Essential Role of Elmo1 in Dock2-Dependent Lymphocyte Migration

Catherine Stevenson, Gonzalo de la Rosa, Christopher S. Anderson, Patrick S. Murphy, Tara Capece, Minsoo Kim and Michael R. Elliott
J Immunol June 15, 2014, 192 (12) 6062-6070; DOI: https://doi.org/10.4049/jimmunol.1303348

Abstract

Elmo1 and Elmo2 are highly homologous cytoplasmic adaptor proteins that interact with Dock family guanine nucleotide exchange factors to promote activation of the small GTPase Rac. In T lymphocytes, Dock2 is essential for CCR7- and CXCR4-dependent Rac activation and chemotaxis, but the role of Elmo proteins in regulating Dock2 function in primary T cells is not known. In this article, we show that endogenous Elmo1, but not Elmo2, interacts constitutively with Dock2 in mouse and human primary T cells. CD4+ T cells from Elmo1−/− mice were profoundly impaired in polarization, Rac activation, and chemotaxis in response to CCR7 and CXCR4 stimulation. Transfection of full-length Elmo1, but not Elmo2 or a Dock2-binding mutant of Elmo1, rescued defective migration of Elmo1−/− T cells. Interestingly, Dock2 protein levels were reduced by 4-fold in Elmo1−/− lymphocytes despite normal levels of Dock2 mRNA. Dock2 polyubiquitination was increased in Elmo1−/− T cells, and treatment with proteasome inhibitors partially restored Dock2 levels in Elmo1−/− T cells. Finally, we show that Dock2 is directly ubiquitinated in CD4+ T cells and that Elmo1 expression in heterologous cells inhibits ubiquitination of Dock2. Taken together, these findings reveal a previously unknown, nonredundant role for Elmo1 in controlling Dock2 levels and Dock2-dependent T cell migration in primary lymphocytes. Inhibition of Dock2 has therapeutic potential as a means to control recruitment of pathogenic lymphocytes in diseased tissues. This work provides valuable insights into the molecular regulation of Dock2 by Elmo1 that can be used to design improved inhibitors that target the Elmo-Dock-Rac signaling complex.

Introduction

Chemokine signaling is an integral component of lymphocyte trafficking, activation, and survival. Rac is a member of the Rho family of GTPases that are central drivers of actin cytoskeleton dynamics downstream of most chemokine receptors (1–4). Rac cycles between inactive (GDP-bound) and active (GTP-bound) states largely because of the action of guanine nucleotide exchange factors (GEFs). Dock2 is a ∼200 kDa Rac-GEF restricted to hematopoietic cells in mice and humans (5). Through the use of Dock2−/− mice, it is now well established that Dock2 is essential for Rac activation and chemotaxis in lymphocytes downstream of multiple chemokine receptors, including CCR7 and CXCR4 (1, 2, 6). Dock2-deficient lymphocytes show greatly reduced entry, egress, and interstitial motility in lymphoid and nonlymphoid tissues. In murine models of cardiac allograft rejection and diabetes, deletion of Dock2 in lymphocytes was found to be protective, pointing to a role for Dock2 in migration of pathogenic T cells (7, 8). Over the past decade, Dock2 has also been found to regulate a range of Rac-dependent functions in neutrophils, dendritic cells, and NKT cells (9–12). However, the molecular regulation of Dock2 is poorly understood, particularly in primary cells and in human lymphocytes.

Elmo1 (75 kDa) is a cytoplasmic adaptor protein that physically associates with members of the Dock-A family of Rac-GEFs, of which Dock1 and Dock2 are the best characterized (5, 13, 14). Extensive structure–function analyses by a number of groups have shown that Elmo binding enhances Dock1 signaling by increasing its Rac-GEF activity, membrane localization, and protein stability (13, 15–21). Studies in invertebrate models and mammalian cell lines have revealed an evolutionarily conserved role for Elmo1 in regulating Dock-Rac signaling in numerous cellular functions, including morphology, motility, and phagocytosis (13, 18, 22–25). Elmo1 has also been shown to interact with Dock2 to promote Rac activation and migration in rodent cell lines (22, 26). More recently, studies in Elmo1−/− mice revealed a critical role for Elmo1 in apoptotic cell clearance during spermatogenesis and hippocampal neurogenesis (27, 28). However, as most of our insights into Elmo function stem from studies of Elmo1 and Dock1 in nonhematopoietic cells, the mechanisms and outcomes of Elmo1-dependent regulation of Dock proteins in leukocytes remain largely unknown.

Elmo1 and Elmo2 are 87% similar at the amino acid level, are widely expressed, and based on Dock1 studies, have largely been considered to be functionally redundant (13). Both proteins contain pleckstrin homology and proline-rich/PxxP domains located in the C-terminal 100 aa (15, 29, 30). These C-terminal regions mediate multiple associations with the N termini of Dock1 and Dock2, as revealed through crystallographic and biochemical analyses (30, 31). Dock1 and Dock2 contain an N-terminal Src homology 3 domain that mediates interaction with the C-terminal polyproline regions of Elmo1 and Elmo2. Interestingly, this PxxP-Src homology 3 association has been reported to be essential for Elmo1 interaction with Dock2, but not Dock1 (31). The PxxP motif is conserved between mouse and human Elmo1 and Elmo2 (PKEP, Elmo1714–717), but whether Elmo2 can interact with or regulate Dock2 has not been reported.

In this study, we used a number of approaches to address the function of Elmo1 and Elmo2 in regulating Dock2 in primary mammalian lymphocytes. Using Elmo1−/− mice and primary human T cells, we demonstrate a previously unknown, nonredundant role for Elmo1 in regulation of endogenous Dock2 that may provide insight toward the development of Dock2-targeting therapeutics.

Materials and Methods

Mice

Animal experiments were approved by the University of Rochester Animal Care and Use Committee. Elmo1-deficient mice have been described elsewhere (27). All mice were 6–12 wk of age and on a C57BL/6J background of at least 10 generations.

Reagents

Commercial reagents were purchased as follows: all chemicals unless noted otherwise were purchased from Sigma-Aldrich; chemokines were from PeproTech; ICAM-1 from R&D Systems; Protein A/G agarose beads from Santa Cruz; 24-well transwell chambers from Corning; TaqMan quantitative RT-PCR (qRT-PCR) probes from LifeTech; cell culture media from Cellgro; and CFSE and TAMRA from Invitrogen. BSA was purchased from Sigma-Aldrich (A4503).

Abs

Flow cytometry Abs used in this study were CD3ε (17A2), CD4 (GK1.5), CD8α (53-6.7), CD11b (M1/70), CD16/32 (93), CD19 (6D5), CD45 (Ly-5), B220 (RA3-6B2), CXCR4 (2B11), CCR7 (4B12), F4/80 (BM8). Commercial Abs for immunoprecipitation (IP) and immunoblotting (IB) were β-actin (Sigma-Aldrich); Dock2 (Millipore); Dock1 (Santa Cruz); and GAPDH, Rac, pERK1/2 (9109), ERK1/2 (9102), phospho-Ser473 AKT (D9E), pan-AKT (40D4), HA, K48 polyubiquitin (D9D5), and DYKDDDDK “flag” tag (Cell Signaling Technology). Elmo1 and Elmo2 Abs were provided by K.S. Ravichandran (15, 27) and were further tested in this study to confirm specificity (see Supplemental Fig. 3).

Cell isolation

Mice were euthanized by CO2 asphyxiation. Spleen and lymph nodes were disaggregated through a 70-μm mesh filter and RBCs lysed using ACK lysis buffer (Sigma-Aldrich). Human leukocytes were obtained from healthy deidentified donors (New York Blood Center), and PBMCs were isolated by Accu-Prep (Accurate Chemical). PBMCs were first depleted of CD14+ cells by positive magnetic selection using MACS separation CD14 MicroBeads (Miltenyi Biotec). Mouse and human CD4 T cells were isolated by negative magnetic selection using the CD4+ T cell Isolation Kit II (Miltenyi Biotec). For isolation of murine macrophages, peritoneal cavities were lavaged with 5 ml cation-free PBS twice and immediately stained and FACS sorted.

