The protein tyrosine phosphatase PTPN22(C1858T) allelic polymorphism is associated with increased susceptibility for development of systemic lupus erythematosus (SLE) and other autoimmune diseases. PTPN22 (also known as LYP) and its mouse orthologue PEP play important roles in antigen and Toll-like receptor signaling in immune cell functions. We demonstrate here that PEP also plays an important inhibitory role in interferon-α receptor (IFNAR) signaling in mice. PEP co-immunoprecipitates with components of the IFNAR signaling complex. Pep−/− hematopoietic progenitors demonstrate increased IFNAR signaling, increased IFN-inducible gene expression, and enhanced proliferation and activation compared to Pep+/+ progenitors in response to IFN-α. In addition, Pep−/− mice treated with IFN-α display a profound defect in hematopoiesis, resulting in anemia, thrombocytopenia, and neutropenia when compared to IFN-α–treated Pep+/+ mice. As SLE patients carrying the PTPN22(C1858T) risk variant have higher serum IFN-α activity, these data provide a molecular basis for how type I IFNs and PTPN22 may cooperate to contribute to lupus-associated cytopenias.
Systemic lupus erythematosus (SLE) is a prototype systemic autoimmune inflammatory disorder characterized by the production of autoantibodies, immune complex formation, and immune complex deposition in end-organs that contribute to skin rash, nephritis, arthritis, cerebritis, and vasculitis. Hematologic abnormalities are also amongst the most common manifestation in SLE and include anemia, thrombocytopenia, neutropenia, and lymphopenia (Bashal, 2013). Autoantibodies against RBCs, platelets (PLTs), lymphocytes, and neutrophils, as well as reduced expression of complement regulatory proteins CD55 and CD59 making cells more susceptible to complement-mediated lysis, account for some cytopenias, but the etiology of many remain unknown (Mittal et al., 1970; Butler et al., 1972; García-Valladares et al., 2006). BM abnormalities, including dyserythropoiesis, hypocellularity, and myelofibrosis, are commonly found and lend support for the contribution of chronic immune activation to BM dysfunction (Voulgarelis et al., 2006).
Genome-wide association studies have identified >40 confirmed genetic loci associated with the development of SLE (Cui et al., 2013). The PTPN22(C1858T) polymorphism, resulting in a missense mutation R620W, is associated with increased risk of developing type I diabetes (Bottini et al., 2004), SLE (Kyogoku et al., 2004), rheumatoid arthritis (Begovich et al., 2004), Graves’ disease (Velaga et al., 2004), and other autoimmune disorders (Stanford and Bottini, 2014; Rawlings et al., 2015). PTPN22/LYP and its mouse orthologue PEP are members of the PEST-domain containing nonreceptor protein tyrosine phosphatase family and play important roles in TCR, BCR, IFN-γ receptor, and Toll-like receptor (TLR) functions (Rhee and Veillette, 2012; Spalinger et al., 2013; Wang et al., 2013; Bottini and Peterson, 2014). Whereas the human PTPN22(C1858T) at-risk SNP has been proposed to have both activating and inhibitory functions, recent studies of knock-in mice expressing the mouse PEP(R619W) orthologue support a loss-of-function phenotype and mimic Pep−/− mice (Vang et al., 2005; Rieck et al., 2007; Zikherman et al., 2009; Zhang et al., 2011; Dai et al., 2013; Rawlings et al., 2015). Pep−/− mice develop splenomegaly and spontaneous germinal center formation as they age and are prone to, but do not develop, systemic autoimmunity (Hasegawa et al., 2004). Consistent with the multifactorial nature of human SLE, development of autoimmunity and clinical disease requires additional genetic and exogenous factors. Whereas Pep−/− mice on a C57BL/6 background do not develop systemic autoimmunity (Hasegawa et al., 2004), Pep−/− mice expressing a CD45(E613R) mutation develop autoantibodies, glomerulonephritis, and increased mortality despite being on a nonautoimmune-prone background (Zikherman et al., 2009).
