Generation ofHem1–deficient mice using chemical mutagenesis

In the course of the ENU program, a flow cytometry screen identified a pedigree (denoted NTB.1 for no T or B cells) in which the mice presented with an increased percentage of CD11b⁺ myeloid cells and a decreased percentage of B220⁺ B lymphocytes in peripheral blood samples (Fig. 1 A). To identify the gene responsible for the NTB.1 phenotype, G1 and G2 NTB.1 carriers were crossed to C3HeB/FeJ mice, and the resulting F1 animals were intercrossed to generate F2 and N2 animals for mapping (see Materials and methods). The levels of B220⁺ B cells in peripheral blood were assessed in the resultant generations. A genome scan of affected and unaffected animals indicated that the NTB.1 mutation mapped to chromosome 15 between 59.8 and 60.1 cM. Candidate genes were sequenced and a single-nucleotide polymorphism was identified in exon 13 of theHem1 gene, which changed a CAG (Gln) to TAG (stop), resulting in premature termination at residue 445 (Fig. 1 B). Immunoblotting of whole cell lysates from normal littermate andHem1 mutant thymocytes, splenocytes, and BM cells with an anti-Hem1 antibody (25) revealed the absence of Hem1 protein in mice homozygous for the NTB.1 mutation (Fig. 1 C; and seeFig. 7 F).

Hem1 is expressed predominantly in hematopoietic cells

A previous Northern blot study using cell lines derived from various tissues suggested thatHem1 expression is restricted to hematopoietic tissues (20). We performed a more extensive survey ofHem1 expression using real-time PCR forHem1 message in cDNA derived from 25 different mouse tissues. We find that althoughHem- is predominantly expressed in hematopoietic tissues, it is also expressed in urogenital tissues including testis (Fig. S1, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1). In contrast,Hem2 is not expressed in hematopoietic tissues but is widely expressed in many other tissues (Fig. S1). Indeed, mice deficient inHem2 die around embryonic day 8.5–10.5 because of morphological abnormalities ranging from severe neurulation defects to complete resorption (26).

We next characterized the expression pattern ofHem1 in different hematopoietic cell lineages and progenitor cells. We find thatHem1 is expressed in lineage-negative (Lin⁻) Sca1⁺cKit⁺ side population (SP)⁺ cells, which are highly enriched for HSCs, as well as Lin⁻Sca1⁺cKit⁺ SP⁻ cells and Lin⁻Sca1⁻cKit⁻ cells, which contain early progenitors (Fig. S1).Hem1 is also expressed in B and T cell progenitors during all stages of maturation and in purified macrophages and neutrophils (Fig. S1). The results collectively indicate thatHem1 is predominantly expressed in developing and mature hematopoietic cells, whereasHem2 is expressed in nonhematopoietic cells.

Hem1-deficient mice exhibit lymphopenia, neutrophilia, and anemia

We performed complete blood counts (CBCs) and differential cell counts to determine the representation of mature hematopoietic cell types in peripheral blood fromHem1^−/− and normal WT littermate mice. We find that loss of Hem1 results in a moderate increase in total white blood cells (WBCs) per microliter of blood, which corresponds to a 25-fold increase in the number of neutrophils and a 3.5-fold decrease in the number of lymphocytes per microliter of blood inHem1^−/− mice compared with WT mice (Figs. 2, A–C).Hem1^−/− mice also exhibit altered RBC indices, such as decreased hematocrit and mean corpuscular volume (MCV;Fig. 2 A), with reticulocytosis (increased RBC progenitors;Fig. 2 D), anisocytosis (variation in RBC size), echinocytosis (membrane spickles), and poikilocytosis (abnormally shaped RBC), which is characterized by keratocytes (horned-shaped cells), dacryocytes (tear-shaped cells), schistocytes (RBC fragmentation), and acanthocytes (thorny cells;Fig. 2 E and not depicted).Hem1^−/− RBCs exhibit increased fragility relative to WT RBCs, as measured by an increased percentage of lysis in response to hypotonic NaCl solutions (Fig. 2 F). Collectively, these RBC indices are suggestive of a microcytic hypochromic hemolytic anemia inHem1^−/− mice, which parallels the anemia phenotype of Rac-deficient mice (27).

