The innate immune system is important for control of infections, including herpesvirus infections. Intracellular DNA potently stimulates antiviral IFN responses. It is known that plasmacytoid dendritic cells sense herpesvirus DNA in endosomes via TLR9 and that nonimmune tissue cells can sense herpesvirus DNA in the nucleus. However, it remains unknown how and where myeloid cells, such as macrophages and conventional dendritic cells, detect infections with herpesviruses. In this study, we demonstrate that the HSV-1 capsid was ubiquitinated in the cytosol and degraded by the proteasome, hence releasing genomic DNA into the cytoplasm for detection by DNA sensors. In this context, the DNA sensor IFN-γ–inducible 16 is important for induction of IFN-β in human macrophages postinfection with HSV-1 and CMV. Viral DNA localized to the same cytoplasmic regions as did IFN-γ–inducible 16, with DNA sensing being independent of viral nuclear entry. Thus, proteasomal degradation of herpesvirus capsids releases DNA to the cytoplasm for recognition by DNA sensors.

The innate immune system represents a first line of defense against infections, including viral infections (1, 2), and uses a limited set of pattern recognition receptors (PRRs) to sense pathogen-associated molecular patterns (PAMP)s, which are either microbe-specific molecules or molecules with abnormal location of chemical modifications (36). A subset of PRRs stimulates expression of the antiviral type I IFNs in response to PAMP recognition (1, 2), and absence of proper IFN responses were shown to lead to severely impaired defense against viral infections in humans and mice (79).

Herpesviruses are a large family of dsDNA viruses, which are the causative agents of disease, including encephalitis, genital herpes (HSV), congenital disorders, and various conditions in immunocompromised individuals (CMV). Following entry of the virus, either by direct fusion with the plasma membrane or via endocytosis, productive infection is initiated by transport of the DNA-containing capsid along microtubules to the nucleus where the viral DNA is delivered (10). However, many virus particles do not lead to productive infection, even in highly permissive cells (11), and cell types differ with respect to permissiveness for herpesvirus infections. Therefore, knowledge of innate immune response to viral infection requires understanding of both productive and nonproductive infections, as well as the issue of cell-type specificity.

DNA represents a potent PAMP stimulating IFN responses in many cell types (1214). About 10 intracellular DNA sensors have been proposed, yet the specific role for most of these sensors remains unclear. TLR9 is predominantly expressed by plasmacytoid dendritic cells (pDCs) and localizes to endosomes where it senses DNA, including herpesvirus DNA (1517). In non-pDCs, DNA is sensed in other subcellular locations. One proposed DNA sensor is IFN-γ–inducible (IFI)16, which is mainly localized to the nucleus, where this protein has long been known to play a role in DNA damage response, p53 signaling, and apoptosis (18). It was recently reported that HSV-1 DNA interacts with IFI16 in the nucleus of the human osteosarcoma cell line U2OS and that induction of IFN-β by HSV-1 in HEK293 cells is dependent on the nuclear localization of IFI16 (19). This was subsequently reported to be counteracted by the HSV-1 protein infected cell protein 0 (20). A previous report demonstrated nuclear sensing of Kaposi’s sarcoma–associated herpesvirus in human microvascular endothelial cells (21). In some cell types, including myeloid cells, a small portion of the cellular pool of IFI16 is localized in the cytoplasm (22), and most of the other proposed DNA sensors, including the helicase DDX41, which is involved in DNA sensing in conventional dendritic cells (cDCs), localize to the cytoplasm (23). However, there is no information about the subcellular site of herpesvirus DNA sensing in myeloid cells or how the viral genomic material is made accessible for DNA sensors in these cells. Myeloid cells, like macrophages and cDCs, play important roles in innate control of virus infections and are important producers of type I IFN during infection (24, 25). In addition, it was reported that cDCs activated by cytosolic DNA sensing potently activate the adaptive immune response (26).

Common for IFI16, DDX41, and other proposed DNA sensors is the requirement for stimulator of IFN genes (STING) for downstream signaling stimulating IFN expression (22, 23, 27, 28). Upon DNA sensing, STING relocalizes to as-yet-uncharacterized cytoplasmic foci, which are believed to serve as assembly platforms for signaling (28). It was recently reported that the C-terminal region of STING is responsible for assembly of the signaling complex activating the transcription factor IRF-3, which drives transcription of IFN-β and IFN-stimulated genes (ISG) (29).

In this study, we demonstrate that HSV-1 and CMV infection induce IFI16-dependent IFN-β expression in human macrophages and that the infections mobilize IFI16 and STING to relocalize to the same subcellular regions. Moreover, IFI16 also associated with the same regions as did the viral DNA genomes. In the macrophages, the ability of HSV-1 to induce IFN responses was independent of the ability of the virus to deliver its DNA into the nucleus, and the IFI16–DNA association was strictly cytoplasmic. Finally, we report that the HSV-1 capsid becomes ubiquitinated in macrophages and describe a vital role for the ubiquitin–proteasome pathway in degradation of HSV-1 capsids, allowing for exposure of viral DNA for immediate sensing by DNA sensors. This study provides a mechanism for how viral DNA is made accessible to cytosolic DNA sensors to stimulate antiviral and inflammatory responses.

