Phosphatidylinositol-4-kinase IIα licenses phagosomes for TLR4 signaling and MHC-II presentation in dendritic cells

Significance

Dendritic cells (DCs) play a key role at the interface between innate and adaptive immunity. DCs continuously sample their microenvironment, respond to microbial cues by signaling through pattern-recognition receptors such as Toll-like receptors (TLRs), and present bacterial antigens to adaptive immune cells. We show that the lipid kinase phosphatidylinositol-4-kinase IIα (PI4KIIα) is required to generate a phosphatidylinositol-4-phosphate pool on DC phagosomes that allows binding of the TLR sorting adaptor TIRAP and promotes TLR4 phagosomal signaling to proinflammatory cytokine production, phagosomal membrane tubule formation, and presentation of phagocytosed antigens. PI4KIIα therefore ensures phagosomal identity and autonomous signaling to initiate antimicrobial immune responses in DCs.

Abstract

Toll-like receptor (TLR) recruitment to phagosomes in dendritic cells (DCs) and downstream TLR signaling are essential to initiate antimicrobial immune responses. However, the mechanisms underlying TLR localization to phagosomes are poorly characterized. We show herein that phosphatidylinositol-4-kinase IIα (PI4KIIα) plays a key role in initiating phagosomal TLR4 responses in murine DCs by generating a phosphatidylinositol-4-phosphate (PtdIns4P) platform conducive to the binding of the TLR sorting adaptor Toll-IL1 receptor (TIR) domain-containing adaptor protein (TIRAP). PI4KIIα is recruited to maturing lipopolysaccharide (LPS)-containing phagosomes in an adaptor protein-3 (AP-3)-dependent manner, and both PI4KIIα and PtdIns4P are detected on phagosomal membrane tubules. Knockdown of PI4KIIα—but not the related PI4KIIβ—impairs TIRAP and TLR4 localization to phagosomes, reduces proinflammatory cytokine secretion, abolishes phagosomal tubule formation, and impairs major histocompatibility complex II (MHC-II) presentation. Phagosomal TLR responses in PI4KIIα-deficient DCs are restored by reexpression of wild-type PI4KIIα, but not of variants lacking kinase activity or AP-3 binding. Our data indicate that PI4KIIα is an essential regulator of phagosomal TLR signaling in DCs by ensuring optimal TIRAP recruitment to phagosomes.

Signaling by pattern-recognition receptors, such as Toll-like receptors (TLRs), is essential to initiate immune responses (1). In addition, TLR signaling is compartmentalized to allow discrimination between myriad self and foreign stimuli that may pose different levels of threat (2). One of the cellular compartments where TLR signaling is particularly important is the phagosome, a lysosome-related organelle formed in phagocytes such as dendritic cells (DCs) upon the capture of a particulate target, such as a bacterium (3, 4). Particularly in DCs, which serve as the main interface between innate and adaptive immunity, phagosomes become autonomous TLR-sensing and -signaling platforms that contain all of the machinery required to process the captured material, load resulting peptides into major histocompatibility complex (MHC) molecules, and form phagosomal tubules (phagotubules) that favor antigen presentation to T cells (5–7). In addition, phagosomes extend proinflammatory responses initiated by plasma-membrane TLRs by the acquisition of an additional pool of intracellular TLRs. This second wave of TLR signaling from phagosomes is focused on a single potentially harmful particulate entity. It is therefore essential to keep phagosomal identity different from the plasma membrane, where sensing reflects a broader spectrum of stimuli (8), and also from endosomes, which can bear soluble cargo that may be less harmful. In addition, preserving phagosomal identity and autonomy is essential in DCs for the initiation of adaptive immune responses.

While TLR signaling pathways have been extensively studied, and TLR4 is recruited to phagosomes from a recycling endosome pool (9), less is known about how TLR localization to phagosomes in DCs is regulated to optimize the immune response. Our current understanding of TLR recruitment to phagosomes in DCs was largely shaped by our analyses of a mouse model of Hermansky–Pudlak syndrome type 2 (HPS2) (10). HPS2 is caused by inactivating mutations in the β3A subunit of adaptor protein-3 (AP-3), an adaptor protein that binds to cytoplasmic signals in transmembrane proteins and mediates their trafficking from early endosomes to lysosomes or lysosome-related organelles (11). HPS2 is characterized by immunodeficiency, among other symptoms (4, 12, 13). While the role of AP-3 in T cell and plasmacytoid DC function (14–16) may explain recurrent viral infections in HPS2, defects in lipid antigen presentation and granulopoiesis (17–19), together with our observations that AP-3 is required for optimal secretion of proinflammatory cytokines, inflammasome activity, and MHC-II presentation of phagocytosed antigen to T cells (10, 20), may explain defective antibacterial immunity in HPS2. The impaired immune responses in AP-3–deficient conventional DCs are, at least in part, due to reduced TLR4 recruitment from an endosomal pool or reduced retention of TLR4 and other TLRs on maturing phagosomes (10). However, it is not known whether AP-3 supports TLR4 localization to phagosomes by direct binding and incorporation into phagosome-bound vesicles or by an indirect mechanism.

We and others showed that TLR4 signaling to a set of downstream immune responses in DCs requires the engagement of the TLR signaling adaptor MyD88 (7, 21). MyD88 binding to TLR4 is facilitated by the TLR sorting adaptor Toll-IL1 receptor (TIR) domain-containing adaptor protein (TIRAP; also known as MAL) (22), which also binds TLR4 to form the multimolecular signaling platform known as the MyDdosome (23). Therefore, TLR4 requires TIRAP to dictate the cellular location of MyD88-dependent signaling. Importantly, TIRAP binding to cellular membranes depends on polyphosphorylated inositide-enriched (or phosphoinositide-enriched) domains, highlighting the importance of phosphoinositides as essential mediators of TLR4 positioning at sites of future signal transduction (24, 25).

Phosphoinositides play an essential role in cellular membrane physiology by regulating membrane curvature, protein recruitment, and vesicular trafficking and, therefore, defining membrane identity (26–29). The type II lipid kinases phosphatidylinositol-4-kinase IIα (PI4KIIα) and IIβ (PI4KIIβ) (29, 30) are distinctly encoded enzymes that phosphorylate phosphatidylinositol (PtdIns) at position 4, generating PtdIns4P at the plasma membrane, Golgi, trans-Golgi network (TGN), and endolysosomes (30). In particular, PI4KIIα is abundant on membranes of the endolysosomal network (31, 32), notably in a vesicular pool rich in AP-3 (33). Importantly, PI4KIIα binds AP-3 directly via a classical dileucine-based sorting signal and is thereby delivered to lysosomes and lysosome-related organelles (34). In macrophages, PI4KIIα is required for TIR-domain-containing adapter-inducing interferon-β (TRIF)-dependent signaling in response to soluble stimuli (35), and both PI4KIIα and its product PtdIns4P are also required for phagosome maturation and resolution (36–38). However, a possible link between PI4KIIα and phagosomal TLR signaling has not been investigated.

