Adult hematopoietic stem/progenitor cell (HSPC) numbers remain stable in the absence of external stressors. After bone marrow (BM) transplant, HSPCs need to expand substantially to repopulate the BM and replenish the peripheral blood cell pool. In this study, we show that a noncanonical Wnt receptor, Frizzled-6 (Fzd6), regulates HSPC expansion and survival in a hematopoietic cell-intrinsic manner. Fzd6 deficiency increased the ratio of Flt3hi multipotent progenitors to CD150+ stem cells in the mouse BM, suggesting defective stem cell maintenance. Competitive transplantation experiments demonstrated that Fzd6/ HSPCs were able to home to the BM but were severely impaired in their capacity to reconstitute a lethally irradiated host. Lack of Fzd6 resulted in a strong activation of caspase-3 and a gradual loss of donor HSPCs and peripheral blood granulocytes. Fzd6 was also necessary for the efficient HSPC expansion during emergency hematopoiesis. Mechanistically, Fzd6 is a negative regulator of Cdc42 clustering in polarized cells. Furthermore, β-catenin–dependent signaling may be disinhibited in Fzd6/ HSPCs. Collectively, our data reveal that Fzd6 has an essential role in HSPC maintenance and survival. Noncanonical Wnt–Fzd6 signaling pathway could thus present an interesting target for promoting HSPC expansion and multilineage hematopoietic recovery after transplant.

Adult hematopoietic stem/progenitor cells (HSPCs) are maintained in a specialized niche in the bone marrow (BM). Multiple signals from the niche regulate HSPC numbers and their ability to self-renew as well as to differentiate into all blood cell types (1, 2). Microenvironmental cues also influence the cell-cycle status of HSPCs and modulate stem cell polarity. Polarity has been suggested to maintain the dynamic balance of HSPC pool by regulating the outcome (self-renewal versus differentiation) of HSPC divisions (36). The molecular mechanisms coordinating the cross-talk between niche-associated signals and HSPC-intrinsic polarity determinants remain unclear.

Wnt signaling is necessary for adult HSPC self-renewal in the BM, but it needs to be tightly regulated (79): mild increases in β-catenin–dependent signaling enhance HSPC function, whereas stronger signals would favor myeloid expansion to the detriment of stem cell self-renewal. Although noncanonical, β-catenin–independent, Wnt signaling pathways are known to modulate cell polarity, cell motility, and tissue patterning (1017), their role in hematopoietic cells is much less well established. Exogenous Wnt5a has been shown to activate β-catenin–independent signaling in LinSca1+c-Kithi (LSK) hematopoietic progenitors and to improve HSPC maintenance and function by promoting their quiescence (1821). By contrast, hematopoietic Wnt5a expression is augmented during HSPC aging and decreasing Wnt5a levels in aged HSPCs resulted in the restoration of their polarity and self-renewal (22). Wnt4 enhances the expansion of fetal liver LSKs, most notably Flt3hi LSKs, and improves thymic recovery after irradiation and hematopoietic cell transplant through mechanisms that depend on Jnk2 but do not require β-catenin or its binding partner Tcf1 (23, 24). Lastly, inhibition of canonical Wnt signaling through the activation of Frizzled (Fzd)-8 and the atypical cadherin Fmi maintained quiescent long-term hematopoietic stem cells (HSCs) in the adult mouse BM (25). Most polarity genes are expressed in fetal HSPCs (23); however, the direct role of most core noncanonical Wnt signaling effectors and planar cell polarity (PCP) have not been addressed in hematopoietic cells.

We recently reported that the receptor Fzd6 was at least partially required for the Wnt4-mediated expansion of fetal liver HSPCs in culture (23). Furthermore, we observed alterations in the frequency of BM HSPCs in Fzd6−/− mice on a mixed C57BL/6 × 129sv background. Fzd6 expression has been demonstrated in HSPCs and mature blood-forming cells in human and mouse (26) with the strongest expression levels corresponding to more immature cell types (27, 28). Fzd6 is generally associated with PCP signaling in epithelial cells (10, 11, 13, 14), and it has been previously proposed to act as a negative regulator of the β-catenin–dependent canonical Wnt pathway (15, 29). Very little is known about the functional role of Fzd6 signaling in the hematopoietic lineage, except for its being involved in the initiation and progression of chronic lymphocytic leukemia in the Eμ-TCL1 mouse model (30).

In this study, we have examined the impact of Fzd6 in the regulation of HSPC function using mice on C57BL/6 background. We report a cell-intrinsic requirement for Fzd6 in competitive short- and long-term hematopoietic reconstitution after irradiation and transplant. We show that Fzd6 deficiency impaired the expansion and survival of HSPCs, resulting in the activation of caspase-3 and inefficient HSPC engraftment in the first week after BM transplant. We further show that Fzd6 is necessary for efficient emergency responses during systemic inflammation. Overall, we demonstrate a crucial role for Fzd6 in HSPC self-renewal and negative regulation of Cdc42/Jnk signaling in the BM and suggest that Fzd6 could be an interesting target for stimulating HSPC expansion in culture or in situ posttransplant.

C57BL/6 (B6; CD45.2+) and B6.SJL-PtprcaPep3b/BoyJ (Ly5a) (B6.SJL; CD45.1+) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Mice deficient in Frizzled-6 (Fzd6−/−) are described elsewhere (14) and were originally a kind gift from J. Nathans (Johns Hopkins, Baltimore, MD). Fzd6+/− mice from mixed C57BL/6 × 129sv background were backcrossed to C57BL/6 for 10 generations and then maintained as Fzd6+/− to Fzd6+/ intercrosses. Fzd6−/− mice were compared with sex-matched Fzd6+/+ littermates. For competitive transplant experiments, first-generation progeny from B6.SJL × Fzd6+/+ intercrosses were used as recipients, and their littermates or B6.SJL mice were used as competing donors. For noncompetitive short-term transplants and homing experiments, the recipient mice were B6.SJL. All mice were bred and housed under specific pathogen-free conditions in sterile ventilated racks at the animal facility of Institut National de la Recherche Scientifique–Institut Armand-Frappier (Centre National de Biologie Expérimentale). All procedures were in accordance with the Canadian Council on Animal Care guidelines.

