Endothelial IL-33 Expression Is Augmented by Adenoviral Activation of the DNA Damage Machinery

Tor Espen Stav-Noraas, Reidunn J. Edelmann, Lars La Cour Poulsen, Olav Sundnes, Danh Phung, Axel M. Küchler, Fredrik Müller, Amine A. Kamen, Guttorm Haraldsen, Mari Kaarbø and Johanna Hol
J Immunol April 15, 2017, 198 (8) 3318-3325; DOI: https://doi.org/10.4049/jimmunol.1600054

Abstract

IL-33, required for viral clearance by cytotoxic T cells, is generally expressed in vascular endothelial cells in healthy human tissues. We discovered that endothelial IL-33 expression was stimulated as a response to adenoviral transduction. This response was dependent on MRE11, a sensor of DNA damage that can also be activated by adenoviral DNA, and on IRF1, a transcriptional regulator of cellular responses to viral invasion and DNA damage. Accordingly, we observed that endothelial cells responded to adenoviral DNA by phosphorylation of ATM and CHK2 and that depletion or inhibition of MRE11, but not depletion of ATM, abrogated IL-33 stimulation. In conclusion, we show that adenoviral transduction stimulates IL-33 expression in endothelial cells in a manner that is dependent on the DNA-binding protein MRE11 and the antiviral factor IRF1 but not on downstream DNA damage response signaling.

Introduction

A member of the IL-1 family, IL-33 (1, 2) appears to be crucially involved in establishing a successful antiviral CD8 T cell response in the mouse (3). Viral infection also drives expression of IL-33 in many contexts. For example, murine lungs infected with influenza A show a dramatic increase in IL-33 expression (4), and patients with chronic viral hepatitis have elevated serum levels of IL-33 (5). These observations triggered interest in understanding how IL-33 expression is regulated at the cellular level. For example, transcription of IL-33 in murine macrophages partially depends on activation of IRF3 via the RNA sensor retinoic acid inducible gene I (6). IL-33 synthesis can also be triggered by detection of polyinosinic-polycytidylic acid (a synthetic analog of viral dsRNA) by TLR3 in murine hepatocytes (7) and human fibroblasts (8). In addition, synthesis of IL-33 is strongly boosted in human fibroblasts when polyinosinic-polycytidylic acid acts in concert with TGF-β (8).

Host recognition of viral infection involves several classes of sensors, including TLRs, C-type lectins, cytosolic RNA or DNA sensors, and the nuclear MRN complex (consisting of MRE11, NBS1, and RAD50) (9–11). This complex is well characterized as an initiator of the DNA damage response. The DNA damage response is crucial to prevent replication of damaged genomic host material, but it also serves to recognize foreign DNA. Human adenovirus 5 (Ad5) has a 36-kb dsDNA genome that is replicated concomitantly with cellular DNA. Thus, the discovery that Ad5 early proteins interfere with DNA damage response mediators elicited great interest, suggesting that the cellular DNA damage response also plays an antiviral role (discussed in Ref. 11). Indeed, adenovirus targets the MRN complex for proteasomal degradation by expressing the early proteins E1b55k/E4orf6 and E4orf3, thus limiting activation of the DNA damage machinery in response to adenoviral DNA (12). In the absence of adenoviral E4 proteins, MRN associates with viral DNA and initiates repair processes that result in tethering of viral linear DNA (concatemer formation) and prevent viral replication (12, 13).

Although the in vivo importance of IL-33 in antiviral defense was highlighted experimentally in mice (3), significant differences in the distribution of IL-33 between mouse and human may point to species-specific functions. For example, although IL-33 is nearly absent from vascular endothelial cells in the mouse, most IL-33 in healthy human tissues is found in the vasculature (14–16). It is unclear whether this vascular pool of IL-33 has a function in antiviral immune defense that cannot be accounted for in murine models.

We report that nonreplicative Ad5 (nrAd5) increases endothelial expression of IL-33 and initiates a DNA damage response. Depletion of MRE11 or IRF1 [essential transcriptional regulator of the DNA damage response (17)] prevented the observed stimulation of IL-33, implying that sensing of viral DNA by MRE11 boosts endothelial IL-33 expression.