Considering the connection between PI4KIIα and AP-3, and the requirement for phosphoinositides in TIRAP/MyD88-dependent TLR4 signaling, we investigated whether phagosomal TLR4 signaling in mouse DCs is dependent on the lipid kinase PI4KIIα and its product PtdIns4P. We show herein that PI4KIIα, but not the related PI4KIIβ, is required for the production of PtdIns4P on phagosomes in DCs, for optimal TIRAP binding to phagosomes and for downstream TLR4 responses. PI4KIIα recruitment to phagosomes is, in turn, dependent on AP-3 function, which may, therefore, at least partly explain the defective antibacterial responses in AP-3–deficient mouse DCs and HPS2 patients. Finally, we show that PI4KIIα plays a major role in DC phagosomes that reflects DC specialization in antigen presentation and ensures the preservation of phagosome identity and autonomous signaling.

Results

PI4KIIα Is Recruited to DC Phagosomes in an AP-3–Dependent Manner.

AP-3 recognizes accessible cytosolic targeting sequences on transmembrane proteins and mediates their trafficking to lysosomes or lysosome-related organelles. The consensus sequences YXXϕ or [DE]XXXL[LI] are recognized by the μ3 and σ3–δ subunits pf AP-3, respectively (39, 40). TLR4 bears a YDAF sequence in the cytoplasmic TIR domain. However, the TLR4 crystal structure suggests that the tyrosine is not accessible for intermolecular interactions (41). Consistent with this, we could not detect direct binding of the TLR cytoplasmic domain to the AP-3 μ3 subunit by yeast two-hybrid analysis (SI Appendix, Fig. S1) under the same conditions that allowed binding of the cytoplasmic tail of human TGN38 to the μ subunits of AP-1, -2, and -3, as expected (42, 43) (SI Appendix, Fig. S1). Although these data do not exclude a direct interaction through other subunits, the absence of the most likely μ3–TLR4 interaction led us to investigate whether additional effectors might favor AP-3–dependent TLR4 recruitment to phagosomes in DCs.

TLR4 signaling complexes are found on lipid microdomains enriched in phosphoinositides. Because PtdIns4P and PI4KIIα are present on maturing phagosomes in macrophages (36, 37), and AP-3 directly interacts with PI4KIIα in neurons and other cell types (33, 34), we investigated the role of PI4KIIα and its product PtdIns4P in TLR4 recruitment to DC phagosomes. We first asked if PI4KIIα and PtdIns4P were present on phagosomes in DCs and whether PI4KIIα recruitment was dependent on AP-3. Bone-marrow-derived DCs (BMDCs) from wild-type (WT) and Ap3b^pe/pe mice that lack AP-3 in nonneuronal cell types (AP-3^−/−) were pulsed with magnetic beads coated with the TLR4 ligand lipopolysaccharide (LPS) and chased over time after phagocytosis. Phagosomes were then isolated and analyzed by immunoblotting for PI4KIIα (Fig. 1 A and B). Phagosomes from WT DCs showed an increased recruitment of endogenous PI4KIIα over 2 h, concomitant with the acquisition of markers of phagosome maturation, such as the late endosomal/lysosomal proteins LAMP-1 and Rab7 (SI Appendix, Fig. S2 A and B), and of proteolytic activity as assessed by flow-cytometry analysis of ovalbumin (OVA) degradation on phagocytosed beads (SI Appendix, Fig. S2C). In contrast, whereas cellular expression of PI4KIIα in WT and AP-3^−/− DCs was similar, PI4KIIα recruitment to phagosomes in AP-3^−/− DCs was significantly reduced (Fig. 1 A and B). PI4KIIα recruitment to mature phagosomes, measured 2 h after engulfment, was also impaired in AP-3^−/− tissue-resident DCs isolated from spleen compared to WT splenic DCs (SI Appendix, Fig. S3), as determined both on isolated phagosomes (SI Appendix, Fig. S3 A and B) or in cells pulsed with LPS-coated polystyrene beads (88 ± 3% in WT DCs vs. 6 ± 2% in AP-3^−/− DCs; SI Appendix, Fig. S3 C and D).

Fig. 1.

PI4KIIα is recruited to BMDC phagosomes in an AP-3–dependent manner. WT and AP-3^−/− BMDCs that had been nontransduced (A and B) or transduced with retroviruses encoding PI4KIIα-GFP (C–G) were pulsed with LPS-coated magnetic beads (A and B), LPS-coated polystyrene beads (C and D), or LPS/OVA-TxR–coated polystyrene beads (E–G) and chased as indicated. (A) Purified phagosomes (Left) or whole-cell lysates (WCL) (Right) were analyzed by SDS/PAGE 10% and immunoblotting for endogenous PI4KIIα and actin. Shown are the relevant bands for each blot. (B) Quantification of band intensities for phagosomal PI4KIIα from three independent experiments, showing fold change relative to WT time 0 and normalized to actin (mean ± SD). (C and D) Phagosomes from nontransduced or PI4KIIα–GFP-expressing WT and AP-3^−/− BMDCs were isolated and analyzed by flow cytometry. GFP-positive cells were not previously sorted. (C) Shown are histogram plots of a representative experiment with the percentages of gated GFP-positive phagosomes (signal over phagosomes from nontransduced cells) indicated. Solid black lines, WT; solid orange lines, AP-3^−/−; dashed lines, nontransduced controls. (D) Data (mean ± SD) from three independent experiments performed in duplicate were normalized to the percent of transduced BMDCs (SI Appendix, Fig. S4A). (E–G) WT and AP-3^−/− BMDCs expressing PI4KIIα-GFP were analyzed by live cell imaging. (E) Representative images at indicated times after phagocytosis. Dotted white lines, cell outlines; arrowheads, PI4KIIα–GFP-positive phagosomes; asterisks, PI4KIIα–GFP-negative phagosomes; arrows, phagotubules. (Scale bar, 9 μm.) (F) Data from three independent experiments, 20 cells per experiment, are presented as percent of PI4KIIα–GFP-positive phagosomes per cell. Black dots, WT; gray dots, AP-3^−/−; solid red lines, means. (G) Quantification was performed as shown on representative images, drawing a line across the phagosomes (Left) and analyzing the line plot with ImageJ (Right; +, positive signal). Phagosomes containing GFP-positive puncta, even partially in the total phagosomal membrane surface (mostly in the case of AP-3^−/− BMDCs) were considered positive. **P < 0.01; ***P < 0.001.