BM was harvested by flushing tibias and femurs with PBS/0.1% BSA/0.5 mmol EDTA using a 25-gauge needle. To analyze BM HSPCs, the following Abs were used: biotin-conjugated anti-lineage mAbs anti-CD3ε (145-2C11), anti-CD11b (M1/70), anti-CD45/B220 (RA3-6B2), anti-GR1 (RB6-8C5), and anti-Ter119; streptavidin conjugated to BD Horizon-V500 (BD Biosciences, Mississauga, ON, Canada) or FITC (BD Biosciences); PE anti-CD117 (c-Kit, 2B8; BD Biosciences), PE-Cy7 anti–Sca-1 (Ly6A/E, D7; BD Biosciences), PerCP–eFluor 710 anti-CD135 (Flt3, A2F10), and Alexa Fluor 647 anti-CD150 (TC15-12F12.2; BD Biosciences). PE-Cy7 anti-CD3, PE anti-CD19 (1D3), allophycocyanin or FITC anti-CD45.1 (A20), eFluor 450 or FITC anti-CD45.2 (104), and allophycocyanin-Cy7 anti-GR1 (BD Biosciences) were used for the analysis of peripheral blood chimerism. For intracellular staining, surface-stained BM cells were fixed and permeabilized using the Foxp3 staining kit (eBioscience, San Diego, CA) and then incubated with PE anti–β-catenin (15B8), Alexa Fluor 488 active Caspase-3 (Cell Signaling Technologies), PE–eFluor 610 anti-Ki67 (SoIA15), PE anti–p-JNK1/2 (pT183/pY185; BD Biosciences), unconjugated anti-active β-catenin (D13A1; Cell Signaling Technologies), unconjugated anti-Celsr1 (Abcam), or appropriate isotype controls. Unconjugated Abs were detected with a PE-conjugated F(ab′)2 fragment against rabbit IgG (Molecular Probes). All Abs were purchased from eBioscience unless indicated otherwise. For cell-cycle analysis, BM cells were first incubated for 30 min at 37°C with Hoechst #33342 (Sigma-Aldrich, Oakville, ON, Canada) in DMEM supplemented with 10% Premium FBS (Wisent Bioproducts, St-Bruno, QC, Canada) and 1 mmol HEPES (Life Technologies, Burlington, ON, Canada), followed by staining with surface Abs and intracellular anti-Ki67 as described above. The FluoReporter lacZ Flow Cytometry kit (Life Technologies) was used for the detection of β-galactosidase expression from the Fzd6 mutant allele. Samples were acquired with a four-laser LSR Fortessa flow cytometer (BD Biosciences, Mountain View, CA) and analyzed using BD FACS Diva software (BD Biosciences) or FlowJo (for histogram overlays; Tree Star). For cell sorting, BM cells were stained as above. For fetal liver HSCs, CD11b was removed from the lineage panel, and cells were further stained with Alexa Fluor 647 anti-CD150 and PE-Cy7 anti–Sca-1, and allophycocyanin–Alexa Fluor 780 CD11b samples were sorted for purity with a three-laser FACSAria II (BD Biosciences).

BM cells were harvested as above and enriched for c-Kit+ cells using biotinylated anti-mouse CD117 (BD Biosciences) and EasySep biotin selection kit (Stem Cell Technologies, Vancouver, BC, Canada). HSPCs were further identified by PE-Cy7 anti–Sca-1 and Alexa Fluor anti-CD150. Flow cytometry analysis conducted in parallel confirmed that all Sca-1+ CD150+ cells were also c-Kit+. Cells were washed in PBS, fixed, and permeabilized at 4°C for 90 min, washed, and blocked with 3% BSA for 30 min, followed by staining with anti-Cdc42 (EMD Millipore) for 1 h. Staining was detected with a PE-conjugated F(ab′)2 fragment of goat anti-rabbit IgG (Molecular Probes) in blocking buffer for 1 h. Cells were counterstained with DAPI (Life Technologies). Stained cells were washed in PBS and acquired with Amnis Imagestream Mark II imaging flow cytometer (EMD Millipore) and analyzed with IDEAS v6.1 software. A cell was considered polarized when Cdc42 protein showed a distinctly asymmetrical distribution when a line was drawn across the middle of the cell.

Single-cell suspensions were prepared in IMDM containing 10% Premium FBS (Wisent Bioproducts), and cells were seeded into 35-mm nonadherent petri dishes at a density of 104 cells/dish in methylcellulose medium containing stem cell factor, IL-3, IL-6, and erythropoietin (Methocult GF M3434; Stem Cell Technologies). The cultures were incubated at 37°C in 5% CO2 for 7–10 d, and hematopoietic colonies were counted and identified based on morphology under an inverted microscope. Harvested colony-forming cells were further stained with Abs against CD11b, CD11c, CD117/c-Kit, GR1, CD45.1 and CD45.2. Stained cells were analyzed as described above.

For competitive long-term reconstitution experiments, 5 × 105Fzd6−/− or Fzd6+/+ BM cells (CD45.2+) were mixed with 5 × 105 competitor cells (CD45.1+ or CD45.1+/CD45.2+) and injected into the lateral tail vein of lethally irradiated (two doses of 450 rad, 16 h apart) congenic recipient mice. For secondary transplants, an equal number (2 × 106) of total BM cells from two primary recipients were pooled and injected into lethally irradiated secondary recipients. For short-term analysis, 10 × 106 donor cells were transferred without competition. To determine peripheral blood chimerism, blood samples were collected through the mandibular vein from recipient mice at 4, 8, 12, and 16 wk and analyzed by flow cytometry. Sixteen weeks after transplantation, mice were euthanized and analyzed for reconstitution in BM and spleen.

To determine the homing efficiency of hematopoietic cells, 20 × 106 BM cells from Fzd6−/− or Fzd6+/+ donors (CD45.2+) were injected into sublethally irradiated (500 rad) CD45.1+ recipient mice. Mice were euthanized and their BM harvested 16 h later for flow cytometry and functional analysis of donor HSPCs.

