential to give rise to B and T lymphocytes, both essential for an effective acquired immune response. CLPs generate early thymic progenitors called double negative or “DN” cells that differentiate from DN1 to DN4 stages and undergo genetic rearrangement of T-cell receptor (TCR) genes and selection to finally give rise to TCR-expressing CD4 and CD8 T cells. CLPs in the BM give rise to B cells lineage progenitors that undergo Ig gene rearrangements to generate a repertoire of mature IgM+ B cells that leave the BM to settle in secondary lymphoid organs to form germinal centers (GC) where somatic hypermutation (SHM) and class switch recombination (CSR) take place. B cells that have successfully passed these two steps carry high-affinity antibodies of different isotypes on their surface and then differentiate into both memory B cells, responsible for a recall of the immune response, and into plasma-cells releasing soluble antibodies.

Many B-cell lymphomas such as Burkitt lymphoma or diffuse large B-cell lymphoma (DLBCL) emerge from GC cells (1). Burkitt lymphoma is a highly aggressive lymphoma mainly driven by the MYC oncogene, which becomes transcriptionally activated by chromosomal translocations to the immunoglobulin (Ig) μ heavy or λ light chain loci (2). Genome-wide sequencing of patients with Burkitt lymphoma revealed that the DDX3X gene is mutated in approximately 30% of Burkitt lymphoma and 5% of DLBCL (3–9). Other types of B-cell lymphoma such as pre-mediastinal B-cell lymphoma, Hodgkin's lymphoma (8) or chronic lymphocytic leukemia (10, 11); as well as natural killer T-cell lymphoma (12, 13) or various T-cell lymphoma subtypes (8, 14–16) also harbor DDX3X mutations. Whether and how DDX3X mutations affect lymphomagenesis remains to be fully investigated, but one study suggests that DDX3X loss-of-function (LOF) mutations buffer the proteotoxic stress induced by translocated MYC in human B-cell lymphoma and thus, facilitates the regulation of global protein synthesis and lymphomagenesis (9).

The DDX3X gene encodes an RNA-binding protein member of the DEAD-box helicase family having an RNA-remodeling activity (17, 18). DDX3X is involved in transcriptional regulation, ribosomal biogenesis, RNA nuclear export, and mRNA translation (19, 20). Several studies also link DDX3X to different cellular processes such as the regulation of cell-cycle progression, innate immune response, apoptosis, stress response, Wnt and NF-κB pathways, and embryonic development (21). DDX3X genetic alterations have been identified in various cancers, but its role as an oncogene or a tumor-suppressor remains unclear (22–24) and probably depends on cellular context. A constitutive Ddx3x knockout in the mouse is lethal at early embryonic stages owing in part to pronounced DNA damage and cell-cycle arrest (25, 26). The DDX3X gene is located on the X chromosome and is one of the genes escaping X-chromosome inactivation in humans as well as in mice (27, 28). Consequently, females have two active copies of DDX3X, whereas males carry only one, although the Y chromosome harbors the DDX3X homolog called DDX3Y encoding a similar protein (>90% homology). However, whether DDX3Y is expressed concomitantly with DDX3X or whether it can compensate for DDX3X LOF is still unclear. In humans, a post-transcriptional mechanism restrains DDX3Y translation to the male germ line cells (29), where DDX3Y plays an essential role in spermatogenesis (30, 31). DDX3Y can be aberrantly expressed in leukemic cells (32) and lymphoma cells where it could balance the effects caused by DDX3X mutations (9). However, murine DDX3Y is dispensable for spermatogenesis (33) and its translation does not seem to be restricted to germ cells (34).

Here, we show that Ddx3x deletion in hematopoietic cells is lethal in mice due to defects in prenatal erythropoiesis and that both DDX3X and DDX3Y proteins are expressed in normal murine lymphocytes. Both hemizygous and homozygous deletion of Ddx3x in adult hematopoietic cells affects progenitors, early steps of lymphoid differentiation, the GC formation and marginal zone (MZ) B cells. In addition, we show that MYC-driven lymphomagenesis is delayed when both Ddx3x alleles are deleted in females, but this effect can be reversed in male KO, suggesting sex-dependent differences of DDX3X LOF mutations in malignant transformation of B cells and a primordial compensatory role of DDX3Y in B cells.

Animal models

Ddx3x-conditional knockout mice were generated by Ingenious Targeting Laboratory. Vav-cre, Cd19-cre, R26-creER, Cγ1-cre, CD45.1, R26mT/mG, Trp53, and Eμ-Myc mice were purchased from The Jackson Laboratory; λ-Myc mice were a gift from Dr. Siegfried Janz (Medical College of Wisconsin, Milwaukee, Wisconsin). Mice were held in a C57BL/6 genetic background in a Specific-Pathogen-Free+ environment at the Institut de recherches cliniques de Montréal (IRCM) animal facility. Experimental procedures and mouse maintenance were approved by the Animal Care Committee (ACC#2013–04) of the IRCM in compliance with the Canadian Council on Animal Care guidelines (www.ccac.ca).

Embryo analysis

Embryos were prepared from euthanized pregnant females counted from the day of the plug (E0.5) and were fixed in formalin and stained with hematoxylin and eosin according to the manufacturer's protocol. For blood analysis, embryos were washed with PBS and blood cells were collected from the umbilical cord and cytocentrifuged onto slides (Thermo Fisher Scientific Cytospin), stained with May–Grünwald–Giemsa (Sigma-Aldrich) and imaged with a DM4000b microscope (Leica) and CellSens Entry software (Olympus).

Flow cytometry

Spleens and thymi were extracted from euthanized mice and single-cell suspensions were obtained by mechanical dissociation and flushed from tibia and femurs to obtain BM single-cell suspension. Cell solutions were filtered and depleted of red blood cells (RBC) by 10 minutes incubation in RBC Lysing Buffer Hybri-Max (Sigma). Cells were incubated with indicated fluorochrome-labeled antibodies (Supplementary Fig. S1). Intracellular staining was done with Cyto-Fast Fix/Perm kit (BioLegend) according to the manufacturer's instructions. Data were recorded on a SA3800 Spectral Cell Analyzer (Sony) or a BD LSRFortessa (BD Biosciences) and analyzed with FlowJo software.

BM transplantation

CD45.1 mice were treated with trimethoprim and sulfamethoxazole (TMS, Chiron pharmaceutique) 3 days pre-irradiation and 7 days post-irradiation. They received an additional enrofloxacin (Baytril, CDMV) treatment on day 8 to 14 postirradiation. CD45.1 mice were irradiated at 9,5 Gy and intravenously transplanted with 1×106 RBC-depleted BM cells from CD45.2 mice. BM reconstitution was validated by flow cytometry analysis of blood sample 8 weeks post-transplantation. The R26-creER was activated with two successive intraperitoneal injections of tamoxifen (Sigma) at 100 mg/kg on day 0 and 50 mg/kg on day 1.

Tumor analysis

Eμ-Myc and λ-Myc mice were crossed with Vav-cre/Ddx3x-floxed or Cd19-cre/Ddx3x-floxed animals and offspring was checked regularly until any endpoint was detected as defined by palpable tumor, respiratory discomfort, weight loss, impaired activity, hunched posture, or any other sign of suffering. Blood was collected by cardiac puncture and analyzed on an Advia 120 cell analyzer (Bayer) using the mouse archetype of multi-species software v.2.2.06. Tumor masses were harvested for genotyping and analyzed by flow cytometry. Pre-tumor stage is defined as 6-week-old mice not presenting any sign of disease.

