Protein Tyrosine Phosphatase PTPRO Signaling Couples Metabolic States to Control the Development of Granulocyte Progenitor Cells

Neutrophils are the largest subset of granulocytes and act as the first responders of defense against invading various microorganism infections caused by bacteria, fungi, and parasites (1–3). Because they are short lived, neutrophils must be continuously replenished from hematopoietic stem cells (HSCs) and strict regulation of granulopoiesis at a basal or emergent stage (3). Neutrophils originate from hematopoietic progenitors that differentiate along a neutrophil-destined lineage in response to both extracellular stimuli (e.g., growth factors and cytokines) and intracellular regulators, such as transcription factors, controlling the timely expression of lineage-specific genes (4). Granulopoiesis can be divided into neutrophil lineage determination and committed granulopoiesis. The classical mouse model of neutrophil lineage determination has been described and different developmental steps are characterized based on expressions of their surface marker (5, 6). Multipotent long-term HSCs (LT-HSCs) are prevalently quiescent cells and occasionally divide to self-renew and give rise to multipotent short-term HSCs (ST-HSCs). ST-HSCs give rise to the multipotent progenitor (MPP) that develops further into either a common lymphoid progenitor or a common myeloid progenitor. Granulocyte–monocyte progenitors are part of common myeloid progenitors and give rise to either granulocyte precursor cells or monocytes/macrophages. Granulocyte precursor cells further differentiate into mature segmented neutrophils, which are released to peripheral blood as mature polymorphonuclear leukocytes (6, 7). However, the developmental regulation from progenitor cells of granulocytes to functionally mature neutrophils remains poorly defined.

Protein phosphorylation is essential for governing the effects of growth factors on stem cell development and differentiation (8, 9). Regulation of tyrosine phosphorylation is partially mediated by protein tyrosine phosphatase (PTPase) (9–11). Relatively little is known about the repertoire of signal transducing molecules in HSCs, especially for tyrosine phosphatases (12). A novel receptor PTPase, PTPRO, was first isolated from murine bone marrow (BM) HSCs and showed some lineage-specific patterns in regulating myeloid cell differentiation in BM cells (13–15), but its role in HSCs remains unclear. Emerging studies highlight the critical roles of metabolic reprogramming in immune cell development and function (16–18). Studies on the metabolic regulation of myeloid cells are largely limited in myeloid immune cell function, whereas little is known about the metabolic processes driving neutrophil cell fate decisions among physiological steady-state or pathological emergent granulopoiesis.

In this study, we used a loss-of-function allele in mice and show that PTPRO signaling is a crucial negative determinant of granulopoiesis by regulating granulocyte progenitor development, as well as granulocyte differentiation and activation during infection of Listeria monocytogenes, under homeostatic conditions and after G-CSF or GM-CSF stimulation with the lineage-specific pattern. A mechanism study showed that mTOR-HIF1α–dependent glycolytic activities are essential for the granulocyte differentiation from progenitor cells. Thus, the receptor PTPase PTPRO shows novel insights into how metabolic reprogramming of PTPRO directed the development and differentiation of granulocyte progenitor cells into mature neutrophils to mediate antibacterial immunity and homeostatic control of hematopoiesis.

All animal experiments were performed with the approval of the Animal Ethics Committee of Beijing Normal University (Beijing, China) and Fudan University (Shanghai, China). C57BL/6, GFP⁺ C57BL/6, and CD45.1⁺ mice were obtained from Beijing Weitonglihua Experimental Animal Center (Beijing, China). Ptpro^−/− mice were provided by Dr. Xingxu Huang from the School of Life Science and Technology, ShanghaiTech University (Shanghai, China). mTor^fl/fl, Hif1a^fl/fl, and Lyz-Cre mice have been described previously (19, 20). Ptpro^−/−mTor^fl/flLyz-Cre and Ptpro^−/−Hif1a^fl/flLyz-Cre mice were mated in our laboratory. All mice were bred and maintained in specific pathogen–free conditions. Sex-mated littermate mice 6–12 wk of age were mainly used for experiments unless noted in the figure legends. Complete or mixed chimeras were generated by transferring 1–2 × 10⁷ BM cells from wild-type (WT) and/or Ptpro^−/− mice into lethally irradiated recipient mice. Mice reconstitution was determined by flow cytometry analysis of blood samples. Mice were used 8 wk after chimera generation for experiments unless noted in the figure legends.

L. monocytogenes bacterial cultures were grown overnight from glycerol stocks at 30°C in Luria-Bertani (LB) medium under constant mixing. Before infection, overnight cultures were subcultured in L staining of infected mouse liver and spleen B medium at 37°C for constant mixing for 3 h. Subcultures were counted at 600-nm wavelength and diluted in ice-cold sterile PBS. Then, 3 × 10⁵ CFU of L. monocytogenes were i.v. injected into mice using 200 µl of sterile PBS. Mice were sacrificed 48 h after injection, after which the target organs, liver and spleen, were aseptically harvested, weighed, and homogenized for CFU analysis as before (20, 21). Formalin-fixed liver and spleen were processed and embedded in paraffin, sectioned at 5 μm, mounted on positively charged glass slides (Thermo Fisher Scientific), and dried at 60°C for 30 min, as described previously (22).