THP1 cells, cultured as nonadherent monocyte-like cells, Vero cells, and U2OS cells were grown in RPMI 1640 (Invitrogen), with 10% FCS, 600 μg/ml glutamine, 200 IU/ml penicillin, and 100 μg/ml streptomycin (Life Technologies). Human foreskin fibroblasts (HFFs) were grown in DMEM supplemented with 15% FCS and antibiotics, as described above. THP1 cells were differentiated into macrophage-like cells by addition of 100 nM PMA (Sigma-Aldrich). All presented data with THP1 cells were based on PMA-differentiated cells. Buffy coats from Aarhus University Hospital blood bank were used to collect PBMCs by Ficoll-Paque (GE Healthcare) gradient centrifugation. Monocytes were purified by plastic adherence using 1 × 108 PBMCs seeded in 6-cm UpCell plates (Nunc, Thermo-Scientific) precoated with poly-l-lysine (0.01% w/v; Cultrex) and allowed to stabilize overnight in IMDM supplemented with 10% FCS, 600 μg/ml glutamine, 200 IU/ml penicillin, and 100 μg/ml streptomycin. The following day, unattached cells were washed away, and adherent cells were differentiated into monocyte-derived macrophages (MDMs) by culturing for an additional 5 d in culture media: IMDM supplemented with 10% (v/v) pooled AB+ human sera (Invitrogen), 600 μg/ml glutamine, 200 IU/ml penicillin and 100 μg/ml streptomycin (Life Technologies), and 20 ng/ml M-CSF (Sigma-Aldrich). A total of 0.5 × 106 MDMs was reseeded on glass coverslips for immunofluorescence studies. MDMs for IFI16 knockdown were generated as previously described (30). Bone marrow–derived macrophages (BMMs) were generated as previously described (31).

The Abs used were anti-IFI16 N-terminal (sc-8023), anti-ubiquitin (sc-34870), anti-RCC1 (sc-1161) (all from Santa Cruz Biotechnology), anti-K48 polyubiquitin (Cell Signaling Technology; #4289), anti-STING (IMGENEX; IMG-6485A), anti-Vp5 (Ab6508), anti-proteasome p20S subunit (ab3325), anti–β-actin (Abcam; Ab49900), anti-CMV minor capsid protein p28 (GenWay; 20-251-400023), anti-DDX41 (Sigma-Aldrich; SAB-2100554), peroxidase-conjugated F(ab′)2 donkey anti-mouse IgG (H+L) and peroxidase-conjugated Affinipure F(ab′)2 donkey anti-rabbit IgG (H+L) (both from Jackson ImmunoResearch), and polyclonal rabbit anti-goat Ig (DakoCytomation). Anti-IFI16 Ab raised against the C-terminal region was produced as described (32). The proteasome inhibitors used were MG132 (Merck Millipore), PI-083 (Merck Millipore), and PS-341 (LifeSensors). The proteasome inhibitors were shown to block LPS-induced IκBα degradation at the concentrations used in this study (data not shown). Leptomycin B (LMB) was obtained from Tocris Bioscience and was shown to block nucleocytoplasmic shuttling of IκBα (33) (data not shown). The synthetic dsDNA 60-mer derived from the HSV-1 genome nt 144107–144166 was obtained from DNA Technology (Aarhus, Denmark) (22).

HSV-1 strains 17+, HFEM, and the HFEM-derived mutant TsB7 were grown as previously described (34). Viruses were used at multiplicity of infection (MOI) 3–10 for HSV-1 and MOI 5 for CMV. Human CMV strain AD169 was cultivated as previously described (30). HSV-1 and CMV were titrated on Vero cells and HFFs (30). The virus preparations were tested for endotoxin content, and all were found to contain similar low levels (∼0.75 endotoxin units/ml). Virus was inactivated with UV-C light irradiation for 1 min (wavelength = 253.7 nm) with an intensity of 450 μW/cm2, which we showed previously to reduce HSV replication by >106-fold (35).

For transient knockdown of IFI16 in MDMs, 100 nM IFI16 small interfering RNA (siRNA) (Stealth RNAi mix; HSS105205, HSS105206, and HSS105207) and recommended controls (Stealth RNAi control; both from Invitrogen) were transfected using HiPerfect (QIAGEN), according to the manufacturer’s instructions, at day 5. The cells were incubated with the HiPerfect siRNA mix for 4 h at 37°C before changing the media to fresh media including GM-CSF. Forty-eight hours after siRNA transfections, the macrophages were used for infection experiments. Two hours before infection, macrophages were supplied with fresh media excluding GM-CSF.

The lentiviral short hairpin RNA (shRNA) expression plasmid pLKO.1 was used for generating stable gene expression knockdown in THP1 cells (Open Biosystems, Waltham, MA). The targeting shRNA sequence was IFI16 clone ID# TCRN0000019079, ddx41-1 clone ID# TRCN0000104010, and ddx41-4 clone ID# TRCN0000104013. The control shRNA vector was an empty vector pLKO.1 with an 18-nt shuttle sequence instead of the hairpin sequences. shRNA plasmids were amplified in TOP10 Escherichia coli (Invitrogen/Life Technologies: Paisley, U.K.) and purified using QIAGEN plasmid Plus kits (QIAGEN, Germantown, MD). Virus-like particles were produced using Fugene 6 (Roche) transfection of HEK293T cells with the packaging system pMDIg/p-RRE, pRSV-rev, pMG.2, and shRNA vector plasmid. Virus supernatants were harvested after 48 h and filtrated through a 0.45-μm membrane. Undifferentiated THP1 cells were infected with titrating amounts of virus supernatant and 2 d postinfection were placed under selection with puromycin at 1 μg/ml (Sigma-Aldrich). Level of knockdown was determined on differentiated THP1 cells by Western blot analysis after 7–10 d.

IFNβ and IFI16 gene expression were determined by real-time PCR, using TaqMan and SYBR green detection systems, respectively (Applied Biosystems, QIAGEN). Expression levels were normalized to β-actin or GAPDH expression, and data are presented as the fold induction over untreated controls for each phenotype. Data represent the mean ± SD from either biological replicates or technical replicates. The following PCR primers were used: IFI16: forward, 5′-TAGGCCCAGCTGAGAGCCATCC-3′ and reverse, 5′-TGAGGTCACTCTGGGCACTGTCTT-3′; ICP27: forward, 5′-AGACCAGACGGATCCCCTGGGAAACCT-3′ and reverse, 5′-AAACACGAAGGATCCAATGTCCTTAAT-3′; GAPDH: forward, 5′-CGACCACTTTGTCAAGCTCA-3′ and reverse, 5′-GGTGGTCCAGGGGTCTTACT-3′ (all from DNA Technology), IFNβ Applied Biosystems TaqMan Assay Hs01077958_s1, DDX41 Applied Biosystems TaqMan Assay Hs00169602_m1, AIM2 Applied Biosystems TaqMan Assay Hs00915710_m1, b-Actin Applied Biosystems TaqMan Assay Hs99999903_m1.