Cell line

The 40LB cells were obtained from Dr. Di Noia's laboratory in December, 2021. The cells tested negative for Mycoplasma by PCR in May, 2022 and were not maintained for more than a month in culture.

Data availability

The data generated in this study are available within the article and its Supplementary Data Files.

Loss of DDX3X arrests development at mid gestation

Using gene targeting, we generated mice carrying Ddx3x alleles with two loxP sites flanking exon 2 containing the translation initiation codon. The cre-mediated deletion of the floxed exon 2 leads to an excision of Ddx3x (Supplementary Fig. S2A and S2B). Ddx3fl/fl females were crossed with the pan-hematopoietic Vav-cre deleter strain to enable deletion in all hematopoietic cells starting from early stages. Female Vav-cre/Ddx3fl/fl pups harboring a full Ddx3x-KO were never obtained, indicating that a developmental arrest must have occurred. However, Vav-cre/Ddx3fl/fl embryos could be analyzed at E14.5. They were smaller, showed an abnormal fetal liver with decreased cellularity and a decrease of enucleated erythrocytes in blood smears (Fig. 1AD). To investigate this further, we generated R26mT/mG/Vav-cre/Ddx3fl/fl embryos and stained blood cells with Hoechst. Both green and red fluorescence could be detected in nucleated and enucleated erythrocytes, albeit with a significant reduction of GFP+ cells in the R26mT/mG/Vav-cre/Ddx3fl/fl compared with controls (Supplementary Fig. S2C and S2D), suggesting that Ddx3x deletion impairs fetal erythropoiesis by blocking the transition from primitive to definitive erythropoiesis.

Figure 1.

DDX3X is essential for erythrocyte development. A, Comparison of a control embryo and Vav-cre/Ddx3fl/fl at stage E14.5. B, Hematoxylin and eosin staining of an E14.5 embryo. Black arrows, RBCs; white arrows, abnormal cellular debris. HRT, heart; LVP, liver parenchyma; S, stomach. C, May–Grünwald–Giemsa staining of fetal blood from embryos at the indicated genotypes at stages E14.5 and E15.5. D, Quantification of enucleated cells in fetal blood samples from animals with the indicated genotypes and developmental stages; t test was used to determine significance. E, Representative flow cytometric analyses and quantification of early erythroid (PreCFUe) and pre-megakaryocyte/erythroid (PreMegE) progenitors in adult BM relative to the number of live erythrocytes from animals with the indicated genotypes. F, Representative flow cytometric analyses and quantification of erythroid developmental stages from proerythroblasts (Pro-E) to basophilic (Baso-E), polychromatic (Poly-E), and orthochromatic (Ortho-E) erythroblast stages in adult BM from animals with the indicated genotypes. Populations are defined as follows: Pro-E (CD71+Ter119low), Baso-E (CD71+Ter119+), Poly-E (CD71lowTer119+), Ortho-E (CD71Ter119+). G, Percentages of RBC in blood of mice with indicated genotypes. t test with Welsh correction was used to compare female heterozygotes with female controls and male KO with male controls. **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 1.

DDX3X is essential for erythrocyte development. A, Comparison of a control embryo and Vav-cre/Ddx3fl/fl at stage E14.5. B, Hematoxylin and eosin staining of an E14.5 embryo. Black arrows, RBCs; white arrows, abnormal cellular debris. HRT, heart; LVP, liver parenchyma; S, stomach. C, May–Grünwald–Giemsa staining of fetal blood from embryos at the indicated genotypes at stages E14.5 and E15.5. D, Quantification of enucleated cells in fetal blood samples from animals with the indicated genotypes and developmental stages; t test was used to determine significance. E, Representative flow cytometric analyses and quantification of early erythroid (PreCFUe) and pre-megakaryocyte/erythroid (PreMegE) progenitors in adult BM relative to the number of live erythrocytes from animals with the indicated genotypes. F, Representative flow cytometric analyses and quantification of erythroid developmental stages from proerythroblasts (Pro-E) to basophilic (Baso-E), polychromatic (Poly-E), and orthochromatic (Ortho-E) erythroblast stages in adult BM from animals with the indicated genotypes. Populations are defined as follows: Pro-E (CD71+Ter119low), Baso-E (CD71+Ter119+), Poly-E (CD71lowTer119+), Ortho-E (CD71Ter119+). G, Percentages of RBC in blood of mice with indicated genotypes. t test with Welsh correction was used to compare female heterozygotes with female controls and male KO with male controls. **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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Adult heterozygous female Vav-cre/Ddx3X/fl mice were obtained at a mendelian ratio, indicating that one Ddx3x allele is sufficient to prevent the developmental arrest caused by defect in fetal erythropoiesis. Similarly, male Vav-cre/Ddx3fl/Y mice carrying a Ddx3x-floxed allele along with the intact Ddx3y allele also overcame the erythroid developmental block and developed into adulthood, indicating that the Ddx3y gene can compensate for the loss of Ddx3x. We confirmed that Vav-cre/Ddx3fl/Y adult mice lacked DDX3X but expressed DDX3Y in spleen and thymus using a newly generated antibody specific to murine DDX3Y (Supplementary Fig. S2E–S2H). However, adult Vav-cre/Ddx3fl/Y males still exhibit increased proerythroblasts along a decrease of orthochromatic erythroblast (Fig. 1E and F) and a decreased percentage of RBC in their peripheral blood even if this reduction is not life-threatening (Fig. 1G). This suggests that DDX3X is necessary for proper prenatal and adult erythropoiesis in male mice. Because DDX3X is not expressed in the different subsets from proerythroblasts to orthochromatic erythroblast (Supplementary Fig. S2I), the defects observed in Vav-cre/Ddx3fl/Y are likely a consequence of defects occurring in early erythroid progenitors. These mild erythropoietic defects are not seen in Vav-cre/Ddx3X/fl females (Fig.1EG) expressing similar levels of DDX3X protein compared with controls (Supplementary Fig. S2G and S2H).

Hematopoietic progenitors require DDX3X

A comparison of female Vav-cre/Ddx3X/fl and male Vav-cre/Ddx3fl/Y mice by flow cytometry revealed a decrease of Linc-kit+ cells (LK), Linsca1+c-kit+ cells (LSK), multipotent progenitors (MPP), lymphoid-primed MMPs (LMPP), and CLPs only in male animals compared with sex-matched controls (Fig. 2A and B). Long-term hematopoietic stem cells, granulocyte/monocyte progenitors, and megakaryocyte progenitors were not affected by DDX3X loss regardless of sex (Supplementary Fig. S3A–S3C), but pre-megakaryocyte/erythrocyte progenitors were reduced in Vav-cre/Ddx3fl/Y male mice (Fig. 1E). DDX3X protein expression was detected in LK and LSK progenitors (Supplementary Fig. S3D). Ddx3x-depleted LSKs from male Vav-cre/Ddx3fl/Y mice were unable to differentiate into B or T cells in vitro and produced less colonies compared with controls (Fig. 2CE). These data indicate that DDX3X is required to maintain the cellularity and lymphoid lineage potential of adult hematopoietic progenitor cells.

Figure 2.