Cell culture

Primary murine CD4 T cells were cultured at 37°C/5% CO2 in RPMI 1640, 20% FBS, 40 U/ml rIL-2, 10 mM HEPES, 1% penicillin-streptomycin (pen-strep)/l-glutamine. Jurkat T cells (clone E6.1) were grown at 37°C/5% CO2 in RPMI 1640, 10% FBS, 10 mM HEPES, 1% pen-strep/l-glutamine.

Cell staining for flow cytometry

For flow cytometry, cells were resuspended in cold FACS buffer (cation-free PBS, 0.5% BSA, 0.05% NaN3), incubated with 1:100 FcR blocking Abs on ice for 10 min before addition of fluorescently labeled Abs for 25 min on ice. Staining with CXCR4 and CCR7 Abs was carried out at 25°C. Cells were washed once and resuspended in FACS buffer before analysis or sorting.

IP and IB

Cell lysates were prepared using lysis buffer (20 mM Tris-HCl pH 7.5, 1% Triton X-100, 150 mM NaCl, 1 mM EDTA, 1× final protease inhibitor mixture III [Calbiochem]) or Laemmli buffer. Protein quantification of lysates was done using bicinchoninic acid reagent (Pierce). Lysates were boiled for 10 min, and protein separation was carried out on 4–15% SDS-PAGE mini gel (Bio-Rad). After transfer to polyvinylidene difluoride at 100 V for 1 h, membranes were blocked for 1 h with 5% nonfat dry milk/TBST before overnight incubation with indicated Abs at 4°C. SuperSignal Pico or Dura ECL reagents were used per manufacturer’s instructions (Pierce). For IP, cell pellets were resuspended in lysis buffer and rotated at 4°C for 10 min before centrifugation at 12,000 × g, 5 min, 4°C. Cleared lysates were incubated with anti-Dock2 (1:250), normal rabbit IgG (1:250), or anti-Elmo1 (1:100), in total volume of 500 μl and rotated 18 h at 4°C. A total of 25 μl protein A/G beads was then added to each sample and rotated for 2 h at 4°C. Beads were washed four times in lysis buffer and boiled in Laemmli buffer. For coimmunoprecipitation (co-IP) analysis of endogenous Elmo1 and Elmo2, IPs were split equally after boiling, loaded in duplicate lanes, and blotted with anti-Elmo1 or anti-Elmo2 Abs separately. For co-IP of Elmo-Flag with Dock2 in 293T cells, all of each anti-Dock2 IP was loaded in a single lane and blotted for Flag using HRP-conjugated anti-DYKDDDDK. For detection of K48-ubiquitinated Dock2 under denaturing conditions, anti-Dock2 IPs from wild-type (WT) CD4+ T cells were boiled for 5 min in 50 μl denaturing buffer (50 mM Tris pH 7.5, 70 mM 2-ME, 1% SDS) followed by addition of 350 μl IP lysis buffer and a second round of IP with anti-Dock2 for 2 h at 4°C before IB analysis.

qRT-PCR

RNA was isolated using DNase I–treated RNeasy columns (Qiagen), and cDNA was synthesized from 10–100 ng RNA using iScript (Bio-Rad). qRT-PCR was performed on a 7300 Real Time Thermocycler (Life Technologies) using SensiFast Probe Hi-ROX polymerase (Bioline) and the following gene-specific TaqMan probes from Life Technologies: Elmo1 (Mm00519109_m1); Elmo2 (Mm00475454_m1); Dock2 (Mm00473720_m1); Actb (Mm00607939_s1). Values were obtained using a relative standard method. In brief, a 2-fold dilution standard curve of total cDNA was used to determine expression levels of each gene for each specimen. Expression levels were then normalized to Actb levels. For comparisons across genes, a calibrator sample was used to account for varying relative levels of each gene in the standard curve sample.

Time-lapse video microscopy

T cell motility experiments were carried out on Δ T dishes (Bioptechs) coated first with Protein A (10 μg/ml; Invitrogen), then ICAM-1 Fc (10 μg/ml; R&D) and 4 μg/ml CCL21 or CXCL12. Splenic CD4+ T cells were labeled with either 0.5 μM CFSE or 1 μM TAMRA-SE (Invitrogen) for 1 h at 37°C/5% CO2. Cells were washed and resuspended at 5 × 105/ml in Leibovitz’s L-15 media supplemented with glucose (2 mg/ml) and cultured at 37°C for 20 min before being added to the microscopy dish. Dish was secured on a heated stage, and imaging was done with an epifluorescence Nikon Eclipse Ti microscope. Images were acquired every 15 s for 15 or 30 min using a 20× objective.

Migration assays

Transwell chemotaxis assays were performed using 24-well plates with 5-μm pore size inserts (Corning). Cells were equilibrated at 37°C/5% CO2 in migration medium (RPMI 1640, 1% BSA, 10 mM HEPES, 1% pen-strep/l-glutamine) at 1 × 106 cells/ml for 30 min before use. A total of 500 μl chemoattractant in migration medium was applied to the lower chamber and 100 μl cells applied to the upper chamber. After 1 h at 37°C/5% CO2, inserts were discarded and 50 μl Accucount beads (5.1-μm diameter; Spherotech) were added to each lower chamber and input samples (100 μl cells plus 400 μl medium) for quantitation by flow cytometry. For postmigration Ab staining, 250 μl cells from the lower chamber was removed before adding beads and stained with indicated Abs. Percent migration was determined as follows: 100 × ([cell events in lower chamber/bead events in lower chamber]/[input cell events/input bead events]). Staining and quantitation were carried out with two to three replicates per condition.

Determination of Rac-GTP, phospho-AKT, and phospho-ERK levels

Active Rac levels were determined using GST-PAK beads (Cytoskeleton) according to manufacturer’s instructions, with the following modifications. CD4+ cells were incubated in migration medium at 1 × 106/ml for 30 min at 37°C/5% CO2. Cells were pelleted and resuspended at 2–3 × 106 cells per 200 μl stimulation medium (RPMI 1640, 10 mM HEPES, 1% pen-strep/l-glutamine). Cells were incubated for 10 min in 37°C water bath and stimulated by addition of 200 μl of 500 ng/ml chemokine in stimulation medium for 15 s. After stimulation, cells were immediately placed on ice, and 400 μl ice-cold TBST was added to each sample. Cells were then pelleted at 4,000 × g, 1 min, 4°C and lysed in 165 μl recommended lysis buffer, and lysates cleared at 10,000 × g, 1 min, 4°C. Cleared lysates were transferred to fresh tubes containing 15–30 μg GST-PAK beads and samples were rotated for 1 h at 4°C. Beads were washed two to three times with recommended wash solution and pellets were boiled 10 min in Laemmli buffer, separated on 12% SDS-PAGE and analyzed by IB. For phospho-protein analysis, T cells were stimulated as described earlier except they were immediately lysed in 1× Laemmli buffer before SDS-PAGE and IB.

Transfection

Jurkat T cells were transfected as previously described using the ECM 830 Square Wave Electroporation system (BTX) (32). The following SMARTpool ON-TARGET Plus small interfering RNA (siRNA) duplexes were purchased from Thermo Scientific: nontargeting pool (D-001810-10-05) and human Elmo1 (L-012851-00-0005). HEK 293T cells were transfected with 1 μg empty pEBB- Flag (vector) or Dock2-Flag (from M. Matsuda) (5) plus 4 μg pEBB-Elmo-Flag plasmids by calcium phosphate (Profection; Promega). Primary T cells were transfected using the Mouse T cell Nucleofector Kit (Amaxa), with 2 × 106 CD4+ T cells, 1 μg pMAX-GFP (Amaxa), and 4 μg expression vectors. Elmo expression plasmids and the pEBB-Flag backbone were provided by K.S. Ravichandran. MT123-HA-ubiquitin vector was provided by Dirk Bohmann (33).