Type I IFNs have long been implicated in human SLE pathogenesis as a subset of patients treated with IFN-α for chronic HCV or cancers develop drug-induced lupus (Ho et al., 2008). As with PTPN22, polymorphisms in transcription factors that regulate type 1 IFN—IRF5 (OR = 1.3–1.8), IRF7 (OR = 1.3), IRF8 (OR = 1.17–1.28; Harley et al., 2008; Hom et al., 2008; Gateva et al., 2009; Han et al., 2009; Chung et al., 2011; Cunninghame Graham et al., 2011; Lessard et al., 2012), and TYK2 (OR = 1.29), a protein tyrosine kinase (PTK) requisite for IFNAR function (Cunninghame Graham et al., 2011)—are associated with increased SLE susceptibility. A type 1 IFN-inducible gene signature has been reported in SLE and a higher magnitude of the IFN-induced gene signature is associated with greater propensity to develop renal, central nervous system, and hematologic manifestations (Baechler et al., 2003). SLE patients carrying the PTPN22(C1858T) at-risk polymorphism have higher IFN-α levels and higher IFN-α levels correlate inversely with leukocyte numbers (Bengtsson et al., 2000; Kariuki et al., 2008). Finally, treatment of patients with an anti–IFN-α mAb (rontalizumab) results in improvement in the Systemic Lupus Responder Index and enables more patients to reduce their concomitant corticosteroid use (Kalunian et al., 2015).
Mouse models of SLE also support a pathogenic role for type I IFNs. Administration of IFN-α recombinant protein (Heremans et al., 1978; Adam et al., 1980), infection with viral-mediated IFN-α expression (Mathian et al., 2005) or administration of poly(I:C) (Carpenter et al, 1970) accelerate autoimmunity in NZB/W F1 mice. NZB mice lacking IFNAR do not develop anti-RBC autoantibodies and are protected from the development of autoimmune hemolytic anemia (Santiago-Raber et al., 2003). Similarly, lack of IFNAR attenuates lupus manifestations in other models of lupus, including NZM2328 and B6.Nba.2 (Jørgensen et al., 2007; Agrawal et al., 2009). Finally, administration of an anti-IFNAR mAb in BXSB male mice improves serologic, cellular, and histologic parameters of lupus, and also extends survival (Baccala et al., 2012).
Type I IFNs are rapidly up-regulated after infection and serve as a first line of innate immune defense. Type I IFNs have pleiotropic effects on hematopoietic and nonhematopoietic cells to control immunity, induce viral inhibitory activities and regulate hematopoiesis. In immune cells, type 1 IFNs induce monocyte differentiation, augment antigen presentation by dendritic cells, promote T cell survival, increase T cell cytolysis, and enhance B cell differentiation and antibody production (Hall and Rosen, 2010). IFN-α has also recently emerged as an important regulator of hematopoiesis. In addition to immunomodulatory effects, IFN-α mobilizes hematopoietic stem cells (HSCs) to proliferate from a quiescent state to a renewal state. However, chronic activation of the IFN-α pathway results in HSC exhaustion and impairs HSC functions (Essers et al., 2009).
Given the association of PTPN22 and IFN-α in SLE (Kariuki et al., 2008), we investigated the effects of IFN-α in Pep−/− mice. These studies reveal a previously unrecognized role for PEP as a negative regulator of IFNAR-mediated activation in hematopoietic progenitors and provides a mechanistic basis by which PEP and IFN-α may contribute to hematopoietic dysfunction in SLE.
PEP deficiency accelerates IFN-α–induced lupus-like disease
Given the genetic links implicating both IFN-α and PTPN22 in human SLE, we tested whether PEP deficiency may affect the ability of IFN-α to induce SLE. Systemic administration of IFN-α5 (109 PFU/mouse) by adenoviral delivery resulted in 40% mortality in Pep+/+ mice (C57BL/6 background) 2 mo after IFN-α administration (Fig. 1 A). In contrast, IFN-α (109 PFU/mouse) treated Pep−/− mice (C57BL/6) demonstrated 40% mortality by 2 wk and ∼90% mortality by 2 mo. Corresponding with increased mortality, IFN-α5–treated Pep−/− mice developed increased proteinuria, a mild glomerulopathy, antinuclear autoantibodies. and cytopenias when compared to IFN-α5–treated Pep+/+ mice (Fig. 1, B–G). Affected glomeruli of IFN-α5–treated Pep−/− mice have segmental or global thickening of capillary basement membranes and increased mesangial matrix (Fig. 1 D, arrows), resulting in markedly narrowed capillary profiles. Using an arbitrary scoring system (described in Materials and methods), the pathology score for IFN-α5–treated Pep+/+ mice was 0.0 + 0, whereas IFN-α5–treated Pep−/− mice was 0.67 + 0.2 (P = 0.03). Cytopenias, including peripheral neutropenia, anemia, and thrombocytopenia, as well as reductions in splenic neutrophils and monocytes, were also more severe in IFN-α5–treated Pep−/− mice (Fig. 1, E–I). In contrast, whereas IFN-α5 reduced peripheral lymphocytes, there were no differences between IFN-α5–treated Pep+/+ and Pep−/− mice (Fig. 1 J). As IFN-α increased serum BAFF/BLys (Mathian et al., 2005), a key pathogenic factor in SLE, BAFF/BLys levels were elevated in IFN-α5–treated mice compared to LacZ controls, but no differences were observed between IFN-α5–treated Pep+/+ and Pep−/− mice (Fig. 1 K). Together, these data suggest that PEP deficiency accelerates IFN-α5–induced lupus-like disease with increased mortality, proteinuria, glomerulopathy, autoantibody production, and cytopenias.