To evaluate HSPC expansion during emergency hematopoiesis, Fzd6−/− and Fzd6+/+ mice were given two i.p. injections of γ-irradiated LPS (E. coli 0111:B4; Sigma-Aldrich) at a dose of 1 mg/kg body weight, 48 h apart (31). Their BM was harvested 24 h after the second injection for flow cytometry and functional analysis.

Each value represents at least three independent experiments. Two-tailed Student t test was used to determine statistical significance unless otherwise noted. A p value <0.05 was considered significant.

Fzd6 mRNA is abundantly expressed by fetal liver (23) as well as BM HSPCs (28). However, given the different methods of detection and the variable markers used for cell identification, we wanted to validate these results and directly compare Fzd6 expression levels in fetal liver and adult mouse BM CD150+ LSK cells, which are enriched in HSCs (32), as well as different HSPC subpopulations in the BM. Fzd6 mRNA levels were comparable between adult and fetal liver CD150+ HSPCs as detected by quantitative RT-PCR on sorted cells (Fig. 1A). To evaluate Fzd6 expression in different BM HSPC subpopulations, we took advantage of the LacZ construct inserted into the Fzd6 mutant allele (14). Using fluorescein di-V-galactoside as the substrate for β-galactosidase, only CD150+ HSPCs displayed detectable levels of Fzd6 promoter activity (Fig. 1B). These results correlate well with previously published data (28) as well as data found in the Immunological Genome project database (26) (http://www.immgen.org) and support the hypothesis that Fzd6 play a key role in the regulation of fetal and adult hematopoiesis.

FIGURE 1.

Noncanonical Fzd6 is expressed on CD150+ HSCs and influences the ratio of Flt3+ versus CD150+ progenitors. (A) Analysis of Fzd6 expression by quantitative RT-PCR in the embryonic day (E) 13.5 fetal liver and 6-wk-old adult BM HSCs. Histogram represents mean + SEM from three independent experiments. (B) β-Galactosidase expression from the Fzd6-nLacZ allele was determined by its ability to cleave the fluorescent substrate FDG in Fzd6−/− (knockout [KO]) and BM stem/progenitor cells. Fzd6+/+ (wild-type [WT]) cells were used as negative control. Similar results were obtained from three independent experiments. (C) Intracellular active β-catenin levels in BM CD150+Fzd6−/− and Fzd6+/+ HSCs. Similar results were obtained from four independent experiments. (D) Flow cytometry analysis of the proportion of CD150+ CD135 (Flt3) HSCs, CD150CD135 MPPs, and CD150CD135+/hi LMPPs. Representative flow cytometry data are shown for 6-wk-old Fzd6−/− and Fzd6+/+ BM. Numbers within the flow cytometry panels represent the mean percentages of total LSKs as well as the different LSK subsets over total LSKs from eight animals per group. Histogram represents the absolute numbers of BM LSK subpopulations HSC, MPP, and LMPP in 3-wk (n = 6) and 6-wk-old (n = 8) Fzd6−/− and Fzd6+/+ mice (mean + SEM). (E) Cell-cycle analysis of CD150+ HSCs. Ki-67/Hoechst costaining was used to distinguish the G0, G1, and S/G2/M cell cycle phases. Histogram represents mean + SEM from three animals per group. *p < 0.05, **p < 0.005 (two-tailed, unpaired Student t test).

FIGURE 1.

Noncanonical Fzd6 is expressed on CD150+ HSCs and influences the ratio of Flt3+ versus CD150+ progenitors. (A) Analysis of Fzd6 expression by quantitative RT-PCR in the embryonic day (E) 13.5 fetal liver and 6-wk-old adult BM HSCs. Histogram represents mean + SEM from three independent experiments. (B) β-Galactosidase expression from the Fzd6-nLacZ allele was determined by its ability to cleave the fluorescent substrate FDG in Fzd6−/− (knockout [KO]) and BM stem/progenitor cells. Fzd6+/+ (wild-type [WT]) cells were used as negative control. Similar results were obtained from three independent experiments. (C) Intracellular active β-catenin levels in BM CD150+Fzd6−/− and Fzd6+/+ HSCs. Similar results were obtained from four independent experiments. (D) Flow cytometry analysis of the proportion of CD150+ CD135 (Flt3) HSCs, CD150CD135 MPPs, and CD150CD135+/hi LMPPs. Representative flow cytometry data are shown for 6-wk-old Fzd6−/− and Fzd6+/+ BM. Numbers within the flow cytometry panels represent the mean percentages of total LSKs as well as the different LSK subsets over total LSKs from eight animals per group. Histogram represents the absolute numbers of BM LSK subpopulations HSC, MPP, and LMPP in 3-wk (n = 6) and 6-wk-old (n = 8) Fzd6−/− and Fzd6+/+ mice (mean + SEM). (E) Cell-cycle analysis of CD150+ HSCs. Ki-67/Hoechst costaining was used to distinguish the G0, G1, and S/G2/M cell cycle phases. Histogram represents mean + SEM from three animals per group. *p < 0.05, **p < 0.005 (two-tailed, unpaired Student t test).

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Fzd6 has been associated with noncanonical Wnt signaling (13, 14) with the potential to inhibit β-catenin–dependent canonical Wnt pathway in nonhematopoietic cells (15, 29). In contrast, Fzd6 expression correlated with increased levels of intracellular β-catenin in a mouse model of chronic B lymphocytic leukemia although there was no evidence of a direct functional relationship (30). To determine whether lack of Fzd6 modulated β-catenin in normal BM HSPCs, we quantified intracellular active β-catenin levels by flow cytometry using an Ab specific for the stable, nonphosphorylated form of the protein. We observed no difference in β-catenin staining between Fzd6+/+ and Fzd6−/− CD150+ HSPCs (Fig. 1C), indicating that Fzd6 was not involved in the maintenance of intracellular β-catenin levels in HSPCs at steady state and thus appeared to signal through a noncanonical pathway.