Data quantitation and statistical analysis

Densitometry values were determined by area under the curve analysis on ImageJ software (National Institutes of Health). Quantitative analysis of time-lapse images was carried out using Nikon software (Nikon). Flow cytometry data were analyzed using FlowJo software (Tree Star). Data shown are the average ± SEM. Statistical significance was determined using the two-tailed Student t test, where p < 0.05 was considered significant.

Results

Impaired migration of Elmo1−/− lymphocytes to CCR7 and CXCR4 ligands

We first analyzed Elmo1 levels in WT and Elmo1−/− mice by IB. Splenocytes were FACS-sorted based on expression of B220 and CD3 surface markers, and cell lysates were analyzed by IB with anti-Elmo1. Elmo1 protein was readily detected in WT but not Elmo1−/− CD3+ (T cells) and B220+ (B cells) splenocytes (Fig. 1A and 1B). CD3−/B220− splenocytes, which are comprised largely of CD11b+ myeloid cells, also expressed Elmo1, albeit at reduced levels compared with lymphocytes (Fig. 1B). The frequency and total numbers of T and B cells in the spleen and lymph nodes of Elmo1−/− mice were not significantly different from WT (Fig. 1A and Supplemental Fig. 1). Also, the frequency of naive splenic T cells (CD62Lhi/CD44lo) was similar between WT and Elmo1−/− mice (Supplemental Fig. 1C).

FIGURE 1.
FIGURE 1.

Impaired migration of Elmo1−/− splenocytes. (A) Splenocytes from WT and Elmo1−/− mice were labeled with anti-B220 and anti-CD3, and analyzed by flow cytometry. The percentage of B220+ and CD3+ cells among all live splenocytes (negative for 7-amino-actinomycin D) is shown. (B) Splenocytes were labeled as in (A), FACS-sorted, and analyzed by IB with Abs indicated (to the left of each blot). Relative molecular masses are shown (right). One representative experiment of five is shown for (A) and (B). (C) Transwell migration of splenocytes from WT, Elmo1+/−, and Elmo1−/− mice to indicated concentrations of CXCL12. The percent of CD3+ and B220+ splenocytes that migrated to the lower chamber was determined by flow cytometry (n = 6 mice/genotype, ± SEM). *p < 0.05, ***p < 0.001, ****p < 0.0001.

Splenocytes from WT, Elmo1+/−, and Elmo1−/− were tested for migration to a range of CXCL12 concentrations in transwell chambers. At all doses tested, WT and Elmo1+/− splenocytes migrated similarly, whereas Elmo1−/− splenocytes showed significantly reduced migration (Fig. 1C). Using flow cytometry, we calculated the fraction of total B220+ (B cells) and CD3+ (T cells) splenocytes that migrated through the transwell. We observed a near-complete loss of T cell migration to CXCL12 by Elmo1−/− T cells, whereas Elmo1−/− B cells showed an intermediate but significant decrease in migration compared with WT (Fig. 1C). The ex vivo survival of unstimulated lymphocytes was similar between WT and Elmo1−/− after 24 h in culture (data not shown). Thus, the loss of Elmo1 in primary lymphocytes results in defective migratory responses to CXCL12.

Based on the strong effect of Elmo1 deletion on T cell migration, we further examined migration responses of CD4+ T cells from WT and Elmo1−/− mice. CD4+ T cells were isolated from lymph nodes and spleen, and analyzed for transwell migration to CXCL12, CCL19, and CCL21. Elmo1−/− CD4+ T cells from both tissues were significantly reduced in their capacity to migrate to these chemokines, although some migration of Elmo1−/− T cells was observed, particularly at higher chemokine doses (Fig. 2A and 2B). To determine whether loss of Elmo1 specifically affected T cell polarization and migration, we used time-lapse microscopy to measure motility patterns of CD4+ T cells plated on ICAM-1 plus CCL21 or CXCL12. CD4+ T cells isolated from WT and Elmo1−/− spleens were labeled with CFSE or TAMRA-SE cell-permeable dyes, mixed in equal numbers, and plated for microscopic analysis. T cells from WT and Elmo1−/− mice adhered to ICAM-1, but although the majority of WT cells adopted a polarized morphology and migrated along the surface, Elmo1-deficient cells largely failed to do so (Fig. 2C–E and Supplemental Videos 1 and 2). Together, these results show that Elmo1 is required for normal T cell motility responses to CCR7- and CXCR4-dependent chemokines.

FIGURE 2.
FIGURE 2.

Defective chemotaxis of Elmo1−/− T cells to CCR7 and CXCR4 ligands. (A) CD4+ T cells isolated from lymph nodes (mediastinal and axillary, left) or spleen (right) were analyzed for transwell migration to CXCL12 (25 ng/ml) and CCL19 (10 ng/ml). One representative experiment of three is shown. (B) Transwell migration of CD4+ T cells from WT and Elmo1−/− mice to CCL21. (C–E) CD4+ splenic T cells labeled with CFSE (WT, green) or TAMRA (Elmo1−/−, red) were plated on a glass dish coated with rmICAM-1 (10 μg/ml) and rmCCL21 (4 μg/ml). Epifluorescence images were acquired every 15 s with 20× objective. A representative series of time-lapse images from a single field containing one WT and one Elmo1−/− cell is shown. The speed (C) and polarization (D) of individual cells were calculated for all cells in one field over 30 min. One representative experiment of two to three for each condition is shown.

T cells require Elmo1 for CCR7- and CXCR4-mediated Rac activation

The failure of Elmo1−/− T cells to polarize and migrate suggested a requirement for Elmo1 in chemokine signaling. Because we found that the surface expression of CCR7 and CXCR4 on splenic lymphocytes was comparable between WT and Elmo1−/− (Fig. 3A), we focused on activation of key molecular pathways downstream of CCR7 and CXCR4. The GTPase Rac is rapidly activated upon chemokine stimulation and is critically required for actin polymerization and polarization during T cell migration (4). To determine whether loss of Elmo1 affects chemokine-induced Rac activation, we measured Rac-GTP levels in CD4+ T cells by GST-PAK pull-down and anti-Rac IB of lysates from WT and Elmo1−/− T cells after CXCL12 or CCL19 stimulation. In WT cells, Rac was clearly activated by both chemokines, whereas Elmo1−/− T cells showed no Rac activation (Fig. 3B). Under the same stimulation conditions, we observed robust phosphorylation of AKT (Ser473) and ERK1/2 (Thr202/Tyr204) in both WT and Elmo1−/− T cells (Fig. 3C). These results indicate that Elmo1 is specifically required for CXCR4 and CCR7 activation of Rac, but not PI3K and Ras/Raf/MEK pathways.

FIGURE 3.
FIGURE 3.

CCR7 and CXCR4 signaling in Elmo1−/− T cells. (A) CXCR4 and CCR7 surface levels on splenic lymphocyte populations from WT and Elmo1−/− mice were determined by flow cytometry. Filled histograms show isotype staining of WT cells. One representative experiment of four is shown. (B and C) CD4+ T cells isolated from WT and Elmo1−/− spleens were stimulated 15 s with 250 ng/ml CXCL12 or CCL19 and lysed. (B) Lysates were analyzed for Rac-GTP levels by GST-PAK pull-down and anti-Rac IB. Bottom graph, Rac-GTP levels normalized to total Rac and expressed as relative to medium control for n = 4 mice/genotype ± SEM. (C) Phospho-Ser473-AKT and phospho-Thr202/Tyr204-ERK1/2 levels were determined by IB. Blots were stripped and reprobed for total AKT and ERK1/2. One representative experiment of three is shown. *p < 0.05.