Peripheral blood smears were stained with a May–Grunwald–Giemsa protocol as described before (23). Peripheral blood smears were photographed on a Zeiss Axio Imager.M1 microscope equipped with an AxioCam MR5 camera (Photometrics) with an EC Plan-Neofluar ×100/0.30 numeric aperture oil immersion lens using ZEN blue software (version 2012).

BM cells were obtained as described before (24). Lineage (Lin)⁻ BM cells and granulocyte–monocyte progenitor cells were sorted as described previously (25, 26). For granulocyte–monocyte progenitor cell isolation, BM cells were stained with lineage markers (anti-CD3, anti-CD4, anti-CD8α, anti-CD45R, anti-Gr1, anti-CD19, anti-CD11b, and anti-TER119) as well as anti–c-Kit, anti-Sca1, anti-CD16/32, and anti-CD34 mAbs, and Lin⁻Sca1⁻c-Kit⁺CD34^hiCD16/32^hi cells were sorted. The purity of the sorted cells was >96%. Whole or sorted Lin⁻ BM cells or granulocyte–monocyte progenitor cells were cultured in DMEM (Life Technologies) supplemented with 10% (v/v) FBS and 1% (v/v) penicillin-streptomycin in the presence of GM-CSF (10 ng/ml), M-CSF (10 ng/ml), or G-CSF (10 ng/ml) for the indicated times. Cells were cultured at 2 × 10⁶ cells/ml in 48-well plates or 1 × 10⁶ cells/ml in 24-well plates in DMEM supplemented with 10% FBS and 1% penicillin plus streptomycin. G-CSF (10 ng/ml), M-CSF (10 ng/ml), or GM-CSF (10 ng/ml) was added into the cultures and cells were cultured for the indicated times. 2-Deoxy-d-glucose (2-DG) and rapamycin (both from Sigma-Aldrich) were dissolved in aqueous media. Recombinant mouse G-CSF, GM-CSF, or M-CSF (all from PeproTech) were dissolved in PBS containing 0.1% (v/v) BSA. Aliquots of 1000× were stored at –80°C and used within fewer than three freeze-thaw cycles.

A total of 5 × 10⁴ BM cells were cultured in methylcellulose (methylcellulose base medium, R&D Systems) supplemented with recombinant murine G-CSF (10 ng/ml), GM-CSF (10 ng/ml), or M-CSF (10 ng/ml). Cells were cultured in triplicate for each concentration at 37°C and 5% CO₂ for 6 d, after which colony numbers were counted and differentiated neutrophils (CD11b⁺Ly6G⁺F4/80⁻), monocytes/macrophages (CD11b⁺F4/80⁺Ly6G⁻), or other cells (CD11b⁻F4/80⁻Ly6G⁻) were determined and calculated. For experiments with sorted Lin⁻ BM cells or 2000 granulocyte–monocyte progenitor cells were placed in MethoCult M3231 supplemented with 50 ng/ml SCF, 10 ng/ml IL-3, and 20 ng/ml GM-CSF or 20 ng/ml G-CSF (STEMCELL Technologies). Colonies were counted and analyzed phenotypically after 8 d of cultures as described (25, 27).

The cell surface markers were analyzed by flow cytometry. The living cells were stained with the buffering which included PBS containing 0.1% (w/v) BSA and 0.1% NaN₃ for 30 min on ice. The following Abs were obtained from BioLegend: anti-CD11b (M1/70), anti-F4/80 (BM8), anti-CD11c (HL3), anti-CD48 (HM48-1), and anti-CD45 (30-F11). The following Abs were obtained from BD Biosciences: anti-CD4 (GK1.5), anti-Ly6G (1A8), and anti-Gr1 (RB6-8C5). The Lin⁻ mixture kit (B220, CD3, Gr1, CD11b, TER119) was obtained from BD Biosciences. The following Abs were obtained from eBioscience: anti-CD34 (RAM34), anti-CD8a (53-6.7), anti-CD45R/B220 (RA3-6B2), anti-CD16/32 (93), anti-CD45.1 (A20), and anti-CD45.2 (104).

Intracellular C/EBPα (14AA) from eBioscience was analyzed with flow cytometry according to the manufacturer’s instructions. For the BrdU in vivo incorporation assay, 150 μl of a 10 mg/ml BrdU solution was i.p. injected into mice. Mice were then bled and BM cells or sorted granulocyte–monocyte progenitor cells were isolated at the indicated time points. BrdU⁺ cells were determined with a BrdU flow kit from BD Biosciences Pharmingen. All flow cytometry data were acquired on an ACEA NovoCyte (ACEA Biosciences) or a FACSCalibur (BD Biosciences), and data were analyzed with NovoExpress or FlowJo (Tree Star, San Carlos, CA). Cell numbers of various populations were calculated by multiplication of the total cell number by the percentages of each individual population from the same mouse, followed by averaging.