To confirm these results, we expressed PI4KIIα–green fluorescent protein (GFP) in DCs by recombinant retroviral transduction of bone marrow (transduction efficiency was similar in WT and AP-3^−/− DCs; SI Appendix, Fig. S4A), pulsed differentiated DCs with LPS-coated polystyrene beads, and tested for PI4KIIα–GFP recruitment to DC phagosomes by flow cytometry of isolated phagosomes (Fig. 1 C and D and SI Appendix, Fig. S4B) or by live cell imaging (Fig. 1 E–G). Like endogenous PI4KIIα, PI4KIIα–GFP was increasingly recruited to phagosomes over 2 h after phagocytosis in WT DCs (Fig. 1 C–F), concomitant with LAMP-1 (SI Appendix, Fig. S4C). Additionally, endogenous LAMP-1 and expressed PI4KIIα–GFP were also detected on phagosomes by immunofluorescence microscopy 60 and 120 min after pulsing WT DCs with LPS-coated polystyrene beads (SI Appendix, Fig. S4D). In contrast to WT DCs, PI4KIIα–GFP recruitment to phagosomes in AP-3^−/− DCs was significantly impaired (Fig. 1 C and D); after normalizing to the percent of PI4KIIα-GFP–positive DCs (SI Appendix, Fig. S4A; note that GFP-positive cells were not previously sorted), PI4KIIα-GFP was recruited to 90% of phagosomes in WT DCs, but only 20% in AP-3^−/− DCs at 120 min (Fig. 1D). Consistent with these results, PI4KIIα–GFP recruitment to phagosomes containing polystyrene beads coated with LPS and Texas Red-conjugated OVA (LPS/OVA-TxR) was significantly impaired in AP-3^−/− DCs, as observed by live cell imaging on transduced DCs (97 ± 3% in WT DCs vs. 12 ± 3% in AP-3^−/− DCs after 120 min; Fig. 1 E–G and Movies S1 and S2; note the absence of a continuous fluorescent pattern in AP-3^−/− DC phagosomes).

Together, these data indicate that AP-3 is required for optimal recruitment of PI4KIIα to phagosomes in both BMDCs and splenic DCs. Noteworthy, PI4KIIα–GFP was also recruited to OVA-containing phagotubules in maturing phagosomes from WT DCs (arrows, Fig. 1E, time 120 min; Movies S3 and S4).

PI4KIIα Is Required for the Accumulation of PtdIns4P on Early and Maturing Phagosomes and Phagotubules.

To test whether PtdIns4P was produced on DC phagosomes concomitantly with PI4KIIα recruitment, we expressed GFP-tagged P4M-SidMx2 (P4Mx2)—a probe for PtdIns4P derived from the SidM domain of Legionella pneumophila (44)—in DCs and analyzed cells over time after phagocytosis of LPS/OVA-TxR beads using live imaging. In addition, we assessed the contribution of PI4KIIα and the related PI4KIIβ—which does not bind AP-3—to the production of PtdIns4P on phagosomes by transducing bone marrow precursors with short hairpin RNA (shRNA) targeted to each PI4KII isoform or a nontarget control shRNA (SI Appendix, Fig. S5A and Fig. 2A). Knockdown was specific for each PI4KII isoform. PI4KIIα shRNA #2 was more effective than #1 and was used in subsequent experiments. Retroviral transduction did not significantly affect DC differentiation (SI Appendix, Fig. S5B), and phagocytic capacity was not affected by the protein knockdowns (SI Appendix, Fig. S5C). DC maturation in response to LPS was reduced by shRNA transduction, but comparable between the different shRNA treatments (SI Appendix, Fig. S5D). The GFP–P4Mx2 probe detected PtdIns4P on the plasma membrane and in an intracellular pool [as described in other cells (44)] (Fig. 2A). In addition, after engulfment of LPS/OVA-TxR beads, PtdIns4P was found on nascent phagosomes in all DC types. In addition, PtdIns4P was present in early and maturing phagosomes (arrowheads) and in phagotubules (arrows) in shRNA control DCs (Fig. 2A). However, DCs knocked down for PI4KIIα showed significantly reduced accumulation of PtdIns4P on early and maturing phagosomes (98% in control DCs vs. 0% in PI4KIIα knockdown DCs between 30 and 120 min; Fig. 2B), while the plasma membrane signal was not affected (Fig. 2A). Conversely, DCs knocked down for PI4KIIβ showed reduced binding of GFP–P4Mx2 to the plasma membrane, while accumulation of PtdIns4P on phagosomes (99 ± 1% between 30 and 120 min) and phagotubules was unaffected (Fig. 2A). Moreover, and consistent with the reduced recruitment of PI4KIIα to phagosomes in AP-3^−/− DCs (Fig. 1), PtdIns4P on phagosomes was significantly reduced in AP-3–deficient DCs (2 ± 2% between 30 and 120 min; Fig. 2B). These data suggest that AP-3 and its cargo protein PI4KIIα are required for PtdIns4P formation on phagosomes, while PI4KIIβ primarily contributes to the plasma membrane PtdIns4P pool in DCs.

Fig. 2.

PI4KIIα is needed to accumulate PtdIns4P on early and maturing phagosomes. WT BMDCs were transduced with retroviruses encoding GFP–P4Mx2 and with lentiviruses encoding nontarget (control), PI4KIIβ or PI4KIIα shRNAs, and AP-3^−/− BMDCs were transduced only with retroviruses encoding GFP–P4Mx2. DCs were pulsed with LPS/OVA-TxR–coated beads, chased as indicated, and analyzed by live cell imaging. (A) Representative images. Dotted white lines, cell outlines; arrowheads, GFP–P4Mx2-positive phagosomes; asterisks, GFP–P4Mx2-negative phagosomes; arrows, phagotubules. (B) Data from three independent experiments, 20 cells per experiment, are presented as percent of GFP–P4Mx2-positive phagosomes per cell. Black dots, nontarget control shRNA; blue dots, PI4KIIβ shRNA; red dots, PI4KIIα shRNA; gray dots, AP-3^−/−; solid color lines, means. (Scale bar, 9 μm.) ***P < 0.001; n.s., not significant.

PI4KIIα Is Required for Optimal Phagosomal TLR4 Signaling and MHC-II Presentation of Phagocytosed Antigen in DCs.

We and others showed that TLR4 activation via its signaling adaptor MyD88 on phagosomes triggers a signaling cascade that leads to proinflammatory cytokine production, phagotubule formation, and MHC-II presentation of phagocytosed cargo in DCs (7, 10, 45). To test whether these responses require PI4KIIα, we probed for TLR4/MyD88 responses in DCs derived from bone marrow transduced with nontarget shRNA or shRNA to PI4KIIα, PI4KIIβ, or the SNARE protein Sec22b (46) as an additional control (SI Appendix, Fig. S5A). We first analyzed the production of the proinflammatory interleukin 6 (IL-6) by DCs pulsed with LPS-coated beads (relative to uncoated beads as a negative control) by enzyme-linked immunosorbent assay (ELISA) analysis of cell supernatants. LPS bead-induced IL-6 levels were significantly reduced in supernatants of PI4KIIα knockdown DCs compared to those from PI4KIIβ knockdown DCs or DCs transduced with control shRNAs (Fig. 3A). To test for phagotubule formation, we analyzed DCs by live cell imaging 2.5 h after exposure to LPS/OVA-TxR beads. Whereas PI4KIIβ knockdown DCs or DCs treated with control shRNAs exhibited robust phagotubule formation, PI4KIIα knockdown severely impaired the formation of phagotubules, as also observed in AP-3^−/− DCs (Fig. 3 B and C and Movies S5–S7; see also Figs. 1E and 2A, time 120 min). These data show that optimal phagosomal TLR-induced proinflammatory cytokine secretion and phagotubule formation require PI4KIIα. Noteworthy, like OVA-containing phagotubules (7), PI4KIIα-positive phagotubules were also positive for MHC-II (SI Appendix, Fig. S6).

Fig. 3.