Materials and Methods

Cell culture and reagents

Umbilical cords were obtained from the Department of Gynecology and Obstetrics at the Oslo University Hospital, according to a protocol approved by the Regional Committee for Research Ethics (S-05152a). HUVECs were isolated as described by Jaffe et al. (18) and cultured in MCDB 131 medium (Life Technologies) containing 7.5% FCS, 5 mM l-glutamine (Invitrogen), 10 ng/ml epidermal growth factor, 1 ng/ml basic fibroblast growth factor (both from R&D Systems), 1 μg/ml hydrocortisone (Sigma-Aldrich), 50 μg/ml gentamycin, and 250 ng/ml amphotericin B (both from Lonza), unless otherwise stated. Cells were used at passages one to six, maintained at 37°C in 95% humidity/5% CO2 atmosphere, and split at a 1:3 ratio. The γ-secretase inhibitor N-[N-(3,5-difluorophenacetyl-l-alanyl)]-S-phenylglycine t-butyl ester (DAPT; EMD Chemicals) was dissolved in DMSO (Sigma-Aldrich) at 25 mM and used at a final concentration of 5–25 μM. Cycloheximide and the MRN inhibitor mirin were purchased from Sigma-Aldrich and used at 3 μg/ml and 1–10 μM, respectively.

Abs

The Abs used in this study included Ad5 Hexon (8C4) and β-tubulin (both from Abcam) ATM pSer1981 (D6H9), ATM (D2E2), ATR pSer428, Chk1 pSer345 (133D3), Chk2 pThr68 (C13C1), H2A.X pSer139 (20E3), histone 3 (D1H2), IFI-16, IRF1 (D5E4), IRF3 (D6I4C), IRF7, MRE11, NICD1 Val1744 (D3B8), STAT1 pTyr701 (58D6), RAD50, STING (D2P2F), and TLR9 (D9MH9) (all from Cell Signaling Technologies); GAPDH (Santa Cruz Biotechnology); IFN-α/β receptor chain 2 (MMHAR-2) and STAT1 (STAT1-79) (both from Thermo Fisher Scientific); DLL4 (YW152F) and NOTCH1 (YW169.60.79) (both from Genentech); and IL-33 (Nessy-1) (Enzo Life Sciences). Secondary Abs were from Jackson ImmunoResearch. Additional details about Abs are in Supplemental Table I.

Amplification of Ad5ΔE1ΔE3-GFP, Ad5ΔE1ΔE3, and Ad5ΔE1 in mammalian cells

Human embryonic kidney 293T cells, transformed with adeno E1 and SV40 large T Ag, were used to amplify nrAd5. Ad5ΔE1ΔE3-GFP and Ad5ΔE1ΔE3 (AdEasy system; Stratagene) were transfected into 293T cells 24 h after seeding (8.0 × 104 cells per square centimeter) (i.e., at 50–70% confluence). A total of 4 μg of nrAd5 plasmid DNA (linearized with PacI) was transfected using Lipofectamine 2000 (Thermo Fisher Scientific), according to the manufacturer’s recommendation. Viral plaques were observed 7–10 d after transfection, and floating and adherent cells were collected by scraping. The cells were pelleted by centrifugation at 300 × g for 10 min at 4°C and resuspended in 2 ml of the supernatant. After three cycles of freezing/thawing (dry ice/methanol bath and rapid thawing at 37°C) and vortexing, the cell debris was pelleted by centrifugation at 3000 × g for 20 min at 4°C. The viral lysates were used to infect 293T cells (70% confluent, 1 ml lysate per T25 tissue culture flask). Three to five days postinfection, when cytopathic effects were observed in 30–50% of the cells, viruses were harvested as described above. The viral titers were determined by infecting 293T cells with 10-fold dilutions (from 10−2 to 10−9) of virus stocks, culturing the cells for 48 h, and harvesting by fixation in 100% methanol (−20°C for 15 min). Endogenous peroxidase was quenched with 0.3% H2O2 in water for 30 min. Cells were washed in 1% BSA (Sigma-Aldrich) diluted in PBS. Plaques were stained with murine mAb specific for the adenovirus hexon protein (Supplemental Table I, using 1 μg/ml in 1% BSA diluted in PBS) for 1 h at 37°C. After washing, the cells were incubated with HRP-conjugated goat anti-mouse IgG (Supplemental Table I, 0.8 μg/ml in 1% BSA diluted in PBS) for 1 h at 37°C. The cells were washed prior to 3,3′-diaminobenzidine staining using Fast DAB tablets, according to the manufacturer’s recommendation (Sigma-Aldrich). Positive plaques were counted in a minimum of three fields per well per dilution in duplicate wells. The average titer was determined as PFU per milliliter. Helper-dependent nrAd5 lacking all adenoviral genes was produced as described by Dormond et al. (19).

Viral transduction

For transduction, HUVECs were seeded in complete medium at 3.8 or 1.9 × 104 cells per square centimeter, 24 or 48 h prior to infection, respectively. On the day of infection, when cells were subconfluent (70–80%) or confluent (90–100%), the complete medium was replaced with fresh complete medium, and viral stocks were added to obtain the desired multiplicity of infection (moi). Viral UV inactivation was performed by diluting the virus stock in 150 μl of complete medium in 24-well plates, followed by irradiation on ice, using different doses of UV (ultraviolet) light to a maximum of 7 J (760 μW/cm2, up to a maximum of 2 h and 30 min).