Glycolysis of Lin⁻ BM cells from WT and Ptpro^−/− mice was determined by measuring the detritiation of [3-³H]glucose. In brief, the assay was initiated by adding 1 μCi of [3-³H]glucose (PerkinElmer) and, 2 h later, the medium was transferred to microcentrifuge tubes containing 50 μl of 5 N HCl. The microcentrifuge tubes were then placed in 20-ml scintillation vials containing 0.5 ml of water, and the vials were capped and sealed. ³H₂O was separated from unmetabolized [3-³H]glucose by evaporation diffusion for 24 h at room temperature as described before (19).

RNA was extracted with TRI reagent (Sigma-Aldrich), and cDNA was synthesized using a PrimeScript RT reagent kit (Takara). An ABI 7500 quantitative real-time PCR system was used for detecting the mRNA expression. Primers and probes were obtained from Applied Biosystems (Carlsbad, CA). Results were analyzed with ABI 7500 software, and the expression of each target gene is presented as the fold change relative to the WT control samples (2^−ΔΔCt), as described previously (28).

Lin^– BM cells were sorted by flow cytometry and cultured in a 48-well plate in DMEM supplemented with 10% FBS, 1% penicillin plus streptomycin. The cells were treated with G-CSF (10 ng/ml) and harvested at the indicated time. The cells were washed twice with cold PBS and then lysed in RIPA buffer (50 mM Tris-HCl, pH 7.4, 1% NP-40, 0.25% Na-deoxycholate, 150 mM NaCl, 1 mM EDTA, pH 7.4) with protease and phosphatase inhibitor mixtures (Sigma) for 10 min on a rocker at 4°C. The protein concentration was determined via BCA (Beyotime). The protein samples were separated by 10% SDS-PAGE and then transferred onto 0.22 µm polyvinylidene fluoride membranes (Merck Millipore). The membranes were blocked with 5% nonfat dried milk for 1 h at room temperature and incubated with primary antibodies overnight on a shaker at 4°C. Subsequently, HRP-coupled secondary antibody (Beyotime) was added for 1 h at room temperature. After sufficient washing, protein samples were detected by chemiluminescence (Millipore) using AllDoc-x software of the Tanon 5200 Imager (Tanon). The following primary Abs were obtained from Cell Signaling Technology: p-S6 (Ser^240/244; D68F8; catalog no. 5364), p-Erk (Thr²⁰²/Tyr²⁰⁴; D13.14.4E; catalog no. 4370), p-p38 (T180/Y182; D3F9; catalog no. 4511), p-JNK (Thr¹⁸³/Tyr¹⁸²; 81E11; catalog no. 4668), p-mTOR (Ser²⁴⁴⁸; D9C2; catalog no. 5536), C/EBPα (D56F10; catalog no. 8178), HIF1α (D2U3T; catalog no. 14179), and β-actin (13E5; catalog no. 4970).

All data are presented as the mean ± SD. A Student unpaired t test was used for parametric data and a Mann–Whitney U test was used for nonparametric data when comparing two samples, and a one-way ANOVA with a Dunnett post hoc test was for parametric data and a Kruskal–Wallis test was used for nonparametric data when comparing more than two samples. Difference between groups were considered statistically significant with a p value (α value) <0.05.

Granulopoiesis is a hallmark of acute infectious inflammation in experimental and clinical settings (29). We first observed granulopoiesis alterations of kinase PTPRO signaling in mice that were challenged with L. monocytogenes. To bypass any potential effects of nonhematopoietic loss of PTPRO signaling, we used Ptpro^−/− complete chimeric mice, which were generated by transplanting CD45.2⁺ Ptpro^−/− BM cells (called Ptpro^−/−→WT mice) or WT (WT→WT mice) BM cells into syngeneic lethally irradiated CD45.1⁺ C57BL/6 recipient mice, respectively (Supplemental Fig. 1A). WT and Ptpro^−/− chimeric mice were challenged with L. monocytogenes to determine whether loss of these genes affected the antibacterial immune response. Ptpro^−/− BM chimeras had much more severe pathological inflammation in the spleen and liver (Fig. 1A), the primary target organs of L. monocytogenes infection, which was further verified by markedly decreased bacterial burden versus WT BM chimeras (Fig. 1B, 1C). Thus, PTPRO deficiency potentiates antibacterial immunity.

