
==== Front
Nucleic Acids Res
Nucleic Acids Res
nar
Nucleic Acids Research
0305-1048
1362-4962
Oxford University Press

39087523
10.1093/nar/gkae654
gkae654
AcademicSubjects/SCI00010
Genome Integrity, Repair and Replication
Loss of the DNA repair protein, polynucleotide kinase/phosphatase, activates the type 1 interferon response independent of ionizing radiation
https://orcid.org/0000-0003-3529-9383
Kate Wisdom Deebeke Department of Oncology, University of Alberta, Edmonton, Alberta T6G 2R3, Canada

Fanta Mesfin Department of Oncology, University of Alberta, Edmonton, Alberta T6G 2R3, Canada

https://orcid.org/0000-0002-8773-1425
Weinfeld Michael Department of Oncology, University of Alberta, Edmonton, Alberta T6G 2R3, Canada

To whom correspondence should be addressed. Tel: +1 780 432 8438; Fax: +1 780 432 8428; Email: mweinfel@ualberta.ca
09 9 2024
01 8 2024
01 8 2024
52 16 96309653
12 7 2024
07 6 2024
29 12 2023
© The Author(s) 2024. Published by Oxford University Press on behalf of Nucleic Acids Research.
2024
https://creativecommons.org/licenses/by-nc/4.0/ This is an Open Access article distributed under the terms of the Creative Commons Attribution-NonCommercial License (https://creativecommons.org/licenses/by-nc/4.0/), which permits non-commercial re-use, distribution, and reproduction in any medium, provided the original work is properly cited. For commercial re-use, please contact reprints@oup.com for reprints and translation rights for reprints. All other permissions can be obtained through our RightsLink service via the Permissions link on the article page on our site-for further information please contact journals.permissions@oup.com.

Abstract

DNA damage has been implicated in the stimulation of the type 1 interferon (T1IFN) response. Here, we show that downregulation of the DNA repair protein, polynucleotide kinase/phosphatase (PNKP), in a variety of cell lines causes robust phosphorylation of STAT1, upregulation of interferon-stimulated genes and persistent accumulation of cytosolic DNA, all of which are indicators for the activation of the T1IFN response. Furthermore, this did not require damage induction by ionizing radiation. Instead, our data revealed that production of reactive oxygen species (ROS) synergises with PNKP loss to potentiate the T1IFN response, and that loss of PNKP significantly compromises mitochondrial DNA (mtDNA) integrity. Depletion of mtDNA or treatment of PNKP-depleted cells with ROS scavengers abrogated the T1IFN response, implicating mtDNA as a significant source of the cytosolic DNA required to potentiate the T1IFN response. The STING signalling pathway is responsible for the observed increase in the pro-inflammatory gene signature in PNKP-depleted cells. While the response was dependent on ZBP1, cGAS only contributed to the response in some cell lines. Our data have implications for cancer therapy, since PNKP inhibitors would have the potential to stimulate the immune response, and also to the neurological disorders associated with PNKP mutation.

Graphical Abstract

Graphical Abstract

Cancer Research Institute 10.13039/100000884 CRI3714 Canadian Institutes of Health Research 10.13039/501100000024 PJT168869 Terry Fox Foundation – Strategic Training in Transdisciplinary Radiation Science for the 21st Century STARS21 Alberta Cancer Foundation 10.13039/501100000001 University of Alberta 10.13039/501100000190
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pmcIntroduction