Gross and histological examination ofHem1^−/− mice reveal a spectrum of lesions, including dense amyloid accumulations containing foci of mineral at the margins of the liver (Fig. 2 G), and occasional diffuse inflammation in the liver, epididymis, mesentery, thoracic pleura, heart, and lungs (not depicted). Membranoproliferative glomerulopathy (Fig. 2 H) and diffuse lymphoid hypoplasia are also common lesions, whereas myocardial fibrosis, amyloidosis, and pancreatic atrophy occur less frequently (not depicted).Hem1^−/− spleens are substantially enlarged, even though lymphoid cells are depleted, because of marked proliferation of myeloid and erythroid progenitors (Fig. S2, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1), which is indicative of extramedullary hematopoiesis.Hem1^−/− thymi are hypoplastic, and lack the normal cortical/medullary architecture evident in WT thymi (unpublished data).

We next examined the potential of clonogenic hematopoietic progenitor cells in the BM, spleen, and blood fromHem1^−/− and WT mice. We performed in vitro quantitation of myeloid (CFU-GM), erythroid (burst forming units-erythroid [BFU-E]), and multi-lineage (CFU-granulocyte/erythroid/macrophage/monocyte [CFU-GEMM]) assays using standard techniques. We find a significant reduction in CFU-GM, BFU-E, and CFU-GEMM in BM fromHem1^−/− mice compared with WT BM (Fig. S2). In contrast, there is a dramatic increase in CFU-GM, BFU-E, CFU-GEMM colonies in spleens and peripheral blood fromHem1^−/− mice compared with WT mice. This correlates with a fourfold increase in the percentage of Lin⁻Sca1⁺cKit⁻ cells in the spleens ofHem1^−/− mice compared with WT mice, which is consistent with increased mobilization of HSC and progenitors (HSC/Ps) from the BM to peripheral blood and spleen (Fig. S2).

Hem1 is required for migration, phagocytosis, and F-actin polymerization in neutrophils

Hem1^−/− mice exhibit an increase in the percentage (Fig. 3, A and B) and total number (not depicted) of cells bearing Mac1 and Gr1 (Fig. 3, A and B [top]), and CD61 (Fig. 3, A and B [bottom]) in the BM and spleen relative to WT mice, suggesting that myeloid cell homeostasis is altered by loss of Hem1 protein. We next determined whether loss of Hem1 affects neutrophil and macrophage functions. Purified neutrophils fromHem1^−/− and WT mice were loaded into trans-well plates, and the ability of cells to migrate through a filter containing different concentrations of MLP (a tripeptide chemoattractant) was measured.Hem1^−/− neutrophils fail to migrate efficiently across the filter in response to different concentrations of fMLP (Fig. 3 C).Hem1^−/− neutrophils are also deficient in their ability to phagocytose fluorescent-conjugated beads (Fig. 3 D) and bacteria (not depicted) in response to MLP stimulation, as measured by flow cytometry. Likewise,Hem1^−/− macrophages are deficient in their ability to phagocytose fluorescent-labeledEscherichia coli after 2 and 4 h of stimulation, relative to WT macrophages (Fig. 3 E). However,Hem1^−/− macrophages produce normal or increased levels of the cytokines IL1β, TNF-α, Mip2 (macrophage-inflammatory protein 2), and cytokine-induced neutrophil chemoattractant (KC) after stimulation withE. coli or LPS, and the transcription of the corresponding genes, as well asIL-23, are increased (Fig. S3, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1).

