Product# 1081 HIV-1 gp120 (ADA)
Product# 1011 HIV-1 gp120 (subtype C)
Product# 1031 HIV-1 gp120 (YU2)
Human immunodeficiency virus type 1 (HIV-1) impairs major functions of macrophages but the molecular basis for this defect remains poorly characterized. Here, we show that macrophages infected with HIV-1 were unable to respond efficiently to phagocytic triggers and to clear bacteria. The maturation of phagosomes, defined by the presence of late endocytic markers, hydrolases, and reactive oxygen species, was perturbed in HIV-1–infected macrophages. We showed that maturation arrest occurred at the level of the EHD3/MICAL-L1 endosomal sorting machinery. Unexpectedly, we found that the regulatory viral protein (Vpr) was crucial to perturb phagosome maturation. Our data reveal that Vpr interacted with EB1, p150Glued, and dynein heavy chain and was sufficient to critically alter the microtubule plus end localization of EB1 and p150Glued, hence altering the centripetal movement of phagosomes and their maturation. Thus, we identify Vpr as a modulator of the microtubule-dependent endocytic trafficking in HIV-1–infected macrophages, leading to strong alterations in phagolysosome biogenesis.
Macrophages play crucial functions at the interface between innate and adaptive immunity and also represent niches for intracellular pathogens. They are professional phagocytes that take up pathogens and debris through various opsonic and nonopsonic receptors (e.g., Fc receptors [FcRs] for the Fc portion of immunoglobulins; Flannagan et al., 2012; Canton et al., 2013). Interactions between these receptors and their ligands induce signaling cascades, leading to strong and transient actin polymerization, plasma membrane remodeling, and pseudopod extension around the particulate material (Flannagan et al., 2012; Deschamps et al., 2013; Niedergang, 2016). The closed compartment that forms (the phagosome) loses its actin coat, undergoes fusion and fission with compartments of the endocytic machinery (Botelho and Grinstein, 2011; Fairn and Grinstein, 2012), and eventually fuses with lysosomes. This progressive maturation into a phagolysosomal compartment is accompanied by an acidification of the compartment and its enrichment in hydrolases and reactive oxygen species, forming a degradative compartment. The molecular machineries required for fusion and fission are thought to be the same as for endosome maturation (Fairn and Grinstein, 2012; Scott et al., 2014). Concomitantly, there is a motor-based migration on microtubules toward the cell center to reach a perinuclear localization where lysosomes are located (Blocker et al., 1998; Harrison et al., 2003).
Human immunodeficiency virus type 1 (HIV-1) infects and kills T cells, which profoundly damages the host-specific immune response but also integrates into memory T cells and long-lived macrophages, establishing a chronic infection (Carter and Ehrlich, 2008; Koppensteiner et al., 2012b). Because macrophages are thought to retain viruses in an infectious form, and to potentially release them in a delayed manner and in different locations, they are proposed to be important for virus dissemination and pathogenesis. HIV-1 infection impairs the functions of macrophages both in vivo and in vitro (Kedzierska and Crowe, 2002; Collman et al., 2003), which may contribute to the development of opportunistic diseases. Impaired phagocytosis was also reported in a population of small alveolar macrophages in HIV-infected patients (Jambo et al., 2014). We previously showed that HIV-1, via the viral negative factor (Nef), a major virulence factor that is highly expressed early during virus replication (Witkowski and Verhasselt, 2013), indeed affects phagocytosis by inhibiting the membrane remodeling events that are required for efficient phagosome formation (Mazzolini et al., 2010). Another regulatory viral protein (Vpr) is specifically incorporated into virus particles. Vpr has several described activities, including cell cycle arrest, control of the reverse transcription process, and modulation of the HIV-1 mutation rate (Planelles and Benichou, 2009; Kogan and Rappaport, 2011; Guenzel et al., 2014).
Here, we show that the late steps of phagocytosis are impaired in HIV-infected primary human macrophages, leading to an alteration in cell activation, cytokine production, and bacterial clearance. Phagosome maturation was inhibited, as the endocytic sorting elements based on EHD3/MICAL-L1 were hijacked by the viral activity. Using mutant strains of HIV-1, we demonstrate that Vpr is unexpectedly involved in the perturbation of phagosome maturation. We further show that Vpr interacts with EB1, p150Glued, and the dynein heavy chain (DHC). During HIV infection, Vpr is crucial to perturb the localization at the plus ends of microtubules of EB1 and p150Glued. This affects the centripetal movement of phagosomes on microtubules, and thus an efficient maturation. We identify Vpr as a major regulator of microtubule-dependent trafficking.