For visualization of IFI16, following infection with viruses at indicated times, cells were fixed and permeabilized with methanol at −20°C and labeled with Abs against IFI16 or STING. For visualization of ubiquitination and proteasome, BMMs were fixed with 4% formaldehyde, permeabilized with 0.2% Triton X-100, and stained with Abs specific for ubiquitin and the proteasome p20S subunit. Images were acquired on a Zeiss LSM 710 confocal microscope, using a 63× 1.4 oil-immersion objective, or on a DeltaVision RT Core fluorescent microscope using a 100× 1.4 oil-immersion objective, fitted with a Cascade 650 CCD camera. Image processing was performed using Zen 2010 (Zeiss) and ImageJ. Deconvolution of images acquired with DeltaVision RT Core was performed using Softworx software. All images are representative of at least two or three independent experiments.

Visualization of HSV-1 genomic DNA by fluorescence in situ hybridization (FISH) was performed as described (22). CMV genomic DNA was visualized using the same protocol. The fluorescein-labeled CMV-specific probes used were Probe 1: 5′-TAGCGGGGGGGTGAAACTTGGAGTTGCGTGTGTGGACGGCGACTAGTTGCGTGTGGTG-3′ and Probe 2: 5′-TTGGCAGGGTGTGTCAGGGTGTGTCGCGGGCGTGTGCCGGGTGTGTCGTGCCGGGTGTGT-3′ (DNA Technology). Briefly, cells were left untreated or were pretreated with LMB (10 ng/ml) or MG132 (10 μg/ml). Genomic viral DNA was labeled with genome-specific probes, and the capsid was labeled with Abs specific for Vp5 (HSV-1) or p28 CMV. In addition, cells were stained with Abs specific for IFI16 (32). Images were acquired as described above. All images are representative of at least three independent experiments.

Cells were infected as described above; supernatants were harvested and CXCL10 levels were measured by ELISA (R&D Systems). Phospho-IκBα levels were determined by Luminex technology using kits from Bio-Rad.

To isolate nuclear proteins, cells were washed twice in ice-cold PBS, scraped into 5 ml PBS, and centrifuged for 1 min at 2000 × g. The cells were resuspended in hypotonic lysis buffer (20 mM HEPES [pH 7.9], 1.5 mM MgCl2, 10 mM KCl, 0.2 mM EDTA, 0.5 mM DTT, and protease inhibitors) and left on ice for 15 min, after which Nonidet P-40 was added to 0.6%, and the suspension was vortexed vigorously for 15 s. The nuclei were recovered by centrifugation (10,000 × g for 1 min) and resuspended in 40 μl extraction buffer (20 mM HEPES [pH 7.9], 20% glycerol, 1.5 mM MgCl2, 420 mM NaCl, 0.2 mM EDTA, 0.5 mM DTT, 0.2% Nonidet P-40, and protease inhibitors). Supernatants containing nuclear proteins were isolated after 30 min of rocking at 4°C.

Cells were washed twice in PBS and lysed in 850 μl lysis buffer (50 mM HEPES [pH 7.5], 100 mM NaCl, 1 mM EDTA, 10% [v/v] glycerol, 0.5% [v/v] Nonidet P-40 containing Complete Protease Inhibitor mixture [Roche], and 1 mM sodium orthovanadate). For immunoprecipitation (IP), anti-Vp5 was precoupled to protein A–Sepharose beads (Sigma-Aldrich) overnight at 4°C. The beads were then washed twice in lysis buffer and incubated with 1.5 mg cell lysate/sample overnight at 4°C. The immune complexes were washed three times in lysis buffer, boiled, and analyzed by standard SDS-PAGE and Western blotting. Blots were visualized using an ImageQuant LAS 4000 mini Luminescent Image Analyzer (GE Healthcare).

The Student t test was used for all statistical analyses.

All data presented are representative of at least three independent experiments.

Type I IFNs are important for control of herpesvirus infections (4, 79). In the human monocyte-like cell line THP1 differentiated into a macrophage-like phenotype with PMA, we found that both HSV-1 and CMV induced IFN-β expression, as well as activation of signal transduction to the NF-κB pathway and induction of the ISG, CXCL10 (Fig. 1A–E). To explore the role of DNA sensors in induction of this response, we generated a THP1-derived cell line stably transfected with control shRNA or shRNA targeting IFI16 (Supplemental Fig. 1A, 1B). Infection of these cells with HSV-1 or CMV revealed a requirement for IFI16 for IFN-β induction by these viruses (Fig. 1F, 1G). Interestingly, knockdown of DDX41 also affected the IFN-β response to HSV-1 infection in the PMA-differentiated THP1 cells (Supplemental Fig. 1C–E), which might indicate that these two proposed DNA sensors act sequentially, in distinct pathways, or cooperatively in the same pathway. Finally, we wanted to validate the above data by testing the role of IFI16 in primary human MDMs. Importantly, similar to what we found in the cell line, we observed virus-induced IFN-β expression, which was decreased following siRNA knockdown of IFI16 (Fig. 1H, 1I, Supplemental Fig. 1F).

FIGURE 1.

Herpesviruses induce IFN responses in macrophages that are dependent on IFI16. PMA-differentiated THP1 macrophages were infected with HSV-1 (A, C) or CMV (B, D). (A and B) Total RNA was harvested 6 h postinfection, and IFN-β mRNA was determined by RT-PCR. (C and D) Supernatants from cells treated for 16 h as indicated were analyzed for CXCL10 protein levels by ELISA. (E) Whole-cell extracts from PMA-differentiated THP1 macrophages, treated as indicated, were isolated, and p-IκBα was determined by Luminex. PMA-differentiated THP1 macrophages stably transfected with control vector (EV) or IFI16 shRNA (sh-IFI16) (F, G) or MDMs transfected with si-CTRL or si-IFI16 RNA (H, I) were infected with HSV-1 (F, H) or CMV (G, I). Total RNA was harvested 6 h postinfection, and IFN-β mRNA was determined by RT-PCR. Data represent mean ± SD of duplicates. *p < 0.05.