Since a type I IFN gene signature in peripheral blood mononuclear cells is associated with cytopenias in human lupus patients (Baechler et al., 2003), we further dissected the underlying pathogenesis and analyzed the effects of IFN-α on the hematopoietic progenitor compartment. Consistent with the previous description that Sca-1 is an IFN-α–induced activation marker (Essers et al., 2009), we observed increased Sca-1 expression on Pep+/+ cKit+ progenitors after IFN-α treatment (unpublished data). Although no differences in HSC and progenitor populations were observed between Pep+/+ and Pep−/− mice under homeostatic conditions (unpublished data), a significant reduction in HSC, multipotent progenitor (MPP2 and MPP3), megakaryocyte/erythrocyte progenitor (MEP), and myeloid progenitor (CMP and GMP) populations was observed in IFN-α5–treated Pep−/− mice compared to IFN-α5–treated Pep+/+ mice (Fig. 2, A and B). This effect was dose dependent, as administration of 10-fold less (108 PFU/mouse) Ad5-IFN-α5 did not affect the HSC population in Pep−/− mice, though the more differentiated multipotent progenitors (MPP2 and MPP3) and myeloid progenitors (CMP/GMP) populations were still reduced in IFN-α5–treated Pep−/− mice (Fig. 2, C and D). There was also a trend in the reduction of LS−K and MEP cells that did not reach statistical significance. Treatment with Ad5-IFN-α5 (108 PFU/mouse) did not result in mortality in either Pep+/+ or Pep−/− mice within a 60-d period (unpublished data). Similar effects were observed with administration of poly(I:C) with reduced cellularity in the more differentiated myeloid progenitors (MEP, CMP and GMP) in poly(I:C)-treated Pep−/− mice (Fig. 2, E and F). Consistent with enhanced progenitor activation, we observed increased Sca-1 expression on poly(I:C)-treated cKit+ Pep−/− progenitors compared to Pep+/+ progenitors (Fig. 2 G). Together, these observations are consistent with increased activation and potential exhaustion of hematopoietic progenitors in Pep−/− mice after IFN-α treatment.
Pep−/− hematopoietic progenitors demonstrated increased activation and proliferation after IFN-α administration
Previous studies have shown that chronic type I IFN stimulation induces hematopoietic progenitors to proliferate and lose self-renewal capacity, resulting in exhaustion and HSC dysfunction (Essers et al., 2009). To investigate the mechanism leading to decreased numbers of progenitor cells in Pep−/− BM after IFN-α treatment, we assessed BrdU incorporation by progenitor cells as a measure of progenitor proliferation. Consistent with increased activation observed in poly(I:C)-treated mice (Fig. 2 G), Pep−/− myeloid progenitors incorporated BrdU at a higher rate compared to Pep+/+ progenitors in poly(I:C)-treated mice (Fig. 3 A). We observed a similar increase in BrdU in Pep−/− compared to Pep+/+ myeloid progenitors after low-dose Ad5-IFN-α5 (107 PFU/mouse) administration (Fig. 3 B). This increased proliferation in Pep−/− LS−K, MEP, and CMP/GMP progenitors were associated with increased entry into the cell cycle after poly(I:C) treatment (Fig. 3, D–F). In contrast, no differences in BrdU incorporation or cell cycling were detected between Pep+/+ and Pep−/− HSC progenitors (LSK) after poly(I:C) treatment (Fig. 3 C).
To analyze the in vivo consequences of increased proliferative capacity of the Pep−/− hematopoietic progenitors, mice were treated with the chemotherapeutic agent 5-fluorouracil (5-FU) to kill actively proliferating cells. Previous work has demonstrated that repeated poly(I:C) infection to induce chronic systemic IFN-α, followed by 5-FU administration results in BM failure and death due to loss of HSC self-renewal (Essers et al., 2009). Pep−/− mice administered a single dose of poly(I:C), followed by 5-FU, had 33% mortality (Fig. 3 G). In contrast, all Pep+/+ mice treated with poly(I:C) were able to compensate and recover after the chemotherapeutic challenge. The greater sensitivity of Pep−/− progenitors to IFN-α was not due to an intrinsic proliferation defect in Pep−/− progenitors, since both Pep+/+ and Pep−/− mice, in the absence of poly(I:C), had similar mortality after chronic 5-FU treatment (Fig. 3 H). Thus, our data suggests that PEP regulates the proliferation of hematopoietic progenitors during IFN-α–mediated stress hematopoiesis.