The polymerization of F-actin is critical for forming phagosomes, as well as lamellipodia and filopodia, which are fingerlike projections responsible for polarizing the cytoskeletal machinery during cell migration. As shown inFig. 3 F, >25% of WT neutrophils polymerize F-actin after 15 s of MLP treatment, as determined by increased phalloidin staining, whereas <5% ofHem1^−/− cells polymerize F-actin even after 120 s of MLP stimulation. Deconvolution images reveal that polymerized F-actin inHem1^−/− cells is distributed diffusely in the cytoplasm, whereas F-actin in WT cells is clearly polarized toward the leading edge (Fig. 3 G). Transfection ofHem1 cDNA into BM-derived neutrophils (28) from WT orHem1^−/− mice resulted in restoration of F-actin polymerization in response to MLP stimulation (Fig. S4, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1). The results collectively indicate that Hem1 is required in innate immune cells for a variety of functions dependent on F-actin polymerization and cytoskeletal reorganization, including phagocytosis and migration.

Lymphocyte development is impaired in Hem1-deficient mice

BecauseHem1-deficient mice are lymphopenic, we examined whether T and/or B cell development are impaired. The development of lymphocytes can be broken down into many distinct stages based on the presence or absence of particular intracellular and surface proteins, as determined by flow cytometry (29,30). As shown inFig. 4 A,Hem1^−/− mice have a 30-fold mean reduction in thymocyte number with an impairment in αβ T cell development within the CD4⁻CD8⁻ double-negative (DN) to CD4⁺CD8⁺ double-positive (DP) cell transition at the CD25^midCD44⁻ (DN3) to CD25⁻CD44⁻(DN4) cell stage, which is mediated by formation of the preTCR. The number of γδ thymocytes is also decreased, albeit to a lesser extent than αβ T cells (Fig. 4, A and B). The total numbers of peripheral CD4, CD8, and γδ T cells (Fig. 4 B) and B lymphocytes (not depicted) are reduced by at least 75%. The ratio of Foxp3⁺CD4⁺ T regulatory cells (T_reg) to Foxp3⁻CD4⁺ T cells is significantly higher in both thymus (Fig. 4 A) and spleen (Fig. 4 B), suggesting that Hem1 regulates both lymphocyte development and homeostasis.

Hem1^−/− hematopoietic stem and progenitor cells compete poorly with WT cells in BM chimeras

To determine if the defects in T and B cell development are autonomous to hematopoietic cells or are caused by a defective microenvironment, we transferred marked Lin⁻ (lineage negative;Fig. 5) or total BM cells (Fig. S5, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1) from WT (CD45.1⁺ marker),Hem1^−/− (CD45.2⁺), or mixed WT andHem1^−/− Lin⁻ cells into unirradiated (Fig. 5), or lethally irradiated (Fig. S5),Rag2^−/−γc^−/− (recombinase activating gene 2/common γ chain double-null) hosts (CD45.2⁺), and lymphocyte development was analyzed 8–20 wk after transplant. As shown inFig. 5 A,Rag2^−/−γc^−/− mice are completely blocked in T cell development at the CD4⁻CD8⁻ DN stage (columns 1 and 2). Injection of WT Lin⁻ cells intoRag2^−/−γc^−/− mice completely reconstitutes T cell development in the thymus, leading to normal representation of CD4 and CD8 cells in the spleen (Fig. 5 A, column 3). In contrast,Hem1^−/− Lin⁻ cells poorly reconstitute the thymi ofRag2^−/−γc^−/− mice, with the majority of cells residing at the DN stage (Fig. 5 A, column 4).Hem1^−/− CD4 and CD8 SP cells can mature and expand inRag2^−/−γc^−/− hosts, as determined by the presence of CD4⁺CD45.2⁺ and CD8⁺CD45.2⁺ cells in the spleen of transplanted mice. However, whenHem1^−/− Lin⁻ cells are mixed with WT Lin⁻ cells at 1:1, 10:1, or 20:1 (Hem1^−/−/WT) ratios, they fail to reconstitute the thymus or spleen of transplanted mice, as determined by the presence of CD4 and CD8 cells bearing the CD45.1 (WT) but not CD45.2 markers (Fig. 5 A column 5; Fig. S5; and not depicted). The inability ofHem1^−/− hematopoietic cells to compete with WT hematopoietic cells in BM chimeras is not caused by inadequate space in the BM niche because we obtain similar results after transplantation of WT,Hem1^−/−, or mixed 1:10 WT:Hem1^−/− Lin⁻ or total BM cells into lethally irradiatedRag2^−/−γc^−/− hosts (Fig. S5 and not depicted).Hem1^−/− BM cells can contribute to the generation of GR-1⁺ myeloid cells when mixed 1:10 WT/Hem1^−/− BM cells and transplanted into lethally irradiatedRag2^−/−γc^−/− hosts, indicating that HSC/Ps are able to home to the BM niche in mixed BM chimeras (Fig. S5). In addition,Hem1 heterozygous (+/−) BM cells compete normally with WT BM cells to produce CD4⁺, CD8⁺, GR1⁺, and B220⁺ when mixed 1:10 WT/Hem1^+/−, indicating that the inability ofHem1^−/− cells to contribute is not because of allorejection by WT cells (Fig. S5).