The type-I interferon (T1IFN) response is a conserved anti-viral and anti-bacterial defense mechanism initiated by the host cells’ pattern recognition receptors, such as cyclic guanosine monophosphate (GMP)–adenosine monophosphate (AMP) synthase (cGAS) or toll-like receptors, following detection of danger-associated molecular patterns (DAMPs) (1–5). Once engaged, cGAS catalyzes the synthesis of the second messenger molecule, 2′3′-cGAMP, which binds stimulator of interferon genes (STING) (6). Through a series of downstream pathways, involving the ordered activation of STING by 2′3′-cGAMP, recruitment and phosphorylation of Tank Binding Kinase (TBK)-1 by activated STING, and subsequent phosphorylation and nuclear translocation of the transcription factor, interferon regulatory factor 3 (IRF3), cGAS-STING pathway activation promotes the production of type I interferon α or β and other inflammatory cytokines (1,7). The secreted interferon (IFN) is important for the recruitment of dendritic cells, which are involved in antigen presentation and subsequent priming and activation of CD8+ T cells (8). In addition, secreted IFN also leads to the production of several interferon-stimulated genes (ISGs) via the Janus Kinase (JAK)-signal transducer and activator of transcription 1 (STAT1) signalling pathway (8,9). Several reports indicate that inhibiting DNA repair proteins – including ATM (10), ATR (11) and PARP1 (12) – either alone or in combination with radiation and immune checkpoint inhibitors can lead to mitochondrial and/or nuclear DNA damage followed by generation of cytosolic DNA and the consequent activation of the T1IFN response. Activation of the T1IFN response is required for the recruitment, priming and homing of CD8+ T cells to tumours for targeted cancer cell killing. The cGAS–STING pathway thus directly connects inhibition of the DNA damage response (DDR) and resultant persistent DNA damage to the activation of the T1IFN response to produce a systemic immunogenic response against cancer cells.

In addition to radiation, reactive oxygen species (ROS) can also induce DNA damage, thereby contributing to the induction of T1IFN response (13,14). ROS are a group of short-lived oxygen-containing molecules such as superoxide, hydrogen peroxide (H2O2) and hydroxyl radicals that are endogenously produced by the mitochondria and by nicotinamide adenine dinucleotide phosphate (NADPH) oxidases (15). Both ROS and ionizing radiation (IR), which also generates ROS by radiolysis of water, cause damage to DNA mostly in the form of oxidative damage to bases, yielding products such as 8-hydroxy-2′-deoxyguanosine (8-OHdG) as well as strand breaks (16,17). These DNA strand breaks are often associated with some kind of terminal modifications such as 3′-phosphate or 3′-phosphoglycolate groups and 5′-hydroxyl groups (18,19). Because DNA strand breaks with a 3′-phosphate group and/or a 5′-hydroxyl group block the activities of DNA polymerases and ligases, and are potentially cytotoxic, they must be restored to their conventional 3′-hydroxyl and 5′-phosphate configurations (20). A particularly important enzyme in this regard is polynucleotide kinase/phosphatase (PNKP) (21).

PNKP is a bifunctional enzyme that possesses both DNA 5′-kinase and 3′-phosphatase activities (22–24). It serves an important role in several DNA repair pathways by catalyzing the restoration of 5′-phosphate and 3′-hydroxyl (OH) on the ends of damaged DNA, thereby allowing DNA polymerases and ligases to rejoin single- and double-strand (ss/ds) breaks (24). Depletion of PNKP renders cells sensitive to a range of genotoxic agents, including IR and H2O2 (25). Moreover, PNKP has been shown to localize to the nucleus and to the mitochondria, and its depletion has been reported to increase the levels of both mitochondrial and nuclear DNA damage (23,26). PNKP plays an important role in maintaining progenitor cell populations (27), and the clinical consequences of PNKP mutation have been observed in neural progenitor cells in which PNKP loss manifests as a number of neurodevelopmental and neurodegenerative disorders (28–30). In addition, mutant huntingtin and mutant ataxin-3 proteins have been shown to impair PNKP repair activity (31).

Given PNKP’s important role in repair of endogenous DNA damage, we sought to determine if loss of PNKP promotes T1IFN response via generation of DNA damage that is detected by DNA sensors within the cytosol. To this end, the present study demonstrates that loss of PNKP in certain cell lines causes a dramatic increase in STAT1 phosphorylation (an indication of T1IFN response), an upregulation of ISGs, increased secretion of pro-inflammatory cytokines and generation of cytosolic DNA. We provide further proof that the increased STAT1 phosphorylation and pro-inflammatory gene induction is dependent on the two DNA sensors, cGAS and ZBP1. Furthermore, our results illustrate that ROS co-operate with loss of PNKP to cause mtDNA damage, and the consequent leakage of mtDNA fragments into the cytosol to activate the T1IFN response. These results lend further support for considering PNKP as a potential therapeutic target, possibly in combination with immune checkpoint inhibitors. They also have a bearing on the possibility that PNKP deficiency may play a role in type I interferonopathies giving rise to neurological disorders (32,33)