Elmo1, but not Elmo2, interacts with Dock2 in primary lymphocytes

Dock2 is the primary GEF responsible for Rac activation downstream of CCR7 and CXCR4 in T cells (1, 2). Previous work has shown that Elmo1 and Dock2 can interact upon overexpression in cell lines, but the relevance of this interaction in primary lymphocytes is not known (26, 31). We tested for endogenous Elmo-Dock2 complexes in T cells by co-IP. Elmo1 and Elmo2 were present in anti-Dock2 IPs of Jurkat lysates, whereas Dock2, but not Elmo2, was present in anti-Elmo1 IPs of these cells (Fig. 4A). This confirmed that our co-IP approach was sufficient to detect native Elmo1-Dock2 and Elmo2-Dock2 complexes, and also indicated that endogenous Dock2 may interact exclusively with either Elmo1 or Elmo2. Using this approach, we tested for Elmo-Dock2 complexes in normal mouse splenocytes and primary human CD4+ T cells. Surprisingly, we could detect Elmo1, but not Elmo2, in Dock2 IPs from these primary cells (Fig. 4B). We then tested whether Dock2 may bind more readily to Elmo1 than Elmo2. Differences in the sensitivities of our anti-Elmo Abs prevented us from testing this with endogenous proteins, so we expressed 1× Flag-tagged Elmo1 or Elmo2 along with Dock2 in 293T cells and examined Elmo-Dock2 complex formation by anti-Dock2 IP and anti-Flag IB. As shown in Fig. 4C, Elmo1 and Elmo2 were expressed at equivalent levels in 293T cells and, as was seen in Jurkat cells (Fig. 4A), both were detectable in anti-Dock2 IPs. However, significantly more Elmo1 coprecipitated with Dock2 (∼2.5-fold) compared with Elmo2 (Fig. 4C). We attempted to directly compare binding of Elmo1 and Elmo2 to Dock2 in the same cells, but we were unable to resolve distinct bands because of their similar molecular weights (Fig. 4C). Together, these data show that Dock2 can interact with either Elmo1 or Elmo2, but that in normal lymphocytes, Elmo1-Dock2 complexes are the most prevalent. This difference may, in part, explain the failure of residual Elmo2 to fully compensate for loss of Elmo1 in T cell migration.

FIGURE 4.
FIGURE 4.

Dock2 interaction with Elmo1 and Elmo2. (A) IP of Jurkat lysates with control rabbit IgG or anti-Dock2 followed by IB with Abs indicated (to the left of each blot). IPs were divided equally and loaded in separate wells for blotting with either anti-Elmo1 or anti-Elmo2. (B) IP of lysates from WT mouse splenocytes (left panel) and primary human CD4+ T cells (right panel) as in (A). One representative experiment of three for (A) and (B) is shown. (C) 293T cells were transfected with vector, Dock2-Flag, Elmo1-Flag, or Elmo2-Flag plasmids, followed by lysis and IP with anti-Dock2 or control IgG. IPs and total lysates (input) were analyzed by IB with anti–Flag-HRP. Right panel, The level of Elmo1-Flag and Elmo2-Flag present in anti-Dock2 IPs was normalized to input levels, and the relative level of each in Dock2 IPs was calculated. The average of n = 3 ± SEM is shown (right panel). Relative molecular masses (kDa) are indicated to the right of blots. h.c. H chain of IgG. **p < 0.01.

C terminus of Elmo1 is essential for T cell migration

The earlier data suggested that Dock2 may preferentially associate with Elmo1 to promote T cell migration. To determine whether Elmo1 interaction with Dock2 is required for Dock2-dependent migration, we attempted to rescue defective migration of Elmo1−/− T cells by transient transfection using full-length Elmo1 (aa 1–727) or C-terminal truncation mutant of Elmo1 (Elmo1T629, aa 1–629). In accord with a previous study, we found that the C-terminal tail of Elmo1 is essential for interaction with Dock2 (Fig. 5A) (26). CD4+ splenic T cells from Elmo1−/− mice were then cotransfected with Elmo plasmids and a GFP reporter plasmid, and tested for transwell migration to CXCL12. The number of GFP+ transfected cells migrating to the lower chamber was quantified by flow cytometry and compared with the total number of GFP+ cells (Supplemental Fig. 2). Transfection with full-length Elmo1, but not Elmo1T629, restored migration of Elmo1−/− T cells to WT levels (Fig. 5B). In the same experiments, Elmo2 was unable to rescue migration of Elmo1−/− T cells (Fig. 5B). These results show that Elmo1 interaction with Dock2 is required for Dock2-dependent T cell migration.

FIGURE 5.
FIGURE 5.

Elmo1–Dock2 interaction is required for CD4+ T cell migration. (A) Lysates from 293T cells transfected with vector, Dock2-Flag, Elmo1-Flag, or mutant Elmo1T629-Flag were immunoprecipitated with anti-Dock2 and analyzed by IB with anti-Flag. One representative experiment of three is shown. Relative molecular masses (kDa) are indicated to the right. (B) Splenic CD4+ T cells from WT and Elmo1−/− mice were nucleofected with GFP plasmid along with either empty vector, Elmo1, Elmo1T629, or Elmo2 plasmids and assayed for transwell migration to CXCL12 (50 ng/ml). The number of GFP+ cells migrating to the lower chamber was determined by flow cytometry and normalized to the total number of GFP+ cells for each transfected population and used to calculate the percent migration for each transfected population (n = 3, ± SEM). *p < 0.05.

Elmo1 selectively regulates Dock2 levels

To gain mechanistic insight into the migration defect of Elmo1−/− T cells, we examined expression of Elmo1, Elmo2, and Dock2 levels in WT and Elmo1−/− lymphocytes. Surprisingly, we found that CD3+, CD4+, and B220+ lymphocytes from Elmo1−/− mice showed an ∼4-fold reduction in Dock2 levels compared with WT (Fig. 6A and 6B). Lymphocytes from all Elmo1−/− mice tested to date (>25) have shown a similar reduction in Dock2 levels compared with WT controls (data not shown). Interestingly, Elmo2 levels were increased ∼2-fold in Elmo1-deficient lymphocytes (Fig. 6A and 6B). Similar results were seen upon acute depletion of Elmo1 in human Jurkat cells by siRNA, further supporting the regulation of Dock2 and Elmo2 levels as a conserved function of Elmo1 in lymphocytes (Fig. 6C). To determine whether Elmo1 deficiency similarly affects Dock1 levels, we FACS-sorted F4/80hi resident peritoneal macrophages from WT and Elmo1−/− mice, and tested for Dock1 and Dock2 levels by IB. Similar to lymphocytes, Dock2 levels were significantly reduced in Elmo1−/− F4/80hi macrophages compared with WT, whereas the level of Dock1 in these macrophages was not significantly different (Fig. 6D). These results show that Elmo1 plays a nonredundant and specific role in regulating the level of endogenous Dock2 protein.

FIGURE 6.
FIGURE 6.

Elmo1 control of Dock2 levels in leukocytes. (A) Splenocytes from WT and Elmo1−/− mice were FACS sorted by anti-B220 and anti-CD3 staining, lysed, and analyzed by IB with the Abs indicated to the left. Bottom graph, Fold change in Dock2 and Elmo2 levels in Elmo1−/− cells relative to WT as determined by IB for n = 3 mice/genotype, ± SEM. (B) Splenic CD4+ T cells were analyzed by IB as in (A). Bottom graph, Average of n = 8 mice/genotype, ± SEM. (C) Jurkat cells transfected with a nontargeting control (Ctl) or Elmo1 targeting siRNA were analyzed by IB with indicated Abs. (D) F4/80hi resident peritoneal macrophages were FACS sorted from WT and Elmo1−/− mice, and analyzed by IB with indicated Abs. Right graph, Relative levels of Dock1 and Dock2 in F4/80hi RPM for n = 4 mice/genotype, ± SEM. (E and F) qRT-PCR analysis of splenic CD3+ (E) and CD4+ T cells (F) for n = 3–5 mice/genotype, ± SEM. *p < 0.05, **p < 0.01, ***p < 0.001.