We next determined the ability of Hem1-deficient progenitor cells to reconstitute the B cell lineage in either unirradiated (Fig. 5 B) or lethally irradiatedRag2^−/−γc^−/− hosts (Fig. S5). As shown inFig. 5 B,Rag2^−/−γc^−/− mice are completely blocked in B cell development at the B220⁺IgM⁻ pro–B cell stage (columns 1 and 2). Lin⁻CD45.1⁺ WT progenitors reconstitute B220⁺IgM⁺ cells in the BM and spleen ofRag2^−/−γc^−/− hosts (Fig. 5 B, column 3). In contrast, Lin⁻CD45.2⁺Hem1^−/− progenitors fail to generate mature B cells in unirradiatedRag2^−/−γc^−/− hosts (Fig. 5 B, column 4), despite partial reconstitution of mature T cells (Fig. 5 A). However,Hem1^−/− progenitors can generate a small population of mature B cells when transplanted into lethally irradiatedRag2^−/−γc^−/− hosts (Fig. S5). WT, but notHem1^−/− progenitors, develop normally into B cells when mixed 1:1 or 1:10 with Hem1-deficient progenitors before transplantation intoRag2^−/−γc^−/− hosts (Fig. 5 B, columns 5 and 6; and Fig. S5). These results suggest that Hem1 plays an important role in the development and/or homeostasis of lymphocytes.

Loss of Hem1 inhibits T cell activation and proliferation and enhances IL-17 production

We next examined whether loss of Hem1 affects T cell activation and T helper differentiation. We find that Hem1 deficiency results in impaired T cell activation based on reduced CD69 expression (Fig. 6 A and Fig. S6, available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1), decreased cell division as assessed by limited CFSE dye dilution of live-gated cells (Fig. 6 B and Fig. S6), and reduced tritiated thymidine incorporation (Fig. S6) after TCR stimulation. However, IL-2 and IFN-γ are produced normally byHem1^−/− T cells after TCR stimulation (Fig. 6, C and E), and exogenous IL-2 does not rescue proliferation ofHem1^−/− T cells after α–CD3-ε/CD28 stimulation (Fig. 6 B). These results suggest that the inability ofHem1^−/− T cells to divide is not because of the inability to make IL-2. Surprisingly,Hem1^−/− T cells produce 15-fold more IL-17 and twofold more IL-6 and TNF-α after α-CD3/CD28 stimulation, compared with their WT counterparts (Fig. 6 C), and contain increased serum IL-17 and IL-6 in vivo (Fig. S6). Purified CD4 T cells also contain a greater percentage of Th17 effector cells after in vitro stimulation, as assessed by intracellular staining with α–IL-17 and α–IL-2 antibodies (Fig. S7). To assess whether Hem1 might modulate Th17 differentation, we purified naive WT andHem1^−/− CD4 T cells and subjected them to Th1 or Th17 polarizing conditions in the presence of irradiated APCs. We find that Th1 differentiation occurs normally in the absence of Hem1 based on an equivalent percentage of IFN-γ–producing cells (Fig. S7) and equivalent IFN-γ production (Fig. 6 E) fromHem1^−/− and WT CD4 T cells after Th1 culture conditions. In contrast, the generation of Th17 cells is enhanced after IL-17 polarizing conditions with the addition of TGF-β (Figs. 6 D). It is unlikely that the skewing toward Th17 cells is caused by secondary infections because we only occasionally find histological evidence of infection inHem1^−/− mice, andHem1^−/− mice treated with antibiotics throughout their lives still produce increased IL-17 (Fig. S7).