Hematopoietic progenitors are decreased in Ddx3x-KO males. A and B, Flow cytometry analysis of BM from animals with the indicated genotype (A) and quantification (B) of absolute number of cells. Populations were defined as follows: LK (Linc-kit+sca-1), LSK (Linc-kit+sca-1+), MPP (Linc-kit+sca-1Flt3low), LMPP (Linc-kit+sca-1Flt3+). t test with Welsh correction was used to compare female heterozygotes with female controls and male KO mice with male controls. C and D, LSKs were sorted from the BM of male KO and control mice and cocultured on OP9 layer with IL7 and Flt3 cytokines for B-cell differentiation (C) or cocultured on OP9-DL1 layer with SCF, Flt3 and IL7 for T-cell differentiation (D). Two experiments were performed in triplicate. E, Sorted LSK cells were grown in Methocult GF-M3434 media at 37°C and colonies were counted and identified 7 days later. This experiment was performed three times and paired t test was used to assess statistical significance. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001. CFU, colony-forming unit; CFU-GM, CFU-granulocyte, monocyte; CFU-GEMM, CFU-granulocyte, erythrocyte, monocyte, megakaryocyte; BFU-E, burst forming unit, erythrocyte; LK, Linc-kit+; LSK, LinSca+c-kit+; MPP, multiple pluripotent progenitor; LMPP, lymphoid multiple pluripotent progenitor; CLP, common lymphoid progenitor.

Figure 2.

Hematopoietic progenitors are decreased in Ddx3x-KO males. A and B, Flow cytometry analysis of BM from animals with the indicated genotype (A) and quantification (B) of absolute number of cells. Populations were defined as follows: LK (Linc-kit+sca-1), LSK (Linc-kit+sca-1+), MPP (Linc-kit+sca-1Flt3low), LMPP (Linc-kit+sca-1Flt3+). t test with Welsh correction was used to compare female heterozygotes with female controls and male KO mice with male controls. C and D, LSKs were sorted from the BM of male KO and control mice and cocultured on OP9 layer with IL7 and Flt3 cytokines for B-cell differentiation (C) or cocultured on OP9-DL1 layer with SCF, Flt3 and IL7 for T-cell differentiation (D). Two experiments were performed in triplicate. E, Sorted LSK cells were grown in Methocult GF-M3434 media at 37°C and colonies were counted and identified 7 days later. This experiment was performed three times and paired t test was used to assess statistical significance. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001. CFU, colony-forming unit; CFU-GM, CFU-granulocyte, monocyte; CFU-GEMM, CFU-granulocyte, erythrocyte, monocyte, megakaryocyte; BFU-E, burst forming unit, erythrocyte; LK, Linc-kit+; LSK, LinSca+c-kit+; MPP, multiple pluripotent progenitor; LMPP, lymphoid multiple pluripotent progenitor; CLP, common lymphoid progenitor.

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Ddx3x deletion affects cells of the lymphoid but not the myeloid lineage

Although early multipotential hematopoietic progenitors such as MPPs and LMPPs were decreased in Ddx3x-deficient male mice, more mature cells of the myeloid lineage such as monocytes, macrophages or granulocytes were not affected by DDX3X loss, correlating with the lack of protein expression (Supplementary Fig. S3E and S3F). However, Ddx3x deletion in Vav-cre/Ddx3fl/Y mice correlated with a significantly reduced number of cells at the DN2 and DN3 stages of pre–T-cell differentiation (Fig. 3A and B), the same stages that were blocked in the in vitro T-cell differentiation assay (Fig. 2D). Despite the defects in pre–T-cell differentiation, the numbers of CD4/CD8 DP and CD4+ or CD8+ T cells remained at WT levels upon Ddx3x deletion in male even though DDX3X is expressed in all thymic T-cell subsets (Supplementary Fig. S4A and S4B). Whereas expression of DDX3X in pre–T-cell stages was evident (Supplementary Fig. S4B), DDX3Y was not readily detectable in DN2-DN4 pre–T-cell stages (Supplementary Fig. S4C and S4D), suggesting absence or low level of expression in these cells. Ddx3x deletion in Vav-cre/Ddx3fl/Y mice also impaired several stages of B-cell differentiation: pro-B cells, pre-B cells, immature and recirculating B cells, expressing DDX3X protein (Supplementary Fig. S4E–S4H).

Figure 3.

Ddx3x deficiency affects pre–T- and pre–B-cell development in male mice. A and B, Thymi were extracted from Vav-cre/Ddx3x-floxed mice and analyzed by flow cytometry (A), with extracellular markers for T-cell progenitors (DN stages from 1 to 4) and population quantification in absolute numbers is represented in B. Populations are defined as follows: DN1 (LinCD44+CD25), DN2 (LinCD44+CD25+), DN3 (LinCD44CD25+), and DN4 (LinCD44CD25). The t test with Welsh correction was used to compare female heterozygous mice with female controls and male KO mice with male controls. C and D, Flow cytometric analysis quantification of absolute cell numbers of BM extracted from Cd19-cre/Ddx3x-floxed mice, including B-cell progenitors and Hardy fractions from A to F. Populations are defined as follows: Fr.A or Pre–Pro-B-cell (B220+CD43+HSABP1), Fr.B or Pro-B-cell (B220+CD43+HSA+BP1), Fr.C-C’ or large Pre-B-cell (B220+CD43+HSA+BP1+), Fr.D or Pre-B-cell (B220+CD43IgMIgD), Fr.E or Immature B-cell (B220+CD43IgM+IgD), Fr.F or mature or recirculating B-cell (B220+CD43IgM+IgD+). The Kruskal–Wallis test was used to compare female KO with female controls and heterozygotes and Mann–Whitney test was used to compare male KO with male controls. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 3.

Ddx3x deficiency affects pre–T- and pre–B-cell development in male mice. A and B, Thymi were extracted from Vav-cre/Ddx3x-floxed mice and analyzed by flow cytometry (A), with extracellular markers for T-cell progenitors (DN stages from 1 to 4) and population quantification in absolute numbers is represented in B. Populations are defined as follows: DN1 (LinCD44+CD25), DN2 (LinCD44+CD25+), DN3 (LinCD44CD25+), and DN4 (LinCD44CD25). The t test with Welsh correction was used to compare female heterozygous mice with female controls and male KO mice with male controls. C and D, Flow cytometric analysis quantification of absolute cell numbers of BM extracted from Cd19-cre/Ddx3x-floxed mice, including B-cell progenitors and Hardy fractions from A to F. Populations are defined as follows: Fr.A or Pre–Pro-B-cell (B220+CD43+HSABP1), Fr.B or Pro-B-cell (B220+CD43+HSA+BP1), Fr.C-C’ or large Pre-B-cell (B220+CD43+HSA+BP1+), Fr.D or Pre-B-cell (B220+CD43IgMIgD), Fr.E or Immature B-cell (B220+CD43IgM+IgD), Fr.F or mature or recirculating B-cell (B220+CD43IgM+IgD+). The Kruskal–Wallis test was used to compare female KO with female controls and heterozygotes and Mann–Whitney test was used to compare male KO with male controls. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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To further investigate DDX3X role in B-cell differentiation, we used the B-cell–specific Cd19-cre deleter (35), which enables us to delete both Ddx3x alleles and to circumvent the early lethality caused by the Vav-cre deleter. The cellularity of total BM or the B220CD19 fraction was not affected by Cd19-cre–mediated deletion (Fig. 3C and D). In contrast, B220+CD19+ B-cell fraction was almost absent in Cd19-cre/Ddx3fl/fl mice (Fig. 3C and D). A closer analysis showed that Cd19-cre/Ddx3fl/Y animals had similar phenotypes compared with Vav-cre/Ddx3fl/Y mice with reduced cell numbers in Hardy Fractions B, E, and F (Supplementary Fig. S4E–S4G; Fig. 3C and D), but female Cd19-cre/Ddx3fl/fl mice showed more pronounced reduction of cell numbers in Hardy Fractions B, C/C’, E and F compared with male Cd19-cre/Ddx3fl/Y animals. These findings demonstrated that DDX3X is required for the lymphoid lineage, especially for early steps of B- and T-cell development and is most critical for the mature B-cell fraction but is dispensable for the myeloid lineage.