To determine whether Fzd6 was required for resting hematopoiesis, we examined 3- and 6-wk-old Fzd6−/− and sex-matched littermate Fzd6+/+ mice. The frequency and the numbers of BM LSKs, CD150 multipotent progenitors (MPPs), and CD150+CD135 HSCs were not altered in Fzd6−/− weanling or young adult mice. Interestingly, 6-wk-old adult Fzd6−/− BM presented a modest, ∼2-fold increase in the number of LSKs with high cell-surface Flt3 (CD135; lymphoid-primed MPPs [LMPPs], whereas no significant differences were observed at 3 wk (Fig. 1D). This was largely due to an overall increase in Flt3 staining intensity in Fzd6−/− BM, suggesting defective HSC maintenance. Given that a relative increase in more differentiated, Flt3+ subsets could correlate with HSC activation (25, 33, 34), our observation led us to speculate that Fzd6 might be involved in the regulation of HSC turnover.

During resting hematopoiesis, the majority of long-term HSCs remain in a quiescent state with only a small fraction that are actively cycling. Adult HSC self-renewal has been frequently linked to HSC quiescence because disruption of quiescence leads to defects in HSC self-renewal (25, 3436). More specifically, noncanonical Wnt5a signaling favors HSC maintenance via quiescence (19, 21, 25). To assess whether Fzd6 regulated HSPC cycling, we stained Fzd6+/+ and Fzd6−/− BM cells with the DNA dye Hoechst and an Ab against Ki-67. Somewhat unexpectedly, we found no significant difference in the proportion of resting (G0) HSPCs (Fig. 1E). However, we did observe a slight but reproducible decline in the proportion of dividing (S-G2-M phase) Fzd6−/− HSCs (Fig. 1E). These results suggested that Fzd6 signaling might promote cell-cycle progression and proliferation of HSCs.

To determine whether Fzd6−/− HSPCs were functionally biased toward lymphoid fates given their elevated levels of Flt3 expression (37), we assessed the frequency of early T lineage progenitors in the thymus and mature T and B lymphoid cell subsets in BM and spleen. Although we saw some evidence of an augmented lymphoid output, as shown by an increase in Fzd6−/− thymic cellularity, flow cytometry analysis revealed no significant differences in the proportion of various lymphoid subsets in the BM or lymphoid organs (spleen and thymus) between Fzd6−/− and Fzd6+/+ mice (Supplemental Fig. 1). Our data therefore indicate that Fzd6 deficiency increases the ratio of Flt3hi LMPPs to CD150+ HSCs within the LSK compartment (Fig. 1D), but the potential lymphoid bias appears compensated for during steady-state hematopoiesis.

Noncanonical Wnt signaling can modulate cell divisions (11, 12), HSPC quiescence (19, 21, 25), and polarity (22). In particular, an increase in Wnt5a/Cdc42-dependent signaling has been linked to the apolarization of HSPCs during aging (4, 22), whereas Fzd8/Fmi-dependent suppression of HSPC activation is necessary for the maintenance of long-term HSCs (25). To evaluate the role of Fzd6 in noncanonical signaling, we examined the expression of Celsr1, which has been shown to interact with Fzd6 in other tissues (16), phosphorylated active JNK, and Cdc42 in Fzd6−/− and Fzd6+/+ HSPCs by flow cytometry. We did not observe any difference in Celsr1 expression (Fig. 2A), although we cannot conclude about its localization. However, both p-JNK and Cdc42 were present at higher levels in Fzd6−/− HSPCs (Fig. 2A), suggesting that Fzd6 might negatively regulate Cdc42/JNK signaling.

FIGURE 2.

Fzd6 is a negative regulator of Cdc42/Jnk signaling. (A) Flow cytometry analysis of the expression of Celsr1, p-Jnk (T183/Y185), and Cdc42 in BM CD150+Fzd6−/− and Fzd6+/+ HSCs. Similar results were obtained from at least three independent experiments. (B) Imaging flow cytometry analysis of Cdc42 staining in BM CD150+ Sca1+ cells. The data are pooled from two independent experiments with a total of four mice per genotype. The x-axis of the histogram represents the staining intensity of Cdc42 clusters for Fzd6−/− and Fzd6+/+ cells. The table shows the numbers of cells within the Cdc42 dim gate (left side of the histogram) and the Cdc42 bright gate (right side of the histogram) according to genotype. The numbers in parentheses represent polarized events (polarized Cdc42 staining) in each group. p < 0.0001 (Fisher exact test). (C) Representative images of Fzd6−/− and Fzd6+/+ cells in G0/G1 and G2/M phases of the cell cycle. The bottom images show the staining control (secondary Ab only) for Cdc42.

FIGURE 2.

Fzd6 is a negative regulator of Cdc42/Jnk signaling. (A) Flow cytometry analysis of the expression of Celsr1, p-Jnk (T183/Y185), and Cdc42 in BM CD150+Fzd6−/− and Fzd6+/+ HSCs. Similar results were obtained from at least three independent experiments. (B) Imaging flow cytometry analysis of Cdc42 staining in BM CD150+ Sca1+ cells. The data are pooled from two independent experiments with a total of four mice per genotype. The x-axis of the histogram represents the staining intensity of Cdc42 clusters for Fzd6−/− and Fzd6+/+ cells. The table shows the numbers of cells within the Cdc42 dim gate (left side of the histogram) and the Cdc42 bright gate (right side of the histogram) according to genotype. The numbers in parentheses represent polarized events (polarized Cdc42 staining) in each group. p < 0.0001 (Fisher exact test). (C) Representative images of Fzd6−/− and Fzd6+/+ cells in G0/G1 and G2/M phases of the cell cycle. The bottom images show the staining control (secondary Ab only) for Cdc42.

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The localization of Cdc42 appears essential for its role in HSPC polarity and aging (22). To visualize Cdc42 in Fzd6−/− and Fzd6+/+ HSPCs, we resorted to imaging flow cytometry and evaluated Cdc42 polarization (asymmetrical distribution within the cell) and the staining intensity of Cdc42 clusters (bright detail intensity). There was no difference in the proportion of HSPCs with polarized Cdc42 between Fzd6−/− and Fzd6+/+ BM cells (32 and 32%, respectively). However, Fzd6−/− HSPCs showed a greater proportion of bright Cdc42 clusters (Fig. 2B, 2C), with almost all polarized HSPCs falling under the Cdc42 bright category (96 versus 67% for Fzd6+/+). Collectively, these results suggest that Fzd6 provides a regulatory function to restrict Cdc42 clustering and thus negatively regulates Cdc42/JNK signaling.