PI4KIIα is required for optimal phagosomal TLR4 signaling. WT BMDCs were nontransduced (−) or transduced with lentiviruses encoding nontarget (control), PI4KIIβ, Sec22b, or either of two PI4KIIα shRNAs, and AP-3^−/− BMDCs were untransduced. DCs were pulsed with uncoated or LPS-coated polystyrene beads (A) or LPS/OVA-TxR–coated beads (B and C). (A) IL-6 released into the supernatants was measured by ELISA after a 3-h chase. Results represent mean ± SD of three experiments, each performed in triplicate. (B and C) BMDCs were analyzed by live cell imaging after a 2.5-h chase. (B) Arrows indicate phagotubules. (Scale bar, 9 μm.) (C) The percentage of BMDCs presenting phagotubules (tubules ≥ 1 μm emerging from phagosomes) in three independent experiments, 20 cells per experiment, is shown. ***P < 0.001. Lack of statistical significance is not indicated.

Finally, we assessed MHC-II presentation of phagocytosed antigen using beads coated with the Eα_52–68 peptide; subsequent presentation of this peptide by the MHC-II molecule I-A^b at the cell surface can be detected by the YAe antibody (47). DCs were pulsed with EαGFP-coated beads or with soluble EαGFP (to monitor presentation of soluble antigen) or Eα_52–68 peptide alone (a control that binds to surface I-A^b); all preparations contained LPS to stimulate TLR4. As expected from the reduced DC maturation observed in shRNA-transduced cells (SI Appendix, Fig. S5D), LPS-bead induced MHC-II expression on transduced DCs was reduced compared to nontransduced DCs, but similar between the shRNA treatments (except for Sec22b shRNA, which was reduced more and, thus, not included in these assays; SI Appendix, Fig. S7A). Of note, the LPS-bead-induced expression of MHC-II and of the costimulatory molecules CD40 and CD86 (SI Appendix, Fig. S7B) and phagosomal degradation capacity (decreased OVA labeling from phagocytosed OVA-beads; SI Appendix, Fig. S7C) did not differ between DCs treated with PI4KIIα and control shRNAs. As also observed for AP-3^−/− DCs (9), whereas formation of cell-surface Eα:I-A^b complexes 6 h following exposure to Eα_52–68 peptide or soluble EαGFP was similar among cells treated with either shRNA (Fig. 4 A and B), surface Eα:I-A^b complex levels after phagocytosis of EαGFP-coated beads were significantly reduced by expression of PI4KIIα shRNA relative to PI4KIIβ or nontarget control shRNAs (Fig. 4 C and D). To test if MHC-II presentation correlated with CD4+ T cell responses, DCs pulsed with the OVA-derived OVA_323–329 peptide, soluble OVA/LPS, or OVA/LPS-coated beads were cocultured with OVA_323–329-specific, I-A^b–restricted OT-II T cells. OT-II cell activation, measured by CD69 expression and IL-2 production, by DCs pulsed with peptide alone or soluble OVA was similar whether DCs expressed PI4KIIα, PI4KIIβ, or control shRNAs (Fig. 4 E–H). In contrast, OT-II cell activation by DCs pulsed with OVA/LPS beads was significantly reduced in PI4KIIα shRNA-expressing DCs relative to cells expressing other shRNAs (Fig. 4 I and J). These data indicate that PI4KIIα promotes MHC-II presentation of antigen following phagocytosis, but not endocytosis, and are consistent with our observations that PI4KIIα is required for PtdIns4P formation on phagosomes, but not on the plasma membrane (Fig. 2).

Fig. 4.

PI4KIIα is required for optimal MHC-II presentation of phagocytosed antigen. WT BMDCs transduced with lentiviruses encoding nontarget (control), PI4KIIβ, or PI4KIIα shRNAs were pulsed with Eα_52–68 peptide (A), soluble EαGFP fusion protein (B), EαGFP-coated beads (C and D), OVA_323–329 peptide (E and F), soluble OVA (G and H), or OVA:BSA-coated beads (I and J) and chased for 6 h. (A–D) Surface expression of Eα_52–68:I-A^b complexes was analyzed by flow cytometry using YAe antibody. (A–C) Shown are the percentages of CD11c⁺ BMDCs that were Eα_52–68:I-A^b+. (D) Shown is the YAe mean fluorescence intensity (MFI) normalized to MHC-II MFI (SI Appendix, Fig. S5D). (E–J) Pulsed BMDCs were fixed and cocultured with preactivated OT-II cells for 18 h. (E, G, and I) CD69 expression by OT-II cells was assessed by flow cytometry. Shown are the percentages of vb5⁺ T cells that expressed CD69. (F, H, and J) IL-2 production by OT-II cells was measured by ELISA on the coculture supernatants. Data in all panels represent mean ± SD of three experiments performed in duplicate. *P < 0.05; **P < 0.01; ***P < 0.001. Lack of significance is not indicated.

PI4KIIα Phagosomal Function Requires the AP-3–Sorting Motif and Kinase Activity.

In order to assess if PI4KIIα recruitment and consequent phagosomal TLR signaling required AP-3 binding and kinase activity, we tested whether PI4KIIα mutants could rescue these phenotypes in PI4KIIα knockdown cells. Bone marrow cells were transduced with retroviruses encoding either human WT PI4KIIα-GFP, the AP-3 sorting mutant (L61,62A—which does not bind AP-3), or the kinase-inactive mutant (D308A), all previously characterized (34), and subsequently transduced with lentiviruses encoding murine PI4KIIα shRNA; control cells were transduced with nontarget shRNA lentiviruses (Fig. 5A). Transduced DCs were then pulsed with LPS/OVA-TxR beads and analyzed by live cell imaging after 2.5 h. Phagosomal recruitment of both the AP-3 sorting mutant and the kinase-inactive mutant was significantly impaired compared to WT PI4KIIα–GFP (WT, 98 ± 1%; D308A, 24 ± 3%; L61,62A, 10 ± 2%; Fig. 5 B and C), consistent with previous reports for PI4KIIα–GFP recruitment to lysosomes and lysosome-related organelles in other cells (33, 34). Moreover, only WT PI4KIIα–GFP restored phagotubule formation in knockdown DCs and was itself recruited to phagosomal tubules (Fig. 5B). WT PI4KIIα–GFP, but not the kinase-inactive or AP-3 binding mutants, also restored control levels of proinflammatory cytokine production (Fig. 5D) and surface MHC-II presentation of phagocytosed Eα (Fig. 5E). Thus, both AP-3 binding and kinase activity are required for PI4KIIα recruitment to phagosomes and consequent phagosomal function in TLR4 signaling.

Fig. 5.