Inducible Ddx3x deletion in adult female mice causes BM failure

Next, we generated a conditionally inducible Ddx3x-KO by crossing Ddx3fl/fl mice with the R26-creER animals allowing cre-mediated deletion of floxed alleles upon tamoxifen administration. BM cells from R26-creER/Ddx3fl/fl mice (CD45.2 cells) were transplanted into irradiated CD45.1 recipients and full reconstitution of the hematopoietic system was validated in recipients eight weeks after transplantation (Supplementary Fig. S5A–S5C). After the tamoxifen injection activating the R26-creER, animals that received R26-creER/Ddx3fl/fl BM cells and had fully reconstituted hematopoiesis died very rapidly within 9 to 10 days after tamoxifen induction unlike recipients transplanted with BM from R26-creER/Ddx3X/X or R26-creER/Ddx3X/fl mice (Fig. 4A). R26-creER/Ddx3fl/fl mice developed symptoms of anemia such as white paws, gray coat, as well as low body temperature and low hematocrit (Fig. 4B). LK and LSK progenitors were almost completely lost, and the amount of live BM cells was significantly lower in KO mice (Fig. 4C and D). The MZ B-cell population was clearly increased, and GC B-cell numbers were decreased in KO mice compared with controls and heterozygotes (Fig. 4E and F).

Figure 4.

Ddx3x deletion in hematopoietic cells of adult female mice induces death-related anemia and severe loss of hematopoietic progenitors. A, Kaplan–Meier curves representing survival of recipient CD45.1 mice transplanted with BM from CD45.2 mice: R26-creER; R26-creER/Ddx3X/fl; or R26-creER/Ddx3fl/fl. The BM transplantation efficiency was validated 8 weeks post-transplantation, the tamoxifen was injected at day 0, and the survival was measured since this injection. B, Hematocrit was measured either at day 9 (gray dots) or day 10 (white dots) from the blood of moribund KO mice. C and D, Flow cytometry analysis of hematopoietic progenitors from the BM of transplanted mice at day 6 (C) and relative quantification in D. E and F, Flow cytometry analysis of splenic B-cell populations from transplanted mice at day 6 (E) and relative quantification in F. Populations are defined as follows: NF (B220+CD21CD23), FO (B220+CD21CD23+), MZ (B220+CD21+CD23), GC (B220+IgDCD95+GL7+). G and H, Flow cytometry analysis of T-cell populations extracted from the thymus of transplanted mice at day 6 (G) and relative quantification in H. Statistical significance was measured by one-way ANOVA. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 4.

Ddx3x deletion in hematopoietic cells of adult female mice induces death-related anemia and severe loss of hematopoietic progenitors. A, Kaplan–Meier curves representing survival of recipient CD45.1 mice transplanted with BM from CD45.2 mice: R26-creER; R26-creER/Ddx3X/fl; or R26-creER/Ddx3fl/fl. The BM transplantation efficiency was validated 8 weeks post-transplantation, the tamoxifen was injected at day 0, and the survival was measured since this injection. B, Hematocrit was measured either at day 9 (gray dots) or day 10 (white dots) from the blood of moribund KO mice. C and D, Flow cytometry analysis of hematopoietic progenitors from the BM of transplanted mice at day 6 (C) and relative quantification in D. E and F, Flow cytometry analysis of splenic B-cell populations from transplanted mice at day 6 (E) and relative quantification in F. Populations are defined as follows: NF (B220+CD21CD23), FO (B220+CD21CD23+), MZ (B220+CD21+CD23), GC (B220+IgDCD95+GL7+). G and H, Flow cytometry analysis of T-cell populations extracted from the thymus of transplanted mice at day 6 (G) and relative quantification in H. Statistical significance was measured by one-way ANOVA. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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In the thymus, a loss of DN2 and DN3 pre-T cells was observed, similar to the observed effects in Vav-cre/Ddx3fl/Y mice (Fig. 3A) whereas CD4+ cells were slightly increased and CD8+ remained unchanged (Fig. 4G and H). However, the toxic effects of tamoxifen, which have been described previously (36) and are clearly evident here considering the strong reduction of the thymic DP T-cell compartment (Fig. 4G and H) precluded further analyses of these mice. B-cell progenitors and mature B cells were less affected by Ddx3x deletion in mice with R26-creER/Ddx3fl/fl BM (Supplementary Fig. S5D–S5G) compared with animals in which Ddx3x was deleted with the CD19-cre (Fig. 3C and D). This absence of a B-cell phenotype in mice that received R26-creER/Ddx3fl/fl BM is most likely due to the tamoxifen toxicity that precluded any further analyses at time points later than 6 days after the first tamoxifen injection.

Ddx3x deletion disturbs differentiated B cells and B-cell immune response

Cd19-cre/Ddx3fl/fl females had smaller spleens, a lower number of cells and a strongly reduction of B220+ cells compared with controls (Fig. 5A and B). The comparison of Cd19-cre/Ddx3fl/Y males and Cd19-cre/Ddx3fl/fl females revealed that the cellularity of their transitional and follicular B-cell compartments as well as their mature B220+IgD+ subsets were decreased, whereas their MZ B-cell subsets were increased in percentages (Fig. 5B and C). All phenotypes were similar between the male and female KO mice but were significantly more severe in females lacking both Ddx3x alleles (Fig. 5B and C), similar to observations made in Vav-cre/Ddx3fl/Y males (Supplementary Fig. S6A–S6C). A decrease of transitional B cells T1, T2, T3, and an accumulation of B220+CD93+IgMCD23+ was observed in Vav-cre/Ddx3fl/Y mice (Supplementary Fig. S6B) confirming data from a previous report (37). Histological sections of spleens from Vav-cre/Ddx3fl/Y male mice demonstrated altered structures of follicles and MZs compared with control sections (Supplementary Fig. S6D). Numbers of plasma B cells were unchanged whereas the B220+IgDCD38+ B cells were significantly decreased in Cd19-cre/Ddx3fl/fl mice but not in Cd19-cre/Ddx3fl/Y or Vav-cre/Ddx3fl/Y animals (Supplementary Fig. S7A–S7D), even though DDX3X protein was found to be expressed in all mature B cells subsets included B220+IgDCD38+ cells and plasma B cells (Supplementary Fig. S7E). Splenic resting B cells isolated from the Cd19-cre/Ddx3 mice responded similarly to the BCR activation compared with the controls demonstrating the functionality of the BCR and downstream signaling (Fig. 5D).

Figure 5.

DDX3X is required to maintain mature B-cell populations. A, Calculation of splenic index (the weight of the spleen divided by the weight of the animal multiplied by 100) and quantification of the total number of splenic cells and B220+ splenic cells extracted from Cd19-cre/Ddx3x-KO mice. B and C, Flow cytometry analysis (B) and quantification (C) in absolute number of splenic B cells in Cd19-cre/Ddx3x-KO mice compared with controls. Populations are defined as follows: transitional B cells (B220+CD93+), NF (B220+CD21CD23), FO (B220+CD23+), and MZ (B220+CD21+CD23). The Kruskal–Wallis test was used to compare female KO with female controls and heterozygotes and Mann–Whitney test was used to compare male KO with male controls. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001. D, Resting B cells were isolated from the spleen of Cd19-cre/Ddx3x-KO mice with the MojoSort magnetic cell separation (BioLegend) and stimulated with secreted IgM for 10 minutes at 37°C. Cells lysates were immunoblotted with ERK and phospho-ERK primary antibodies. This experiment was done in triplicate. sIgM, secreted IgM.

Figure 5.