Consistently, immunological analysis showed that Ptpro^−/− mice had markedly recruited distribution of CD11b⁺Ly6G⁺ neutrophils in the spleen and liver (Fig. 1D–F). The spleen and liver were digested in advance, and then the percentage and absolute number of CD11b⁺Ly6G⁺ cells were detected. To obtain the wider view about the dynamic process of neutrophils, we also determined the population of other organ neutrophils and found that PTPRO deficiency had markedly expanded percentages and numbers of CD11b⁺Ly6G⁺ neutrophils in the BM cells, blood, and peritoneal cavity (Fig. 1D–F). These observations showing that the number of neutrophils progressively expanded with granulopoiesis from progenitor cells and circulating cells to local inflammatory sites promoted us to propose that PTPRO might be an important modulator for neutrophil development in the antibacterial immunity.

To test this hypothesis, we first determined the expressions of PTPRO through myeloid development in highly purified cell populations from BM to peripheral blood. PTPRO expressions are at very low levels in pluripotent HSCs and common myeloid progenitor cells (Supplemental Fig. 1B). As granulocytic differentiation proceeds through granulocyte–monocyte progenitor cells, immature BM neutrophils, and subsequently mature peripheral blood granulocytes, the expressions of PTPRO gradually increase. These results show that PTPRO expression leads to a highly lineage-specific pattern during granulopoiesis.

To further test this hypothesis, we used Ptpro^−/− mice in this study (Supplemental Fig. 1C). Morphological analyses of circulating neutrophils by May–Grunwald–Giemsa staining also revealed that unusual hypersegmentation morphology in Ptpro^−/− mice was characterized by nuclear hypersegmentation and blebbing (Fig. 2A), which was further confirmed in BM complete chimeric mice (Fig. 2B, 2C). These data collectively suggest that PTPRO deficiency is probably related with a hypermature phenotype of neutrophils. To access the requirement of PTPRO in granulopoiesis, we analyzed the distribution of neutrophils in mice. Ptpro^−/− mice had markedly expanded percentages and numbers of CD11b⁺Ly6G⁺ neutrophils, but not T cells, B cells, and monocyte/macrophages, of BM cells, blood, and spleen (Fig. 2D, Supplemental Fig. 1D, 1E). These data indicate that Ptpro^−/− mice are severely neutrophilic. To bypass any potential effects of nonhematopoietic loss of PTPRO signaling, we used Ptpro^−/− complete chimeric mice as described above. Ptpro^−/− BM chimeras displayed an expanded neutrophil population in BM cells, blood, and spleen (Supplemental Fig. 1F), which further indicates that PTPRO deficiency from nonhematopoietic cells regulates the distribution of neutrophils. Is PTPRO intrinsic for controlling neutrophil populations? We performed a competitive repopulation and constructed the mixed hematopoietic chimeric mice through transplanting either CD45.1⁺ WT or CD45.2⁺Ptpro^−/− BM cells at the ratio of 1:1 into syngeneic lethally irradiated GFP⁺ recipient mice. By 2–8 wk after BM transplant, most of the neutrophils of BM cells, blood, and spleen in mice receiving WT plus Ptpro^−/− BM cells were of Ptpro^−/− donor origin (Fig. 2E, Supplemental Fig. 1G). The same recipient mouse environment failed to recover the Ptpro^−/−-caused neutrophilia, which strongly suggests that PTPRO is critical for regulating neutrophil maturation and development in a cell-autonomous manner.

The imbalance between death and proliferation can lead to neutrophilia caused by Ptpro^−/−. We determined the cell death of neutrophils by combining 7-AAD and annexin V staining in Ptpro^−/− mice and Ptpro^−/− chimeric mice. Ptpro^−/− showed a similar cell death kinetic course in Ptpro^−/− mice (Supplemental Fig. 2A, 2B) and Ptpro^−/− chimeric mice (Supplemental Fig. 2C). This suggests that PTPRO deficiency had no significant effects on neutrophil cell death. We next determined the cell proliferation of neutrophils by in vivo BrdU incorporation in mice. Ptpro^−/− mice had a significantly enhanced percentage and number of BrdU⁺ neutrophils in BM cells and blood compared with WT mice (Fig. 3A–C). In the competitive repopulation assay of mixed chimeras, Ptpro^−/− caused a significantly enhanced percentage of BrdU⁺ neutrophils compared with the WT compartment in the same recipient mice (Fig. 3D). These data suggest that neutrophilia of Ptpro^−/− mice might be the result of enhancement in the differentiation of neutrophils from progenitors.