PI4KIIα function on phagosomes requires the AP-3 sorting motif and kinase activity. WT BMDCs that were nontransduced (−) or transduced first with retroviruses encoding GFP, human WT PI4KIIα–GFP, PI4KIIα(D308A)–GFP, or PI4KIIα(L61,62A)–GFP were then transduced with lentiviruses encoding mouse nontarget (ctrl) or PI4KIIα shRNAs. Cells were left untreated (A) or pulsed with LPS/OVA-TxR–coated beads (B and C), uncoated or LPS-coated polystyrene beads (D), or EαGFP-coated beads (E). (A) Whole-cell lysates were analyzed by SDS/PAGE and immunoblotting for PI4KIIα, GFP, and tubulin. Shown are the relevant bands for each blot. (B and C) BMDCs were analyzed by live cell imaging after a 2.5-h chase. (B) Representative images. Dotted white lines, cell outlines; arrowheads, PI4KIIα–GFP-positive phagosomes; asterisks, PI4KIIα–GFP-negative phagosomes; arrows, phagotubules. (Scale bar, 9 μm.) (C) Data from three independent experiments, 20 cells per experiment, are presented as percent of PI4KIIα–GFP-positive phagosomes per cell. Black dots, WT; purple dots, PI4KIIα(D308A)–GFP; tidal dots, PI4KIIα(L61,62A)–GFP; solid color lines, means. (D) IL-6 released into the supernatants after 3 h was measured by ELISA. (D, Left) Representative experiment performed in triplicate. (D, Right) IL-6 values from three independent experiments performed in triplicate are shown as percent of values for BMDCs treated with nontarget (ctrl) shRNA, as a representation of phenotypic rescue (mean ± SD). (E) Surface expression of Eα_52–68:I-A^b complexes was analyzed by flow cytometry using YAe antibody. (E, Left) Shown are the percentages of CD11c⁺ BMDCs that were Eα_52–68:I-A^b+ in a representative experiment. (E, Right) YAe values from two independent experiments performed in duplicate are shown as percent of values for BMDCs treated with nontarget (ctrl) shRNA, as a representation of phenotypic rescue (mean ± SD). (D and E) Significance relative to nontarget shRNA-treated WT control (Left) or PI4KIIα shRNA-treated DCs (−) (Right) is indicated. ***P < 0.001. Lack of significance is not indicated.

PI4KIIα Promotes TLR4 Accumulation on Phagosomes in DCs.

To test whether TLR4 localization to phagosomes required PI4KIIα, we pulsed DCs with LPS-coated beads and analyzed TLR4 presence on isolated phagosomes by immunoblotting and flow cytometry using two different antibodies. Immunoblotting showed that cellular TLR4 expression was similar in DCs treated with control, PI4KIIα, or PI4KIIβ shRNAs (Fig. 6 A, Left). In both assays, TLR4 increasingly accumulated on phagosomes over time after phagocytosis in control [as we had shown before (10)] and PI4KIIβ knockdown DCs, but not in cells knocked down for PI4KIIα (Fig. 6 A–D). This was true whether data were analyzed for total TLR4 content on phagosomes by immunoblotting (Fig. 6 A and C) or by percentage of phagosomes harboring TLR4 by flow cytometry (Fig. 6 B and D). These data indicate that efficient TLR4 localization to phagosomes requires PI4KIIα.

Fig. 6.

PI4KIIα promotes TLR4 recruitment to phagosomes in DCs. WT BMDCs transduced with lentiviruses encoding nontarget (control), PI4KIIβ, or PI4KIIα shRNAs were pulsed with LPS-coated magnetic beads (A and C) or LPS-coated polystyrene beads (B and D) and chased as indicated. (A) Purified phagosomes (Right) or whole-cell lysates (WCL) (Left) were analyzed by SDS/PAGE and immunoblotting for TLR4 and actin. Shown are the relevant bands for each blot. (C) Quantification of band intensities for phagosomal TLR4 from three independent experiments, showing fold change relative to each shRNA treatment at time 0 and normalized to actin (mean ± SD). (B and D) Phagosomes were purified and analyzed by flow cytometry using a fluorescein isothiocyanate-conjugated anti-TLR4 antibody. (B) Shown are histogram plots of a representative experiment with the percentages of gated TLR4-positive phagosomes indicated. Solid black lines, nontarget (control) shRNA; solid blue lines, PI4KIIβ shRNA; solid red lines, PI4KIIα shRNA; dashed lines, nontransduced controls. (D) Data from three independent experiments performed in duplicate are shown as fold change of percent of TLR4-positive phagosomes relative to each shRNA treatment at time 0 (mean ± SD). *P < 0.05; **P < 0.01; ***P < 0.001.

Sorting Adaptor TIRAP Recruitment to Phagosomes Requires PI4KIIα.

Formation of the MyDdosome complex and downstream TLR4 signaling is favored by TLR4 binding to its sorting adaptor TIRAP (23, 48). TIRAP contains an N-terminal lysine-rich polybasic motif that promiscuously binds to different phosphoinositide species, including PtdIns(4,5)P₂ and PtdIns4P (25). While PtdIns(4,5)P₂ mostly localizes to the plasma membrane and to nascent phagosomes, PtdIns4P—which is present on late endocytic compartments—was a strong candidate for TIRAP binding to maturing phagosomal membranes. To test whether TIRAP is recruited to phagosomes and whether recruitment requires PI4KIIα, we followed the kinetics of TIRAP–GFP recruitment to phagosomes by live cell imaging (Fig. 7) and flow cytometry on isolated phagosomes (Fig. 8) from TIRAP–GFP-transduced DCs (SI Appendix, Fig. S7 D, Upper; note that GFP-positive cells were not previously sorted). In cells pulsed with OVA-TxR beads, TIRAP–GFP was detected on the plasma membrane and on nascent phagosomes (arrowheads, Fig. 7 A and B), consistent with its ability to bind PtdIns(4,5)P₂ and similar to the localization of the PtdIns(4,5)P₂-sensing probe GFP–PH-PLCδ (pleckstrin homology domain of phospholipase Cδ; SI Appendix, Fig. S8 A and B). Recruitment of TIRAP–GFP and GFP–PH-PLCδ to the plasma membrane was modestly affected by knockdown of PI4KIIβ, but not PI4KIIα, whereas TIRAP–GFP (but not GFP–PH-PLCδ) was detected on many fewer phagosomes in PI4KIIα knockdown cells after the pulse (Fig. 7 A and B, time 0, and SI Appendix, Fig. S8 A and B). Over time, TIRAP–GFP was recruited to phagosomes in control and PI4KIIβ knockdown cells, as quantified both by fluorescence microscopy (Fig. 7B and Movie S8) and flow cytometry on isolated phagosomes (Fig. 8A), normalizing to the percentage of TIRAP–GFP-positive cells (Fig. 8C). TIRAP–GFP was also detected on phagotubules at 120 min (arrows, Fig. 7A and Movie S9). However, TIRAP recruitment to phagosomes was severely reduced in PI4KIIα knockdown cells between 30 and 120 min (on 90 ± 10% of phagosomes in control and PI4KIIβ knockdown cells vs. 1 ± 5% of phagosomes in PI4KIIα knockdown DCs; Fig. 7B and Movie S10), with no detectable phagosomal increase, as measured by flow cytometry on isolated phagosomes (Fig. 8 A and C). Consistent with the role of AP-3 in PI4KIIα recruitment to phagosomes, TIRAP recruitment to phagosomes was also impaired in AP-3–deficient DCs (on 1 ± 5% of phagosomes between 30 and 120 min; Fig. 7B). In contrast to these observations, phagosomal recruitment of the p40-phox domain containing, PX-TIR–GFP construct (SI Appendix, Fig. S7 D, Lower), the TIRAP variant that exclusively binds PtdIns3P (25)—a lipid enriched on early endosomes and early phagosomes (49)—was not affected by PI4KIIα knockdown or AP-3 deficiency (Fig. 8 B and D and SI Appendix, Fig. S8 C and D). PX-TIR–GFP was mainly detected on phagosomes between 30 and 60 min after the pulse (Fig. 8B and SI Appendix, Fig. S8 C and D), at times when the PtdIns4P probe GFP–P4Mx2 and TIRAP–GFP were also detected in control and PI4KIIβ knockdown DCs (Figs. 2 and 7). The observation that PI4KIIα knockdown or AP-3 knockout severely impairs TIRAP, but not PX-TIR, recruitment to phagosomes suggests that TIRAP preferentially binds PtdIns4P on DC phagosomes. Note that phagotubules are labeled by OVA-TxR, but not by PX-TIR-GFP (SI Appendix, Fig. S8C), suggesting that phagotubules, which emanate from mature phagosomes, do not contain PtdIns3P. These results show that PI4KIIα is necessary and sufficient for TIRAP recruitment to phagosomes and suggest that, in contrast to its recruitment to early endosomes, PtdIns3P is not sufficient to recruit TIRAP to early phagosomes in DCs.