To determine the hematopoietic requirement for PEP in IFN-α–mediated hematopoietic progenitor proliferation, we generated mixed BM chimeras by transferring Pep+/+ or Pep−/− donor (CD45.2+) BM competitively with host (CD45.1+) BM at a 1:1 ratio. Reconstituted chimeric mice were treated with poly(I:C) and administered BrdU 24 h later. Donor-derived CD45.2+ Pep−/− myeloid progenitors (LS−K) incorporated more BrdU compared to donor-derived CD45.2+ Pep+/+ myeloid progenitors (Fig. 4 A, left graph). In contrast, host-derived (CD45.1+) myeloid progenitors (LS−K) incorporated BrdU equivalently (Fig. 4 A, right graph).
To assess the contribution of IFNAR for progenitor activation, we generated competitive BM chimeric mice with donor-derived (CD45.2+) Pep−/− or Pep−/−IFNAR−/− at a 1:1 ratio with host-derived (CD45.1+). Chimeric mice were treated with poly(I:C) for 24 h, and up-regulation of Sca-1 on cKit+ progenitors was analyzed by FACS. Although Pep−/− cKit+ progenitors up-regulated Sca-1 after poly(I:C) treatment (Fig. 4 B, compare left and middle), Pep−/−IFNAR−/− progenitors had reduced Sca-1 up-regulation compared to Pep−/− progenitors in poly(I:C)-treated mice (Fig. 4 B, compare middle and right). In addition, whereas Pep−/− mice treated with poly(I:C) had reduced percentage of LS−K progenitors when compared to untreated Pep−/− mice, Pep−/− IFNAR−/− mice treated with poly(I:C) had a similar percentage of LS−K progenitors compared to untreated Pep−/− mice (Fig. 4 C). These data suggest that PEP expression in the hematopoietic compartment is necessary for augmented activation and proliferation in response to IFN-α signaling through the IFNAR.
PEP is a negative regulator of IFNAR signaling in progenitor cells
Our data and that of others (Essers et al., 2009) suggest an intrinsic requirement for IFNAR signaling on progenitor cells during IFN-α–mediated disease and stress hematopoiesis. To determine the role of PEP in IFNAR signaling, lineage-depleted BM cells from Pep+/+ and Pep−/− mice were treated with IFN-α and STAT1 phosphorylation was assessed by Western blot using an anti–phospho-specific STAT1 antibody. Addition of recombinant mouse IFN-α4 (rIFN-α4) to Pep−/− progenitors demonstrated a twofold increase in STAT1 phosphorylation on tyrosine 701 compared to Pep+/+ progenitors (Fig. 5, A and B). Increased STAT1 phosphorylation was also observed using in vitro–generated Pep−/− myeloid progenitors (Fig. 5, C and D). To determine the downstream consequences of enhanced STAT phosphorylation in Pep−/− progenitor cells, we treated Pep+/+ and Pep−/− mice with poly(I:C) for 16 h and sorted lineage-negative progenitor cells for transcriptome analysis. Consistent with increased phosphorylation and activation of STAT1, IFN-responsive genes Isg15, Irf4, Mx1, and Osmr were significantly up-regulated in Pep−/− compared to Pep+/+ progenitor cells (Fig. 5 E and Table S1).
As PEP has been demonstrated to play a positive regulatory, rather than an inhibitory, function for TLR signaling in mouse macrophages (Wang et al., 2013), we analyzed the role of PEP in IFNAR signaling of BMDMs. Addition of rIFN-α4 to in vitro–differentiated BMDMs resulted in a time-dependent increase in STAT1 phosphorylation on tyrosine 701 that was further enhanced in Pep−/− BMDMs (Fig. 5, F-G). Whereas Mx1 and Irf4 mRNAs were not induced or below detection levels (CT > 35) in BMDMs, IFN-responsive genes Irf8, Il7, and Prlr mRNAs (Bovolenta et al., 1994; Martin et al., 2004; Hou et al., 2015) were further augmented in Pep−/− compared to Pep+/+ BMDMs after treatment with rIFN-α4 (Fig. 5 H and Table S2). Together, these data support a negative regulatory role for PEP on IFNAR signaling in HSCs and BMDMs.