Evidence for in vivo posttranslational regulation of Dock2 by Elmo1

Elmo1 has been shown to control Dock1 levels through negative regulation of Dock1 ubiquitination and proteasomal degradation in cell lines, but Dock2 ubiquitination has not been studied (19–21). To address this, we first measured Dock2 mRNA levels in WT and Elmo1−/− lymphocytes by qRT-PCR. As shown in Fig. 6E and 6F, steady-state levels of Dock2 mRNA were comparable between WT and Elmo1−/− splenic CD3+ and CD4+ T cells, although the latter showed a slight but significant increase in Dock2 mRNA. Likewise, levels of Elmo2 mRNA were not significantly different between WT and Elmo1−/− T cells (Fig. 6E and 6F).

We next examined in vivo ubiquitination of Dock2 in CD4+ T cells. Lysine 48 (K48) of ubiquitin serves as a substrate for the formation of covalently attached polyubiquitin chains to lysine residues of target proteins. K48-linked polyubiquitinated proteins are subsequently targeted to the 26S proteasome for degradation (34). Using an Ab specific for K48-linked polyubiquitin, we measured in vivo levels of ubiquitinated endogenous Dock2 in WT CD4+ T cells. Cells were first treated with the proteasome inhibitor MG132 to allow for accumulation of ubiquitinated proteins (Fig. 7A), followed by anti-Dock2 IP of CD4+ T cell lysates and IB with anti-K48. We observed one band corresponding to the m.w. of Dock2 (∼200 kDa) in the anti-Dock2 but not control IgG IPs of WT T cells before and after MG132 treatment (Fig. 7B). To determine whether the K48-ubiquitin band in Dock2 IPs is due to direct ubiquitination of Dock2 or association of Dock2 with ubiquitinated proteins, we heat-denatured Dock2 IPs from WT CD4+ T cells and subjected them to a second round of anti-Dock2 IP before anti–K48-ubiquitin IB. As shown in Fig. 7C, these denaturing conditions successfully disrupted Dock2-Elmo1 association but did not abrogate the K48-ubiquitin Dock2 band. These data show that Dock2 undergoes constitutive polyubiquitination in primary T cells.

FIGURE 7.
FIGURE 7.

Posttranslational regulation of Dock2 in T cells. (A) Splenic CD4+ T cells from WT and Elmo1−/− mice were treated with the proteasome inhibitor MG132 for 3 h, lysed, and analyzed by IB using an Ab that recognizes K48-linked lysines of polyubiquitin chains or β-actin. (B) Splenic CD4+ T cells from WT and Elmo1−/− mice were treated as in (A), followed by IP with anti-Dock2 or control rabbit IgG. IPs were analyzed by IB with anti-K48 ubiquitin (above) and anti-Dock2 (above). Arrows indicate position of Dock2 bands. Graph on the right shows relative levels of K48-Dock2 normalized to total Dock2 in IP. n = 2 mice/genotype, ± SEM. (C) WT splenic CD4+ T cells were treated 3 h with 10 μM MG132 followed by lysis and anti-Dock2 IP. Half of each IP was then denatured (5 min at 95°C) followed by a second round of anti-Dock2 IP before the denatured and nondenatured (“native”) fractions were analyzed by anti-K48 ubiquitin IB. (D) Splenic CD4+ T cells from WT and Elmo1−/− mice were treated with MG132 for 6 h and total cell lysates were analyzed by IB using the indicated Abs. Values below indicate the relative levels of Dock2 protein normalized to β-actin in each sample as determined by densitometry. Results are representative of three independent experiments. (E) 293T cells were transfected with indicated plasmids for 24 h followed by anti-Dock2 IP of lysates and IB with indicated Abs. Results are representative of three independent experiments. Relative molecular masses (kDa) are indicated to the right of blots.

We next addressed the role of Elmo1 in regulating Dock2 ubiquitination. Unlike WT, K48-ubiquitinated Dock2 was only detectable in Elmo1−/− CD4+ T cells after MG132 treatment (Fig. 7B). When normalized to the total amount of Dock2 present in each IP, the level of K48-ubiquitin Dock2 in Elmo1−/− cells was 3-fold higher than WT (Fig. 7B, graph). MG132 treatment of Elmo1−/− T cells partially restored total Dock2 levels (Fig. 7D), whereas treatment with bafilomycin, an inhibitor of lysosome-mediated proteolysis, had no effect on Dock2 levels in Elmo1−/− cells (data not shown). In time-course experiments, we found that Dock2 levels in Elmo1−/− cells peaked 6 h after MG132 treatment of Elmo1−/− T cells, and that neither extended incubation with MG132 (up to 24 h) nor activation of the T cells (with plate-bound anti-CD3/anti-CD28 or PMA/ionomycin) enhanced Dock2 levels above that seen at 6 h (data not shown). Finally, to determine whether Elmo1 inhibits Dock2 ubiquitination, we cotransfected 293T cells with Dock2 and HA-tagged ubiquitin along with Elmo1 or Elmo1T629 followed by Dock2 IP and anti-HA IB. Expression of Elmo1, but not Elmo1T629, markedly inhibited ubiquitination of Dock2 (Fig. 7E). Together, these data indicate that Elmo1 controls Dock2 protein levels, at least in part, through inhibition of Dock2 ubiquitination and proteasomal degradation.

Discussion

Elmo1 and Elmo2 are expressed in most human and mouse tissues including central and peripheral lymphoid organs (13, 35). Both have been shown to promote actin-dependent phagocytosis and cell migration in a wide range of immortalized mammalian cell lines and invertebrate models. This report is the first, to our knowledge, to examine the function of endogenous Elmo1 in primary lymphocytes. In this article, we show that Elmo1 is essential for normal CCR7- and CXCR4-dependent migration of primary T and B lymphocytes in vitro. Although Elmo1−/− lymphocytes expressed normal surface levels of these chemokine receptors and were able to adhere to chemokine-infused ICAM-1 substrates, Elmo1-deficient T cells failed to effectively polarize and migrate in response to CCR7 and CXCR4 ligands. Rac activation in response to such stimulation was abrogated in Elmo1−/− T cells, although AKT and ERK1/2 phosphorylation were similar to WT. The requirement for Elmo1 in Rac, but not PI3K activation downstream of CCR signaling, is strikingly similar to that seen in Dock2−/− T cells (1, 2). It is interesting to note that although Elmo1 and Dock2 are dispensable for PI3K activation downstream of CCR activation in lymphocytes, there are conditions where the Elmo-Dock module is important for Rac-dependent PI3K activation. Notably, recent findings from Fritsch et al. (36) show that the Elmo1-Dock1 complex is required for normal PI3K activation in fibroblasts stimulated with lysophosphatidic acid or sphingosine 1-phosphate (S1P). In this study, Elmo1 was found to interact with the Gβγ complex upon lysophosphatidic acid and S1P receptor activation, and to promote Rac-dependent activation of the p110β type I PI3K (36). In another study using breast cancer cell lines, Elmo1 interaction with the Gαi subunit of CXCR4 was shown to be necessary for Dock1-dependent Rac activation, although CXCR4-dependent AKT phosphorylation was not affected by Elmo-Dock depletion (18). Interestingly, although PI3K activation downstream of CXCR4 appears to be Elmo1 and Dock2 independent in lymphocytes, Dock2 is required for S1P receptor–mediated AKT phosphorylation in lymphocytes (6). Thus, although Elmo is clearly important for recruitment of Dock proteins to multiple G protein–coupled receptors (18, 23, 36), it appears that the specific mechanisms of Elmo–G protein–coupled receptor association can regulate the pathways, including PI3K, that are activated by Dock-Rac signaling. Nevertheless, the data presented in this article show that in lymphocytes, a principal function of Elmo1 in CCR signaling is the regulation of Dock2 levels.