DDX3X is required to maintain mature B-cell populations. A, Calculation of splenic index (the weight of the spleen divided by the weight of the animal multiplied by 100) and quantification of the total number of splenic cells and B220+ splenic cells extracted from Cd19-cre/Ddx3x-KO mice. B and C, Flow cytometry analysis (B) and quantification (C) in absolute number of splenic B cells in Cd19-cre/Ddx3x-KO mice compared with controls. Populations are defined as follows: transitional B cells (B220+CD93+), NF (B220+CD21CD23), FO (B220+CD23+), and MZ (B220+CD21+CD23). The Kruskal–Wallis test was used to compare female KO with female controls and heterozygotes and Mann–Whitney test was used to compare male KO with male controls. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001. D, Resting B cells were isolated from the spleen of Cd19-cre/Ddx3x-KO mice with the MojoSort magnetic cell separation (BioLegend) and stimulated with secreted IgM for 10 minutes at 37°C. Cells lysates were immunoblotted with ERK and phospho-ERK primary antibodies. This experiment was done in triplicate. sIgM, secreted IgM.

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GC B cells were significantly decreased in Vav-cre/Ddx3fl/Y male KO mice as well as in both Cd19-cre/Ddx3fl/Y and Cd19-cre/Ddx3fl/fl mice (Fig. 6A and B); a phenotype already observed in transplanted R26-creER/Ddx3fl/fl (Fig. 4E and F). Given the critical role of GC B cells in the initiation of B-cell malignancies, we investigated this further by generating mice with a GC-specific deletion using a Cγ1-cre allele that can be induced by immunization with sheep red blood cells (SRBC). Under these conditions, GCs failed to expand in Cγ1-cre/Ddx3fl/Y males and Cγ1-cre/Ddx3fl/fl homozygous females compared with controls or heterozygous females (Fig. 6C), demonstrating that the GC defect in intrinsic and not a consequence of an impaired B-cell differentiation in Ddx3x-KO mice. Using the R26mT/mG reporter allele, we observed that GFP+ GC B cells were significantly decreased in Cγ1-cre/Ddx3fl/Y males and very low in Cγ1-cre/Ddx3fl/fl females 10 days after SRBC immunization (Fig. 6D), suggesting that GC B cells are eliminated in absence of DDX3X. Although GC cells are significantly decreased in KO males, GFP+ GC B cells were clearly detectable, suggesting that DDX3Y can partially compensate for Ddx3x loss in these cells. Similar to other cell populations, the GC compartment was intact in heterozygous Cγ1-cre/Ddx3X/fl females, excluding a haploinsufficiency of the Ddx3x locus in mice. GFP+ GC B cells were almost completely absent in Cγ1-cre/Ddx3fl/fl female mice at different time points after the SRBC immunization (Fig. 6E), suggesting that GC critically require DDX3X or DDX3Y for expansion.

Figure 6.

DDX3X is necessary for GC B-cell maintenance and T-cell–dependent expansion. A, Flow cytometry analysis of GC (B220+IgDCD95+GL7+) in the spleen of Vav-cre/Ddx3x-floxed mice. The t test with Welsh correction was used to compare female heterozygotes mice with female controls and male KO mice with male controls. B, Flow cytometry analysis of GC (B220+IgDCD38CD95+GL7+) in spleen of Cd19-cre/Ddx3x-floxed mice compared with controls and quantification in absolute number of cells. The Kruskal–Wallis test was used to compare female KO with female controls and heterozygous and Mann–Whitney test was used to compare male KO with male controls. C, Flow cytometry analysis of GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mice 10 days after immunization with 1 × 108 sheep RBCs injected into the lateral tail vein. For females, medians were compared using a Kruskal–Wallis test, whereas for males, medians were compared using the Mann–Whitney U test. D, Absolute number of GFP+ GC (B220+IgD CD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mice crossed with the R26mT/mG reporter mice 10 days after immunization. In females, GC GFP+ was compared using a Dunnett T3 test, whereas in males, a Mann–Whitney U test was used. E, Absolute number of GFP+ GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed female mice crossed with the R26mT/mG reporter mice analyzed at days 3, 5, and 7 after injection. Data from same time point were compared with an unpaired t test. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 6.

DDX3X is necessary for GC B-cell maintenance and T-cell–dependent expansion. A, Flow cytometry analysis of GC (B220+IgDCD95+GL7+) in the spleen of Vav-cre/Ddx3x-floxed mice. The t test with Welsh correction was used to compare female heterozygotes mice with female controls and male KO mice with male controls. B, Flow cytometry analysis of GC (B220+IgDCD38CD95+GL7+) in spleen of Cd19-cre/Ddx3x-floxed mice compared with controls and quantification in absolute number of cells. The Kruskal–Wallis test was used to compare female KO with female controls and heterozygous and Mann–Whitney test was used to compare male KO with male controls. C, Flow cytometry analysis of GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mice 10 days after immunization with 1 × 108 sheep RBCs injected into the lateral tail vein. For females, medians were compared using a Kruskal–Wallis test, whereas for males, medians were compared using the Mann–Whitney U test. D, Absolute number of GFP+ GC (B220+IgD CD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mice crossed with the R26mT/mG reporter mice 10 days after immunization. In females, GC GFP+ was compared using a Dunnett T3 test, whereas in males, a Mann–Whitney U test was used. E, Absolute number of GFP+ GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed female mice crossed with the R26mT/mG reporter mice analyzed at days 3, 5, and 7 after injection. Data from same time point were compared with an unpaired t test. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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Several reports have linked DDX3X to p53 and reported a role of this helicase in the DNA damage response (DDR; refs. 21, 25, 38). To test whether absence of DDX3X triggers p53-dependent cell death in GC B cells, we crossed Trp53+/– and Trp53–/– alleles into the Cγ1-cre/Ddx3fl/fl animals. However, Trp53 gene deletion did not rescue GC expansion in Cγ1-cre/Ddx3fl/fl mice after SRBC immunization, excluding a p53-dependent cell death as the underlying cause of this phenotype (Fig. 7A and B). Furthermore, an NP-CGG immunization demonstrated an almost identical impairment of female Cγ1-cre/Ddx3fl/fl and male Cγ1-cre/Ddx3fl/Y mice to generate antigen-specific, NP+ GC or switched memory B cells (Fig. 7C). Although non-immunized male Vav-cre/Ddx3fl/Y or Cd19-cre/Ddx3fl/Y mice also showed a reduction of GC B cells, the B220+IgDCD38+ B cells decrease is not observed in these animals (Supplementary Fig. S7D).

Figure 7.

DDX3X is required for GC B cells expansion in vivo and in vitro on a p53-independent manner. A and B, Relative percentages of GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mouse females in A and males in B crossed with the Trp53-KO 10 days after sheep RBC immunization. A Kruskal–Wallis test was used to assess statistical significance in this experiment. C,Cd19-cre/Ddx3x-floxed mice were immunized with NP-CGG and analyzed 39 days post-injection for NP-specific immune response. NP+ gate was assessed using mice that received alum without NP-CGG. Populations are defined as follows: switched memory B cells (B220+IgDCD38+IgG1+NP+), GC (B220+IgDCD95+GL7+NP+). The Mann–Whitney U test was used to compare male KO with male controls and female KO with female controls. D, Schematic representation of the in vitro coculture system allowing iGB expansion. E, Trypan blue was used to count the iGBs and follow their proliferation. Day 0 marks the day when primary B cells isolated from R26mT/mG/Cγ1-cre/Ddx3x-floxed mice were plated on 40LB feeder cells. Indicated statistics were calculated using a two-way ANOVA test at the pic of the iGB expansion (day 5). F, Micrographs of iGBs expansion 5 days after primary B cells isolated from indicated mice were cocultured on 40LB feeder cells. G, iGBs isolated from were collected at day 5 and submitted to flow cytometry analysis to evaluate GFP and dTomato emissions. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 7.