To further explore the biochemical nature by which PEP regulates IFNAR signaling, we expressed PEP in 3T3 cells that are IFN-α responsive. 3T3 cells expressing transfected PEP demonstrated reduced STAT1 and STAT2 phosphorylation after rIFN-α4 stimulation compared to vector control cDNA-transfected cells (Fig. 6, A and B). The phosphatase activity of PEP was required for maximal reduction of STAT1 phosphorylation because expression of a catalytically inactive PEP(C227S) did not reduce STAT1 phosphorylation to the same extent as wild-type PEP (Fig. 6, C and D). Finally, we investigated whether PEP could co-localize with the IFNAR signaling complex. In 3T3 cells transfected with a Pep cDNA, PEP co-immunoprecipitated with multiple components of the IFNAR signaling complex, including JAK1, TYK2, SOCS1, IFNAR, and STAT1 (Fig. 6 E). Hence, PEP is physically positioned to regulate IFNAR signaling.
Our studies here provide a mechanistic basis by which PTPN22 and IFN-α pathways may cooperate to manifest cytopenias in SLE. Pep−/− mice treated with IFN-α or poly(I:C) demonstrate greater morbidity/mortality than treated Pep+/+ mice. Although IFN-α has pleiotropic effects on both immune and nonimmune cells, our studies here reveal a novel role for PEP as a negative regulator of IFNAR signaling in hematopoietic progenitors. IFN-α induced a greater degree of activation as assessed by STAT1 phosphorylation and increased transcription of type I IFN-dependent genes and cell cycle entry of Pep−/− progenitors. In turn, these compound effects of IFN-α and PEP deficiency in mice result in severe anemia, thrombocytopenia, and neutropenia. IFN-α has recently emerged as an important regulator of hematopoiesis. IFN-α induces dormant HSCs to proliferate, and chronic IFN-α stimulation results in HSC dysfunction and exhaustion (Essers et al., 2009). Similarly, mice deficient in IRF2, a critical negative regulator of type I IFN signaling, have greater numbers of proliferating as opposed to dormant HSCs (Sato et al., 2009). As SLE patients with the PTPN22 at-risk variant have higher serum levels of IFN-α (Kariuki et al, 2008), these synergistic effects of PTPN22 and IFN-α may provide an additional mechanistic basis for the BM dysfunction observed in SLE.
PEP was first demonstrated to regulate TCR signaling through association with CSK (Cloutier and Veillette, 1996), a PTK that represses SRC-family PTKs. Overexpression of PEP in T cells inhibits T cell functions (Cloutier and Veillette, 1999; Gjörloff-Wingren et al., 1999). Generation of Pep−/− mice on a nonautoimmune prone background (C57BL/6) revealed an essential function for PEP in effector/memory, but not naive, T cells. Pep−/− effector T cells demonstrate enhanced TCR signaling and, as Pep−/− mice age, they develop splenomegaly, spontaneous germinal center formation, and elevated serum IgG1, IgG2a, and IgE levels, though without any clinical manifestations of autoimmunity (Hasegawa et al., 2004). PEP also plays an important role in regulating weak agonist antigens in naive OT-1 TCR transgenic T cells on a Rag1−/− background or when competitively transferred into a profoundly lymphopenic environment by regulating TCR signaling and by modulating LFA-1–dependent inside-out signaling and adhesion (Salmond et al., 2014). Whereas autoantibody production was not observed in Pep−/− mice on a nonautoimmune prone strain (C57BL/6; Hasegawa et al., 2004), Pep−/− mice expressing a wedge CD45(E613R) mutation, that ablates dimerization-induced inhibition of CD45 phosphatase activity, on a C57BL/6 background developed systemic autoimmunity with autoantibody production, glomerulonephritis, and increased mortality greater than either single-mutant mice (Zikherman et al., 2009). Hence, PEP plays an important role in determining signaling thresholds in autoreactive T cells to maintain immunologic tolerance.
Studies of peripheral human T and B cells expressing the at-risk PTPN22(C1858T) allele suggested a gain-of-function phenotype with accumulation of circulating memory T cells, decreased numbers of circulating memory B cells, decreased TCR signaling, and impaired BCR signaling (Vang et al., 2005; Rieck et al., 2007; Arechiga et al., 2009). Further analysis of autoreactive B cells indicate that the PTPN22(C1858T) allelic variant interferes with deletion of autoreactive B cells (Menard et al., 2011; Habib et al., 2012).
Two groups have recently generated mice expressing the orthologous PEP(R619W) mutation that differ in their genetic backgrounds. In both cases, PEP(R619W) knock-in (KI) mice phenocopy Pep−/− mice with development of enhanced positive and negative T cell selection, enhanced TCR signaling, splenomegaly, lymphadenopathy, and spontaneous germinal center formation (Zhang et al., 2011; Dai et al., 2013). PEP(R619W) KI mice on a C57BL/6 background also exhibited enhanced antigen presentation by dendritic cells and modest increases in BCR signaling (Zhang et al., 2011). PEP(R619W) KI mice on a mixed C57BL/6J and 129/Sv genetic backgrounds differed from Pep−/− or PEP(R619W) KI C57BL6/J mice as they also demonstrated enhanced TCR signaling in naive T cells, altered B cell selection, and enhanced BCR signaling in mature B cells. Moreover, PEP(R619W) knock-in mice on a mixed background develop systemic autoimmunity with high-titer autoantibodies, vasculitis, and increased mortality (Dai et al., 2013).