IFN-α/β neutralization

In vitro blocking of the IFN-α/β receptor was performed with a murine mAb to human IFN-α/β receptor chain 2 (MMHAR-2, 10 μg/ml). A species-, isotype-, and concentration-matched mAb against the E-tag epitope (Supplemental Table I, 10 μg/ml) was used as a negative control. Ad5ΔE1ΔE3-GFP (50 moi) was added to the culture, and the cells were incubated for 24 h. The results were compared with those from nontreated and nontransduced cells. To test the efficacy of the neutralizing Ab, CXCL10 was measured in isotype- and MMHAR-2–treated cells stimulated with 1000 U/ml IFN-α.

Quantitative RT-PCR

Upon harvest, total RNA was extracted from cells using TRIzol Reagent (Thermo Fisher Scientific), according to the manufacturer’s recommendations. One microgram of RNA was used for first-strand cDNA synthesis with the SuperScript III Reverse Transcriptase cDNA system (Thermo Fisher Scientific), oligo(dT) primers, and dNTPs (GE Healthcare). Quantitative RT-PCR (qRT-PCR) was carried out on a Stratagene Mx3005P instrument and analyzed by Stratagene MxPro software (both from Agilent Technologies). The PCR reaction consisted of 5 μl of 10-times diluted cDNA in a 20-μl qPCR reaction consisting of 5000 U of HotStarTaq DNA polymerase (QIAGEN), dNTP (GE Healthcare), 20× EvaGreen (Biotium), and the individual primer sets. The PCR was run up to 40 cycles and included a melting curve analysis to ensure amplification of single products. Standard curves were made from serial dilutions of cDNA to calculate primer efficiencies. HPRT was used to normalize for sample-to-sample variation. The relative gene expression levels were calculated using the Pfaffl method (when primer efficiencies differed) (20) or the comparative Ct method (21) relative to nontreated controls. The primer sets used were: HEY1: F, 5′-GCTGGTACCCAGTGCTTTTGAG-3′, R, 5′-TGCAGGATCTCGGCTTTTTCT-3′; HES1: F, 5′-ACGTGCGAGGGCGTTAATAC-3′, R, 5′-CATGGCATTGATCTGGGTCA-3′; HPRT: F, 5′-AATACAAAGCCTAAGATGAGAGTTCAAGTTGAGTT-3′, R, 5′-CTATAGGCTCATAGTGCAAATAAACAGTTTAGGAAT-3′; IL-33: F, 5′-GCAGCTCTTCAGGGAAGAAATC-3′, R, 5′-TGTTGGGATTTTCCCAGCTTGA-3′; NOTCH1: F, 5′-CGGGTCCACCAGTTTGAATG-3′, R, 5′-GTTGTATTGGTTCGGCACCAT-3′; and DLL4: F, 5′-GAAGTGGACTGTGGCCTGGACAAGT-3′, R, 5′-TCGCTGATATCCGACACTCTGGCT-3′.

Small interfering RNA transfection

HUVECs were seeded at a density of 3.8 or 1.9 × 104 cells per square centimeter in complete medium 24 h prior to transfection. The transfection was carried out in medium without antibiotics. Lipofectamine/small interfering RNA (siRNA) mix (Lipofectamine 2000 or Lipofectamine RNAiMAX, respectively; Thermo Fisher Scientific) was prepared according to the manufacturer’s instructions, added to the cells, and subsequently incubated for 6 h. The following Ambion Silencer Select siRNAs were purchased from Thermo Fisher Scientific: IRF1 s4502, IRF-3 s7507, IL-33 s40521, NOTCH1 s9633, DLL4 s29213, JAG1 s1175, IFI-16 s7136, TLR9 s28872 and s28873, MRE11 s8960, NBS1 (NLRP2) s31177, RAD50 s793, STING s50644 (STING1) and s50645 (STING2), and ATM s1710. Silencer select Scrambled #1 and Scrambled #2 (Thermo Fisher Scientific) were used to control for nonspecific effects of transfection.