FIGURE 1.

Herpesviruses induce IFN responses in macrophages that are dependent on IFI16. PMA-differentiated THP1 macrophages were infected with HSV-1 (A, C) or CMV (B, D). (A and B) Total RNA was harvested 6 h postinfection, and IFN-β mRNA was determined by RT-PCR. (C and D) Supernatants from cells treated for 16 h as indicated were analyzed for CXCL10 protein levels by ELISA. (E) Whole-cell extracts from PMA-differentiated THP1 macrophages, treated as indicated, were isolated, and p-IκBα was determined by Luminex. PMA-differentiated THP1 macrophages stably transfected with control vector (EV) or IFI16 shRNA (sh-IFI16) (F, G) or MDMs transfected with si-CTRL or si-IFI16 RNA (H, I) were infected with HSV-1 (F, H) or CMV (G, I). Total RNA was harvested 6 h postinfection, and IFN-β mRNA was determined by RT-PCR. Data represent mean ± SD of duplicates. *p < 0.05.

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IFI16-induced IFN responses are dependent on STING (22), which relocates to discrete punctuate structures after DNA recognition (28). Using MDMs and confocal microscopy, we were able to demonstrate that both IFI16 and STING relocated to such structures postinfection with HSV-1 or CMV, with a clear colocalization between IFI16-positive and STING-positive structures (Fig. 2). Over the course of studying two donors we found that 20 and 51% of STING foci were also positive for IFI16 postinfection with HSV-1 and human CMV, respectively. The constitutive expression of IFI16 in MDMs and PMA-differentiated THP1 cells (Fig. 2, Supplemental Fig. 2A, 2B) suggests a role for IFI16 as an early sensor of DNA. This was further underscored by colocalization of synthetic dsDNA and IFI16 as early as 30 min after DNA transfection and prior to the colocalization of dsDNA and STING, which was observed 60 min after transfection (Supplemental Fig. 2C). Thus, HSV-1 and CMV mobilize STING in human macrophages and induce IFN-β expression in an IFI16-dependent manner.

FIGURE 2.

IFI16 and STING colocalize following infection with herpesviruses. MDMs were mock infected or infected with HSV-1 or CMV for 4 h. Subcellular distribution of IFI16 and STING was determined by confocal microscopy. White box indicates area displayed in zoom column (zoom is magnified 5× from original image). Scale bar, 10 μm.

FIGURE 2.

IFI16 and STING colocalize following infection with herpesviruses. MDMs were mock infected or infected with HSV-1 or CMV for 4 h. Subcellular distribution of IFI16 and STING was determined by confocal microscopy. White box indicates area displayed in zoom column (zoom is magnified 5× from original image). Scale bar, 10 μm.

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To look for the subcellular localization of cellular DNA sensors in macrophages, we isolated nuclear and cytosolic fractions and performed Western blotting. We blotted for IFI16 and DDX41, as well as RRC1 (regulator of chromosome condensation 1, a nuclear protein) and β-actin (a cytoplasmic protein).

IFI16 was mainly found in the nuclear fraction, but a small pool of the cellular IFI16 localized to the cytoplasm in macrophages (Fig. 3A). In contrast, DDX41 was found exclusively in the cytoplasm (Fig. 3A). To look for the subcellular localization of viral DNA in the cells, HSV-1– and CMV-infected macrophages were probed with genome-specific probes and visualized by fluorescence. In the infected cells, the HSV-1 and CMV genomes were detected in the cytoplasm (Fig. 3B, 3C). In the case of HSV-1, most of the viral DNA (29 ± 5 genome-positive spots/100 cells) colocalized with the capsid protein Vp5 2 h postinfection (data not shown) but not at 4 h postinfection (Fig. 3B, 3C). At 6 h postinfection, very little staining for HSV-1 DNA was observed (data not shown). The cytoplasmic localization of herpesvirus DNA in the macrophages was in contrast to the predominantly nuclear localization of the viral genome in Vero cells (Fig. 3D), consistent with potent viral transcription and replication in Vero cells (Supplemental Fig. 3A, 3B). Importantly, in macrophages, IFI16 colocalized with the HSV-1 and CMV genomic DNA in the cytoplasm (Fig. 3E, 3F), with >40% colocalization between DNA and IFI16-positive cytoplasmic foci (42 ± 7%). The colocalization of IFI16 and HSV-1 DNA in the cytoplasm was not inhibited by LMB (Fig. 4A), which inhibits CRM1-mediated nuclear export signal–mediated export of proteins from the nucleus. This treatment also did not inhibit the detection of capsid-free HSV-1 DNA spots in the cytoplasm of infected macrophages (Fig. 4B, 4C), as determined by the application of a Hi-Lo look-up table to each image to allow for the observation of individual foci. These data indicate that sensing of herpesvirus DNA occurs in the cytoplasm in macrophages and is independent of nucleus-to-cytoplasm translocation of proteins.

FIGURE 3.

Herpesvirus DNA is present in the cytosol of infected macrophages and colocalizes with IFI16. (A) PMA-differentiated THP1 cells were mock infected or infected with HSV-1 at MOI 3 for 6 h. Cytosolic and nuclear extracts were isolated and analyzed for IFI16, DDX41, RCC1, and β-actin by Western blotting (WB). PMA-differentiated THP1 cells were infected with HSV-1 (B) or CMV (C) and Vero cells were infected with HSV-1 (D) for 4 h. Viral DNA was visualized by virus-specific FISH probes (green) and costained with anti-capsid–specific Abs. White box indicates area displayed in zoom column. Scale bar, 10 μm. (E and F) MDMs were infected with HSV-1 or CMV for 4 h, and viral DNA was visualized by virus-specific FISH probes (green) and costained with anti-IFI16–specific Abs. White box indicates area displayed in zoom column (zoom is magnified 5× from original image). Scale bar, 10 μm.