Fig. 7.

Sorting adaptor TIRAP recruitment to phagosomes is severely impaired by PI4KIIα knockdown. WT BMDCs were transduced with retroviruses encoding TIRAP–GFP, and lentiviruses encoding nontarget (control), PI4KIIβ or PI4KIIα shRNAs, and AP-3^−/− BMDCs were transduced only with retroviruses encoding TIRAP–GFP. DCs were pulsed with LPS/OVA-TxR–coated beads, chased as indicated and analyzed by live cell imaging. (A) Representative images. Dotted white lines, cell outlines; arrowheads, TIRAP–GFP-positive phagosomes; asterisks, TIRAP–GFP-negative phagosomes; arrows, phagotubules. (B) Data from three independent experiments, 20 cells per experiment, are presented as percent of TIRAP–GFP-positive phagosomes per cell. Black dots, nontarget control shRNA; blue dots, PI4KIIβ shRNA; red dots, PI4KIIα shRNA; gray dots, AP-3^−/−; solid color lines, means. (Scale bar, 9 μm.) ***P < 0.001; n.s., not significant.

Fig. 8.

Impaired IL-6 production upon TIRAP knockdown is restored by expression and phagosomal recruitment of TIRAP–GFP, but not of the PtdIns3P-binding PX-TIR. (A–D) WT BMDCs transduced with retroviruses encoding TIRAP–GFP or PX-TIR–GFP and lentiviruses encoding nontarget (control), PI4KIIβ, or PI4KIIα shRNAs, were pulsed with LPS-coated polystyrene beads and chased as indicated. Phagosomes were purified and analyzed by flow cytometry. GFP-positive cells were not previously sorted. (A and B) Shown are histogram plots of a representative experiment with the percentages of gated phagosomes that were GFP-positive indicated. Solid black lines, nontarget (control) shRNA; solid blue lines, PI4KIIβ shRNA; solid red lines, PI4KIIα shRNA; dashed lines, nontransduced controls. (C and D) Data (mean ± SD) from three independent experiments performed in duplicate were normalized to the percent of transduced BMDCs (SI Appendix, Fig. S7D). (E) WT BMDCs were pulsed with LPS-coated polystyrene beads, and IL-6 released into the supernatants after 5 h was measured by ELISA. (E, Left) Representative experiment performed in triplicates. (E, Right) IL-6 values from three independent experiments performed in triplicate are shown as percent of values (mean ± SD) for BMDCs treated with nontarget (ctrl) shRNA, as a representation of phenotypic rescue. Significance relative to TIRAP shRNA-treated DCs (−) (E, Right) is indicated. **P < 0.01; ***P < 0.001; n.s., not significant.

To further investigate if TIRAP binding to PtdIns4P is required for TLR4 signaling from maturing phagosomes, we knocked down the endogenous TIRAP in BMDCs (SI Appendix, Fig. S9A) and expressed either human TIRAP–GFP or PX-TIR–GFP. TIRAP knockdown did not affect DC differentiation compared to shRNA control-treated DCs (SI Appendix, Fig. S9B), and knocked-down DCs were similarly transduced with TIRAP or PX-TIR constructs (SI Appendix, Fig. S9C). Consistent with our observations in WT DCs (Figs. 7 and 8), TIRAP–GFP was efficiently recruited to phagosomes at all times and to phagosomal tubules in mature phagosomes, while PX-TIR–GFP was recruited to early phagosomes and was no longer detected at 120 min (asterisks, SI Appendix, Fig. S9D), as expected, since PtdIns3P is limited to early phagosomes (49). TIRAP knockdown significantly reduced DC production of IL-6 after LPS-bead stimulation, as expected (Fig. 8E). Remarkably, expression of PX-TIR–GFP did not improve IL-6 production, whereas TIRAP–GFP significantly increased proinflammatory cytokine production (Fig. 8E). This result supports that TIRAP binding to PtdIns3P is not sufficient to promote TLR4 signaling from phagosomes and that binding to PtdIns4P is necessary.

Discussion

Intracellular trafficking pathways play a crucial role in preserving organelle identity. We previously showed that a key mediator of TLR4 localization to phagosomes in DCs is the trafficking adaptor protein AP-3. However, loss of AP-3 expression did not completely abrogate phagosomal TLR accumulation or signaling, suggesting that AP-3 might play a regulatory role in this process. We have now identified the lipid kinase PI4KIIα as an AP-3 cargo that is required to promote TLR4 signaling from phagosomes. PI4KIIα trafficking to phagosomes in DCs requires its kinase activity and binding to AP-3, consistent with the requirements for PI4KIIα delivery to lysosomes, lysosome-related organelles, and synaptic vesicles in other cell types (34). PI4KIIα recruitment to phagosomes and its dependence on AP-3 binding were observed both in BMDCs—which, despite their heterogeneity, may mirror monocyte-derived DCs in vivo (50, 51)—and in tissue-resident DCs isolated from spleen. PI4KIIα, in turn, generates PtdIns4P on phagosomes, and knockdown of PI4KIIα leads to reduced phagosomal PtdIns4P detected by the P4Mx2 probe. The phagosomal effects are specific for PI4KIIα, as knockdown of the genetically distinct PI4KIIβ—which does not bind AP-3—does not impact phagosomal PtdIns4P or TLR4 signaling. Finally, we show that PI4KIIα activity is required to recruit the TLR sorting adaptor TIRAP to phagosomes and that TIRAP binding to PtdIns4P is necessary for optimal proinflammatory TLR4 signaling. Together, the data suggest a model in which AP-3–dependent trafficking of PI4KIIα to phagosomes generates a pool of PtdIns4P that is necessary to recruit TIRAP and sustain TLR4 proinflammatory signaling and MHC-II presentation from phagosomes.