To confirm this conclusion, we therefore directly examined the ability of BM cells from WT and Ptpro^−/− mice to generate maturate myeloid immune cells in response to different myeloid growth factors, that is, GM-CSF, G-CSF, and M-CSF, with the lineage-specific pattern in in vitro liquid culture by differentiating along the monocyte lineage in the generation of monocytes/macrophages (CD11b⁺F4/80⁺Ly6G⁻), neutrophil lineage in the generation of neutrophils (CD11b⁺Ly6G⁺F4/80⁻), and other cell populations (CD11b⁻F4/80⁻Ly6G⁻). Ptpro^−/− BM cells had a significantly specifically expanded neutrophil population, similar monocyte/macrophage population, and a shortened other cell population in the myeloid cell pool compared with WT BM cells stimulated by different myeloid growth factors (Fig. 3E). In addition, Ptpro^−/− significantly specifically enhanced the percentage of BrdU⁺ neutrophils in the developing neutrophils, but not macrophages, derived from BM cells in the presence of GM-CSF (Fig. 3F). Furthermore, to circumvent any potential contributions of an existing myeloid population in the BM, we purified Lin⁻ cells from WT and Ptpro^−/− BM and cultured them in vitro in the presence of GM-CSF or G-CSF. Ptpro^−/− Lin⁻ progenitor cells had enhanced CFU and developed neutrophil number compared with the WT compartment (Fig. 3G). These data collectively indicate that PTPRO is specifically required for the differentiation of neutrophil lineage differentiation from neutrophil precursors in a cell-autonomous manner.

We next determined the effects of PTPRO deletion on the homeostasis of a hematopoietic progenitor. Ptpro^−/− showed a comparable percentage and number of Lin⁻Sca1⁻c-Kit⁺ (LSK) stem cells, LT-HSCs (LSK⁺CD150⁺CD48⁺), MPPs (LSK⁺CD150⁻CD48⁺), megakaryocyte-erythroid progenitors (MEPs, LSK⁺CD34^lowCD16/32^low), and common myeloid progenitor cells (LSK⁺CD34^hiCD16/32^low), but significantly increased the percentage and number of granulocyte–monocyte progenitor cells (LSK⁺CD34^hiCD16/32^hi) compared with WT (Fig. 4A–C). This demonstrates that PTPRO deficiency majorly affects the differentiation processes of granulocyte–monocyte progenitor cells into neutrophils. Furthermore, to circumvent any potential contributions of an existing myeloid population in the BM, we purified granulocyte–monocyte progenitor cells from WT and Ptpro^−/− BM and cultured them in vitro in the presence of G-CSF. Ptpro^−/− granulocyte–monocyte progenitor cells had an enhanced cell number per CFU and a CFU number per 2000 granulocyte–monocyte progenitor cells and a developed neutrophil number per 2000 granulocyte–monocyte progenitor cells compared with the WT compartment (Fig. 4D). Following the recently confirmed markers of early neutrophil precursors in granulocyte–monocyte progenitor cells by single-cell transcriptomics, we used CD81 and CD115 to distinguish early neutrophil progenitor cells from common monocyte progenitor cells (30). Consistently, PTPRO deficiency significantly expanded granulocyte–monocyte progenitor cell populations. It was further found that these populations were mainly expanded from neutrophil progenitor cells, but had little effect on common monocyte progenitor cell populations (Supplemental Fig. 2D). Altogether, these data demonstrate that neutrophilia was induced by Ptpro^−/− just through regulating the differentiation course from granulocyte–monocyte progenitor cells to neutrophils.

To investigate how PTPRO regulates neutrophil differentiation from granulocyte–monocyte progenitor cells, we determined the metabolism features of developing neutrophils and effects of PTPRO during this course. The glycolytic activity of Lin⁻ BM cells was measured by the generation of ³H-labeled H₂O from [3-³H]glucose. PTPRO deficiency enhanced the glycolytic activities in Lin⁻ BM cells stimulated by G-CSF compared with the WT control (Fig. 5A). Glucose utilization depends on a chain of reactions catalyzed by multiple enzymes, eventually leading to the generation of lactate and net production of two ATP molecules as the energy source. Consistently, PTPRO deficiency upregulated the expression of glucose transport 1 (Glut1), glucose-6-phosphate isomerase (GPI), enolase 1 (Eno1), and LDHα (lactate dehydrogenase α) in Lin⁻ BM cells stimulated by G-CSF (Fig. 5B, 5C). Taken together, these data indicate strong upregulation of glucose metabolism in developing neutrophils caused by Ptpro^−/−.

Previous studies showed mTOR signaling is critical for glycolysis metabolism of different types of immune cell differentiation (27, 31, 32). We therefore determined the mTOR signaling effects in the Lin⁻ BM cells. Ptpro^−/− cells had significant upregulation of p-mTOR and p-S6, the downstream target of mTOR, and stimulation by G-CSF compared with the WT control (Fig. 5D). Although MAPK signaling is essential for granulopoiesis (25), PTPRO deficiency displays comparable alteration compared with the WT compartment in G-CSF–treated Lin⁻ BM cells (Fig. 5D). These data suggest that mTOR signaling is probably related with granulopoiesis and metabolism regulated by Ptpro^−/−.