Elmo binding has been shown to regulate Dock1 localization, relief of autoinhibition, and protein stability (18, 23). The relative contribution and/or integration of these distinct mechanisms in optimal Dock1 signaling remains unsettled and is likely dependent on the specific tissue and function being studied. Although it was previously shown that Elmo1 interaction inhibits Dock1 ubiquitination and proteasomal degradation, our data show for the first time, to our knowledge, that endogenous Dock2 undergoes K48-linked polyubiquitination in normal primary T cells, and that Elmo1 plays an important role in controlling overall Dock2 levels through regulation of Dock2 polyubiquitination. However, it is important to note that despite a residual pool of Dock2 in Elmo1−/− T cells, Rac activation and migration were almost completely impaired. This may indicate a threshold requirement for Dock2 levels in T cells, but may also reflect a requirement for Elmo1 in proper targeting and activation of Dock2 in response to chemokine stimulation. Indeed, Elmo1 was recently reported to be required for localization of Dock1 at the CXCR4 receptor in migrating human breast cancer cells (18). A better understanding of the molecular regulation of Dock2 ubiquitination by Elmo1, most notably the identity of lysines in Dock2 that are targeted for ubiquitination, the E3 ligase complex responsible for ubiquitination, and how Elmo1 binding controls Dock2 modifications will be necessary to distinguish these modes of Elmo1-dependent Dock regulation.

Despite the homology and presumed redundancy of Elmo1 and Elmo2, we present several lines of evidence that show Elmo1 is a key regulator of Dock2-dependent migration of primary lymphocytes (13, 15). Despite being upregulated 2-fold in Elmo1−/− T cells, Elmo2 failed to compensate for Elmo1 in maintaining normal migration and Dock2 levels in vitro. Certainly this outcome could be a consequence of inadequate expression of Elmo2. Based on qRT-PCR analyses, Elmo2 mRNA levels were 3-fold lower than Elmo1 in WT CD4+ T cells (data not shown), which is in agreement with published microarray analyses (35). However, Elmo2 mRNA levels in Elmo1−/− T cells were comparable with WT, indicating that mRNA levels may not accurately reflect relative levels of Elmo proteins. Moreover, transfection of Elmo1, but not Elmo2, rescued defective migration of Elmo1−/− T cells, further suggesting that inadequate Elmo2 expression was unlikely to fully account for the lack of compensation. We then focused on potential differences in Dock2 binding. Quite surprisingly, we found that endogenous Elmo1-Dock2, but not Elmo2-Dock2, complexes were present in primary mouse and human lymphocytes. In a direct comparison of Dock2 binding in 293T cells, Elmo1 showed a 2.5-fold greater ability to complex with Dock2 than did Elmo2. Our finding that Dock2, but not Dock1, levels were significantly reduced in Elmo1−/− peritoneal macrophages indicates that Elmo1 is an essential regulator of Dock2 levels, whereas Elmo1 and Elmo2 may function redundantly in regulating Dock1 levels. Along these lines, it is interesting that peripheral lymphocyte populations of Elmo1−/− mice appear normal up to 12 wk of age. This is in contrast with Dock2−/− mice, which display a dramatic reduction in peripheral T and B cell populations because of the inability of Dock2-deficient lymphocytes to properly home to peripheral lymphoid organs (1, 2, 6). Dock2−/− T cells show a near-complete loss of in vitro migration to CXCL12 and CCL21 even at high concentrations of ligand (1), whereas in our study, Elmo1−/− T cells showed some residual ability to migrate to these chemokines. Taken together, these data suggest that under homeostatic conditions, the residual pool of Dock2 protein in Elmo1−/− T cells is sufficient to allow for the normal accumulation of peripheral T cells. Whether this homeostatic migration occurs through Elmo2-dependent or Elmo-independent mechanisms remains to be determined. Moreover, the requirement for Elmo1 in regulating Dock2-dependent migration in nonlymphoid tissues or under nonhomeostatic conditions is currently being investigated. Thus, it appears that Elmo1 plays a preferential role in interacting with and regulating Dock2 levels, but that as yet unidentified compensatory mechanisms exist to allow for normal homeostatic migration in vivo.

Recruitment of activated lymphocytes is a major cause of tissue pathology associated with many disease states including asthma, allograft rejection, chronic inflammation, and autoimmunity. Chemokines play an integral role in lymphocyte trafficking, and thus targeting these signaling pathways holds promise for curbing lymphocyte-mediated tissue damage. However, complex expression patterns and ligand–receptor promiscuity have hampered attempts to directly target chemokines or their receptors (37). Dock2 inhibition is an attractive approach for controlling T cell migration in vivo for a number of reasons. First, Dock2 regulates in vivo lymphocyte migration downstream of multiple receptors (1, 2, 6). Second, Dock2 inhibition does not appear to directly affect survival of peripheral T cells (1). Finally, Dock2 expression is largely restricted to hematopoietic cells (5). For these reasons, targeted inhibition of Dock2 could be used to specifically and reversibly block migration of lymphocytes to a broad range of chemokines in tissues without broadly depleting the repertoire of Ag-specific lymphocytes. Toward this goal, Nishikimi and colleagues (38) recently reported the development of a small-molecule inhibitor of Dock2 that acts at the Rac exchange domain to block Rac activation and T cell migration. Unfortunately, this inhibitor was also found to strongly inhibit Dock1 function probably because of the high degree of conservation of the Rac exchange domains of Dock-A proteins (38). Because Dock1 is widely expressed and essential for life, any strategy to disrupt Dock2 must avoid Dock1 inhibition (5, 39). In this study, we describe important and previously unknown features of Elmo-Dock2 interaction in primary cells that can guide efforts to design more effective strategies to selectively disrupt Dock2 function and T cell migration.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Jim Miller and Edward Stites for reviewing the manuscript, Taeg Kim for helpful discussions, and Nathan Laniewski and the University of Rochester Medical Center Flow Cytometry Core for technical assistance.

Footnotes

  • This work was supported by the Ellison Medical Foundation (to M.R.E.) and National Institutes of Health Grants AI027767-24 (to M.R.E, through a Creative and Novel Ideas in HIV Research award from the University of Alabama – Birmingham Center for AIDS Research), HL087088 (to M.K.), and T32AI007285 (to P.S.M. and T.C.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    co-IP
    coimmunoprecipitation
    GEF
    guanine nucleotide exchange factor
    IB
    immunoblotting
    IP
    immunoprecipitation
    pen-strep
    penicillin-streptomycin
    qRT-PCR
    quantitative RT-PCR
    siRNA
    small interfering RNA
    S1P
    sphingosine 1-phosphate
    WT
    wild-type.

  • Received December 17, 2013.
  • Accepted April 10, 2014.