Materials and methods

Chemicals/inhibitors, plasmids, primer sequences, siRNAs and antibodies

Inhibitors of cGAS (G140, Inh-g140), cyclosporin A (CSA, 12088), JAK1/2 inhibitor (Ruxolitinib, 11609), STING inhibitor (C-178, 25860) were purchased from Cayman Chemical (Ann Arbor, Michigan, USA). VDAC (VBIT-4, HY-129122) inhibitor was purchased from MedChemExpress (Princeton, New Jersey, USA). N-Acetyl-l-cysteine (NAC, A9165-5G), uridine (U30003-5G), tiron (172553–25G), Leptomycin B (L2913), CDK1 inhibitor (RO-3306), ethidium bromide (E1385) and TBK1 inhibitor (MRT67307, 506306, Calbiochem) were purchased from Millipore Sigma (Burlington, MA, USA). 2′,3′-Dideoxycytidine (ddC, L10619.MF) and digitonin (BN20061) were purchased from Thermo Fisher Scientific (Waltham, MA, USA). All inhibitors were dissolved in DMSO unless stated otherwise. The final DMSO concentration was less than or equal to 0.2% in all experiments.

Small interfering RNAs targeting human PNKP (M-006783-02-0010), cGAS (M-015607-01-0005 and D-015607-04-0005, L-015607-02-0005), MAVS (J-024237-05-0005), STING (D-024333-04-0005), ZBP1 (D-014650-04-0002), and non-targeting siControl (D-001206-14-20) were all purchased from Horizon Discovery (Dharmacon, Cambridge, UK). Other siRNAs used included: single targeting siRNA against PNKP: AGAGAUGACGGACUCCUCU (25), or single targeting siRNA to the 3′-untranslated region of PNKP: CACAAUAAACGCUGUUUCUCC (Dharmacon) (34). All primers for PCR, RT-qPCR and short DNA sequences were purchased from Integrated DNA Technology (IDT, Iowa, USA).

Primary antibodies to the following proteins were used: PNKP (1:1000, sc-365724 Santa Cruz, Texas, USA; PA5-82263, Invitrogen, Massachusetts, USA), pSTAT1 (1:1000–1:3000, sc-136229, Santa Cruz; and 9167S, Cell Signaling), total STAT1 (1:1000, sc-462,1:3000, Santa Cruz; MA5-15129, 1:2000, Invitrogen), pIRF3 (1:1000, ab76493, Abcam, Cambridge, UK), total IRF3 (1:1000, 11904S, Cell Signaling, Massachusetts, USA), ISG15 (1:1000, sc-166755, Santa Cruz), STING (1:1000, 13647S, Cell Signaling), pTBK1(1:1000, ab109272, Abcam; and MA5-35869, Invitrogen), total TBK1 (1:1000, MA1-20344, Invitrogen), β-actin (1:2000, sc-47778, Santa Cruz), GAPDH (1:3000, NBP2-27103, Novus, Colorado, USA; and MA5-15738-D680, Invitrogen), β-tubulin (1:3000, 926–42211, Li-Cor, Nebraska, USA; and ab6046, Abcam), MTCO1 (1:1000, 459600, Invitrogen), MAVS (1:1000, sc-166583, Santa Cruz), TFAM (1:2000, PA5-29571, Invitrogen), TREX1 (1:1000, 15107S, Cell Signaling; and MA5-34734, Invitrogen), Lamin A/C (1:1000, MA3-1000, Invitrogen), cGAS (1:1000, PA5-76367, Invitrogen; and 15102S, Cell Signaling), Nucleolin (1:2000, ab22758, Abcam), and anti-dsDNA (1:1000, ab27156, Abcam). Secondary antibodies (all diluted 1:5000 in TBST) were purchased from Li-Cor and included goat anti-rabbit HRP (926-80011), goat-anti-mouse HRP (926–80010), goat anti-rabbit IRDye 800 (926-32211), goat anti-mouse IRDye 800 (926-32210). Neutralizing antibodies used in this study were purchased from PBL Assay Science (New Jersey, USA) and included anti-human IFNβ (31410-1), anti-human IFNα (21100-2) and anti-human IFNα/β receptor chain 2 (21385-1). The human IgG isotype control (Cat. #: 31154) for the neutralization experiments was purchased from Invitrogen.