We next determined the expression of C/EBPα, PU.1, C/EBPβ, and GATA1, which are the essential transcriptional factors in regulating the cell fate decision of granulopoiesis (33, 34). It is probable that the subsequent decision between granulocyte and monocyte commitment is determined by the balance between PU.1 and C/EBPα (35, 36). By sorting Lin⁻ BM cells, we determined these transcription factor expressions. Ptpro^−/− had significantly enhanced mRNA expression of C/EBPα and C/EBPβ compared with WT (Fig. 5E). Furthermore, we sorted granulocyte–monocyte progenitor cells (Lin⁻Sca1⁻c-Kit⁺CD34^hiCD16/32^hi) and determined their C/EBPα protein expression with intracellular staining methods (Fig. 5F, 5G). PTPRO deficiency had significantly enhanced C/EBPα expression compared with WT, which strongly indicates that C/EBPα is probably critical for granulopoiesis from granulocyte–monocyte progenitor cells regulated by Ptpro^−/−.

To test the role of mTOR and glycolysis in granulopoiesis induced by PTPRO, purified granulocyte–monocyte progenitor cells from WT and Ptpro^−/− BM cells were cultured in the presence of varying concentrations of rapamycin, an mTOR inhibitor, which can block the mTOR-dependent metabolic pathway, including glycolysis and 2-DG, a nonhydrolyzable glucose analog and a prototypical inhibitor of glycolytic pathway that acts via blocking hexokinase, the first rate-limiting enzyme of glycolysis. As expected, optimized doses of rapamycin and/or 2-DG significantly diminished the glycolytic activity (Supplemental Fig. 2E, 2F). Importantly, rapamycin and/or 2-DG treatment resulted in diminished neutrophil number and C/EBPα expression, an essential transcription factor for granulopoiesis, in an in vitro CFU assay and recovered the changes in the Ptpro^−/− cells (Fig. 6). Taken together, these data demonstrate that mTOR and glycolytic activity are required for PTPRO-directed granulopoiesis.

To determine whether mTOR is involved in regulatory activity, we crossed Ptpro^−/− and Mtor^flox/floxLyz-Cre mice to conditionally delete Ptpro and Mtor gene expression in myeloid cells, including a neutrophil lineage (called Ptpro^−/−mTOR^−/−). The expanded neutrophil populations of BM cells, blood, and spleen in Ptpro^−/− mice are significantly recovered in Ptpro^−/−mTor^−/− mice (Fig. 7A). Consistently, enhanced granulopoiesis of the CFU assay in Ptpro^−/− granulocyte–monocyte progenitor cells is significantly restored to a normal level in Ptpro^−/−mTor^−/− cells (Fig. 7B). Interestingly, PTPRO deficiency caused the significant upregulation of HIF1α (Fig. 7C), a key transcriptional factor of regulating glucose metabolism and glycolytic enzyme activities (37, 38), which suggests that HIF1α and glycolytic activities are involved in PTPRO-directed granulopoiesis. Importantly, the upregulated HIF1α expressions in Ptpro^−/− granulocyte–monocyte progenitor cells were significantly recovered and almost completely blocked in Ptpro^−/−Mtor^−/− cells (Fig. 7C), which revealed that HIF1α is a downstream target of mTOR in PTPRO-directed granulopoiesis. Thus, mTOR-HIF1α and glycolysis are probably related with HIF1α signals in PTPRO-directed granulopoiesis. Consistent with these results, the upregulated neutrophil lineage transcription factor C/EBPα expression and increased glycolytic activities in Lin⁻ BM cells of Ptpro^−/− mice were also significantly restored to a normal level in those cells of Ptpro^−/−Mtor^−/− mice (Fig. 7D–F).

To further differentiate the role of mTOR-HIF1α and glycolysis and determine whether there is reciprocal regulation between mTOR-HIF1α and glycolysis in granulopoiesis induced by PTPRO, we crossed the Ptpro^−/− and Hif1a^flox/floxLyz-Cre mice to conditionally delete Ptpro and Hif1α gene expression in myeloid cells, including a neutrophil lineage (called Ptpro^−/−Hif1α^−/−). The expanded neutrophil populations of BM cells, blood, and spleen in Ptpro^−/− mice are significantly recovered to a normal level in Ptpro^−/−Hif1α^−/− mice (Fig. 8A). Consistently, enhanced granulopoiesis of the CFU assay and lineage transcription factor C/EBPα expressions in Ptpro^−/− granulocyte–monocyte progenitor cells are significantly restored to a normal level in Ptpro^−/−Hif1α^−/− cells (Fig. 8B, 8C). Consistently, the increase glycolytic activities in Lin⁻ BM cells of Ptpro^−/− mice are also significantly restored to a normal level in those cells of Ptpro^−/−Hif1α^−/− mice (Fig. 8D, 8E). Collectively, the above data reveal that HIF1α and glycolytic activities are critically required for PTPRO-directed granulopoiesis.