Our data show that PI4KIIα builds a PtdIns4P platform on the maturing phagosome that allows the binding of the TLR sorting adaptor TIRAP. TIRAP serves as a landmark for the assembly of the MyDdosome complex on phosphoinositide-enriched domains, promoting the initiation of TLR signal transduction. TIRAP was shown to promiscuously bind to different phosphoinositides in macrophages and DCs, including PtdIns(4,5)P₂ on the plasma membrane and PtdIns3P on early endosomes (25). Here, we show that on DC-maturing phagosomes, TIRAP binding is highly dependent on PtdIns4P and that knockdown of PI4KIIα, but not of PI4KIIβ, significantly reduces TIRAP recruitment to phagosomes. In contrast, TIRAP association with the plasma membrane and nascent phagosomes is PI4KIIα-independent—similarly to PH-PLCδ, a probe for PtdIns(4,5)P₂—suggesting that TIRAP binding to the plasma membrane is not absolutely dependent on PtdIns4P and requires PtdIns(4,5)P_2. Consistent with observations in macrophages, PtdIns3P (detected with the TIRAP construct PX-TIR) accumulates on early phagosomes in DCs, and its presence is not dependent on PI4KIIα or PI4KIIβ. However, WT TIRAP is not recruited to phagosomes at this stage in the absence of PI4KIIα, suggesting that TIRAP is preferentially recruited to early phagosomes by PtdIns4P. Furthermore, unlike exogenous WT TIRAP, PX-TIR recruitment to early phagosomes failed to promote proinflammatory signaling in the absence of endogenous TIRAP. Our observation that TIRAP is preferentially recruited to PtdIns4P-rich domains on phagosomes resembles TIRAP preferential binding to PtdIns(4,5)P₂ on the plasma membrane to promote TLR4 signaling (23). This also suggests that additional organelle-specific determinants might limit or promote phosphoinositide-dependent TIRAP recruitment to membranes and highlights differences between the early endosomal and phagosomal systems in DCs. Moreover, these data support the importance of distinguishing TLR signaling from distinct subcellular locations to reflect differences in cargo characteristics or origins—in this case, soluble vs. particulate—consistent with our observations that AP-3 is required for TLR signaling induced by phagosomal, but not endosomal, cargoes (10).

This distinction between the endosomal and phagosomal systems in DCs could also reflect variations between DCs and other phagocyte endolysosomal systems. Consistent with this, while PtdIns(4,5)P₂ and PtdIns3P are present on nascent phagosomes and early phagosomes, respectively, in macrophages (37), both persist longer on phagosomes in DCs than in macrophages. By contrast, PtdIns4P is generated earlier in the phagosomal maturation process in DCs compared to macrophages. Based on differences in the progression of phosphoinositide content shown here for DCs and previously for macrophages (36, 37), and on the acquisition of lysosomal proteins and low pH, the phagosomal maturation process is slowed in DCs compared to that in macrophages, supporting DC specialization in antigen presentation (52, 53). The earlier formation of PtdIns4P on DC phagosomes with the consequent recruitment of TIRAP and assembly of the MyDdosome licenses DC phagosomes for antigen presentation, an activity that is not a primary function in macrophages (54, 55).

Of note, even though knockdown of PI4KIIα abolished TIRAP binding to maturing phagosomes, TLR4 downstream responses were not completely abrogated. The persistence of TLR4 signaling in the absence of TIRAP phagosomal recruitment may result from residual signaling from plasma membrane-engaged TLRs, which are not regulated by PI4KIIα or AP-3. Indeed, we show that PI4KIIα specifically contributes to the phagosomal PtdIns4P pool. In contrast, the plasma-membrane pool of PtdIns4P—while somewhat reduced by knockdown of PI4KIIβ, but not of PI4KIIα—is primarily maintained by PI4KIIIα (56), supporting—together with the large pool of PtdIns(4,5)P₂—TLR4/MyDdosome signaling from the plasma membrane in cells depleted of PI4KIIα. In addition, it is possible that TLR4 could signal on the phagosome independently of TIRAP. Indeed, the TIRAP requirement for TLR signaling could be bypassed by high ligand concentrations that may not reflect physiological conditions (25, 57, 58).

In live cell-imaging experiments, PI4KIIα, PtdIns4P, and TIRAP were also detected on phagosomal tubules that we had previously shown to promote MHC-II presentation from phagosomes. Moreover, knockdown of PI4KIIα or loss of AP-3 expression abrogated phagosomal tubule formation. This could reflect reduced binding of the Rab7 adaptor FYCO1, which modulates kinesin microtubule motor protein required for tubule formation (59). Indeed, Rab7 is present on PtdIns4P enriched compartments (60). Of note, kinesin recruitment to phagosomal tubules in macrophages is mediated by the GTPase Arl8b (38), which is also required for MHC-II presentation in DCs (61). In addition, PtdIns4P may be required for the recruitment of membrane curvature stabilizing proteins (62). Expression of WT PI4KIIα, but not the kinase-inactive or AP-3 sorting mutants, rescued DC defects in phagosomal TLR signaling and MHC-II presentation. These results are in agreement with previous reports showing that both the dileucine sorting motif and the kinase active site are required for PI4KIIα recruitment to lysosomes in neuronal cells and that PI4KIIα acts both as a cargo and as a regulator of AP-3 function by promoting the formation of additional AP-3–recruiting PtdIns4P patches (34). Together, these observations support the conclusion that AP-3–dependent recruitment of PI4KIIα to phagosomes generates a pool of PtdIns4P that is required for the formation of phagosomal tubules, TIRAP recruitment, and concomitant enhancement of MHC-II presentation. Our data also suggest that impaired phagosomal PI4KIIα recruitment may at least, in part, explain defective antibacterial immune responses in AP-3–deficient mice and HPS2 patients.

In summary, our data indicate that AP-3–mediated recruitment of PI4KIIα early in the life cycle of the DC phagosome is a prerequisite for the binding of TIRAP and the promotion of TLR4 signaling and is a key determinant of the fate of the phagosome as an autonomous signaling organelle. Further studies will be required to elucidate the signals that drive TIRAP and TLR4 together on the phagosome to allow MyDdosome formation.

Experimental Procedures

Mice.

Mice were bred under pathogen-free conditions in the Department of Veterinary Resources at the Children’s Hospital of Philadelphia and were euthanized by carbon dioxide narcosis according to guidelines of the American Veterinary Medical Association Guidelines for the Euthanasia of Animals (63). All animal studies were performed in compliance with the federal regulations set forth in the recommendations in the Public Health Service Policy on the Humane Care and Use of Laboratory Animals (64), the National Research Council’s Guide for the Care and Use of Laboratory Animals (65), the NIH Office of Laboratory Animal Welfare, the American Veterinary Medical Association Guidelines on Euthanasia, and the guidelines of the Institutional Animal Care and Use Committees of the Children’s Hospital of Philadelphia. All protocols used in this study were approved by the Institutional Animal Care and Use Committee at the Children’s Hospital of Philadelphia.

See SI Appendix for details on mouse strains, reagents, antibodies, DNA constructs, shRNAs, lentiviral and retroviral production, and transduction of DCs.

Cell Culture.