Modification of activation and clearance activity in HIV-1–infected macrophages
To gain insight into the defect in phagocytic functions in HIV-infected macrophages, we aimed to dissect the signaling cascades downstream of the engagement of surface receptors. Monocytes from healthy donors were differentiated into macrophages (monocyte-derived macrophages [MDMs]) with recombinant macrophage-colony stimulating factor for 11 d and were then infected with HIV-1ADA wild type (WT) for 8 d. MDMs were incubated for various times with IgG-opsonized sheep red blood cells (SRBCs) to induce a phagocytic trigger. After various times of contact, cells were lysed and analyzed by Western blotting to detect activation of the MAPKs p38, extracellular signal regulated-kinase 1/2 (ERK1/2), and stress-activated protein kinase (SAPK)/JNK. These kinases play a role in the maturation process of the phagosomes (Moretti and Blander, 2014) and also in activation of transcription factors, such as nuclear factor-κB, which leads to subsequent induction of secretion of proinflammatory cytokines. We noticed that the basal phosphorylation of SAPK/JNK, ERK1/2, and p38, as well as p65/RelA, was higher in HIV-1–infected macrophages than in noninfected cells, despite the relatively low rate of infection of primary human macrophages by WT viruses (between 10% and 40%; Fig. 1, A–D). After stimulation of FcR, however, the phosphorylation of ERK1/2 was markedly reduced in HIV-1–infected macrophages compared with noninfected cells (Fig. 1, E and F). Quantification of the results indicated that there are two waves of activation of ERK1/2 in control cells with peaks at 10 and 180 min, but no increase in ERK1/2 phosphorylation in HIV-infected macrophages. This was specific to the HIV-1 viral infection, because we did not observe the same defects after other preactivation treatments (Fig. 1 G).
When we used a cytokine array to detect various cytokines and chemokines in a semiquantitative manner in the supernatant of cells 6 h after stimulation (Fig. 1 H), we also noticed reduced production of cytokines in the supernatant of cells preinfected with HIV-1ADAWT, both in resting conditions and after a phagocytic stimulus, compared with noninfected cells. Therefore, our results indicate that the HIV-1 infection of macrophages induced a basal “preactivation” of the cells that dampened the cellular response downstream of the engagement of the phagocytic receptors.
Phagocytosis eventually leads to the degradation of the ingested material. Some pathogens, such as Salmonella typhimurium, are invasive facultative intracellular bacteria that have evolved highly adapted gene expression programs to shape the vacuole in which they reside. We compared the intracellular survival of S. typhimurium in HIV-1ADA-VSV-GWT–infected versus noninfected macrophages using a gentamicin-plating assay (Fig. 1 I). A VSV-G pseudotyped virus was used to achieve higher rates of infection and the bacterial survival was assessed by counting the number of intracellular bacteria 1 and 24 h postincubation. The data are expressed as a ratio (i.e., index of survival, expressed relative to noninfected macrophages). Intracellular S. typhimurium survived 3.2-fold ± 0.2-fold better in HIV-infected macrophages compared with control cells. Such a defect in bacterial clearance is indicative of an altered phagosomal maturation.
HIV infection impairs phagosome maturation
To further characterize the defective clearance activity in macrophages infected with HIV-1, we analyzed the late steps of phagosome maturation and the luminal content of phagosomes using 3-µm beads coated with IgG to target FcRs. These beads were coupled to fluorophores sensitive to the hydrolytic activity (DQ-BSA beads) or to the oxidative burst (dichlorodihydrofluorescein diacetate [H2DCFDA]-OxyBURST beads), as well as a pH-insensitive calibration fluorophore to correct for variation in phagocytosis (Yates and Russell, 2008; Podinovskaia et al., 2013). MDMs infected with HIV-1ADA-VSV-GWT or noninfected controls were incubated for various times at 37°C with beads and then analyzed by flow cytometry focusing on the population of cells associated with beads (Fig. 2, A–D). The oxidative burst was detected as soon as 20–30 min until 3 h after contact with the beads in noninfected MDMs. In HIV-1–infected macrophages, the signal was reduced at each time point, up to 92% of the control condition (Fig. 2 C). The hydrolytic activity was detectable after 1.5 h of contact with beads in control conditions. In HIV-infected macrophages, we observed that the hydrolytic activity was reduced compared with noninfected MDMs (between 23% and 80% depending on the time; Fig. 2 D). Therefore, there is a marked decrease in the production of reactive oxygen species and hydrolytic activity in phagosomes of HIV-1–infected macrophages.