The PTPN22(C1858T) polymorphism has now been described for a variety of autoimmune disorders. PTPN22/LYP associates with CSK in resting T cells and upon TCR activation, PTPN22 dissociates from CSK and translocates to lipid rafts where PTPN22/LYP can dephosphorylate and inactivate the TCR signalsome (Cloutier and Veillette, 1996; Vang et al., 2012). The PEP(R620W) variant alters the proline-rich region (P1) to disrupt its interaction with CSK (Begovich et al., 2004; Bottini et al., 2004). Although some have proposed that the PTPN22(C1858T) at risk variant can provide greater inhibition of antigen–receptor signaling due to its compromised ability to interact with and be sequestered by CSK (Vang et al., 2012), another model suggests that the PTPN22(C1858T) at risk variant may differentially affect downstream effectors than PTPN22 deficiency (Dai et al., 2013).
In addition to T and B cell antigen receptor signaling, PTPN22/PEP can play both positive and negative functions depending on receptor and cellular context. PEP plays a requisite function to dephosphorylate phosphoanandamide to generate anandamide, a lipid second messenger with antiinflammatory effects produced by macrophages upon LPS induction (Liu et al., 2006). In the human THP-1 monocytic cell line, PTPN22 co-immunoprecipitates with SOCS1 and inhibits SOCS1 tyrosine phosphorylation to enhance IFN-γ signaling (Spalinger et al., 2013). In mouse macrophages, PEP interacts with TRAF3 and positively regulates type I IFN production after TLR stimulation. Pep−/− macrophages stimulated with LPS or poly(I:C) produce less IFN-β (Wang et al., 2013). Consistent with this requisite function for PEP in TLR signaling in macrophages, peripheral blood monocytes from PTPN22(C1958T) carriers for the at-risk allele demonstrated reduced STAT1 phosphorylation and reduced type 1 IFN-induced gene expression after LPS challenge (Wang et al, 2013). Contrary to the studies of Wang et al. (2013) using IFN-α and IFN-β, our studies demonstrate an inhibitory role of PEP in IFN-α4–mediated IFNAR signaling in BMDMs (Fig. 5, F–H). The basis of these differences is unclear and may relate to differences in IFN-α subtypes used as the potencies of the 12 IFN-αs differ (Hillyer et al., 2013), stimulation conditions, and/or differences in IFNAR signaling of IFN-α and IFN-β (de Weerd et al., 2013).
Our studies here support an inhibitory function for PEP in IFNAR function in hematopoietic precursors and BMDMs. PEP phosphatase activity is required for optimal IFNAR regulation, though it is likely that additional noncatalytic scaffolding domains can also contribute to PEP function. Additionally, as PEP can be coimmunoprecipitated with the IFNAR signaling complex, PEP is appropriately positioned to down-regulate IFN-α–induced signaling through JAK1, TYK2, SOCS, STATs, and other IFNAR regulators. Additional biochemical studies will be required to better define the molecular nature by which PEP regulates IFNAR signaling.
A similar duality of function has also been demonstrated for the CD45 PTPase. Whereas CD45 is required for T and B cell development, as well as T and B cell antigen receptor signaling (Hermiston et al., 2003), CD45 plays an inhibitory role in cytokine, IFNAR, and α5β1 integrin-mediated signaling (Penninger et al., 2001). CD45 interacts with JAKs and down-regulates IL-3 and cKit-mediated proliferation of BM-derived mast cell lines, erythropoietin-mediated erythropoiesis, IL-3–mediated differentiation of BM precursors to neutrophil/macrophage colonies, and IFN-regulated antiviral responses (Irie-Sasaki et al., 2001). In addition, CD45 is required for α5β1 integrin-mediated adhesion to fibronectin, an activity dependent on both CD45 phosphatase activity and CD45 association with CD45-associated protein (Shenoi et al., 1999).
In summary, our study provides insights by which PEP and IFN-α might cooperate to give rise to dysfunctional hematopoiesis. As a consequence of dysregulated IFN-α expression in SLE, patients carrying the PTPN22(C1858T) risk variant may have enhanced IFN-α–mediated JAK–STAT signaling in HSCs to exhaust their precursor pool that manifest in hematologic abnormalities in SLE.