Bone-marrow cells were isolated and cultured for 7 ± 9 d in Roswell Park Memorial Institute (RPMI) 1640 medium (Gibco, ThermoFisher) supplemented with 10% low endotoxin-defined fetal bovine serum (HyClone), 2 mM l-Gln, 50 μM 2-mercaptoethanol (Invitrogen), and 30% granulocyte-macrophage colony-stimulating factor-containing conditioned medium from J558L cells (kindly provided by former Ralph Steinman laboratory, The Rockefeller University, New York, and Maria Paula Longhi, Queen Mary University of London, London) for differentiation to DCs, as described (66, 67). Maturation was induced by 18-h treatment with LPS (0.1 μg/mL). Splenic DCs were isolated from single-cell suspensions with anti-CD11c (N418) microbeads after depletion of T, B, and natural killer cells with a mixture of biotin-conjugated antibodies (to CD90.2, CD45R, and CD49b) and anti-biotin microbeads (Miltenyi Biotec Inc.). CD4+ T cells were isolated from single-cell suspensions of lymph nodes from OT-II transgenic mice, by positive selection with anti-CD4 microbeads (Miltenyi Biotec Inc.).

Phagosome Purification and Protein Recruitment.

Phagosomes were isolated essentially as described (68). Briefly, BMDCs were incubated for 15 min with LPS-coated latex beads or 3-μm magnetic beads (Dynabeads M-280 streptavidin; Invitrogen) and then chased. Magnetic and nonmagnetic phagosomes were purified after different chase times by means of a magnet or differential centrifugation, respectively, as described (69, 70). Purified LPS-bead phagosomes were fixed and stained with antibodies to TLR-4, lamp-1, or negative controls, or left unstained in the case of the GFP-expressing DCs, and analyzed concurrently by flow cytometry, gating on the bead population (68), and normalizing to the total percentage of GFP-positive cells in the case of the transduced DCs. Flow cytometry was performed by using FACSCalibur and CellQuest software (BD Biosciences). Protein extracts from purified magnetic phagosomes were analyzed by immunoblotting.

Protein-Degradation Assays.

To evaluate intraphagosomal degradation, BMDCs were pulsed with OVA-coated latex beads in a 1:5 ratio of DC:beads for 15 min and chased for the indicated times. Cells were then disrupted in detergent-containing lysis buffer and pelleted by centrifugation at 150 × g for 4 min as described (68). Supernatants containing the latex beads were collected and stained with anti-OVA antibodies (Sigma), followed by phycoerythrin-conjugated anti-rabbit antibodies (Jackson ImmunoResearch) in 96-well V-bottom microplates. Labeling was analyzed by flow cytometry, gating on the latex bead population by forward scatter and side scatter (68).

Immunoblotting.

Immunoblotting was performed essentially as described (10). Briefly, Laemmli sample buffer with 2-mercaptoethanol was added to protein lysates from phagosome purification and whole-cell lysates. Samples were then fractionated by 8%, 10%, or 12% sodium dodecyl sulfate (SDS)/polyacrylamide gel electrophoresis (PAGE) on polyacrylamide gels, transferred to polyvinylidene difluoride membranes (Immobilon-FL, Millipore), and analyzed by using Alexa Fluor 680- or 790-conjugated or horseradish peroxidase-conjugated secondary antibodies (Jackson ImmunoResearch) and Odyssey (LI-COR) or iBright (Invitrogen) imaging systems. Densitometric analyses of band intensity was performed by using NIH ImageJ software, normalizing to control protein levels.

Cytokine Secretion after TLR4 Stimulation.

BMDCs were incubated with LPS-coated beads (1 μg/mL) for 3 or 5 h as described (10). IL-6 concentration in culture supernatants was measured by ELISA (BD Biosciences).

Immunofluorescence Microscopy.

BMDCs on day 7 of culture or freshly isolated splenic DCs were seeded on poly-l-lysine–coated glass-bottom 35-mm culture dishes (MatTek), pulsed with OVA (1 mg/mL) and LPS (100 μg/mL) coupled to 3-μm amino polystyrene beads as described (67), fixed with 3% formaldehyde in phosphate-buffered saline (PBS), permeabilized with Permwash (BD), and labeled with the indicated antibodies. Cells were analyzed by fluorescence confocal microscopy with a DMi8 Leica microscope and VisiView software (Visitron Systems GmbH) for image capture and analyzed by using ImageJ (NIH). Protein recruitment to phagosomes was visualized with the ImageJ plugin three-dimensional (3D) viewer and quantified by using Analyze/Plot profile and Analyze/3D surface plot.

For the detection of phagosomal tubules, BMDCs were seeded and pulsed as indicated, chased for 2 h, fixed for 10 min at 37 °C in a periodate–lysine–paraformaldehyde fixative (71) at 37 °C, washed with Hank’s balanced salt solution, permeabilized with Permwash, and stained with the indicated antibodies.

Live Cell Imaging.

BMDCs expressing retroviral and/or lentiviral constructs were seeded on poly-l-lysine–coated glass-bottom 35-mm culture dishes on day 7 of culture. On day 8, DCs were pulsed for 30 min with 1 mg/mL TxR-conjugated OVA (Invitrogen, ThermoFisher Scientific) and LPS (100 μg/mL) covalently coupled to 3-μm amino polystyrene beads, as described (67). DCs were then washed with RPMI, chased for 0 to 2.5 h, and visualized by spinning-disk confocal microscopy using an Olympus inverted microscope equipped with an environmental chamber at 37 °C and 5% CO₂ at the University of Pennsylvania’s Confocal Microscopy core or a BioVision and DMi8 Leica spinning-disk system using an Andor 888 cooled electron-multiplying charge-coupled device camera equipped with temperature and CO₂ control units and associated VisiView software for image and video capture. Time-lapse microscopy was performed by capturing image streams over 1 to 5 min at 1 frame per s and analyzed by using ImageJ. Protein recruitment to phagosomes was visualized and quantified as explained above.

Antigen Presentation Assays.

DCs were exposed to OVA, OVA:BSA-coated 3-μm latex beads (Polysciences), or OVA-specific MHC-II peptides for 15 to 30 min at 37 °C, then washed in PBS and chased in complete medium at 37 °C. DCs were then fixed with 0.005% glutaraldehyde in PBS for 1 min, washed with 0.2 M glycine in PBS, and cocultured with CD4+ OT-II T cells that had been prestimulated with anti-CD3 and anti-CD8 antibodies (72). T cell activation was monitored 18 h later as CD69 expression by flow cytometry (FACSCalibur, BD Biosciences) and IL-2 secretion in coculture supernatants by ELISA (BD Biosciences). For presentation of Eα_52–68 peptide on I-A^b, BMDCs were incubated with 0.5 mg of soluble EαGFP, Eα_52–68 peptide, or EαGFP-coated (1 mg/mL) beads for 30 min, washed with PBS, and chased. Cells were then fixed with 3% paraformaldehyde in PBS, stained with biotinylated YAe and allophycocyanin–streptavidin (Invitrogen), and analyzed by flow cytometry. YAe labeling was quantified on CD11c+ cells that had taken up one bead as gated on a forward-scatter vs. side-scatter plot (73).

Statistical Analyses.

Statistical analyses and data plots were performed by using Microsoft Excel and GraphPad Prism software. Significance for experimental samples relative to untreated or nontarget shRNA-treated WT control (unless otherwise stated) was determined by using the unpaired Student’s t test and ANOVA. Mean ± SEM values are indicated in Results. Error bars in figures represent mean ± SD.

Data Availability.

All study data are included in the article and SI Appendix.