Together, these results point to a defect in phagosome maturation at the level of sorting/recycling endosomes with hijacking of the EHD3/MICAL-L1 sorting machinery by HIV-1.
The perturbation of phagosome maturation in macrophages requires established infection and expression of the viral factor Vpr
To better understand how HIV perturbs the functions of macrophages, we infected the primary human macrophages with HIV-1ADAWT for various times before assessing LAMP1 recruitment as in Fig. 2. There was no significant inhibition of phagosomal maturation after 2 or 3 d (Fig. 4 A). However, we observed a marked inhibition of the recruitment of LAMP1 on the phagosomes after 6 or 8 d of HIV infection. Moreover, the inhibition of phagosomal maturation by HIV-1 was not observed when macrophages were treated with integrase inhibitor raltegravir, indicating that viral integration was necessary (Fig. 4 B). The capacity of macrophages to ingest IgG-opsonized particles, on the other hand, was progressively reduced with time of infection (Fig. 4 C); using nef-deleted HIV-1 variants, we confirmed that Nef was important for the internalization step (Fig. 4, C and F; Mazzolini et al., 2010). In contrast, Nef was not significantly involved in inhibiting recruitment of LAMP1 on the phagosomes that still did get internalized (Fig. 4 D). Most importantly, infection of macrophages with an HIV-1 strain deleted for the Vpr factor showed a recovery of the recruitment of LAMP1 on phagosomes, indicating that Vpr was essential for the virus to inhibit phagosome maturation (Fig. 4 E). Interestingly, there was no difference between HIV-YU-2WT and HIV-YU-2ΔVpr on the efficiency of the internalization step of phagocytosis (Fig. 4 G), demonstrating that, unlike Nef, Vpr is not involved in phagosome formation but is involved in phagosome maturation.
Together, these data clearly demonstrate that the impairment of phagosome maturation in human macrophages is not a consequence of the perturbation of Nef-dependent early membrane remodeling events. Moreover, they reveal that the virus evolved with two factors to inhibit entry and maturation. Thus, Vpr was identified as a major regulator of phagosome maturation in HIV-infected macrophages.
The centripetal movement of phagosomes is slower in HIV-infected macrophages in a Vpr-dependent manner
Next, we investigated the subcellular localization of phagosomes containing IgG-opsonized SRBCs after uptake (Fig. 5, A and B). In control macrophages, ≈70% of phagosomes reached the cell center within 20 min and ≈90% after 60 min, whereas only 30% of the phagosomes were at the cell center in HIV-infected macrophages after 20 min and 44% after 60 min, with much variability. The movement of the phagosomes to the cell center was delayed in macrophages infected with HIV-1ADAWT, and this delay was at least partially dependent on Vpr because the phagosome distribution was not significantly different between cells infected with HIV-YU-2ΔVpr and noninfected macrophages (Fig. 5 C; P < 0.05).