To directly determine whether Fzd6 was required for cell-intrinsic regulation of HSPC function in vivo, we performed competitive transplantation assays. Fresh BM cells from Fzd6−/− and Fzd6+/+ mice (CD45.2+) were mixed with congenic competitor BM cells at a 1:1 ratio and transferred into lethally irradiated recipients (Fig. 3A). Donor chimerism was assessed by flow cytometry analysis of peripheral blood leukocytes at 4, 8, 12, and 16 wk posttransplant. Approximately half of the mice having received Fzd6−/− cells displayed poor reconstitution already at 4 wk posttransplant (Fig. 3B). The defects in the repopulating ability of Fzd6−/− HSPCs became even more pronounced at 12 and 16 wk (Fig. 3B) with a progressive loss of Fzd6−/− GR1hi granulocytes, which suggested defective long-term HSC self-renewal (Fig. 3C). To further determine the cell-intrinsic role of Fzd6 in myelolymphoid differentiation in vivo, the contribution to B cells, T cells, and GR1hi myeloid cells within CD45.2+ donor cells was evaluated by flow cytometry analysis of peripheral blood at different time points. There was no difference in the distribution of granulocytes, monocytes, and lymphocytes within Fzd6−/− donor cells when compared with Fzd6+/+ donors at 8 wk after transplant. However, the proportion of granulocytes decreased significantly with time, whereas the relative contribution of Fzd6−/− cells to the more long-lived lymphoid lineages and monocytes was not negatively affected (Fig. 3D). To obtain a more quantitative representation of lymphoid and myeloid lineages, we also analyzed the spleen at 16 wk after transplant. The numbers of Fzd6−/− donor-derived cells were significantly decreased for all subsets (Fig. 3D), with an ∼5-fold difference for B lymphocytes and monocytes and an ∼10-fold decrease for T lymphocytes and granulocytes.

FIGURE 3.

Fzd6 is required for short-term and long-term competitive hematopoietic reconstitution. (A) Experimental design of the competitive reconstitution assay. (B) Peripheral blood (PB) chimerism in primary recipient mice at 4, 8, 12, and 16 wk after transplant. Representative flow cytometry data at 4 and 16 wk are shown in the left panel. Pooled data from two independent groups of transplants are shown in the right panel. Dots represent individual mice (open circles for Fzd6+/+ donors; black circles for Fzd6−/− donors), and horizontal lines represent the mean (solid line, Fzd6+/+; dotted line, Fzd6−/−). (C) Flow cytometry analysis of short- and long-term Fzd6+/+ and Fzd6−/− donor cell contribution to peripheral blood SSChi GR1hi granulocytes. (D) Representative flow cytometry data and quantitative analysis of the relative distribution of CD19+ B lymphocytes, CD3ε+ T lymphocytes, CD11b+GR1lo monocytes, and GR1hiSSChi granulocytes among donor-derived peripheral blood cells (top right panel) and donor-derived splenocytes (bottom right panel) in recipient mice. Histograms indicate mean + SEM from two independent experiments (n = 8). *p < 0.05, **p < 0.005 (two-tailed, unpaired Student t test).

FIGURE 3.

Fzd6 is required for short-term and long-term competitive hematopoietic reconstitution. (A) Experimental design of the competitive reconstitution assay. (B) Peripheral blood (PB) chimerism in primary recipient mice at 4, 8, 12, and 16 wk after transplant. Representative flow cytometry data at 4 and 16 wk are shown in the left panel. Pooled data from two independent groups of transplants are shown in the right panel. Dots represent individual mice (open circles for Fzd6+/+ donors; black circles for Fzd6−/− donors), and horizontal lines represent the mean (solid line, Fzd6+/+; dotted line, Fzd6−/−). (C) Flow cytometry analysis of short- and long-term Fzd6+/+ and Fzd6−/− donor cell contribution to peripheral blood SSChi GR1hi granulocytes. (D) Representative flow cytometry data and quantitative analysis of the relative distribution of CD19+ B lymphocytes, CD3ε+ T lymphocytes, CD11b+GR1lo monocytes, and GR1hiSSChi granulocytes among donor-derived peripheral blood cells (top right panel) and donor-derived splenocytes (bottom right panel) in recipient mice. Histograms indicate mean + SEM from two independent experiments (n = 8). *p < 0.05, **p < 0.005 (two-tailed, unpaired Student t test).

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Flow cytometry analysis of BM cells from the primary recipient mice confirmed the absence of Fzd6−/− HSCs in the majority of recipients at 16 wk after transplantation (Fig. 4A). Consistent with the peripheral blood analysis, in vitro colony-forming ability of myeloid progenitor cells of Fzd6−/− origin was also significantly decreased (Fig. 4B). To further test the self-renewal activity of donor cells, we selected primary recipients of Fzd6−/− BM cells with detectable donor contribution at 16 wk (11–32% total chimerism in peripheral blood) together with representative primary recipients of Fzd6+/+ cells for secondary transplants. As expected from the frequency of phenotypic donor HSCs (Fig. 4A), Fzd6−/− donor cells were unable to establish multilineage hematopoietic reconstitution (>1% in all lineages) in secondary recipients, even at short-term (Fig. 4C). In comparison, Fzd6+/+ donor cells were present in secondary recipients at a frequency that was at least equivalent to that detected in primary recipients (Fig. 4C, 4D). These data strongly suggest that Fzd6 is crucial for in vivo repopulating activity and HSC self-renewal capacity, particularly under stress.

FIGURE 4.

Fzd6−/− HSPCs display defective long-term engraftment and self-renewal in vivo. (A) Representative flow cytometry data from peripheral blood (PB; left panel) and BM 16 wk after transplant. Pooled data from two independent groups of transplants are shown in the right panel. Dots represent individual mice (open circles for Fzd6+/+ donors; black circles for Fzd6−/− donors), and horizontal lines represent the mean (solid line, Fzd6+/+; dotted line, Fzd6−/−). n = 8 for both groups. (B) Flow cytometry analysis of cells recovered from CFC assays. Left panel represents the percentage of donor-derived cells among all cells. Right panel shows the distribution GR1hi granulocytes, GR1lo/neg monocytes, and CD11c+ dendritic cells among donor-derived CD11b+ cells. Similar results were obtained from four independent experiments. (C) Representative flow cytometry data from peripheral blood of secondary recipients. Left panel represents total chimerism, and right panel shows the contribution of donor-derived cells to GR1hi granulocytes. Numbers indicate the mean from one experiment (n = 4 for both groups). Similar results were obtained from a second, independent transplant experiment. **p < 0.005 (two-tailed, unpaired Student t test).