DDX3X is required for GC B cells expansion in vivo and in vitro on a p53-independent manner. A and B, Relative percentages of GC (B220+IgDCD95+GL7+) from spleens of Cγ1-cre/Ddx3x-floxed mouse females in A and males in B crossed with the Trp53-KO 10 days after sheep RBC immunization. A Kruskal–Wallis test was used to assess statistical significance in this experiment. C,Cd19-cre/Ddx3x-floxed mice were immunized with NP-CGG and analyzed 39 days post-injection for NP-specific immune response. NP+ gate was assessed using mice that received alum without NP-CGG. Populations are defined as follows: switched memory B cells (B220+IgDCD38+IgG1+NP+), GC (B220+IgDCD95+GL7+NP+). The Mann–Whitney U test was used to compare male KO with male controls and female KO with female controls. D, Schematic representation of the in vitro coculture system allowing iGB expansion. E, Trypan blue was used to count the iGBs and follow their proliferation. Day 0 marks the day when primary B cells isolated from R26mT/mG/Cγ1-cre/Ddx3x-floxed mice were plated on 40LB feeder cells. Indicated statistics were calculated using a two-way ANOVA test at the pic of the iGB expansion (day 5). F, Micrographs of iGBs expansion 5 days after primary B cells isolated from indicated mice were cocultured on 40LB feeder cells. G, iGBs isolated from were collected at day 5 and submitted to flow cytometry analysis to evaluate GFP and dTomato emissions. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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We next used an in vitro system in which primary B cells are cultured with IL-4 on a 40LB feeder layer expressing BAFF and CD40 L allowing the expansion of “induced GC” or iGB cells (Fig. 7D; ref. 39). In vitro culture of murine primary B cells from R26mT/mG/Cγ1-cre/Ddx3x-floxed mice demonstrated that a full Ddx3x deletion almost entirely prevents GC expansion (Fig. 7E), albeit some GFP+R26mT/mG/Cγ1-cre/Ddx3fl/fl cells were detectable by microscopy and flow cytometry (Fig. 7F and G). In contrast, male R26mT/mG/Cγ1-cre/Ddx3fl/Y B cells were able to proliferate and expand to almost to control levels (Fig. 7E), suggesting that under these conditions DDX3Y can almost fully compensate for DDX3X loss.

Loss of DDX3X prevents tumorigenesis in MYC-driven lymphomagenesis

Because a high frequency of DDX3X LOF mutations was reported in B-cell lymphoma where the activation of the MYC oncogene plays a significant role (2), we tested whether Ddx3x deletion could influence MYC-driven lymphomagenesis. We used mice expressing a Myc transgene driven by the IgH (Eμ) enhancer that are prone to develop a spectrum of B-lymphoid tumors ranging from pre-B cells lymphoma to IgM+ B-cell lymphoma (40). Incidences and latency period of B-cell lymphoma were unchanged in female Eμ-Myc/Vav-cre/Ddx3X/fl mice compared with Eμ-Myc/Vav-cre/Ddx3X/X (Supplementary Fig. S8A). In contrast, we observed that the majority of Eμ-Myc/Vav-cre/Ddx3fl/Y males did not develop lymphoma and showed a longer survival compared with controls (Fig. 8A). Prelymphomatous Eμ-Myc/Vav-cre/Ddx3fl/Y mice lacked pre–B-cell expansion and splenic enlargement usually observed in young, lymphoma-free Eμ-Myc mice (Fig. 8B) and opposed to what occurs in female Eμ-Myc/Vav-cre/Ddx3X/fl or female mice with both WT Ddx3x alleles (Supplementary Fig. S8B). Furthermore, B cells from Eμ-Myc/Vav-cre/Ddx3fl/Y mice presented the same phenotypes as those found in Vav-cre/Ddx3fl/Y: A decrease of immature B cells and recirculating B cells in the BM, as well as a general decrease of splenic B cells but a higher percentage of MZ B cells (Supplementary Fig. S8C).

Figure 8.

Sex dependency of Ddx3x deletion in MYC-driven lymphomagenesis. A, Kaplan–Meier curves representing survival of Eμ-Myc/Vav-cre/Ddx3-floxed male mice. Statistics were assessed with the Mantel–Cox test. The median survival is indicated in brackets and is followed by the number of animals in the cohort. Repartition of tumor subtypes was assessed by flow cytometry analysis of tumor samples. B, Flow cytometry analysis of BM and spleens from 6-week-old Eμ-Myc/Vav-cre/Ddx3 male mice (pre-tumor phase) and quantification in absolute number of cells. Splenic index corresponds to the weight of the spleen divided by the weight of the animal multiplied by 100. Statistical significance was assessed with the Kruskal–Wallis test. C, Western blot analysis of nuclear extracts from splenic B220+ cells extracted from male animals with the indicated genotypes during the pre-tumor phase. D, Flow cytometric analysis of splenocytes stained for Annexin V and the indicated surface markers. Significance was assessed by an ordinary one-way ANOVA test. E, Western blot analysis of splenocytes and thymocytes extracted from animals with indicated genotypes and age; old Eμ-Myc/Vav-cre/Ddx3fl/Y mice (<52 weeks) were lymphoma-free animals. F and G, Western blot analysis of whole cell lysates from tumors developed in Eμ-Myc/Vav-cre/Ddx3fl/Y or Eμ-Myc/Vav-cre/Ddx3X/Y as controls. H, Kaplan–Meier curves representing the survival of Eμ-Myc/Cd19-cre/Ddx3x-floxed mice. I, Kaplan–Meier curves representing the survival of λ-Myc/Cd19-cre/Ddx3x-floxed mice. λ-Myc tumors have been classified in three groups according to the intensity of the GL7 marker assessed by flow cytometry of tumor samples. Isolation and PCR genotyping of CD19+ tumor cells from KO-mice allowed to determine if a tumor achieved a complete deletion or not. On the basis of these PCR results, a deletion efficiency percentage was calculated and is indicated in the bottom left of the corresponding survival curves. Statistics for survival curves were assessed with the Mantel–Cox test. The median survival is indicated in brackets and is followed by the number of animals in the cohort. J, Western blot analysis of whole cell lysates from CD19+ tumor cells from λ-Myc/Cd19-cre/Ddx3x-floxed mice. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

Figure 8.