Immunoblotting

Cultured cells were washed with PBS before harvesting samples in a Tris-HCl (pH 6.8) SDS (2.5%)/glycerol (10%) lysis buffer containing a reducing agent (100 mM 2-ME; Sigma-Aldrich or 10 mM DTT), protease inhibitors (1 mM phenylmethylsulfonyl fluoride [Sigma-Aldrich], complete Protease Inhibitor Cocktail [Roche]), and phosphatase inhibitor (2 nM sodium orthovanadate; Sigma-Aldrich). The samples were homogenized using a QIAshredder (QIAGEN), according to the manufacturer’s instructions, and incubated at 95°C for 5 min before loading them onto 10% Mini-PROTEAN TGX Precast Gels, 4–20% Mini-PROTEAN TGX Precast Protein Gels (both from Bio-Rad), or 12.5% SuperSep Phos-tag gels (Wako Pure Chemical Industries). The Phos-tag gels were used according to the manufacturer’s instructions to evaluate the phosphorylation of IRF1. After loading cell lysates onto gels, they were run for 15–25 min at 300 V before blotting to nitrocellulose membranes using the Trans-Blot Turbo Transfer System and Turbo blotter (both from Bio-Rad). Blotted membranes were blocked with 5% Blotting-Grade Blocker (Bio-Rad) or 5% BSA (when using Abs specific for phosphorylated proteins, Supplemental Table I) and were incubated with primary (4°C, overnight) and secondary (room temperatures, 2 h) Abs diluted in 1% Blotting-Grade Blocker (Bio-Rad) in TBST. The protein bands were detected using Pierce Super Signal West Dura Extended Duration Substrate (Thermo Fisher Scientific) and visualized using a ChemiDoc MP System (Bio-Rad). All protein mass indications (kDa) in the figures were assigned according to the protein ladder (Full-Range Rainbow Molecular Weight Markers, GE Healthcare).

Image processing

Figures and images were generated and processed in Adobe Photoshop CS6, Adobe Illustrator CS6, GraphPad Prism 6, or FlowJo Vx. All adjustments were performed on the image as a whole and with equal adjustment of image series.

Results

Endothelial IL-33 expression is enhanced by replication and transcription–deficient Ad5

While using a nonreplicative adenoviral vector as a tool to ectopically express IL-33 in cultured HUVECs, we observed that transduction with a control vector (Ad5ΔE1ΔE3) markedly increased IL-33 expression 48 h posttransduction (hpt) compared with nontransduced cells (Fig. 1A). The stimulation of IL-33 correlated positively with increasing viral titers. In contrast to nontransduced HUVECs that require contact-mediated quiescence to express IL-33 (14, 22), Ad5ΔE1ΔE3-transduced cells expressed IL-33 in confluent and subconfluent cultures (Fig. 1A). The increase in IL-33 expression was evident at 48 hpt and continued until 72 hpt (Fig. 1B). To establish whether the increased endothelial expression of IL-33 was due to viral gene transcription, HUVECs were transduced with a helper-dependent nrAd5 lacking all adenoviral genes but retaining the cis-regulatory elements, including the viral-packaging signals and the inverted terminal repeats (19, 23). We found that helper-dependent nrAd5 also enhanced IL-33 expression (Fig. 1C), concluding that IL-33 can be induced by nrAd5 vectors in human endothelial cells in a manner that is independent of viral transcription.

FIGURE 1.
FIGURE 1.

Endothelial IL-33 expression is enhanced by replication and transcription–deficient Ad5. HUVECs were transduced with Ad5ΔE1ΔE3 (increasing titers and 10 moi) (A and B) or helper-dependent nrAd5 (HdAd5, low to high moi) (C) for 48 h (A and C) or for the indicated time (B) before harvesting cellular lysates for immunoblotting with Abs specific for IL-33 (Nessy-1; Enzo Life Sciences) and tubulin. Net luminescence of bands corresponding to IL-33 in (A) and (B) were quantified and normalized to the loading control. The amount of IL-33 in control cells (mock) was set to 1, and the average fraction of three independent experiments was plotted showing the mean ± SD. The data shown are representative of three independent experiments.

IL-33 upregulation is abrogated by UV irradiation of viral particles

Although elicitation of IL-33 was independent of viral gene transcription, UV irradiation of Ad5ΔE1ΔE3-GFP (before adding the virus to cells) dose dependently abrogated the stimulation of IL-33 expression (Fig. 2A). Detection of virally driven GFP by flow cytometry was used to control for viral inactivation, showing a steady, inverse correlation with the dose of UV light applied (Fig. 2B). This shows that adenoviral entry alone is insufficient to stimulate IL-33 expression in endothelial cells and that the host response involved in IL-33 augmentation is not triggered by UV-inactivated virus particles.

FIGURE 2.
FIGURE 2.

IL-33 upregulation is abrogated by UV irradiation of viral particles. (A) HUVECs were transduced with Ad5ΔE1ΔE3-GFP (10 moi) that had been UV irradiated before adding the viral particles to the cells, harvested after 48 h, and analyzed by immunoblotting with Abs specific for IL-33 and tubulin. Net luminescence of the bands corresponding to IL-33 was quantified and normalized to the loading control. The amount of IL-33 in control cells (mock) was set to 1, and the average fractions of three independent experiments were plotted showing the mean ± SD. (B) Transduced HUVECs from wells parallel to those sampled in (A) were harvested for flow cytometry as a control for UV inactivation of Ad5ΔE1ΔE3-GFP. Gating for HUVECs was performed according to size (forward and side scatter).