FIGURE 3.

Herpesvirus DNA is present in the cytosol of infected macrophages and colocalizes with IFI16. (A) PMA-differentiated THP1 cells were mock infected or infected with HSV-1 at MOI 3 for 6 h. Cytosolic and nuclear extracts were isolated and analyzed for IFI16, DDX41, RCC1, and β-actin by Western blotting (WB). PMA-differentiated THP1 cells were infected with HSV-1 (B) or CMV (C) and Vero cells were infected with HSV-1 (D) for 4 h. Viral DNA was visualized by virus-specific FISH probes (green) and costained with anti-capsid–specific Abs. White box indicates area displayed in zoom column. Scale bar, 10 μm. (E and F) MDMs were infected with HSV-1 or CMV for 4 h, and viral DNA was visualized by virus-specific FISH probes (green) and costained with anti-IFI16–specific Abs. White box indicates area displayed in zoom column (zoom is magnified 5× from original image). Scale bar, 10 μm.

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FIGURE 4.

Induction of type I IFN response and IFI16 colocalization are independent of nuclear delivery of the viral genome. (A) MDMs were treated with vehicle or the nuclear export inhibitor LMB and infected with HSV-1 for 4 h. Viral DNA was visualized by HSV-1–specific FISH probes (green) and costained with anti-IFI16–specific Abs. Scale bar, 10 μm. Zoom is magnified 5× from original image. (B and C) PMA-differentiated THP1 macrophages were pretreated with LMB and infected with HSV-1 for 4 h. HSV-1 DNA foci/100 cells (B) and percentage DNA foci negative for Vp5 (C) were determined. Data represent mean ± SD. (D) PMA-differentiated THP1 macrophages were infected with the HSV-1 mutant TsB7 at 33°C (permissive) or 39°C (nonpermissive) for 4 h, and the percentage of HSV-1 DNA foci negative for Vp5 was determined. Data represent mean ± SD. (E and F) PMA-differentiated THP1 macrophages were infected with TsB7 and HSV-1 for 6 h, and total RNA and whole-cell extracts were isolated for measurement of IFNβ mRNA (quantitative RT-PCR) and p-IκBα (Luminex), respectively. Data represent mean ± SD of duplicates. (G) PMA-differentiated THP1 macrophages and U2OS cells were infected with TsB7 and HSV-1 for 6 h at 33 or 39°C. Live virus was used for THP1 cells, and UV-inactivated virus was used for U2OS cells. Total RNA was isolated for measurement of ISG56 mRNA (quantitative RT-PCR). Data represent mean ± SD of triplicates. (H) MDMs were infected with TsB7 at 33 or 39°C for 4 h, and viral DNA was visualized by HSV-1–specific FISH probes (green) and costained with anti-IFI16–specific Abs. Scale bar, 10 μm. Zoom is magnified 5× from original image. *p < 0.05.

FIGURE 4.

Induction of type I IFN response and IFI16 colocalization are independent of nuclear delivery of the viral genome. (A) MDMs were treated with vehicle or the nuclear export inhibitor LMB and infected with HSV-1 for 4 h. Viral DNA was visualized by HSV-1–specific FISH probes (green) and costained with anti-IFI16–specific Abs. Scale bar, 10 μm. Zoom is magnified 5× from original image. (B and C) PMA-differentiated THP1 macrophages were pretreated with LMB and infected with HSV-1 for 4 h. HSV-1 DNA foci/100 cells (B) and percentage DNA foci negative for Vp5 (C) were determined. Data represent mean ± SD. (D) PMA-differentiated THP1 macrophages were infected with the HSV-1 mutant TsB7 at 33°C (permissive) or 39°C (nonpermissive) for 4 h, and the percentage of HSV-1 DNA foci negative for Vp5 was determined. Data represent mean ± SD. (E and F) PMA-differentiated THP1 macrophages were infected with TsB7 and HSV-1 for 6 h, and total RNA and whole-cell extracts were isolated for measurement of IFNβ mRNA (quantitative RT-PCR) and p-IκBα (Luminex), respectively. Data represent mean ± SD of duplicates. (G) PMA-differentiated THP1 macrophages and U2OS cells were infected with TsB7 and HSV-1 for 6 h at 33 or 39°C. Live virus was used for THP1 cells, and UV-inactivated virus was used for U2OS cells. Total RNA was isolated for measurement of ISG56 mRNA (quantitative RT-PCR). Data represent mean ± SD of triplicates. (H) MDMs were infected with TsB7 at 33 or 39°C for 4 h, and viral DNA was visualized by HSV-1–specific FISH probes (green) and costained with anti-IFI16–specific Abs. Scale bar, 10 μm. Zoom is magnified 5× from original image. *p < 0.05.

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To further investigate the role of the nucleus in sensing HSV-1 DNA for stimulation of IFN-β expression, we used a temperature-sensitive HSV-1 mutant TsB7. TsB7 behaves as a wild-type virus at the permissive temperature of 33°C. However, at the nonpermissive temperature of 39°C, the TsB7 HSV-1 is unable to deliver its genome to the nucleus (36). We confirmed that infection with wild-type HSV-1 was able to stimulate expression of ICP27 at both 33 and 39°C, whereas infection with TsB7 led to ICP27 expression only at 33°C (Supplemental Fig. 3C). Detection of capsid-free TsB7 genomes in the cytosol was not affected following incubation at the nonpermissive temperature compared with the permissive temperature (Fig. 4D), further supporting the contention that HSV-1 DNA does not need to enter the nucleus prior to cytosolic delivery. In addition, nuclear exclusion of HSV-1 DNA at the nonpermissive temperature did not diminish, but in fact increased, the phosphorylation of IκBα, as well as the expression of IFN-β mRNA (Fig. 4E, 4F). At both the permissive and nonpermissive temperatures, no statistically significant differences were observed between the responses to wild-type and TsB7 HSV-1 (Fig. 4E, 4F). The IFN-β responses to the viruses at either temperature were not inhibited by pretreatment with inhibitory oligonucleotides, excluding a role for TLR9 in the innate response at the abnormal temperatures (data not shown).