Hem1 is required for formation of the WAVE complex and F-actin polymerization

To determine how loss of Hem1 results in defective T cell activation and proliferation, we examined whether F-actin polymerization is altered inHem1^−/− T cells. Thymocytes and peripheral T cells were stimulated with α–CD3-ε/CD28 for 24 h, followed by PMA/Ionomycin for 15 min, and then stained with Alexa Fluor 488–conjugated phalloidin. BothHem1^−/− thymocytes (Fig. 7 A) and T cells (not depicted) fail to polymerize F-actin after α–CD3-ε/α-CD28 stimulation, relative to WT thymocytes and T cells.Hem1^−/− T cells also fail to form synapse-like actin cap structures after stimulation with α–CD3-ε-coated beads (Fig. 7 B). Thymocytes and T cells fromHem1^−/− mice exhibit reduced integrin-mediated cell adhesion to fibronectin after TCR stimulation, relative to WT cells (Fig. 7 C). These results suggest an essential role for Hem1 in cytoskeletal reorganization and cell adhesion at the immune synapse.

We next asked at what point Hem1 deficiency impinges on the signaling pathways leading to F-actin polymerization in lymphocytes and neutrophils.Hem1^−/− thymocytes and T cells express normal levels of the membrane-proximal signaling molecules CD3-ε (not depicted), Lck, Zap70, and Rac1 (Fig. 7 D, right). In addition, we observe equivalent levels of total phosphotyrosine, active (phospho) extracellular regulated mitogen-activated protein kinase (Erk), and active Rac1 (Pak1 binding) in WT andHem1^−/− thymocytes, T cells stimulated with α–CD3-ε (Fig. 7 D), and neutrophils stimulated with MLP (Fig. 7 E), suggesting that membrane proximal signaling is intact.Hem1^−/− CD4 T cells also exhibit normal nuclear translocation of NF-κB (active NF-κB) after TCR stimulation and slightly increased basal NF-κB translocation before stimulation relative to WT CD4 T cells, as measured using ImageStream quantitative flow cytometry (Fig. S7) (31,32). However, analysis of whole cell lysates from WT andHem1^−/− thymocytes, T cells, and neutrophils indicates that loss of Hem1 results in the near complete absence of the WAVE complex proteins Wave2, Abi1, Abi2, and Sra1 in thymocytes and purified T cells (Fig. 7 F, left and middle), whereas WAVE members are reduced inHem1^−/− neutrophils (Fig. 7 F, right). The reduction in WAVE complex proteins is not caused by decreased transcription becauseAbi1,Abi2,Sra1, andWave2 mRNA are expressed normally inHem1^−/− T cells relative to WT T cells (Fig. S8). In addition, acute small inhibitory RNA (siRNA) knockdown (33) ofHem1 in WT T cells in vitro results in decreased amounts of Wave2, Abi2, and Sra1 (Fig. S8). These results suggest that Hem1 is required for stabilization and/or translation of the WAVE complex proteins in hematopoietic cells (Fig. 8), and they are consistent with several recent studies indicating that siRNA knockdown of Hem1 in HL-60 cells (25) or Jurkat T cells (34) results in degradation of WAVE complex proteins, which are essential mediators of F-actin polymerization.