Adenoviral upregulation of IL-33 depends on Notch signaling

Our recent finding that Notch signaling drives IL-33 expression in quiescent endothelial cells (22) prompted us to ask whether nrAd5 transduction might drive IL-33 expression via Notch signaling. Transcriptional analysis of HUVECs transduced with Ad5ΔE1ΔE3 revealed an increase in mRNA levels of the Notch ligand DLL4, the Notch receptor NOTCH1, and the direct Notch target genes HES1 and HEY1 (Fig. 3A). In addition, the levels of cleaved NOTCH1 intracellular domain (csNICD1, the signaling mediator of activated NOTCH1) were increased after transduction (Fig. 3B). Moreover, Notch signaling was required for the nrAd5-driven increase in IL-33 to take place, because IL-33 expression could be inhibited by siRNA-mediated knockdown of Notch components (Fig. 3C), by the γ-secretase inhibitor DAPT (Fig. 3D), or by inhibitory Abs to NOTCH1 or DLL4 (Fig. 3D) in nontransduced and Ad5ΔE1ΔE3-transduced cells. These data demonstrate that endothelial Notch signaling is increased by nrAd5 transduction and confirm that NOTCH1 strongly supports IL-33 expression, also when enhanced by nrAd5.

FIGURE 3.
FIGURE 3.

Adenoviral upregulation of IL-33 depends on Notch signaling. HUVECs were transduced with Ad5ΔE1ΔE3 (10 moi) for 0–72 h (A) or for 48 h (B–D) before harvest and analysis by qPCR (A) or immunoblotting (B–D). (A) Transcription levels of Notch components and target genes in Ad5ΔE1ΔE3-transduced HUVECs. Graphs show the mean ± SEM. (B) Levels of active (cleaved) NOTCH1 (csNICD1) in Ad5ΔE1ΔE3-transduced HUVECs. (C) HUVECs were transfected with siRNA targeting NOTCH1, DLL4, and IL-33 (as a positive control) 24 h before transduction with Ad5ΔE1ΔE3 and were analyzed for csNICD1 and IL-33 to assess the effect of Notch inhibition on nrAd5-stimulated IL-33. (D) Neutralizing Abs specific for NOTCH1 (0.3 μg/ml) and DLL4 (0.3 μg/ml), isotype-matched control IgG (0.3 μg/ml), and the γ-secretase inhibitor DAPT (5 μM) were administered 15 min before transduction with Ad5ΔE1ΔE3, and lysates were analyzed for expression of csNICD1 and IL-33. The net luminescence of bands corresponding to csNICD1 and IL-33 were quantified and normalized to the loading control. The amounts of csNICD1 and IL-33 in control cells (mock) were set to 1, and the average fractions of two independent experiments were plotted showing the mean. The data shown are representative of three (A–C) or two (D) independent experiments.

Adenoviral stimulation of IL-33 depends on the antiviral transcription factor IRF1

The ability of adenoviral transduction to stimulate IL-33 expression, even in subconfluent endothelial cell cultures, implicated a mechanism that extends beyond the activation level of Notch signaling. Therefore, we assessed the involvement of transcription factors that are commonly involved in regulating the expression of antiviral genes. We found IRF3 and IRF1 to be constitutively present in the nuclear fraction of HUVECs, whereas IRF7 was undetectable throughout the course of adenoviral stimulation (Fig. 4A). Therefore, we depleted IRF3 and IRF1 using siRNA (Fig. 4B, 4C, Supplemental Fig. 1C); although the reduction of IRF3 did not affect IL-33 levels, reduction of IRF1 abrogated IL-33 expression in nontransduced and Ad5ΔE1ΔE3-transduced HUVECs. We were unable to detect any change in phosphorylation status or half-life of IRF1 following transduction by Ad5ΔE1ΔE3 (Supplemental Fig. 1A, 1B), suggesting that IRF1 is activated by a phosphorylation-independent mechanism by Ad5ΔE1ΔE3 or that its activity is not altered and IRF1 plays a permissive role in IL-33 expression. The dynamics of IRF1 degradation after cycloheximide treatment was similar to that observed by other investigators (24), indicating a half-life ∼ 30 min.

FIGURE 4.
FIGURE 4.