FIGURE 4.

Fzd6−/− HSPCs display defective long-term engraftment and self-renewal in vivo. (A) Representative flow cytometry data from peripheral blood (PB; left panel) and BM 16 wk after transplant. Pooled data from two independent groups of transplants are shown in the right panel. Dots represent individual mice (open circles for Fzd6+/+ donors; black circles for Fzd6−/− donors), and horizontal lines represent the mean (solid line, Fzd6+/+; dotted line, Fzd6−/−). n = 8 for both groups. (B) Flow cytometry analysis of cells recovered from CFC assays. Left panel represents the percentage of donor-derived cells among all cells. Right panel shows the distribution GR1hi granulocytes, GR1lo/neg monocytes, and CD11c+ dendritic cells among donor-derived CD11b+ cells. Similar results were obtained from four independent experiments. (C) Representative flow cytometry data from peripheral blood of secondary recipients. Left panel represents total chimerism, and right panel shows the contribution of donor-derived cells to GR1hi granulocytes. Numbers indicate the mean from one experiment (n = 4 for both groups). Similar results were obtained from a second, independent transplant experiment. **p < 0.005 (two-tailed, unpaired Student t test).

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To understand the mechanisms underlying the observed defects in the engraftment and reconstitution capacity of Fzd6−/− HSPCs, we first examined the role of Fzd6 in BM homing. To assess the homing efficiency of HSPCs, Fzd6−/− or Fzd6+/+ BM cells were transplanted into sublethally irradiated congenic recipient mice, and the numbers of donor-derived HSPCs in the BM were determined by flow cytometry at 16 h (Fig. 5A). Our results revealed no differences in the ability of Fzd6−/− cells to home to the BM when compared with Fzd6+/+ controls (Fig. 5B). Furthermore, the percentage of donor-derived granulocytes generated in a colony-forming assay was similar for both Fzd6+/+ and Fzd6−/− donors (Fig. 5C). Not surprisingly, we saw no major differences in the expression levels of Cxcr4 or CD44 between Fzd6+/+ and Fzd6−/− HSPCs at baseline (Fig. 5D). These results indicate that the presence of Fzd6 is not necessary for migration to the BM. It might rather be required for HSPC expansion and maintenance.

FIGURE 5.

Defective long-term reconstitution of Fzd6−/− HSPCs is not due to altered homing. (A) Experimental design for the homing assay. (B) Representative flow cytometry data showing the percentage donor-derived cells and donor-derived CD150+ HSCs in recipient BM 16 h after transplant. Histogram represents the absolute number of donor-derived HSCs in BM. Similar results were obtained from two independent experiments with three mice per group in each experiment. (C) Frequency of donor-derived granulocytes among cells recovered from mixed colony assays seeded with BM harvested 16 h after transplant. Similar results were obtained from two independent experiments. (D) Representative flow cytometry data depicting CD44 and Cxcr4 expression on Fzd6+/+ (black line) and Fzd6−/− (solid gray) BM HSPCs. Similar results were obtained from four pairs of mice.

FIGURE 5.

Defective long-term reconstitution of Fzd6−/− HSPCs is not due to altered homing. (A) Experimental design for the homing assay. (B) Representative flow cytometry data showing the percentage donor-derived cells and donor-derived CD150+ HSCs in recipient BM 16 h after transplant. Histogram represents the absolute number of donor-derived HSCs in BM. Similar results were obtained from two independent experiments with three mice per group in each experiment. (C) Frequency of donor-derived granulocytes among cells recovered from mixed colony assays seeded with BM harvested 16 h after transplant. Similar results were obtained from two independent experiments. (D) Representative flow cytometry data depicting CD44 and Cxcr4 expression on Fzd6+/+ (black line) and Fzd6−/− (solid gray) BM HSPCs. Similar results were obtained from four pairs of mice.

Close modal

Given the problems with myeloid reconstitution already present at 4 wk posttransplant (Fig. 3C), we hypothesized that the loss of Fzd6−/− donor HSPCs would be an early event. We therefore decided to investigate the initial HSPC expansion to repopulate the recipient BM at 4 and 8 d posttransplant (Fig. 6A). Fzd6−/− HSPCs were present at comparable numbers on day 4 but failed to expand between days 4 and 8 (Fig. 6B). Cell-cycle analysis correlated with cell numbers, with the proportion of Fzd6+/+ donor cells in the proliferating phase increasing between the two time points in contrast to Fzd6−/− HSPCs (Fig. 6D). Fzd6−/− cells showed similar behavior in both noncompetitive (Fig. 6) and competitive settings (Figs. 3, 4), suggesting a decrease in not only relative Fzd6−/− HSPC fitness but also in absolute terms. These results parallel the cell cycle analysis at steady state (Fig. 1E), suggesting that either: 1) Fzd6 is directly implicated in cell-cycle progression; or that 2) dividing Fzd6−/− HSPCs are lost due to differentiation or cell death.

FIGURE 6.

Fzd6−/− HSPCs cannot expand and die by apoptosis in the first week after transplant. (A) Experimental design for the short-term transplants. (B) Numbers of donor-derived HSCs in BM on days 4 and 8 after transplant. Graph represents pooled results from two independent experiments with three to five mice per group. (C) Representative flow cytometry data depicting p-Jnk (T183/Y185) staining in Fzd6+/+ and Fzd6−/− donor HSPCs. Similar results were obtained from four pairs of mice. (D) Cell-cycle analysis of donor-derived HSCs in the recipient BM. Graphs represent pooled results from two independent experiments with two to five mice per group. (E) Representative flow cytometry data depicting β-catenin and cleaved caspase-3 staining in Fzd6+/+ and Fzd6−/− donor HSPCs. Similar results were obtained from four pairs of mice. (F) Representative flow cytometry data depicting the proportion of c-Kit and CD11b staining within Fzd6+/+ and Fzd6−/− donor-derived cells recovered from mixed colony assays. Similar results were obtained from four pairs of mice. *p < 0.05, **p < 0.005.