In cells highly permissive for HSV-1 infection, such as Vero cells and U2OS cells, in which we observed viral DNA in the nucleus (Fig. 3D), we did not observe significant IFN-β induction by the live virus (data not shown). However, using UV-inactivated HSV-1, we observed an induction of the IFN-inducible gene ISG56 in U2OS cells (Fig. 4G). Importantly, the ISG56 response was increased in these cells at 39°C postinfection with HSV-1, but it was significantly inhibited postinfection with TsB7 (p = 0.003) (Fig. 4G).

Finally, HSV-1 DNA was detectable in the cytoplasm of the differentiated THP1 cells colocalizing with IFI16 postinfection with either HSV-1 or TsB7 at 33 or 39°C (Fig. 4H). These results demonstrate that recognition of HSV-1 DNA in the cytoplasm is responsible for IFN-β induction in macrophages.

The mechanism through which herpesvirus DNA is exposed for sensing by PRRs in the cytosol of myeloid cells is unclear. We analyzed the potential role of the ubiquitin–proteasome pathway in this process (37, 38). Prior to infection, ubiquitin was distributed throughout the cytosol and nucleus; the proteasome subunit p20S exhibited a punctuate distribution in the cytosol, with more pronounced staining in the nucleus; and no staining for Vp5 was observed (data not shown). Following infection with HSV-1, ubiquitin and p20S localized to discrete areas of the cytosol, with some HSV-1 capsid staining clearly colocalizing with ubiquitin and the proteasome (Fig. 5A). By counting >100 Vp5-positive foci from three experiments, we found that 18 ± 2.5% of Vp5 foci colocalized with p20S at 2 h postinfection.

FIGURE 5.

Ubiquitination of the HSV-1 capsid. (A) BMMs were infected with HSV-1 (MOI 3) for 2 h and costained with anti-Vp5, anti-ubiquitin (Ub), and anti-proteasome (p20S). White box indicates area displayed in zoom row (zoom is magnified 5× from original image). Arrows indicate the following colocalizations: 1, capsid and p20S; 2, capsid and Vp5; and 3, capsid p20S and Vp5. Scale bar, 20 μm. (B) Lysates from PMA-differentiated THP1 macrophages infected for 30 or 90 min with HSV-1 (MOI 300) were subjected to IP using anti-Vp5–coupled beads. Total ubiquitin and K48-coupled ubiquitin in the precipitate were detected using Western blotting. Levels of Vp5 and β-actin in the input lysate used for IP were determined by Western blotting. (C) Lysates from PMA-differentiated THP1 macrophages infected for 90 min with HSV-1 or the HSV-1 mutant TsB7 (MOI 300) at either 33 or 39°C were subjected to IP using anti–Vp5-coupled beads. Total ubiquitin in the precipitate was detected using Western blotting.

FIGURE 5.

Ubiquitination of the HSV-1 capsid. (A) BMMs were infected with HSV-1 (MOI 3) for 2 h and costained with anti-Vp5, anti-ubiquitin (Ub), and anti-proteasome (p20S). White box indicates area displayed in zoom row (zoom is magnified 5× from original image). Arrows indicate the following colocalizations: 1, capsid and p20S; 2, capsid and Vp5; and 3, capsid p20S and Vp5. Scale bar, 20 μm. (B) Lysates from PMA-differentiated THP1 macrophages infected for 30 or 90 min with HSV-1 (MOI 300) were subjected to IP using anti-Vp5–coupled beads. Total ubiquitin and K48-coupled ubiquitin in the precipitate were detected using Western blotting. Levels of Vp5 and β-actin in the input lysate used for IP were determined by Western blotting. (C) Lysates from PMA-differentiated THP1 macrophages infected for 90 min with HSV-1 or the HSV-1 mutant TsB7 (MOI 300) at either 33 or 39°C were subjected to IP using anti–Vp5-coupled beads. Total ubiquitin in the precipitate was detected using Western blotting.

Close modal

To examine whether the viral capsid was ubiquitinated, we precipitated Vp5 from HSV-1–infected THP1 macrophages and analyzed ubiquitination by Western blotting. Interestingly, although purified HSV-1 virions did not contain detectable levels of ubiquitin, Vp5 precipitated from infected macrophages did give rise to a clear signal for both total ubiquitin and K48-linkage polyubiquitin chains (Fig. 5B). Blotting with anti-K63 ubiquitin did not reveal detectable levels of this form of ubiquitin on either the virion capsid or capsids isolated from infected macrophages (data not shown). We used TsB7 to determine whether the capsid ubiquitination was indeed targeting the incoming capsids in the cytoplasm or empty capsids that had already delivered viral DNA to the nucleus. Importantly, we found that Vp5 ubiquitination, which was higher at 39°C compared with 33°C, was indistinguishable between HSV-1 and TsB7 (Fig. 5C).