Adenoviral stimulation of IL-33 depends on the antiviral transcription factor IRF1. HUVECs were transduced with Ad5ΔE1ΔE3 (10 moi) for the indicated times (A and D) or for 48 h (B, C, E, and F) before harvest of nuclear and cytoplasmic fractions (A) or whole-cell extracts (B–D) and analysis by immunoblotting with Abs as indicated or qPCR (E and F). (A) Levels of IRF7, IRF1, and IRF3 in cytoplasmic and nuclear fractions of HUVECs transduced with Ad5ΔE1ΔE3. Lysates of HUVECs stimulated with TNF-α (10 ng/ml for 2 h) or IFN-α (100 ng/ml for 4 h) were included as positive controls for IRF expression. Levels of IL-33 after siRNA-mediated depletion of IL-33 (as a positive control) and IRF3 (B) or IRF1 (C) 24 h before transduction with Ad5ΔE1ΔE3. (D) Levels of p-STAT1 and STAT1 in Ad5ΔE1ΔE3-transduced HUVECs. Net luminescence of bands corresponding to IL-33 and p-STAT1 were quantified and normalized to the loading control. The amount of IL-33 and p-STAT1 in control cells (mock) were set to 1, and the average fractions of three independent experiments were plotted showing the mean ± SD. Protein mass indications were set according to the protein ladder. The IRF7 band is approximately 55 kDa and runs between the 76 and 52 kDa marker. (E) A neutralizing Ab specific for the INF-α/β receptor (MMHAR-2) or a negative-control Ab was administered to HUVECs 30 min before transduction with Ad5ΔE1ΔE3. IL33 expression was analyzed by qPCR. (F) Supernatants from Ad5ΔE1ΔE3-transduced HUVECs 24 hpt were transferred to nontransduced subconfluent cells. Cells were harvested after another 24 h and analyzed by qPCR. Fresh growth medium was used as control. The bar graph shows the mean ± SD, but the SD is not visible because of low variation. IL33 mRNA expression is presented relative to control cells, with HPRT as a reference gene. The data shown are representative of three independent experiments.

IRF1 can be activated in response to cytosolic DNA in several cell types (25, 26), has powerful cell-intrinsic antiviral properties (25), and can also be activated by type I IFNs (27). Therefore, to test the involvement of IFN and possible auto/paracrine effects, we assessed phosphorylation of the essential IFN-activating transcription factor STAT1; it was phosphorylated at an earlier time point than IL-33 was upregulated after transduction of HUVECs with Ad5ΔE1ΔE3 (Fig. 4D). Considering the possibility of IFN signaling, we exposed cells to an Ab specific for the IFN-α/β receptor chain 2 during infection. This reagent neutralizes the effect of seven type I IFNs (28); in our hands, it reduced IFN-α–driven induction of CXCL10 in control cells (data not shown), yet it failed to reduce the viral stimulation of IL33 expression (Fig. 4E). Finally, we assessed the possible involvement of other soluble factors by exposing nontransduced cells to supernatants harvested from transduced cell cultures (Fig. 4F), again observing no increase in IL33 expression. Taken together, these findings indicate that endothelial expression of IL-33 is supported by the presence of the antiviral transcription factor IRF1 but not by IRF3 or by the auto/paracrine stimulation of soluble mediators, such as type I IFNs.

Nonreplicative adenovirus activates the endothelial DNA damage response

Because IRF1, in addition to its role in innate immune responses, is closely linked to the DNA damage response (17, 29), we next evaluated whether the DNA damage response was activated in our system. Endothelial cells transduced with Ad5ΔE1ΔE3 responded by inducing elements of a DNA damage response at 24 hpt (Fig. 5A). ATM and CHK2 were phosphorylated after transduction with Ad5ΔE1ΔE3 (Fig. 5A, 5B), correlating in time with the upregulation of IL-33. However, phosphorylation of histone H2AX was not observed, in line with a previous report showing that ATM activation by adenoviral DNA is not accompanied by an extensive amplification by pH2AX, most likely due to the limited size of the adenoviral DNA (30).

FIGURE 5.
FIGURE 5.

nrAd5 activates an MRE11-dependent DNA damage response in HUVECs, and MRE11 mediates the nrAd5 stimulation of IL-33. HUVECs were transduced with Ad5ΔE1ΔE3 (10 moi) for 0–72 h (A and B) or for 48 h (C) before harvesting and analyzing by immunoblotting, as designated. (A) DNA damage components are activated in HUVECs in response to nrAd5. (B) The net luminescence of the bands in (A) was quantified and normalized to the loading control. The average amount of luminescence relative to control cells (mock) of three individual experiments was plotted. (C) ATM, RAD50, NBS1, MRE11, or IRF1 was depleted using siRNA 24 h before transduction with Ad5ΔE1ΔE3. The outlined box emphasizes the effect of MRE11 knockdown in Ad5-transduced cells. The asterisk (*) indicates proteins that were reduced after MRE11 depletion (p-ATM, IL-33, RAD50, MRE11). (D) To inhibit MRE11 endonuclease activity, HUVECs were treated with mirin at the indicated concentrations, together with nrAd5, as designated. (E) STING was depleted using two siRNAs (STING1 and STING2) 24 h before transduction with Ad5ΔE1ΔE3 (10 moi). Protein mass indications were set according to the protein ladder. ATM bands run above the top maker (225 kDa). The net luminescence of the bands in (D) and (E) was quantified and normalized to the loading control. The average amount of luminescence relative to control cells (mock) of three individual experiments was plotted. The data shown are representative of three independent experiments.