Centripetal movement of phagosomes is inhibited in HIV-1–infected macrophages. (A and B) Primary human macrophages were noninfected (black bars) or infected with HIV-1ADAWT (red bars) for 8 d. The cells were incubated for different time points with IgG-SRBCs at 37°C and then fixed and permeabilized. They were labeled and analyzed as described in Fig. 2. Peripheral SRBCs, situated at a distance to the nucleus of more than two SRBCs in diameter, were counted for at least 200 phagosomes per condition at each time point. Results are expressed as the percentage of total number of SRBCs; two independent experiments on different donors are shown. (C) Primary human macrophages were noninfected (black bars) or infected with HIV-1YU-2WT (red bars) or HIV-1YU-2ΔVpr (blue bars) for 8 d. Cells were incubated with IgG-SRBCs for 1 h at 37°C and then analyzed as in A. The means ± SEM of three independent experiments are plotted. (D) Primary human macrophages were infected with HIV-1Gag-iGFP for 8 d, then incubated with IgG-SRBCs at 37°C under a spinning disk confocal microscope equipped with 5% CO2 and a heated chamber. Images were recorded every minute for 120 min. The distances covered by internal phagosomes were measured for 53 phagosomes in noninfected cells and 60 phagosomes in HIV-infected macrophages and were plotted against time. The speed was calculated by linear regression. (E) Primary human macrophages were treated as in C. Gallery of phase contrast images of noninfected (top panels; see also Video 1) and HIV-1Gag-iGFP–infected (bottom panels; see also Video 2) cells showing phagosome movement. Bar, 10 µm. *, P < 0.05; **, P < 0.005.
To further characterize the centripetal movement of phagosomes, we followed their movement in living HIV-1–infected macrophages (Fig. 5, D and E) using a spinning disk confocal microscope and HIV-1Gag-iGFP, which generates infectious virions in primary macrophages (Koppensteiner et al., 2012a; Gaudin et al., 2013). Phagosome movements were recorded every minute for 2 h after uptake, the internalization of particles being detected via a transition from bright phase to dark phase. The recorded velocities are of similar magnitude to the speeds already reported (Blocker et al., 1998; Harrison et al., 2003). The movements were slower in HIV-infected macrophages immediately after uptake (Fig. 5, C and D). Therefore, the peripheral location of phagosomes in HIV-infected macrophages appears as a consequence of a slowdown in the intracellular trafficking of the newly formed phagosomes.
Perturbed localization at microtubule plus ends of EB1 and p150Glued in HIV-infected macrophages
Because phagosome movement is a microtubule-dependent process (Blocker et al., 1998; Harrison et al., 2003), we investigated the microtubule network in HIV-1–infected macrophages (Fig. 6). Control MDMs and cells infected with HIV-YU-2WT or HIV-YU-2ΔVpr for 8 d were treated with nocodazole to depolymerize the microtubules (Fig. 6 A). After extensive washes, the cells were placed in medium without nocodazole to allow microtubule repolymerization. Although there was no striking difference between HIV-1–infected or control cells in steady-state conditions (basal), microtubule repolymerization was slower in HIV-1–infected MDMs compared with noninfected cells. The phenotype was intermediate in MDMs infected with a vpr-deleted HIV-1 mutant. Therefore, the infection of macrophages with HIV-1 perturbs the microtubule dynamics.
HIV-1 and Vpr perturb the microtubule dynamics and localization of EB1 and p150Glued. (A) Primary human macrophages were noninfected (top), infected with HIV-1YU-2WT (middle), or infected with HIV-1YU-2ΔVpr (bottom) for 8 d. Cells were treated or not (basal) with nocodazole at 10 µM for 1 h at 37°C. After washing, cells were fixed (time 0) or incubated at 37°C without nocodazole for the indicated times before fixation. Macrophages were then labeled with an anti-p24 antibody, followed by Cy2-labeled anti–mouse IgG and a recombinant anti-tubulin, and then followed by Cy3-labeled anti–human IgG. Images were acquired under a spinning disk microscope. One optical section for tubulin and Z projections of sections for p24 staining, from stacks of representative cells, are shown. Bar, 10 µm. (B) Primary human macrophages were noninfected (top), infected with HIV-1YU-2WT (middle), or infected with HIV-1YU-2ΔVpr (bottom) for 8 d. The cells were then fixed and stained with an anti-p24, followed by Cy2-labeled anti–goat IgG (left) and an anti-EB1, followed by Cy3-labeled anti–mouse IgG (middle). Stacks of images were acquired with a wide-field microscope and analyzed to define (right) and quantify (C) the comet-shaped ellipsoid objects. Bar, 5 µm; magnification in insets is 3.3×. (C and D) Macrophages were treated as in B and the number of EB1-positive comets (C; n > 2000 comets per condition) or the total intensity of EB1 staining (D) was calculated in 12 noninfected (black bars), 12 HIV-1YU-2WT–infected (red bars), and 12 HIV-1YU-2ΔVpr (blue bars) cells. Results are expressed as a percentage of control noninfected cells. The means ± SEM of three independent experiments (donors) are plotted. (E) Primary human macrophages were noninfected (top), infected with HIV-1YU-2WT (middle), or infected HIV-1YU-2ΔVpr (bottom) for 8 d. The cells were then fixed and stained with an anti-p24 antibody, followed by Cy2-labeled anti–goat IgG (not depicted) and an anti-p150Glued, followed by Cy3-labeled anti–mouse IgG. Stack of images were acquired, and Z projection of images is shown after TopHatFilter treatment. Bar, 10 µm; magnification in insets is 2.2×. *, P < 0.05; ***, P < 0.0005.