Sex dependency of Ddx3x deletion in MYC-driven lymphomagenesis. A, Kaplan–Meier curves representing survival of Eμ-Myc/Vav-cre/Ddx3-floxed male mice. Statistics were assessed with the Mantel–Cox test. The median survival is indicated in brackets and is followed by the number of animals in the cohort. Repartition of tumor subtypes was assessed by flow cytometry analysis of tumor samples. B, Flow cytometry analysis of BM and spleens from 6-week-old Eμ-Myc/Vav-cre/Ddx3 male mice (pre-tumor phase) and quantification in absolute number of cells. Splenic index corresponds to the weight of the spleen divided by the weight of the animal multiplied by 100. Statistical significance was assessed with the Kruskal–Wallis test. C, Western blot analysis of nuclear extracts from splenic B220+ cells extracted from male animals with the indicated genotypes during the pre-tumor phase. D, Flow cytometric analysis of splenocytes stained for Annexin V and the indicated surface markers. Significance was assessed by an ordinary one-way ANOVA test. E, Western blot analysis of splenocytes and thymocytes extracted from animals with indicated genotypes and age; old Eμ-Myc/Vav-cre/Ddx3fl/Y mice (<52 weeks) were lymphoma-free animals. F and G, Western blot analysis of whole cell lysates from tumors developed in Eμ-Myc/Vav-cre/Ddx3fl/Y or Eμ-Myc/Vav-cre/Ddx3X/Y as controls. H, Kaplan–Meier curves representing the survival of Eμ-Myc/Cd19-cre/Ddx3x-floxed mice. I, Kaplan–Meier curves representing the survival of λ-Myc/Cd19-cre/Ddx3x-floxed mice. λ-Myc tumors have been classified in three groups according to the intensity of the GL7 marker assessed by flow cytometry of tumor samples. Isolation and PCR genotyping of CD19+ tumor cells from KO-mice allowed to determine if a tumor achieved a complete deletion or not. On the basis of these PCR results, a deletion efficiency percentage was calculated and is indicated in the bottom left of the corresponding survival curves. Statistics for survival curves were assessed with the Mantel–Cox test. The median survival is indicated in brackets and is followed by the number of animals in the cohort. J, Western blot analysis of whole cell lysates from CD19+ tumor cells from λ-Myc/Cd19-cre/Ddx3x-floxed mice. ns, nonsignificant, P > 0.05; *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

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Transgenic MYC expression was lower in Eμ-Myc/Vav-cre/Ddx3fl/Y males than normally seen in Eμ-Myc transgenic mice and was even comparable with the endogenous MYC levels of animals not carrying the Eμ-Myc transgene (Fig. 8C). Because the pre–B-cell expansion that is typically observed in Eμ-Myc mice (40), was not observed in young Eμ-Myc/Vav-cre/Ddx3fl/Y males (Fig. 8B) and given the decreased numbers of B cells previously observed in male KO mice, this suggested that the concomitant activation of c-MYC and the deletion of Ddx3x had eliminated the B cells that could potentially transform and generate a lymphoma. This notion is supported by the increase of Annexin V+ pre-B cells detected in Eμ-Myc/Vav-cre/Ddx3fl/Y mice compared with Eμ-Myc/Ddx3fl/Y and Vav-cre/Ddx3fl/Y (Fig. 8D). Moreover, the analysis of Eμ-Myc/Vav-cre/Ddx3fl/Y lymphoma-free animals (<52 weeks) revealed the absence of DDX3X but also the absence of DDX3Y, although DDX3Y was clearly present in splenic B cells from WT mice of similar age or younger (Fig. 8E). DDX3Y was also detected in thymic cells, where the Eμ-Myc transgene is not expressed, regardless of age or whether Ddx3x was deleted or not (Fig. 8E), again indicating that simultaneous transgenic expression of c-Myc and Ddx3x deletion is incompatible with cell survival.

In the rare cases where Eμ-Myc/Vav-cre/Ddx3fl/Y animals developed a lymphoma (<15% of cases), we detected an upregulation of DDX3Y at levels that were higher than in control tumors (Fig. 8F and G), indicating that the loss of DDX3X can be rescued in cells by an upregulation of DDX3Y expression that are then selected to generate a B-cell lymphoma.

Next, we generated Eμ-Myc/Cd19-cre/Ddx3fl/fl animals to investigate the role of DDX3X in B-cell lymphomagenesis in which both Ddx3x alleles can be deleted specifically in B cells of female mice avoiding the pan-hematopoietic effects of the Vav-cre deleter. Ddx3x deletion significantly delayed lymphomagenesis in Eμ-Myc/Cd19-cre/Ddx3fl/fl animals compared with mice carrying Eμ-Myc/Cd19-cre transgenes and two intact Ddx3x alleles (Fig. 8H). Genotyping analysis of CD19+ tumor cells revealed that all tumors that developed in female Eμ-Myc/Cd19-cre/Ddx3fl/fl mice were clearly generated from cells that had escaped the Ddx3x deletion, demonstrating that DDX3X is critically required for lymphomagenesis. A small, but significant delay in lymphoma development was observed in male Eμ-Myc/Cd19-cre/Ddx3fl/Y but most of the tumors (62.5%) also escaped the Ddx3x deletion (Fig. 8H), which is not the case in tumors appearing in Eμ-Myc/Vav-cre/Ddx3fl/Y mice (Fig. 8A, F, and G) explaining the difference in survival curves between Eμ-Myc/Cd19-cre/Ddx3fl/Y mice and Eμ-Myc/Vav-cre/Ddx3fl/Y animals: Those curves differ most likely because of the impossibility to escape the Vav-cre deletion. However, 37.5% of tumors from Eμ-Myc/Cd19-cre/Ddx3fl/Y mice achieved a complete Ddx3x deletion, suggesting that DDX3Y can compensate for DDX3X loss in lymphomagenesis.

We confirmed these data using λ-Myc transgenic mice, which develop spontaneous, monoclonal mature B-cell lymphoma with Burkitt lymphoma characteristics owing to the deregulation of the MYC transgene by the λ light-chain enhancer (41). λ-Myc/Cd19-cre/Ddx3fl/fl female mice had a significant delay in lymphoma initiation compared with λ-Myc/Cd19-cre/Ddx3X/X (Fig. 8I). Genotyping of CD19+ tumor cells revealed that all female tumors without exception emerged from cells that had escaped Ddx3x deletion, again confirming an absolute requirement of Ddx3x for lymphomagenesis. In males, tumors were developed in both λ-Myc/Cd19-cre/Ddx3fl/Y and λ-Myc/Cd19-cre/Ddx3X/Y mice (Fig. 8I) and a full Ddx3x deletion was observed in all λ-Myc/Cd19-cre/Ddx3fl/Y tumors. The absence of DDX3X protein and elevated level of DDX3Y protein expression in Ddx3x-KO tumors was confirmed by Western blot (Fig. 8J), demonstrating the ability of DDX3Y to compensate for DDX3X loss in mature B-cell lymphomagenesis, which is more efficient in these Burkitt lymphoma–like tumors than in the malignancies from earlier B-cell stages generated in Eμ-Myc/Cd19-cre/Ddx3fl/Y mice (Fig. 8H). Moreover, the λ-Myc/Cd19-cre/Ddx3fl/Y tumors may appear more mature compared with control tumors (Fig. 8I), again in favor of a more efficient compensatory effect of DDX3Y in mature B-cell tumors. This efficient compensatory effect may explain why the tumorigenesis is accelerated in λ-Myc/Cd19-cre/Ddx3fl/Y animals.

The high frequency of mutations in the X-linked gene DDX3X in GC-derived lymphoma has raised interest in understanding its role in hematopoietic cells and their malignant transformation. Because a large proportion of DDX3X mutations found in malignancies generate LOF variants (12, 42–44), we generated conditionally deficient mice to investigate DDX3X role in hematopoiesis and lymphomagenesis. We present new evidence from these mouse models supporting a sex-dependent, critical role of DDX3X for specific steps erythropoiesis as well as lymphoid differentiation (summarized in Supplementary Fig. S8D). Most significantly, we show that male and female mice lacking Ddx3x behave differently in MYC-driven lymphomagenesis.