MATERIALS AND METHODS
B6.SJL-Ptprca Pepcb/BoyJ mice were purchased from The Jackson Laboratory. Pep−/− mouse generation was previously described (Hasegawa et al., 2004). IFNAR−/− mice were purchased from The Jackson Laboratory for breeding with Pep+/+ and Pep−/− mice. All mice were maintained under specific pathogen–free conditions unless otherwise noted. Poly(I:C) (LMW) was purchased from InvivoGen, and mice were administered 200 µg/mouse i.p. for the indicated times. 5-FU was purchased from APP Pharmaceuticals. For 5-FU studies, mice were administered 200 µg poly(I:C) i.p. and, 48 h later, injected with a single dose of 5-FU (150 mg/kg), or mice were injected once weekly for the length of the experiment. All animal experimentation protocols were approved by the Laboratory Animal Resources Committee at Genentech, Inc. (South San Francisco, CA).
8-wk-old (CD45.1+) Pep+/+ (B6.SJL-Ptprca Pepcb/BoyJ) mice were lethally irradiated (2 doses of 500 cGy with 137Cs γ-irradiator) at 3-h intervals and ∼3 h before injection. BM from Pep+/+ (CD45.1+) mice were isolated and mixed at a 1:1 ratio with either Pep+/+ or Pep−/− (CD45.2+) or Pep−/−IFNAR−/− (CD45.2+) BM in Dulbecco’s Modification of Eagle’s Medium. Irradiated mice (CD45.1+) were injected in the lateral tail vein with approximately 1 × 106 total BM cells. All irradiated mice were given antibiotic-supplemented water (1.1 g/l neomycin and 110 mg/l polymycin B; Sigma-Aldrich) for 2 wk.
Adenoviral and recombinant IFN-α.
Mouse IFN-α5 was cloned into a pShuttleX vector and transferred into Ad5 viral vector by the Baylor College of Medicine Vector Development Lab (VDL). Ad5-IFN-α and Ad5-LacZ control viruses were purified on CsCl gradients and titered per VDL specification. Virus was administered by i.v. injection into 16-20-wk-old female mice. For in vitro experiments, recombinant mouse IFN-α4 (rIFN-α4) was purchased from PBL Assay Sciences.
BM-derived myeloid progenitor cells were generated from hematopoietic progenitor cells isolated from BM using a Miltenyi MACS lineage depletion kit. Cells were stimulated for 3 d in IL-3 (25 ng/ml), IL-6 (25 ng/ml), and SCF (50 ng/ml), washed, and stimulated for indicated times with rIFN-α4 (2,000 U/ml, PBL Assay Science). BMDMs were differentiated by incubating total BM cells in 15-cm tissue culture plates with recombinant M-CSF (100 ng/ml; Genentech) in DMEM for 7-10 d. M-CSF (100 ng/ml) was replenished every 3 d during culturing. BMDMs were plated overnight in M-CSF, rested for 4 h and stimulated for the indicated times with rIFN-α4 (2,000 U/ml; PBL Assay Science). For cell transfection experiments, 3T3 cells were transfected using Lipofectamine LTX with plus reagent (Life Technologies) according to manufacturer’s instruction.
Complete blood count.
Blood was collected from IFN-α treated Pep+/+ and Pep−/− mice in Microtainer tubes (BD) containing EDTA. Cell counts were analyzed on a XT-V Automated Hematology Analyzer (Sysmex).
Renal histologic evaluation.
Kidneys from IFN-α administered Pep+/+ (n = 5) and Pep−/− (n = 3) mice were fixed in 10% neutral buffer formalin, processed, and embedded in paraffin for sectioning. ∼3-µm sections were de-paraffinized and stained with hematoxylin and eosin or Periodic acid-Schiff (PAS). Renal lesions were scored on arbitrary severity scales from 0 to 3 for each of the features: glomerulopathy, interstitial nephritis, and arteritis. Proteinuria was evaluated in urine collected weekly. Urine protein levels were scored with Multistix 10 SG (Siemens) dipsticks on a Clinitek Status Analyzer (Siemens). Protein levels are scored as 1 (30 mg/dl), 2 (100 mg/dl), or 3 (>300 mg/dl).
ANA and BAFF ELISA.
Serum IgG antinuclear antibodies (ANA) were measured using an Alpha Diagnostic International per the manufacturer’s instruction. BAFF detection was determined using an anti-BAFF soluble mouse detection kit (Enzo Life Sciences) per manufacturer’s instruction.