In the current study, we show that the novel receptor PTPase PTPRO negatively regulates progenitor cell proliferation and granulocyte differentiation and activation at a basal or emergent stage. Generation of mature neutrophils during granulopoiesis requires sequential progression from HSCs to precursor cell populations before terminal differentiation. The progression of granulopoiesis increases the greater need for protecting against microorganism infection, and Ptpro^−/− chimeric mice are more likely to survive. Therefore, in response to L. monocytogenes infection, Ptpro^−/− mice have more inflammatory neutrophils generated from BM precursors and play a crucial roles in clearance of bacterial infection compared with WT mice. Although more granulopoiesis causes a stronger inflammatory response that is conducive to bacterial clearance, it will significantly aggravate the inflammatory damage of the target organ liver and spleen in the short term, but with the increase of bacterial clearance, it will alleviate the disease and help solve the infectious inflammation. Interestingly, the expression of PTPRO protein increased gradually with the development and maturation of granulocytes. This suggests that the expression of PTPRO protein may be critical for the process of neutrophil maturation, and it may play an important role in maintaining the function of granulopoiesis or maturation. However, the specific regulatory effect is that the deletion of PTPRO promotes the development and maturation of granulocytes. These suggest that tyrosine phosphatase PTPRO has a negative regulatory effect on granulopoiesis, although it shows a gradually increasing expression during development and maturation. Additionally, at the normal physiological status, we performed systemic analysis with Ptpro^−/− mice, complete or mixed chimeras, and in vitro CFU experiments, and we showed that Ptpro^−/− mice result in neutrophilia with an expanded granulocytic compartment resulting from a cell-autonomous increase in the number of granulocyte progenitors under homeostasis and potentiated responses to anti-bacterial infection. In our results, granulocyte–monocyte progenitor cells isolated from Ptpro^−/− mice preferentially developed into granulocytes at the expense of undifferentiated cells, but not monocytes/macrophages when cultivated in the GM-CSF or G-CSF. Consistent with the in vitro results, there was no significant alteration in cell percentage and number of LT-HSCs, ST-HSCs, and MPPs, as well as the myeloid progenitor cell population, including MEPs and common myeloid progenitor cells in the Ptpro^−/− mice compared with the WT control. However, the subpopulation of common myeloid progenitor cells and granulocyte–monocyte progenitor cells in the BM cells have an expanded cell population. Consistently, Ptpro^−/− mice have a higher BrdU⁺ cell percentage in Lin⁻ BM cells, granulocyte–monocyte progenitor cells, and mature neutrophils. In addition, Ptpro^−/− mice or complete chimeric mice also show the mature characteristic of a higher percentage of hypersegmented neutrophils with Giemsa staining or a higher percentage of mature neutrophils (CD11b^hiLy6G^hi). Collectively, these data confirmed that PTPRO negatively directs granulopoiesis and maturation from its progenitor cells in an intrinsic manner (Fig. 8F).

PTPs play a critical role in counteracting the activity of tyrosine kinases, many of which are critical in HSC differentiation (10, 34, 39). Regulation of tyrosine phosphorylation is partially mediated by PTPase (9). These unique proteins are also involved in a variety of signal transduction pathways that regulate cell proliferation, differentiation, metabolism, cell survival, and gene transcription (40, 41). Defective or inappropriate operation of these networks leads to aberrant tyrosine phosphorylation, which contributes to the development of many diseases, including inflammation and cancer (41–43). PTPRO was first characterized and identified in BM stem cells (9, 13). The murine and full-length human PTPRO cDNAs, which share 89% homology, indicate that PTPRO is highly conserved between these species. In adult tissue, PTPTO expression was less restricted and was observed in the lung, heart, skeletal muscle, prostate, testis, and in various areas of the brain, indicating that PTPRO expression is developmentally regulated (14). Expression of PTPRO was also observed in human CD34⁺ BM cells. PTPRO is a receptor-type PTP. For the single gene encoding PTPRO, one of two promoters is active in various hematopoietic tissues (14, 44). Studies to date have suggested a potential cell differentiation property of PTPRO. PTPRO is expressed quite strongly in osteoclasts and positively regulates osteoclast differentiation and activity (45). Suppression of PTPRO in cultured rabbit osteoclasts by antisense oligodeoxynucleotides reduced their bone resorbing activity, whereas overexpression of PTPRO in U937 monocytes enhanced their differentiation into osteoclast-like cells and increased their ability to resorb bone. In the present studies, to dissect fully the role of PTPRO in hematopoietic differentiation, we evaluated PTPRO expression throughout myeloid development in highly purified cell populations from BM and peripheral blood. Expression of PTPRO was detected at low levels in pluripotent HSCs and common myeloid progenitor cells. As granulocytic differentiation proceeds through granulocyte–monocyte progenitor cells, immature BM neutrophils, and subsequently mature peripheral blood granulocytes, expression of PTPRO steadily increases. Conversely, as granulocyte–monocyte progenitor cells adopt the alternative monocyte fate, PTPRO levels are repressed. These data collectively reveal a highly lineage-specific pattern of expression for PTPRO. Furthermore, Lin⁻ BM progenitor cells were cultured with GM-CSF, M-CSF, and G-CSF with in vitro liquid culture by differentiating along the monocyte lineage in the generation of monocytes/macrophages (CD11b⁺F4/80⁺Ly6G⁻), the neutrophil lineage in the generation of neutrophils (CD11b⁺Ly6G⁺F4/80⁻), and other cell populations (CD11b⁻F4/80⁻Ly6G⁻). Ptpro^−/− BM cells had a significantly specifically expanded neutrophil population, similar to the monocyte/macrophage population and shortened other cell population in the myeloid cell pool compared with WT BM cells stimulated by different myeloid growth factors cultured by G-CSF, even in GM-CSF. These data further demonstrate its characteristic of a granulocytic lineage differentiation marker. The specifically accelerated differentiation to neutrophils was also supported by the enhanced BrdU incorporation into mature neutrophils from BM cells or granulocyte–monocyte progenitor cells in vivo. These differentiated neutrophils in Ptpro^−/− mice also display segmented or hypermature phenotypes (CD11b^hiLy6G^hi). When combining these data, we arrived at the conclusion that PTPRO functions as a cell type–specific negative regulator of granulocytic lineage differentiation during granulopoiesis and homeostasis.