To examine the importance of Vp5 ubiquitin and proteasomal colocalization for capsid degradation, the proteasome was inhibited with MG132. Readouts used were Vp5 levels in cytosolic extracts and quantification of Vp5-negative HSV-1 DNA spots in macrophages (as a measure of capsid-free DNA). Following infection of PMA-differentiated THP1 macrophages with HSV-1, intact Vp5 expression steadily decreased over 6 h (Fig. 6A). This decrease in expression could be halted by pretreatment of the cells with MG132 (Fig. 6A) and was also seen using two different proteasome inhibitors (Supplemental Fig. 4A). MG132 also extended the half-life of the minor CMV capsid protein p28 (Fig. 6B). Importantly, the ubiquitination of Vp5 was amplified if proteasome activity was inhibited by MG132 (Fig. 6C). The consequence of proteasomal degradation likely would be the release of viral DNA into the cytosol, and this was visualized using HSV-1–specific FISH probes (22, 39). To allow for the quantification of HSV-1 DNA release, the number of HSV-1 DNA foci was determined and expressed as HSV-1 DNA foci/100 cells, as above (Fig. 4B, 4C). Chemical inhibition of the proteasome did not affect the efficiency of viral entry, as determined by total DNA foci/100 cells (Fig. 6D), but it led to a significant decrease in the amount of capsid-free viral DNA in primary macrophages (Fig. 6E) and THP1 cells (data not shown). Importantly, MG132 treatment of macrophages prevented HSV-1–induced expression of ISG56 (Fig. 6F), which is driven by IRF-3 (40). This was observed postinfection at MOI 3 or 300 (Fig. 6F, data not shown). Inhibition of the proteasome did not lead to accumulation of viral DNA in the nucleus and did not affect HSV-1 ICP27 gene expression (data not shown). In contrast to the observations in macrophages, inhibition of the proteasome had only a minor effect on HSV-1–induced ISG56 expression in HFFs and U2OS cells (Fig. 6F, Supplemental Fig. 4B) in which ISG induction was dependent on delivery of the viral genome to the nucleus (Fig. 4G) (20).

FIGURE 6.

The HSV-1 capsid is degraded by the proteasome. (A and B) PMA-differentiated THP1 macrophages were untreated or pretreated with MG132 (10 μg/ml) and infected with HSV-1 or CMV (MOI 3). Total-cell lysates were isolated at the indicated time points postinfection, and the capsid protein Vp5 (A) or CMV p28 (B) was determined by Western blot. (C) Lysates from PMA-differentiated THP1 macrophages infected for 90 min with HSV-1 (MOI 300) in the presence or absence of MG132 were subjected to IP using anti-Vp5–coupled beads. Total and K48 ubiquitin in the precipitate were detected using Western blotting. (D and E) BMMs were pretreated with MG132 and infected with HSV-1 (MOI 3) for 4 h. The cells were fixed and probed with anti-Vp5 and a HSV-1–specific FISH probe. The data are presented as number of HSV-1 DNA foci/100 cells (D) and percentage HSV-1 DNA foci negative for Vp5 (E). Data represent mean ± SD. (F) PMA-differentiated THP1 macrophages or HFFs were treated with vehicle or pretreated with MG132 and infected with live (THP1 cells) or UV-inactivated (HFFs) HSV-1 (MOI 3) for 6 h. Total RNA was isolated for measurement of ISG56 mRNA (quantitative RT-PCR). Data represent mean ± SD of triplicates. *p < 0.05.

FIGURE 6.

The HSV-1 capsid is degraded by the proteasome. (A and B) PMA-differentiated THP1 macrophages were untreated or pretreated with MG132 (10 μg/ml) and infected with HSV-1 or CMV (MOI 3). Total-cell lysates were isolated at the indicated time points postinfection, and the capsid protein Vp5 (A) or CMV p28 (B) was determined by Western blot. (C) Lysates from PMA-differentiated THP1 macrophages infected for 90 min with HSV-1 (MOI 300) in the presence or absence of MG132 were subjected to IP using anti-Vp5–coupled beads. Total and K48 ubiquitin in the precipitate were detected using Western blotting. (D and E) BMMs were pretreated with MG132 and infected with HSV-1 (MOI 3) for 4 h. The cells were fixed and probed with anti-Vp5 and a HSV-1–specific FISH probe. The data are presented as number of HSV-1 DNA foci/100 cells (D) and percentage HSV-1 DNA foci negative for Vp5 (E). Data represent mean ± SD. (F) PMA-differentiated THP1 macrophages or HFFs were treated with vehicle or pretreated with MG132 and infected with live (THP1 cells) or UV-inactivated (HFFs) HSV-1 (MOI 3) for 6 h. Total RNA was isolated for measurement of ISG56 mRNA (quantitative RT-PCR). Data represent mean ± SD of triplicates. *p < 0.05.

Close modal

Cumulatively, these results indicate that HSV-1 capsids are subjected to ubiquitination in macrophages and degraded by the proteasome, leading to the release of viral DNA to the cytosol for recognition by PRRs.

DNA is a potent PAMP demonstrated to be involved in stimulation of both protective immunity to infections and excess inflammation (41, 42). Herpesviruses are DNA viruses that stimulate innate immune responses mainly through nucleic acids, and these events are important for prevention of disease (17, 22, 23, 41, 43, 44). Although pDCs sense herpesvirus DNA in endosomes via TLR9 (1517), recent reports demonstrate a role for IFI16-mediated sensing of herpesvirus DNA in the nucleus of nonimmune tissue cells (19, 21). However, it remains unresolved how herpesvirus DNA is exposed to PRRs for sensing in myeloid cells. Our present study demonstrates that the HSV-1 capsid becomes ubiquitinated in the cytoplasm of macrophages and degraded via the ubiquitin–proteasome pathway, hence exposing viral genomic DNA for recognition by DNA sensors. Thus, HSV-1 and CMV infection of macrophages leads to viral DNA recognition by DDX41 and IFI16, STING mobilization, and IFN-β expression.