ENU mutagenesis, screening, and mutation mapping.

Mutagenesis with ENU was performed on C57BL/6J male mice according to the protocol used by Justice et al. (56). Two mating schemes were used. G2 NTB.1 carriers were crossed to C3HeB/FeJ and the resulting F1 animals were intercrossed to generate F2 animals for mapping. In addition, the G1 NTB.1 carrier was mated to C3HeB/FeJ mice and the resulting F1 animals were backcrossed to the G1 carrier to generate N2 mice for mapping. F2 and N2 animals were screened for the NTB.1 trait and DNA was collected. DNA was analyzed with 85 microsatellite markers spaced a mean of 15 cM apart. Linkage analysis was performed using MapManager QTX13b software (K. Manly, Roswell Park, Buffalo, NY). The ∼360-kbNTB.1 candidate interval was narrowed between 59.8 and 60.1 cM on chromosome 15. Computational analysis identified 11 genes, whose exons were PCR amplified from genomic DNA and sequenced.

Mice.

Hem-1 mutant mice were housed under specific pathogen-free conditions. Mouse procedures were approved by the Institutional Animal Care and Use Committees of Celltech R&D, Inc., and the University of Washington. The NTB.1 line was screened and maintained by DNA sequencing after amplification withHem1 forward (5′-TGCAGAGCTTCTGTTCCTGTTGGA-3′) and reverse (5′-AGAAGGAACTGCTGGCTACCCAAA-3′) primers and by flow cytometry comparing the ratio of B220^low to Gr-1^high cells. The majority of the phenotypic analysis ofHem1^−/− mice was performed on intercrosses between mice backcrossed three generations onto C57BL6J (from the original C57BL6J × C3H hybrids).

Generation of neutrophils by in vitro cell culture.

Single-cell suspensions of BM cells were cultured in 6-well culture dishes at 2 × 10⁶ cells/ml. Cells were grown with DMEM/20% FBS medium in the presence of IL-3/GM-CSF (28). After 3 d, nonadherent cells were removed, and adherent cells were continuously grown with IL-3/GM-CSF. After an additional 10 d, nonadherent cells were used in the experiments after initiation of the culture. Nonadherent cells were stained with Gr-1/Mac1 and analyzed by flow cytometry.

Erythrocyte osmotic fragility assay.

Blood was obtained from WT andHem1^−/− mice and diluted 1:1,000 in PBS containing EDTA for the cell counting. 20 × 10⁶ erythrocytes in 100 μl (0.9% NaCl) were dropped into solutions containing either 0.9% saline or a hypotonic solutions to generate final NaCl concentrations of 0.86, 0.74, 0.69, 0.47, or 0.26% NaCl. In brief, erythrocyte samples were slowly dropped (without mixing) into 250-μl NaCl solutions, incubated for 10 min, and then centrifuged for 1 min at 1,300 rpm. The supernatant was analyzed for released hemoglobin by measuring the absorbance at 540 nm. Samples were normalized to a standard curve generated from complete lysis (100% lysis) and zero lysis (0.9% NaCl) for each of the mice (57).

Generation of BM chimeras.

Rag2^−/−γc^−/− mice (Taconic) were reconstituted withHem1^−/− (KO; CD45.2) or C57BL/6J WT (CD45.1; Jackson ImmunoResearch Laboratories) Lin⁻ cells or mixed populations of Lin⁻ or total BM cells (WT:Hem1^−/−, 1:1, 1:10, or 1:20). Lineage⁺ cells were depleted using the Mouse lineage panel kit (BD) according to manufacturer's instructions. Lin⁻ cells (2 × 10⁴ cells/each) were injected i.v. into nonirradiated or 950 rad–irradiatedRag2^−/−γc^−/− recipients. In other experiments, total BM cells depleted of CD3⁺ cells were used for BM chimera studies. Thymus, BM, and spleen cells were collected from the recipient mice 6–20 wk after reconstitution.