References

  1. ↵
    1. Fukui Y.,
    2. O. Hashimoto,
    3. T. Sanui,
    4. T. Oono,
    5. H. Koga,
    6. M. Abe,
    7. A. Inayoshi,
    8. M. Noda,
    9. M. Oike,
    10. T. Shirai,
    11. T. Sasazuki
    . 2001. Haematopoietic cell-specific CDM family protein DOCK2 is essential for lymphocyte migration. Nature 412: 826–831.
    OpenUrlCrossRefPubMed
  2. ↵
    1. Nombela-Arrieta C.,
    2. R. A. Lacalle,
    3. M. C. Montoya,
    4. Y. Kunisaki,
    5. D. Megías,
    6. M. Marqués,
    7. A. C. Carrera,
    8. S. Mañes,
    9. Y. Fukui,
    10. C. Martínez-A,
    11. J. V. Stein
    . 2004. Differential requirements for DOCK2 and phosphoinositide-3-kinase gamma during T and B lymphocyte homing. Immunity 21: 429–441.
    OpenUrlCrossRefPubMed
    1. Tybulewicz V. L. J.,
    2. R. B. Henderson
    . 2009. Rho family GTPases and their regulators in lymphocytes. Nat. Rev. Immunol. 9: 630–644.
    OpenUrlCrossRefPubMed
  4. ↵
    1. Faroudi M.,
    2. M. Hons,
    3. A. Zachacz,
    4. C. Dumont,
    5. R. Lyck,
    6. J. V. Stein,
    7. V. L. J. Tybulewicz
    . 2010. Critical roles for Rac GTPases in T-cell migration to and within lymph nodes. Blood 116: 5536–5547.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Nishihara H.,
    2. S. Kobayashi,
    3. Y. Hashimoto,
    4. F. Ohba,
    5. N. Mochizuki,
    6. T. Kurata,
    7. K. Nagashima,
    8. M. Matsuda
    . 1999. Non-adherent cell-specific expression of DOCK2, a member of the human CDM-family proteins. Biochim. Biophys. Acta 1452: 179–187.
    OpenUrlPubMed
  6. ↵
    1. Nombela-Arrieta C.,
    2. T. R. Mempel,
    3. S. F. Soriano,
    4. I. Mazo,
    5. M. P. Wymann,
    6. E. Hirsch,
    7. C. Martínez-A,
    8. Y. Fukui,
    9. U. H. von Andrian,
    10. J. V. Stein
    . 2007. A central role for DOCK2 during interstitial lymphocyte motility and sphingosine-1-phosphate-mediated egress. J. Exp. Med. 204: 497–510.
    OpenUrlAbstract/FREE Full Text
  7. ↵
    1. Jiang H.,
    2. F. Pan,
    3. L. M. Erickson,
    4. M.-S. Jang,
    5. T. Sanui,
    6. Y. Kunisaki,
    7. T. Sasazuki,
    8. M. Kobayashi,
    9. Y. Fukui
    . 2005. Deletion of DOCK2, a regulator of the actin cytoskeleton in lymphocytes, suppresses cardiac allograft rejection. J. Exp. Med. 202: 1121–1130.
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Nishikimi A.,
    2. M. Kukimoto-Niino,
    3. S. Yokoyama,
    4. Y. Fukui
    . 2013. Immune regulatory functions of DOCK family proteins in health and disease. Exp. Cell Res. 319: 2343–2349.
    OpenUrlCrossRefPubMed
  9. ↵
    1. Kunisaki Y.,
    2. A. Nishikimi,
    3. Y. Tanaka,
    4. R. Takii,
    5. M. Noda,
    6. A. Inayoshi,
    7. K.-I. Watanabe,
    8. F. Sanematsu,
    9. T. Sasazuki,
    10. T. Sasaki,
    11. Y. Fukui
    . 2006. DOCK2 is a Rac activator that regulates motility and polarity during neutrophil chemotaxis. J. Cell Biol. 174: 647–652.
    OpenUrlAbstract/FREE Full Text
    1. Kunisaki Y.,
    2. Y. Tanaka,
    3. T. Sanui,
    4. A. Inayoshi,
    5. M. Noda,
    6. T. Nakayama,
    7. M. Harada,
    8. M. Taniguchi,
    9. T. Sasazuki,
    10. Y. Fukui
    . 2006. DOCK2 is required in T cell precursors for development of Valpha14 NK T cells. J. Immunol. 176: 4640–4645.
    OpenUrlAbstract/FREE Full Text
    1. Gotoh K.,
    2. Y. Tanaka,
    3. A. Nishikimi,
    4. A. Inayoshi,
    5. M. Enjoji,
    6. R. Takayanagi,
    7. T. Sasazuki,
    8. Y. Fukui
    . 2008. Differential requirement for DOCK2 in migration of plasmacytoid dendritic cells versus myeloid dendritic cells. Blood 111: 2973–2976.
    OpenUrlAbstract/FREE Full Text
  12. ↵
    1. Ippagunta S. K.,
    2. R. K. S. Malireddi,
    3. P. J. Shaw,
    4. G. A. Neale,
    5. L. Vande Walle,
    6. D. R. Green,
    7. Y. Fukui,
    8. M. Lamkanfi,
    9. T.-D. Kanneganti
    . 2011. The inflammasome adaptor ASC regulates the function of adaptive immune cells by controlling Dock2-mediated Rac activation and actin polymerization. Nat. Immunol. 12: 1010–1016.
    OpenUrlCrossRefPubMed
  13. ↵
    1. Gumienny T. L.,
    2. E. Brugnera,
    3. A. C. Tosello-Trampont,
    4. J. M. Kinchen,
    5. L. B. Haney,
    6. K. Nishiwaki,
    7. S. F. Walk,
    8. M. E. Nemergut,
    9. I. G. Macara,
    10. R. Francis,
    11. et al
    . 2001. CED-12/ELMO, a novel member of the CrkII/Dock180/Rac pathway, is required for phagocytosis and cell migration. Cell 107: 27–41.
    OpenUrlCrossRefPubMed
  14. ↵
    1. Côté J.-F.,
    2. K. Vuori
    . 2007. GEF what? Dock180 and related proteins help Rac to polarize cells in new ways. Trends Cell Biol. 17: 383–393.
    OpenUrlCrossRefPubMed
  15. ↵
    1. Brugnera E.,
    2. L. Haney,
    3. C. Grimsley,
    4. M. Lu,
    5. S. F. Walk,
    6. A. C. Tosello-Trampont,
    7. I. G. Macara,
    8. H. Madhani,
    9. G. R. Fink,
    10. K. S. Ravichandran
    . 2002. Unconventional Rac-GEF activity is mediated through the Dock180-ELMO complex. Nat. Cell Biol. 4: 574–582.
    OpenUrlCrossRefPubMed
    1. Lu M.,
    2. J. M. Kinchen,
    3. K. L. Rossman,
    4. C. Grimsley,
    5. M. Hall,
    6. J. Sondek,
    7. M. O. Hengartner,
    8. V. Yajnik,
    9. K. S. Ravichandran
    . 2005. A Steric-inhibition model for regulation of nucleotide exchange via the Dock180 family of GEFs. Curr. Biol. 15: 371–377.
    OpenUrlCrossRefPubMed
    1. deBakker C. D.,
    2. L. B. Haney,
    3. J. M. Kinchen,
    4. C. Grimsley,
    5. M. Lu,
    6. D. Klingele,
    7. P. K. Hsu,
    8. B. K. Chou,
    9. L. C. Cheng,
    10. A. Blangy,
    11. et al
    . 2004. Phagocytosis of apoptotic cells is regulated by a UNC-73/TRIO-MIG-2/RhoG signaling module and armadillo repeats of CED-12/ELMO. Curr. Biol. 14: 2208–2216.
    OpenUrlCrossRefPubMed
  18. ↵
    1. Li H.,
    2. L. Yang,
    3. H. Fu,
    4. J. Yan,
    5. Y. Wang,
    6. H. Guo,
    7. X. Hao,
    8. X. Xu,
    9. T. Jin,
    10. N. Zhang
    . 2013. Association between Gαi2 and ELMO1/Dock180 connects chemokine signalling with Rac activation and metastasis. Nat. Commun. 4: 1706.
    OpenUrlCrossRefPubMed
  19. ↵
    1. Makino Y.,
    2. M. Tsuda,
    3. S. Ichihara,
    4. T. Watanabe,
    5. M. Sakai,
    6. H. Sawa,
    7. K. Nagashima,
    8. S. Hatakeyama,
    9. S. Tanaka
    . 2006. Elmo1 inhibits ubiquitylation of Dock180. J. Cell Sci. 119: 923–932.