FIGURE 6.

Fzd6−/− HSPCs cannot expand and die by apoptosis in the first week after transplant. (A) Experimental design for the short-term transplants. (B) Numbers of donor-derived HSCs in BM on days 4 and 8 after transplant. Graph represents pooled results from two independent experiments with three to five mice per group. (C) Representative flow cytometry data depicting p-Jnk (T183/Y185) staining in Fzd6+/+ and Fzd6−/− donor HSPCs. Similar results were obtained from four pairs of mice. (D) Cell-cycle analysis of donor-derived HSCs in the recipient BM. Graphs represent pooled results from two independent experiments with two to five mice per group. (E) Representative flow cytometry data depicting β-catenin and cleaved caspase-3 staining in Fzd6+/+ and Fzd6−/− donor HSPCs. Similar results were obtained from four pairs of mice. (F) Representative flow cytometry data depicting the proportion of c-Kit and CD11b staining within Fzd6+/+ and Fzd6−/− donor-derived cells recovered from mixed colony assays. Similar results were obtained from four pairs of mice. *p < 0.05, **p < 0.005.

Close modal

To investigate the second possibility, we stained posttransplant BM cells with an Ab against the active form of caspase-3 together with β-catenin. Sustained β-catenin expression has been shown to activate caspase-3 and result in loss of HSPCs shortly after transplant (38). Whereas Fzd6+/+ donor HSPCs were mostly caspase-3 negative, 35–75% Fzd6−/− HSPCs stained positive for active caspase-3 (Fig. 6E). Caspase-3–positive cells also costained strongly for β-catenin (relative fluorescent intensity of 21–50 for Fzd6−/− cells and 8.5–11 for Fzd6+/+). Furthermore, colonies generated by Fzd6−/− HSPCs contained a smaller proportion of cKit+ cells, which are indicative of colony-forming cells with high proliferative potential (Fig. 6F), suggesting that a larger proportion of the remaining Fzd6−/− HSPCs had initiated myeloid differentiation. In contrast to steady-state, we did not observe any difference in p-JNK activity (Fig. 6C). Collectively with the data presented in Fig. 5, these results clearly demonstrate that Fzd6−/− HSPCs are fully able to reach the BM, but, once at destination, fail to expand due to a strong activation of caspase-3 and consequently decreased HSPC survival.

To determine if Fzd6−/− HSPCs fail to expand also in response to other types of proliferative stress, we induced emergency myelopoiesis using a sublethal dose of LPS (Fig. 7A) (31). Although both Fzd6+/+ and Fzd6−/− HSPCs expanded in both proportion and number (Fig. 7B), BM from Fzd6−/− mice contained significantly fewer LSKs and CD150+ HSCs. This difference did not appear to stem from decreased cell-cycle entry (Fig. 7C) but would be rather due to loss of cells either through cell death or differentiation. Similar numbers of HSCs were found in the spleen (on average, 5.0 × 103 for Fzd6+/+ versus 5.9 × 103 for Fzd6−/−), supporting the hypothesis of increased differentiation and BM exit. BM Fzd6−/− HSPCs also generated proportionately fewer c-Kit+ cells in a colony-forming assay (Fig. 7D), similar to what we saw shortly after transplant. Therefore, Fzd6 is necessary for efficient BM HSPC expansion and self-renewal, not only after transplant but also under proliferative stress such as emergency myelopoiesis.

FIGURE 7.

Fzd6−/− HSPCs exhibit poor emergency hematopoiesis. (A) Experimental design for the emergency response. (B) Flow cytometry analysis of the BM HSPC compartment. Representative flow cytometry data are shown in the left panel. The numbers within flow cytometry plots show the percentage of HSPC subpopulations within LSKs. Histograms represent the proportion and absolute numbers of HSPCs per BM. Data are derived from three independent experiments with three to five mice per group. (C) Cell-cycle analysis of HSCs in the BM. Histogram represents pooled results from three independent experiments with two to five mice per group. (D) Representative flow cytometry data depicting the proportion of c-Kit and CD11b staining within Fzd6+/+ and Fzd6−/− cells recovered from mixed colony assays. Similar results were obtained from four pairs of mice. *p < 0.05, **p < 0.005.

FIGURE 7.

Fzd6−/− HSPCs exhibit poor emergency hematopoiesis. (A) Experimental design for the emergency response. (B) Flow cytometry analysis of the BM HSPC compartment. Representative flow cytometry data are shown in the left panel. The numbers within flow cytometry plots show the percentage of HSPC subpopulations within LSKs. Histograms represent the proportion and absolute numbers of HSPCs per BM. Data are derived from three independent experiments with three to five mice per group. (C) Cell-cycle analysis of HSCs in the BM. Histogram represents pooled results from three independent experiments with two to five mice per group. (D) Representative flow cytometry data depicting the proportion of c-Kit and CD11b staining within Fzd6+/+ and Fzd6−/− cells recovered from mixed colony assays. Similar results were obtained from four pairs of mice. *p < 0.05, **p < 0.005.

Close modal

We have examined in this study the role of the Wnt signaling receptor Fzd6 in the expansion and differentiation of mouse BM HSPCs at steady state and under replicative stress. At steady state, the proportion of CD150+ HSCs was normal in the Fzd6−/− BM but Fzd6−/− HSPCs expressed a higher level of Flt3 than their Fzd6+/+ counterparts. Furthermore, we show that although Fzd6−/− HSPCs were able to reach the recipient BM, they failed to reconstitute an irradiated host due to impaired expansion and survival. Fzd6−/− HSPCs also failed to expand in response to LPS. Thus, Fzd6 was necessary for normal HSPC function in a hematopoietic cell-intrinsic manner.