Rac activation assay.

The EZ-Detect Rac1 activation kit (Thermo Fisher Scientific) was used to pull down active Rac1. The PAK1-bound active Rac1 was detected by Western blot analysis (52) using α-Rac1 antibody (BD) and visualized using the ODYSSEY infrared imaging system (LI-COR Biosciences) located at the Fred Hutchinson Cancer Research Center (Seattle, WA).

Neutrophil purification and chemotaxis assays.

Neutrophils were purified from WT andHem1^−/− BM using Histopaque 1119 and 1077 gradients (Sigma-Aldrich). Chemotaxis assays were conducted according to a protocol provided by Neuro Probe, Inc. The number of migrated cells per 400× field were counted for three random spots and mean values were displayed on the y axis (6).

In vitro phagocytosis assay.

Neutrophils or peritoneal macrophages (10⁵ cells) were incubated with fluorescent beads (Polysciences, Inc.) or fluorescentE. coli (Invitrogen) for 0, 2, and 4 h. The intensity of phagocytosed beads orE. coli was then analyzed by flow cytometry.

Measurements of cytokines and chemokines.

Cytokines (IL-2, IFN-γ, IL-17, IL-6, IL-1β, and TNF-α) and chemokine (MIP2 and KC) levels were quantified with capture/detection antibody-based ELISA systems (eBioscience; R&D Systems), according to the manufacturer's instructions. IL-17 in sera from WT andHem1^−/− mice was assessed using a Biosource (Invitrogen) mouse IL-17 antibody bead kit according to the manufacturer's recommendations. In brief, 100 μl of sera were incubated with fluorescent beads conjugated to the capture antibody. After addition of the detection antibody mix and PE-conjugated streptavidin, plates were read on a LUMINEX machine using xMAP technology (QIAGEN).

Cell adhesion assay.

Flat-bottom Maxisorb 96-well plates were coated with 0.5 μg/ml of human fibronectin (BD) and bovine serum albumin (Sigma-Aldrich). Cells were added to the wells, stimulated with α–CD3-ε, washed, and counted under an inverted microscope as previously described (11).

Capping formation, immunofluoresence, and confocal microscopy.

Thymocytes from WT andHem1^−/− mice were stimulated with α-CD3/α-CD28 for 24 h, followed by PMA for 15 min. Cells were fixed, permeabilized, and stained with Alexa Fluor 488 phalloidin. F-actin polymerization and actin capping formation were determined by flow cytometry and confocal microscopy after phalloidin–Alexa Fluor 488 and DAPI staining, as previously described (3,7). Hamster IgG– or α–CD3-ε-coated 5-μm sulfate white microspheres were prepared according to the manufacturer's protocol (Invitrogen).

T cell isolation and Th differentiation.

CD4⁺CD44^lowCD25⁻CD62L⁺ T cells (4 × 10⁴ cells) sorted using FACS Vantage or FACS Aria (BD) were activated with plate-bound 1 μg/ml α-CD3 and irradiated APC (10⁵ cells, 3,000 rads) in the presence of 10 ng/ml IL-12 (R&D Systems), 10 μg/ml α–IL-4 antibody (11B11; eBioscience), and 5 ng/ml IL-2 (R&D Systems) for Th1 differentiation condition, as previously described (4). For Th17 differentiation conditions, different doses of TGF-β (5, 1, 0.2, and 0 ng/ml; R&D Systems), IL-6 (50 ng/ml, R&D Systems), α–IL-4 (10 μg/ml), and α–IFN-γ (10 μg/ml) antibodies were added to culture media (58).