Proteins that localize to growing microtubule plus ends, collectively called plus-end tracking proteins (+TIPs), such as EB1 (Akhmanova and Steinmetz, 2010; Gouveia and Akhmanova, 2010), are important to stabilize microtubules and confer local functions of the microtubule cytoskeleton. We observed a reduced localization of EB1-positive comet-shaped structures at the periphery of the cells in HIV-1–infected macrophages (Fig. 6 C), although the total amount of EB1 was unchanged (Fig. 6 D). There was no such reduction in HIV-ΔVpr–infected cells (Fig. 6 D). The centripetal movement of organelles relies on the dynein/dynactin motor complex (Harrison et al., 2003) and EB1 was recently shown in vitro to recruit p150Glued to target the dynein/dynactin complex to the plus ends of microtubules (Duellberg et al., 2014). We observed that p150Glued was also mislocalized in HIV-infected macrophages (Fig. 6 E) and that the effect was partial in HIV-ΔVpr–infected cells.
Collectively, our data show that HIV-1 infection affects the plus end loading of EB1 and p150Glued in a Vpr-dependent manner in primary human macrophages.
Vpr is sufficient to interact with and perturb the localization of EB1, p150Glued, and DHC
To analyze whether the expression of Vpr alone induces the mislocalization of +TIPs, we transiently transfected primary human macrophages to express HA-Vpr and stained them for EB1 (Fig. 7, A and B) and p150Glued (Fig. 7, C and D). Confocal sections and 3D reconstructions show that expression of Vpr led to the mislocalization of EB1 and p150Glued from the plus ends of microtubules to more perinuclear and also nuclear localization where Vpr is accumulated in these conditions.
To further confirm the interaction between Vpr and microtubule-associated proteins, we performed coimmunoprecipitation experiments to precipitate HA-Vpr and mass spectrometry analysis on lysates from transfected HeLa cells. We found that DHC was part of the proteins precipitated with HA-Vpr but not after control transfection with the empty HA plasmid. We confirmed this result by Western blotting (Fig. 7 E).
Next, we used the proximity ligation in situ assay (Duolink; Söderberg et al., 2006) to assess protein interaction (Fig. 7, F–K). For this, we used HeLa cells with conditions to avoid massive toxic overexpression of HA-Vpr but reaching around 30% transfection efficiency. The mean number of spots detected per cell randomly analyzed on microscopy fields showed that Vpr interacted with EB1, p150Glued, and DHC (Fig. 7, H–K), compared with negative controls obtained by omitting one primary antibody in HA-Vpr–expressing coverslips (Fig. 7 F) or with the results obtained after empty HA plasmid transfection or a positive control combining the detection of α-tubulin and DHC.
In conclusion, Vpr interacted with EB1, p150Glued, and DHC and is sufficient to induce their mislocalization.
Vpr is sufficient to impair phagosome maturation
When human macrophages transiently expressing HA-Vpr were allowed to phagocytose IgG-opsonized SRBCs, we observed that Vpr led to a significant inhibition of LAMP1 recruitment on internalized phagosomes (Fig. 8, A and B), indicating that expression of Vpr alone was sufficient to alter the recruitment of this late phagosomal marker. In addition, the depletion of EB1 induced a 38.1% ± 4.5% defect in phagosome maturation, as measured with the acquisition of LAMP1 on internalized phagosomes (Fig. 8, C and D). Together, these results show that EB1 depletion or Vpr expression was sufficient to lead to a defect in phagosome maturation.
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