How does PTPRO set the “brake” and direct granulopoiesis? Several studies have shown that regulation of glucose availability offers an independent route to control cellular responses in developing myeloid cells and that M-CSF–mediated myelopoiesis is very sensitive to glucose uptake, glycolysis, and downstream anabolic metabolism (46, 47). Also, mTOR signaling (mTORC1) is critical for reprogramming M-CSF–mediated myelopoiesis (27). However, the role of granulopoiesis in progenitor cells of granulocytes remains unclear. In our results, Ptpro^−/− causes significant higher glycolytic activities and glycolytic enzyme expression in developing Lin⁻ BM progenitor cells. mTOR is critically involved in regulating the metabolism pathway. Furthermore, blocking glycolysis with 2-DG or blocking mTOR signaling with rapamycin or in mTor^−/− mice significantly repressed the higher glycolytic metabolic activities and recovered the phenotypes induced by PTPRO deficiency. These data indicate that mTOR signaling and glycolysis are required for PTPRO-directed granulopoiesis.

HIF1α is a key transcriptional factor that orchestrates glucose metabolism and glycolytic enzyme activities (37, 38). HIF1α is induced in hypoxia or normoxia and plays a central role in the proinflammatory cytokine secretion or regulates the function of myeloid-derived suppressor cells in infection or cancer (48, 49). Previous studies have shown that HIF1α is responsible for glycolysis activities and is a downstream target of mTORC1 in regulating myeloid-derived suppressor cell differentiation and innate immune cell function (50). In the current study, by genetic ablation of mTOR or HIF1α in Ptpro^−/− mice, we found that mTOR-HIF1α–dependent glycolytic activities are critically involved in the granulopoiesis induced by PTPRO deficiency. The disruption of mTOR, HIF1α, or glycolytic metabolism signals further highlights their critical roles in supporting PTPRO-directed granulopoiesis.

Development of hematopoietic cells is controlled by lineage-specific transcriptional factors (36). The balance between PU.1, the C/EBPs family, and GATA1 determines differentiation into the granulocytic or monocytic pathway (7). PU.1 is expressed in all stages of myeloid differentiation and has been shown to be critically important for myeloid development (33, 35). Conditional C/EBPα depletion in granulocyte–monocyte progenitor cells led to enhanced granulopoiesis (34), suggesting specific functions of C/EBPα during granulocyte development. Our results show that increased expressions of C/EBPα, but not PU.1 or GATA1, are critically involved in the granulopoiesis induced by Ptpro^−/−. Furthermore, depletion of mTOR or HIF1α or blocking mTOR, HIF1α, or glycolytic activities all significantly diminished the upregulated lineage-specific transcriptional factor C/EBPα expression caused by Ptpro^−/−. These results point to a self-amplifying loop during granulopoiesis, involving glucose metabolism, mTOR activation, HIF1α activation, and lineage-specific transcriptional factor expression.

In summary, PTPRO provides a critical negative feedback loop in regulating the development, maturation, and homeostasis of granulopoiesis and in the immune response to anti-bacterial infection mainly by the mTOR-HIF1α-C/EBPα axis coupling with the glycolytic signaling pathway (Fig. 8F). PTPRO, as a receptor type of PTPases, is identified as an intrinsic negative master for granulopoiesis, perhaps representing a promising target for therapeutic intervention to regulate granulocyte-mediated diseases.

The authors have no financial conflicts of interest.