Because deletion of both Ddx3x alleles in females is incompatible with life past mid-gestation, it was necessary to model a full Ddx3x-KO by transplantation of cells that could be conditionally depleted in adult animals. This allowed to recognize the role of DDX3X in DN2 and DN3 pre-T cells depending on IL7R or Notch1 signaling and on signals delivered through the pre-TCR (45). Whether and how the DDX3X helicase is implicated in the interpretation of these pathways requires further investigation. However, both DN2 and DN3 pre–T-cell stages are similar to the pre–B-cell stages where DDX3X also seems to play an important role because their numbers are altered in deficient mice as shown here and in other studies (37, 46). At these stages of pre–B- and pre–T-cell differentiation, V(D)J recombination takes place to generate either a TCRβ chain in pre-T cells or an Igμ heavy chain in pre-B cells or later in B-cell development κ and λ light chains to produce IgM+ B cells (47). These processes are critical to produce mature T and B cells and require a coordination of DNA double-strand breaks and DNA repair. A role for DDX3X in these critical steps of lymphoid development was previously suggested by a report, indicating that Ddx3x-deleted small pre-B cells may express lower levels of the Bromodomain and WD repeat domain containing protein BRWD1 (37). This BRWD1 protein restricts V(D)J recombination at the Igk locus and Brwd1 mutant mice harbor similar defects to Ddx3x-deficient animals.

CSR and SHM, which occur in GC B cells, require DNA strand breaks that must be repaired without generating an abortive DDR through TP53 activation (48). Our data with Trp53/Ddx3x double KO mice demonstrate that TP53 activation is still intact in Ddx3x-deficient GC B cells, suggesting that DDX3X probably regulates the proliferative expansion of GC cells, but this need to be clarified.

Our study also provides first answers to the question whether male homologue of DDX3, DDX3Y, is expressed during blood cells formation and whether it can exert the same role as DDX3X. The observation that Vav-cre/Ddx3fl/Y pups were produced at a mendelian ratio but live born Vav-cre/Ddx3fl/fl female mice were never obtained would be consistent with the view that DDX3Y can indeed compensate for DDX3X loss at least in fetal erythropoiesis. Moreover, in vitro expansion of induced GC B cells and in vivo stimulation of GC B cells are other examples where DDX3Y can compensate for DDX3X loss.

However, any compensatory effect of DDX3Y is likely context dependent and cell type specific. This context dependency of DDX3Y is most likely due to a variance in expression levels in different cell types, or its subcellular localization, or other unknown mechanism that may differ from DDX3X and remain to be clarified. It may also be a consequence of differences in the enzymatic activity between DDX3X and DDX3Y. A recent report showed that DDX3X and DDX3Y enzymes are functionally redundant in mRNA translation catalyzing protein synthesis (49), but their function may differ on other contexts such as stress response for instance (50).

The discovery of high frequency of somatic LOF mutations in DDX3X in human Burkitt lymphoma has raised interest in the role of DDX3X in lymphomagenesis. Our observation that male Eμ-Myc/Vav-cre mice lacking DDX3X are almost free of lymphoma indicates a strict requirement of DDX3X for MYC-driven lymphomagenesis. Although this was recently suggested by others in human Burkitt lymphoma (9), it was not yet shown in a murine lymphoma model. Here, we provide evidence for a requirement of DDX3X for B-cell lymphomagenesis from both Eμ-Myc and λ-Myc transgenic mice, the latter being a widely accepted model for human Burkitt lymphoma (41).

Combination of the Eμ-Myc transgene and the pan-hematopoietic Vav-cre deleter showed that the pre–B-cell expansion that normally takes place during the pre-lymphomatous phase in Eμ-Myc mice is not detected in male mice when DDX3X is absent, suggesting that the combination of Ddx3x deletion and MYC activation eliminates these cells, as shown by the increased number of apoptotic pre-B cells; possibly through cell death triggered by increased cell stress as previously suggested (9). It is thus conceivable that Ddx3x deletion and activation of MYC eliminates the cells available for MYC-driven tumorigenesis protecting male mice against lymphoma in the Eμ-Myc/Vav-cre model. This is also supported by our finding that only B cells that upregulate DDX3Y expression can tolerate Ddx3x deletion and MYC activation, and therefore can develop a lymphoma, whereas DDX3Y is undetectable in the spleen of Eμ-Myc/Vav-cre/Ddx3fl/Y lymphoma-free mice. Lymphomagenesis in male mice lacking DDX3X occurs, therefore, only when DDX3Y is upregulated, a conclusion supported by two studies that demonstrated that DDX3Y, not present in normal human B lymphocytes, is expressed in a malignant context, in particular in Burkitt lymphoma (9, 32).

The use of the B-cell–specific Cd19-cre deleter in the Eμ-Myc and λ-Myc models showed a clearer picture, because it avoided the effects of a pan-hematopoietic deletion caused by the Vav-cre deleter in females. The data from these two models provided strong support for the notion that B cells do not tolerate Ddx3x loss in the context of an activated Myc, because lymphomas with a full deletion of both Ddx3x alleles were never observed in Eμ-Myc nor in λ-Myc female mice. Moreover, the critical requirement of DDX3 activity in general for B-cell lymphomagenesis was also evident in male mice. Indeed, 37.5% of Eμ-Myc/Cd19-cre/Ddx3fl/Y and all λ-Myc/Cd19-cre/Ddx3fl/Y mice develop tumors that have a full Ddx3x deletion, indicating that DDX3Y can compensate for DDX3X loss, sustains c-Myc activation and allows B-cell lymphoma development. This result is in agreement with a report indicating that DDX3X loss in male Burkitt lymphoma patients is compensated by re-expression of DDX3Y (9).

Another recent study demonstrates that DDX3X and DDX3Y have redundant function in translational regulation confirming that one may compensate the loss of the other (49). It is, therefore, plausible that DDX3Y compensates the deleterious effects caused by DDX3X LOF mutations in Burkitt lymphoma cells allowing MYC-driven lymphomagenesis to take place in patients with Burkitt lymphoma. Although we demonstrate that DDX3Y is expressed in normal murine lymphocytes, which is a fundamental difference compared with humans, our murine lymphoma model still leads to a very similar conclusion: DDX3Y allows lymphoma development in case of DDX3X absence. It is thus well possible that the co-occurrence of DDX3X loss and a gain of MYC represents a synthetic lethal combination that could be exploited for the development of new therapeutical options for human MYC-driven B-cell lymphoma, but also conceivable that DDX3Y represents the interesting target in male patients harboring DDX3X mutations.

M. Lacroix reports grants from the Cole Foundation and IRCM Foundation during the conduct of the study. No disclosures were reported by the other authors.

M. Lacroix: Conceptualization, resources, formal analysis, validation, investigation, visualization, methodology, writing–original draft, writing–review and editing. H. Beauchemin: Conceptualization, supervision, validation, investigation, visualization, methodology, writing–original draft. J. Fraszczak: Formal analysis, investigation, methodology. J. Ross: Formal analysis, investigation, methodology. P. Shooshtarizadeh: Conceptualization, data curation, supervision, investigation, methodology. R. Chen: Resources, methodology. T. Möröy: Conceptualization, resources, supervision, funding acquisition, validation, investigation, visualization, methodology, writing–original draft, project administration, writing–review and editing.

The authors are indebted to Mathieu Lapointe for technical assistance, Pénélope Bergeron, and Jo-Anny Bisson for excellent animal care as well as CPA team for technical assistance, Eric Massicotte and Julie Lord for FACS and cell sorting, Dominique Filion for microscopy, and Simone Terouz for histology platform. The authors thank Dr. S. Janz for sharing the λ-Myc mouse model as well as Dr. J. Di Noia for sharing the 40LB feeder cells and protocols. T. Möröy holds a Foundation grant from the CIHR (FDN-148372) and support from the Cancer Research Society (CRS). M. Lacroix is supported by fellowships from the Cole Foundation and the IRCM Foundation.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Note: Supplementary data for this article are available at Cancer Research Online (http://cancerres.aacrjournals.org/).

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Supplementary data