Self-renewal is a crucial feature of HSCs, which allows them to replenish all hematopoietic lineages under conditions of BM stress (e.g., ablative chemotherapy, irradiation, infections, and injury) (34, 39, 40). Wnt signaling can play a critical role in HSPC self-renewal, although the extent and source of signals are still under debate and likely involve different pathways under different circumstances (22, 25, 41). Canonical Wnt signaling is induced by myeloablative stress (25) and reportedly required for hematopoietic recovery in response to irradiation (41). However, in the absence of negative regulators, such as Fzd6 (29), the response is likely to remain unchecked. Although we did not detect differences in canonical Wnt signaling in Fzd6−/− HSPCs at steady state, in contrast to what was reported for Fmi−/− and Fzd8−/− cells (25), we did observe stronger intracellular β-catenin staining after transplant. Fzd6−/− HSPCs were present at numbers comparable to controls at 4 d posttransplant but showed increased apoptosis and failed to expand over the following days. Sustained stabilization of β-catenin has been previously shown to be directly associated with HSPC exhaustion and apoptosis and could thus result in the loss of Fzd6−/− donor cells after transplant (38, 42, 43).

Fmi and Fzd8 loss-of-function studies suggested a noncanonical requirement for the maintenance of HSPC quiescence, as their absence altered the pool size, self-renewal, and engraftment of HSPCs (25). Namely, Fzd8/Fmi signaling axis inhibited canonical Wnt signaling in HSCs and thus maintained HSC quiescence at steady state. Label-retaining HSCs expressed much higher levels of Fzd8 and Fmi than cycling HSCs, and both Fmi and Fzd8 expression was decreased after activation of HSCs in response to 5-fluoro-uracil. Exogenous Wnt5a is also reported to maintain HSCs by enforcing their resting state (19, 21). Although we did not detect significant differences in the fraction of quiescent (G0) HSCs in the Fzd6−/− BM, we cannot exclude possibility that Fzd6 is also required for the quiescence of a specific subset of CD150+ HSCs. Loss of quiescence would ultimately result in decreased HSC survival and decreased long-term repopulation (34, 35, 44), in agreement with our results after transplant. Further studies using, for example, label-retention approaches will be required to conclusively clarify the issue.

Our results suggest that Fzd6 is a negative regulator of both Cdc42/Jnk and β-catenin–dependent signaling in HSPCs. In contrast to Fmi−/− and Fzd8−/− cells, Fzd6−/− HSPCs displayed normal active β-catenin levels at steady state. There was also no difference in the expression of CD44, a classical Wnt target gene (45). However, we observed enhanced Cdc42 and Jnk activity as indicated by the increased size of Cdc42 clusters and stronger staining with the phospho-specific Jnk Ab. Cdc42 activity can have opposing downstream results: activation of NFAT via the Cdc42/Ca2+ pathway and suppression of NFAT nuclear translocation through the Cdc42/Casein-kinase-1 pathway (46). Fmi/Fzd8 signaling suppressed HSC activation through the inhibition of NFAT/IFN-γ axis and consequent downregulation of β-catenin activity (25), with Fmi−/− and Fzd8−/− HSPCs likely losing their self-renewal capacity due to excessive proliferation. Wnt5a/Cdc42 activity has been shown to inhibit canonical Wnt signaling also in culture (19) and during aging (22). It is thus possible that even though we do not detect altered active β-catenin levels in Fzd6−/− HSPCs, the enhanced Cdc42 activity could interfere with canonical Wnt signaling in our model and contribute to, for example, decreased cell-cycle progression. In contrast, increased phosphorylation of β-catenin by Jnk could promote the translocation of stable β-catenin to the nucleus and thus enhance the potential for canonical Wnt signaling (47).

Altered engraftment of HSPCs in the BM is frequently associated with impaired homing to the BM niche. This is certainly the case for HSPCs mutated for cell adhesion–related genes, such as Rho-family small GTPases Cdc42 and Rac1 (48, 49). Although we cannot completely exclude defective interaction of Fzd6−/− HSPCs with the recipient stromal cells, we detected no impairment in their ability to home to the BM immediately after transplant. There was also no difference in the expression of CD44, a classical Wnt target gene (45) that is involved in BM homing (50). There may have been a subtle decrease in CXCR4 expression in Fzd6−/− HSPCs when compared with their Fzd6+/+ counterparts, and we certainly cannot exclude the possibility of defective CXCR4 signaling in Fzd6−/− cells. However, we saw no significant accumulation of Fzd6−/− HSPCs in the spleen, for example, demonstrating that Fzd6−/− donor cells failed to expand and generate differentiated progeny irrespective of their location.

Taken together, we have shown in this study that Fzd6 plays an essential role in long-term maintenance and self-renewal of mouse HSPCs in a hematopoietic cell–autonomous manner that was likely independent of HSPC homing to the BM but rather related to their expansion, maintenance, and survival. Mechanistically, Fzd6 is a negative regulator of both Cdc42/Jnk and β-catenin–dependent signaling. We propose that the activation of Fzd6 signaling through pharmacologic means would constitute an interesting option for improving HSPC engraftment and the recovery of immune function after transplant.

We thank Dr. Claude Perreault (University of Montreal, QC, Canada) for the gift of Fzd6−/− mice, Jessy Tremblay at the Institut National de la Recherche Scientifique–Institut Armand-Frappier Imaging facility for advice on flow cytometry, Danièle Gagné and Gaël Dulude from the Institute for Research in Immunology and Cancer at the University of Montréal for cell sorting, and the staff of the Centre National de Biologie Expérimentale animal care facility for superb assistance.

This work was supported by the Natural Sciences and Engineering Research Council of Canada (NSERC grant 419226-2012 to K.M.H.) and the Canada Foundation for Innovation (CFI Leaders Fund 31377).

The online version of this article contains supplemental material.

Abbreviations used in this article:

B6

C57BL/6

BM

bone marrow

B6.SJL

B6.SJL-PtprcaPep3b/BoyJ (Ly5a)

Fzd6

Frizzled-6

HSC

hematopoietic stem cell

HSPC

hematopoietic stem/progenitor cell

LMPP

lymphoid-primed MPP

LSK

LinSca1+c-Kithi

MPP

multipotent progenitor cell

PCP

planar cell polarity.

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The authors have no financial conflicts of interest.

Supplementary data