It was reported that the proposed murine ortholog of IFI16, p204, is important for IFN induction during herpesvirus infection (22, 45), as well as that HEK293 cells overexpressing IFI16 induce IFN-β in an IFI16-dependent manner (19). Using siRNA, we provide the first data, to our knowledge, for a role for IFI16 in HSV-induced IFN-β induction in a primary human cell type. We also observed that IFI16 was constitutively expressed to high levels and localized mainly in the nucleus, with a significant subfraction localizing to the cytosol. IFI16 colocalized with viral DNA and relocated to the STING-positive foci, proposed to be key signaling platforms after DNA sensing (28). Using synthetic DNA, we observed IFI16 to associate with DNA very early after transfection and prior to colocalization with STING. These data strongly support the conclusion that IFI16 is a bona fide IFN-inducing DNA sensor involved in early detection of foreign DNA in myeloid cells. The data also argue against the previously proposed role for IFI16 as a secondary amplifier of the IFN response (23). It is of note that HSV-1–induced IFN-β expression was potently reduced by knockdown of either DDX41 or IFI16. This raises the question as to how these two DNA sensors interact and advocates for more knowledge on the mechanism of action of the DNA sensors with respect to both actual DNA recognition and signal transduction. In particular, it will be interesting to learn whether IFI16 and DDX41 work in the same pathway or in separate pathways merging on STING.

Two studies recently addressed the role of IFI16 in recognition of HSV-1 and Kaposi’s sarcoma–associated herpesvirus infections, and both reported that the nucleus was the site of herpesvirus DNA recognition (19, 21). These studies were both conducted in cells permissive for the viruses used, with accompanying nuclear localization of viral DNA and activation of viral gene expression. In this work, we were able to confirm the nuclear location of HSV-1 DNA in the permissive cell line Vero, which was in contrast to the cytoplasmic localization of viral DNA in the macrophages. Moreover, we found that nuclear delivery of viral DNA was essential for induction of ISGs in these cells. In contrast, in the macrophages, IFI16 associated with DNA in the cytoplasm in a manner not inhibited by the nuclear export inhibitor LMB and induced IFN-β expression in a manner independent of delivery of viral DNA to the nucleus. Thus, it seems that IFI16 can recognize DNA both in the nucleus and the cytoplasm and that the subcellular site of DNA recognition depends on the location to which DNA is delivered. In the case of macrophages, this is primarily the cytoplasm due to an active cytoplasmic system to detect and degrade herpesvirus capsids. It remains to be determined why permissive cells, such as Vero cells, U2OS cells, and HFFs, do not sense viral DNA in the cytosol after capsid degradation. One possibility is the lack of a capsid-sensing system or efficient viral evasion in these cell types. It also will be interesting to learn whether there are qualitative differences in the cellular response to DNA sensing in the nucleus versus the cytoplasm.

A key finding of the current study is the ubiquitination of the HSV-1 capsid and degradation of the major capsid protein Vp5 in a proteasome-dependent manner, leading to exposure of viral DNA, recognition by DNA sensors, and induction of innate responses, such as IFN-β. It was reported that the capsids of adeno-associated virus type 2 and 5 are ubiquitinated and that proteasome inhibitors increased transduction efficiency with these viruses (37). This indicates that the ubiquitin–proteasome pathway may represent an important antiviral pathway that both degrades viral capsids and exposes viral nucleic PAMPs to cytosolic PRRs for induction of IFNs. With the data from the present work and previous reports by other investigators, there is now information about how DNA derived from viruses, bacteria, and the host genome may be delivered to the cytoplasm for detection by DNA sensors and induction of protective and pathological responses (42, 46).

Pertel et al. (47) recently demonstrated that the cytosolic protein Trim5α, which has long been known to detect retroviral capsids and target them for degradation (38), also acts as a cytosolic PRR, stimulating ubiquitination of host proteins and establishment of an antiviral state. Our present data demonstrating that herpesvirus capsids become ubiquitinated and subject to proteasomal degradation strongly suggest that the herpesvirus capsid is also recognized within the cytosol. Interestingly, Trim5α was recently reported to inhibit HSV-1 and -2 replication at an early stage of the infection cycle (48), suggesting a role for this or a related protein in cytosolic sensing of herpesvirus capsids. Early work by Sodeik et al. (49) demonstrated that empty HSV-1 capsids (indicating delivery of DNA to the nucleus) start to accumulate inside Vero cells ∼1 h postinfection and account for 60% of all intracellular capsids at 4 h postinfection. Such data suggest that the cytosolic innate immune surveillance pathway has a time window of between 1 and 4 h to detect and degrade viral capsids to avoid nuclear entry and productive replication.

In conclusion, we propose that during herpesvirus infection in myeloid cells, the viral DNA is released into the cytosol after degradation of the viral capsid via the ubiquitin–proteasome pathway. This leads to cytosolic herpesvirus DNA sensing by IFI16 and IFN-β expression through the STING pathway. Together with the knowledge on herpesvirus DNA sensing in endosomes of pDCs and recent work on innate sensing of herpesviruses in the nucleus of nonimmune tissue cells (19), this work demonstrates that the subcellular localization of innate DNA sensing is highly cell-type specific and most likely is determined by the cellular sensing and trafficking systems, as well as the ability of microbes to interfere with these systems.

We thank Kirsten Stadel Petersen for technical assistance.

This work was funded by the Danish Medical Research Council (Grant 09-072636), the Novo Nordisk Foundation, the Velux Foundation, the Lundbeck Foundation (Grant R34-3855), Elvira og Rasmus Riisforts almenvelgørende Fond, and Fonden til Lægevidenskabens Fremme (all to S.R.P.), as well as by grants from the National Institutes of Health (AI083713 and AI083215 to K.A.F; DE018281 and CA019014 to B.D.). K.A.H. was funded by a Marie Curie Incoming International Fellowship.

The online version of this article contains supplemental material.

Abbreviations used in this article:

BMM

bone marrow–derived macrophage

cDC

conventional dendritic cell

FISH

fluorescence in situ hybridization

HFF

human foreskin fibroblast

IFI

IFN-γ–inducible gene

IP

immunoprecipitation

ISG

IFN-stimulated gene

LMB

Leptomycin B

MDM

monocyte-derived macrophage

MOI

multiplicity of infection

PAMP

pathogen-associated molecular pattern

pDC

plasmacytoid dendritic cell

PRR

pattern recognition receptor

shRNA

short hairpin RNA

siRNA

small interfering RNA

STING

stimulator of IFN genes.

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The authors have no financial conflicts of interest.