The DNA damage machinery component and dsDNA sensor MRE11 is required for viral IL-33 stimulation

Activation of the DNA damage response in adenovirally transduced cells is initiated by MRE11, the DNA binding component of the MRN complex, which, in the absence of the early adenoviral protein E4 (not expressed by replication-deficient viral vectors), is reported to associate with viral DNA in nuclear-replication centers (13, 30, 31). Therefore, we targeted the MRN components MRE11, NBS1, and RAD50, as well as the downstream kinase ATM, via siRNA-mediated knockdown before transduction with nrAd5, observing that depletion of MRE11 abrogated stimulation of IL-33 expression and reduced the basal expression of IL-33 in nontransduced cells (Fig. 5C). In accordance with previous studies, MRE11-depleted cells also showed reduced levels of RAD50 and p-ATM (32, 33). Depleting RAD50 reduced phosphorylation of ATM, but did not affect IL-33 expression. In addition, inhibition of MRE11 nuclease activity by mirin reduced IL-33 expression, confirming the role of MRE11 in IL-33 stimulation (Fig. 5D). Mirin also inhibited IL-33 expression in the absence of nrAd5 (Fig. 5D). Together with the reduction of IL-33 observed in nontransduced cells when MRE11 was depleted by siRNA (Fig. 5C), this suggests a low-level activation of MRE11 in nontransduced confluent endothelial cell cultures that also contributes to the constitutive expression of IL-33. When MRE11 acts as a cytoplasmic sensor of dsDNA (32), it activates IRF3 via the endoplasmic reticulum–resident protein STING. However, siRNA-mediated knockdown of STING did not affect IL-33 expression in response to nrAd5 transduction (Fig. 5D). Likewise, knockdown of another nuclear sensor of foreign DNA, IFI-16, did not affect IL-33 expression (Supplemental Fig. 1C). Taken together, our observations suggest that MRE11-mediated sensing of nuclear adenoviral DNA promotes IRF1-driven stimulation of IL-33 in primary human endothelial cells.

Discussion

This study shows that expression of IL-33 in human endothelial cells is upregulated by the nuclease activity of DNA-binding MRE11 in response to transduction of adenoviral vectors. Our observation that IL-33 expression was enhanced by transduction with Ad5ΔE1ΔE3, as well as by helper-dependent nrAd5 that lacks all viral genes, implicates the involvement of a viral structure common to these constructs. Adenoviral entry and the cytoplasmic presence of viral capsid proteins may, in itself, trigger cellular responses, even when viral DNA is absent (34). However, UV irradiation of virus particles before transduction abrogated IL-33 stimulation, suggesting that viral entry is insufficient to trigger the response and that intact viral DNA is required. Interestingly, although most viral DNA has been removed from helper-dependent nrAd5, the construct still contains viral packaging signals and inverted terminal repeats (23). Such terminal repeats are also expressed by adeno-associated viral vectors and are believed to represent a favored recognition site for the MRN complex (35). Therefore, the ability of helper-dependent nrAd5 to enhance IL-33 expression is in line with our finding that MRE11, the DNA-binding component of MRN, is crucial for the stimulation of IL-33 production observed in endothelial cells transduced with adenoviral vectors.

We also discovered that the well-known antiviral transcription factor IRF1 is essential for maintaining basal and adenovector-induced expression of IL-33 in endothelial cells. IRF1 was shown to inhibit a wide range of viruses in a manner preserved in STAT1-deficient fibroblasts (25, 36); therefore, the response appears to be cell intrinsic rather than driven by IFN production. Interestingly, other IRFs are capable of inducing IL-33 in nonendothelial cells: IRF3 is required for transcription of IL-33 in murine macrophages upon nucleic acid ligand transfection and viral infection (6), IRF7 is required for induction of IL-33 by serum amyloid protein in human and murine monocytes (37), and IRF4 is essential for IL-33 induction in mice exposed to house dust mite allergen (38). Indeed, IRF4 was shown to bind within the first intron of the IL-33 gene in murine dendritic cells (38), supporting the concept that IL-33 can be regulated by IRF binding. Although all of these IFN regulatory factors have similar DNA-binding properties (39) and may regulate IL-33 gene transcription in a similar manner, they appear to differ with respect to cell type specificity and milieus.