Western blot analysis.

Western blot analyses were performed as previously described (59) using rabbit polyclonal antibodies specific for Hem1 (gift from O.D. Weiner, University of California, San Francisco, San Francisco, CA), β-actin (Santa Cruz Biotechnology, Inc.), Wave2 (Santa Cruz Biotechnology, Inc.), Abi1 (Santa Cruz Biotechnology, Inc.), Abi2, and Sra1 (gifts from A.M. Pendergast [Duke University, Durham, NC] and J.D. Scott [University of Washington, Seattle, WA]), Lck, Zap70 (Millipore), and donkey α–goat IgG HRP (Santa Cruz Biotechnology, Inc.), goat α–rabbit IgG HRP (Bio-Rad Laboratories), goat α–mouse IgG HRP (Bio-Rad Laboratories), 4G10 antibody (Millipore), Phospho-Erk, and total Erk (Cell Signaling Technology) (38).

Histology and hematology.

Tissues were embedded in paraffin and stained with hematoxylin and eosin (Histology Consultation Services, Inc.). CBCs were performed by Phoenix Central Laboratory (Everett, WA).

T cell proliferation.

T cells (10⁶ cells/ml) were labeled with CFSE dye (10 μM; Invitrogen) and stimulated with α-CD3/α-CD28 in the presence of 10 ng/ml IL-2 (R&D Systems) for 4 d. Cell divisions were determined in 1 nM of live TO-PRO-3 (Invitrogen) and negative cells were determined by flow cytometry, as previously described (59). In addition, cells were labeled with³H thymidine for 18 h, and cell proliferation was determined as previously described (60).

Flow cytometry and cell sorting.

Hem1^−/− and WT tissues (BM, thymus, spleen, and LN) were stained with fluorescent-conjugated antibodies specific for CD8, GR1, B220 (Invitrogen), CD62L, Sca1, cKit, Foxp3, IL-2, IL-17, IFN-γ (eBioscience), CD4, CD3-ε, CD69, CD43, ICOS, IgM, MAC1, GR1, Ter119, CD61, CD45.1, CD45.2, Annexin V, CD44, CD25, β-TCR, γδ-TCR, BP1, and heat stable antigen (BD) and analyzed by flow cytometry as previously described (59,60). To purify enriched HSC/Ps, Lin⁻ cells were then sorted based on expression of ckit (CD117), Sca1 (Ly-6A/E), and Hoechst 33342 (Invitrogen) to define SP cells (61).

Statistical analysis.

Data were analyzed using the Student's one-tailedt test. P-values < 0.05 were considered as significant values.

Online supplemental material.

Fig. S1 shows that theHem1 gene is preferentially expressed in lymphoid tissues and HSC. Fig. S2 shows decreased clonogenic hematopoietic cell potential in the BM and increased clonogenic hematopoietic cell potential in the blood and spleen inHem1^−/− mice. Fig S3 shows thatHem1^−/− peritoneal macrophages produce normal or increased levels of proinflammatory cytokines in response to toll-like receptor signaling. Fig. S4 shows that transfection ofHem1 plasmid rescues F-actin formation inHem1^−/− granulocytes. Fig. S5 shows thatHem1^−/− BM cells compete poorly with WT BM cells after transfer into irradiatedRag2^−/−/γc^−/− mice. Fig. S6 shows that loss of Hem1 impairs T cell proliferation and survival, whereas proinflammatory cytokine production is increased. Fig. S7 shows thatHem1^−/− mice produce increased IL-17 and exhibit normal or increased NF-κB activation. Fig. S8 shows that components of the WAVE complex are transcribed normally inHem1^−/− mice, and siRNA knockdown ofHem1 in WT T cells results in decreased amounts of WAVE complex proteins. Online supplemental material is available athttp://www.jem.org/cgi/content/full/jem.20080340/DC1.

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