Single-cell suspensions from spleen, blood, BM, or BALF were washed in PBS containing 5 mM ethylenediaminetetraacetic acid and 0.5% BSA. Cells were incubated for 15 min with Fc receptor block (clone 2.4G2; BD) before antibody staining. The following antibodies were used for FACS staining and sorting: biotinylated antibodies for lineage-negative staining-biotin lineage staining cocktail includes CD3, B220, GR-1, CD11b, and Ter119, as well as NK1.1, CD11c, and DX-5 (BD), were added to the staining cocktail; CD45.2 (104), cKit (2B8), and CD34 (Ram34) were purchased from BD; CD16/32 (93), Sca-1 (D7), and CD135 (A2F10) were purchased from eBioscience; CD45.1 (A20), CD48 (HM48-1), and CD150 (TC15-12F12.2) were purchased from BioLegend. Aqua Fluorescent reactive dye (Life Technologies) was used to discriminate viable from dead cells. Intracellular BrdU and 7AAD for cell cycle analysis using the FITC BrdU Flow kit (BD) was performed per the manufacturer’s protocol. Flow cytometry data was analyzed using FlowJo software (Tree Star).
3T3 cells were lysed in 100 mM Tris buffer containing 1% Triton X-100, protease inhibitor, phosphatase inhibitor, NaF (10 mM), and PMSF (1 mM) for 30 minutes with rotation at 4oC, and cleared by centrifugation at 12,000 rpm for 5 min. Antibodies used for immunoprecipitation included IFNAR1 (Santa Cruz Biotechnology, Inc.), STAT1 (Cell Signaling Technology), SOCS1 (Abcam), TYK2 (Abcam), and JAK1 (Santa Cruz Biotechnology, Inc.). Protein/antibody conjugates were precipitated with protein A Dynabeads (Life Technologies) and washed four times in lysis buffer. Beads were boiled in LDS buffer (Life Technologies) and loaded on 8% Tris-glycine gel (Novex; Life Technologies).
Western blot analysis.
Whole-cell lysates were generated with RIPA lysis buffer (Thermo Fisher Scientific) with protease and phosphatase inhibitors, DTT (1 mM), NaF (10 mM), and PMSF (1 mM). Total protein from each fraction was determined by BSA quantification method (Thermo Fisher Scientific). Cell lysates were loaded on a NuPAGE 4-12% Bis-Tris gel (Life Technologies) and transferred to PVDF membrane using an iBlot Gel transfer device (Invitrogen). Membranes were blocked with 5% milk and 0.2% Tween in PBS. Antibodies specific for ACTIN, STAT1, and STAT2 were purchased from Cell Signaling Technology. Antibodies for pSTAT1, pSTAT2, SOCS1, and TYK2 were purchased from Abcam. Anti-PEP (P1) antisera to detect PEP by Western blotting have been previously described (Hasegawa et al., 2004). All antibodies were diluted in blocking buffer and incubated overnight at 4oC. Secondary antibodies included HRP (Millipore)-conjugated goat anti–rabbit or goat anti–mouse antibodies in conjunction with Amersham ECL prime Western blotting detection reagent (GE Healthcare) to visualize bands using x-ray film (Kodak). Bands were measured by densitometry analysis using ImageJ analysis software (National Institutes of Health).
Mouse IFN and receptor PCR array.
RNA was prepared using RNeasy Mini Plus kits (QIAGEN) and reverse-transcribed with RT2 First Strand kit (QIAGEN), in accordance with the manufacturer’s instructions. Mouse IFN and receptor PCR Array was performed according to manufacturer’s instruction (QIAGEN). PCR array (PAMM-064ZE) was performed and gene expression measured on a 7900HT Fast Real-Time PCR System (Applied Biosystems).
Online supplemental material.
Table S1 shows increased IFN-responsive gene transcription in Pep−/− BM progenitors. Table S2 shows IFN-responsive genes in BMDMs.
All authors are or were (D.A. Holmes and H.R.C. Smith) employees of Genentech, Inc. The authors declare no additional competing financial interests.
Author contributions: D.A. Holmes designed experiments, generated, and interpreted data, and prepared the manuscript. Q. Ou and Q. Gong designed, generated, and interpreted data presented in Fig. 5 (F–H) and Table S2. H.R.C. Smith contributed to the initial design and execution of the experiments represented in Fig. 1. A.C. Chan designed experiments, interpreted data, and prepared the manuscript. E. Suto and W.P. Lee performed injections of Ad5 virus. P. Caplazi performed pathology analysis.
B cell activating factor
B lymphocyte stimulator
common myeloid progenitor
granulocyte myeloid progenitor
hepatitis C virus
hematopoietic stem cell
protein tyrosine kinase
systemic lupus erythematosus
single nucleotide polymorphism
signal transducer and activator of transcription