Although IRF1 was required for the IL-33 response induced by nrAd5, the total levels of IRF1 remained constant during the course of viral transduction. Furthermore, we could not detect phosphorylated forms or an increase in the t1/2 of IRF1. Therefore, it possible that IRF1 is constitutively active in endothelial cells and permits IL-33 expression without further activation. In contrast, IRF1 activity can also be influenced by factors not addressed in this study, including antagonistic action by IRF2 (40) and posttranslational modifications different from phosphorylation; hence, our results do not fully eliminate the possibility that IRF1 activity is altered upon adenovirus transduction in endothelial cells. It should also be noted that our approach to detect phosphorylation of IRF1 lacked a positive control. The involvement of Notch signaling in the stimulation of IL-33 also tempts us to speculate that IRF1 may form a transcriptional complex with the canonical Notch transcription factor RBP-jK in an enhanceosome similar to that described for IRF1 and NF-κB (41). Such interactions deserve further investigations.

The involvement of IRF1 and the observation that adenoviral DNA in the absence of the adenoviral protein E4 elicits a cellular DNA damage response (12, 30), led us to explore the DNA damage response pathway in endothelial cells. Indeed, we observed that transduction with nrAd5 induced the activation of ATM but prevented the phosphorylation of H2AX, in line with recent findings in small airway epithelial cells (30). Furthermore, the dynamics of IL-33 stimulation coincided with phosphorylation of ATM, which, in the context of adenoviral transduction, represents a downstream event to recognition of viral DNA by the MRN complex. We next depleted the DNA-binding MRN component MRE11 by siRNA and observed a reduction in DNA damage response signaling and IL-33 expression (Fig. 5C). To confirm our data in a siRNA-independent manner, we also treated cells with mirin, an inhibitor of MRE11 nuclease activity, and observed a similar attenuation of IL-33 expression as when cells were treated with siRNA targeting MRE11. In contrast to recently described STING/IRF3-dependent MRE11 signaling in response to cytoplasmic dsDNA (32), IL-33 stimulation by adenoviral DNA did not require STING or IRF3.

Cellular secretion of IL-33 is still not fully understood (42). Considering our findings in light of the recent discovery that IL-33 is required for a successful host response to viral infections (3, 43) and the fact that most IL-33 in the human body is found within nuclei of vascular endothelial cells (14, 15), the question of whether IL-33 can be released to the extracellular space from vascular endothelial cells in the absence of cell death appears more relevant than ever. Full-length IL-33 can be detected in supernatants of cultured endothelial monolayers after scratching (44) or in response to in vitro cold ischemia and reperfusion (45). However, both of these experimental approaches presumably bring about a significant degree of cell death, and the detected IL-33 could be a result of passive release from necrotic cells. In contrast, efforts to demonstrate active IL-33 secretion from cultured endothelial cells have been unfruitful. Interestingly, endothelial cells appear to be an important source of extracellular IL-33 in the mouse heart during pressure overload, where it engages in cardioprotective mechanisms (46), thus supporting a model in which IL-33 can be released extracellularly from endothelial cells under some circumstances. Therefore, it is urgent to address whether IL-33 can undergo regulated secretion from human endothelial cells in a viral context.

The novel connection between IL-33 and the DNA damage response, together with its conserved relationship with IRFs, also makes it tempting to speculate whether IL-33 may possess cell-intrinsic properties that could influence the outcome of viral infections. The nuclear effects of IL-33 remain ill-defined; however, IL-33 is predicted to bind an acidic pocket of the nucleosome that can also harbor the latency-associated nuclear Ag of Kaposi sarcoma–associated herpesvirus (47). Similar to other proteins that dock into this pocket (48), IL-33 appears to modulate chromatin condensation (47, 49) and was reported to associate with the transcriptional repressor and histone methyltransferase SUV39H1 (50). Chromatin-remodeling factors play an active role in the fine-tuning of DNA damage responses (51) and contribute significantly to host–viral interactions that ultimately determine the outcome of viral infections (52). Therefore, further experiments should be designed to determine whether IL-33 associates with the DNA damage machinery and/or viral replication centers and whether IL-33 expression affects viral replication or persistence.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank Veit Hornung and Magnar Bjørås for scientific guidance and discussions. We also thank Eirill Ager-Wick, Linda Solfjell, Kathrine Hagelsteen, and Filip Nicolaysen for excellent technical assistance.

Footnotes

  • This work was supported by grants from Helse Sør-Øst (2010051, 2010019, 2013115, 2014032), the Research Council of Norway (221929/F20), and the University of Oslo (131406).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    Ad5
    adenovirus 5
    csNICD1
    cleaved NOTCH1 intracellular domain
    DAPT
    N-[N-(3,5-difluorophenacetyl-l-alanyl)]-S-phenylglycine t-butyl ester
    hpt
    h posttransduction
    moi
    multiplicity of infection
    nrAd5
    nonreplicative Ad5
    qRT-PCR
    quantitative RT-PCR
    siRNA
    small interfering RNA.

  • Received January 12, 2016.
  • Accepted February 6, 2017.

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