    OpenUrlAbstract/FREE Full Text
    1. Sanematsu F.,
    2. M. Hirashima,
    3. M. Laurin,
    4. R. Takii,
    5. A. Nishikimi,
    6. K. Kitajima,
    7. G. Ding,
    8. M. Noda,
    9. Y. Murata,
    10. Y. Tanaka,
    11. et al
    . 2010. DOCK180 is a Rac activator that regulates cardiovascular development by acting downstream of CXCR4. Circ. Res. 107: 1102–1105.
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Jarzynka M. J.,
    2. B. Hu,
    3. K. M. Hui,
    4. I. Bar-Joseph,
    5. W. Gu,
    6. T. Hirose,
    7. L. B. Haney,
    8. K. S. Ravichandran,
    9. R. Nishikawa,
    10. S. Y. Cheng
    . 2007. ELMO1 and Dock180, a bipartite Rac1 guanine nucleotide exchange factor, promote human glioma cell invasion. Cancer Res. 67: 7203–7211.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Grimsley C. M.,
    2. J. M. Kinchen,
    3. A. C. Tosello-Trampont,
    4. E. Brugnera,
    5. L. B. Haney,
    6. M. Lu,
    7. Q. Chen,
    8. D. Klingele,
    9. M. O. Hengartner,
    10. K. S. Ravichandran
    . 2004. Dock180 and ELMO1 proteins cooperate to promote evolutionarily conserved Rac-dependent cell migration. J. Biol. Chem. 279: 6087–6097.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Park D.,
    2. A.-C. Tosello-Trampont,
    3. M. R. Elliott,
    4. M. Lu,
    5. L. B. Haney,
    6. Z. Ma,
    7. A. L. Klibanov,
    8. J. W. Mandell,
    9. K. S. Ravichandran
    . 2007. BAI1 is an engulfment receptor for apoptotic cells upstream of the ELMO/Dock180/Rac module. Nature 450: 430–434.
    OpenUrlCrossRefPubMed
    1. Van Goethem E.,
    2. E. A. Silva,
    3. H. Xiao,
    4. N. C. Franc
    . 2012. The Drosophila TRPP cation channel, PKD2 and Dmel/Ced-12 act in genetically distinct pathways during apoptotic cell clearance. PLoS ONE 7: e31488.
    OpenUrlCrossRefPubMed
  25. ↵
    van Ham, T. J., D. Kokel, and R. T. Peterson. 2012. Apoptotic cells are cleared by directional migration and elmo1-dependent macrophage engulfment. Curr. Biol. 22: 830–836.
  26. ↵
    1. Sanui T.,
    2. A. Inayoshi,
    3. M. Noda,
    4. E. Iwata,
    5. J. V. Stein,
    6. T. Sasazuki,
    7. Y. Fukui
    . 2003. DOCK2 regulates Rac activation and cytoskeletal reorganization through interaction with ELMO1. Blood 102: 2948–2950.
    OpenUrlAbstract/FREE Full Text
  27. ↵
    1. Elliott M. R.,
    2. S. Zheng,
    3. D. Park,
    4. R. I. Woodson,
    5. M. A. Reardon,
    6. I. J. Juncadella,
    7. J. M. Kinchen,
    8. J. Zhang,
    9. J. J. Lysiak,
    10. K. S. Ravichandran
    . 2010. Unexpected requirement for ELMO1 in clearance of apoptotic germ cells in vivo. Nature 467: 333–337.
    OpenUrlCrossRefPubMed
  28. ↵
    1. Lu Z.,
    2. M. R. Elliott,
    3. Y. Chen,
    4. J. T. Walsh,
    5. A. L. Klibanov,
    6. K. S. Ravichandran,
    7. J. Kipnis
    . 2011. Phagocytic activity of neuronal progenitors regulates adult neurogenesis. Nat. Cell Biol. 13: 1076–1083.
    OpenUrlCrossRefPubMed
  29. ↵
    1. Lu M.,
    2. J. M. Kinchen,
    3. K. L. Rossman,
    4. C. Grimsley,
    5. C. deBakker,
    6. E. Brugnera,
    7. A.-C. Tosello-Trampont,
    8. L. B. Haney,
    9. D. Klingele,
    10. J. Sondek,
    11. et al
    . 2004. PH domain of ELMO functions in trans to regulate Rac activation via Dock180. Nat. Struct. Mol. Biol. 11: 756–762.
    OpenUrlCrossRefPubMed
  30. ↵
    1. Komander D.,
    2. M. Patel,
    3. M. Laurin,
    4. N. Fradet,
    5. A. Pelletier,
    6. D. Barford,
    7. J.-F. Côté
    . 2008. An alpha-helical extension of the ELMO1 pleckstrin homology domain mediates direct interaction to DOCK180 and is critical in Rac signaling. Mol. Biol. Cell 19: 4837–4851.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    Hanawa-Suetsugu, K., M. Kukimoto-Niino, C. Mishima-Tsumagari, R. Akasaka, N. Ohsawa, S.-I. Sekine, T. Ito, N. Tochio, S. Koshiba, T. Kigawa, et al. 2012. Structural basis for mutual relief of the Rac guanine nucleotide exchange factor DOCK2 and its partner ELMO1 from their autoinhibited forms. Proc. Natl. Acad. Sci. U.S.A. 109: 3305–3310.
  32. ↵
    1. Elliott M. R.,
    2. F. B. Chekeni,
    3. P. C. Trampont,
    4. E. R. Lazarowski,
    5. A. Kadl,
    6. S. F. Walk,
    7. D. Park,
    8. R. I. Woodson,
    9. M. Ostankovich,
    10. P. Sharma,
    11. et al
    . 2009. Nucleotides released by apoptotic cells act as a find-me signal to promote phagocytic clearance. Nature 461: 282–286.
    OpenUrlCrossRefPubMed
  33. ↵
    1. Treier M.,
    2. L. M. Staszewski,
    3. D. Bohmann
    . 1994. Ubiquitin-dependent c-Jun degradation in vivo is mediated by the delta domain. Cell 78: 787–798.
    OpenUrlCrossRefPubMed
  34. ↵
    1. Finley D.
    2009. Recognition and processing of ubiquitin-protein conjugates by the proteasome. Annu. Rev. Biochem. 78: 477–513.
    OpenUrlCrossRefPubMed
  35. ↵
    1. Heng T. S. P.,
    2. M. W. Painter,
    3. Immunological Genome Project Consortium
    . 2008. The Immunological Genome Project: networks of gene expression in immune cells. Nat. Immunol. 9: 1091–1094.
    OpenUrlCrossRefPubMed
  36. ↵
    1. Fritsch R.,
    2. I. de Krijger,
    3. K. Fritsch,
    4. R. George,
    5. B. Reason,
    6. M. S. Kumar,
    7. M. Diefenbacher,
    8. G. Stamp,
    9. J. Downward
    . 2013. RAS and RHO families of GTPases directly regulate distinct phosphoinositide 3-kinase isoforms. Cell 153: 1050–1063.
    OpenUrlCrossRefPubMed
  37. ↵
    1. Schall T. J.,
    2. A. E. I. Proudfoot
    . 2011. Overcoming hurdles in developing successful drugs targeting chemokine receptors. Nat. Rev. Immunol. 11: 355–363.
    OpenUrlCrossRefPubMed
  38. ↵
    1. Nishikimi A.,
    2. T. Uruno,
    3. X. Duan,
    4. Q. Cao,
    5. Y. Okamura,
    6. T. Saitoh,
    7. N. Saito,
    8. S. Sakaoka,
    9. Y. Du,
    10. A. Suenaga,
    11. et al
    . 2012. Blockade of inflammatory responses by a small-molecule inhibitor of the Rac activator DOCK2. Chem. Biol. 19: 488–497.
    OpenUrlCrossRefPubMed
  39. ↵
    1. Laurin M.,
    2. N. Fradet,
    3. A. Blangy,
    4. A. Hall,
    5. K. Vuori,
    6. J.-F. Côté
    . 2008. The atypical Rac activator Dock180 (Dock1) regulates myoblast fusion in vivo. Proc. Natl. Acad. Sci. USA 105: 15446–15451.
    OpenUrlAbstract/FREE Full Text
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