The primer sequences for qPCR, PCR and RT-qPCR experiments are provided in Supplementary Table S1.

Cell culture

The human luminal A breast cancer cell lines (MCF7 and T47D), the triple-negative breast cancer cell lines (MDA-MB-231, MDA-MB-468 and MDA-MB-436), the pancreatic cancer cell line PANC-1 and the human pancreatic cancer cell line PC-3 were all obtained from the American Type Culture Collection (ATCC). The human embryonic kidney 293T (HEK293T) cell line was obtained from Dharmacon. All cell lines were cultured and maintained in Dulbecco's Modified Eagle's (DMEM)-F12 media supplemented with 5–10% foetal bovine serum (FBS), 2 mM l-glutamine with or without 50 U/ml penicillin and 50 μg/ml streptomycin, except the triple-negative cell lines and HEK293T, which were cultured in high glucose DMEM media with the same supplements as above.

The human mammary epithelial cell line MCF10A cells (sourced from ATCC) were cultured in mammary epithelium basal medium (Cat. #: CC-3151, Lonza, Switzerland) supplemented with growth factors and other supplements from the SingleQuots Kit (Cat. #: CC-4136, Lonza, Switzerland). All cell lines were maintained at 37°C under 5% CO2 in a humidified incubator. Cells were routinely screened for mycoplasma contamination. All cell lines were validated by ATCC cell line authentication service using short tandem repeat (STR) analysis.

Protein depletion by transient siRNA transfection

For transient transfection, all cell lines were seeded at the appropriate cell number and reverse transfected with the non-targeting siRNA or siRNA targeting PNKP, cGAS, MAVS, STING or ZBP1 or a combination of both siPNKP and sicGAS or siZBP1 all at a final concentration of 20 nM for ∼24 h using 0.15% lipofectamine RNAiMAX (Cat. #: 3778-075, Invitrogen) following the manufacturer's instructions. Following reverse transfection, transfection media were replaced with the appropriate fresh growth media and cells were allowed to grow for an additional 18 hours before any downstream treatment/experimental procedures were applied. For experiments not requiring further treatment, cells were left in fresh growth media up to 66 h before trypsinization, cell lysis and western blotting or RT-qPCR.

RNA extraction, reverse transcription and real-time quantitative PCR

Total RNA was extracted from cells using the RNeasy mini kit (Cat. #: 74104; Qiagen) following the manufacturer's instructions. The concentration and quality of the purified total RNA was determined using a NanoDrop UV/Vis spectrophotometer (Thermo Scientific). Subsequently, 1 μg of total RNA was used for reverse transcription to synthesize cDNA in a 20 μl reaction using the High-Capacity cDNA reverse transcription kit (Cat. #: 4368814; Applied Biosystems) according to the manufacturer's instructions, after which the synthesized cDNA was diluted 10-fold with RNase- and DNase-free deionised water. RT-qPCR was performed on 4 μl cDNA using the BrightGreen 2X qPCR MasterMix (Cat. #: G892, ABMGood, British Columbia, Canada) in a 10 μl reaction in triplicate for each primer. Thermal cycling conditions (on QuantStudio 6; Applied Biosystems; fast protocol) included a holding stage at 95°C for 20 s for enzyme activation followed by 40 cycles of each PCR step of denaturation at 95°C for 3 s and annealing/extension at 60°C for 30 s. A melt curve analysis was also done to ensure the specificity of the corresponding qPCR reactions, and the absence of primer dimers was confirmed by the absence of amplification in the no-template control (NTC) wells. For data analysis, the cycle threshold (Ct) values were exported to an Excel file for subsequent calculations of relative gene expression (by the comparative ΔΔCt method) using ARHGDIA or GAPDH as reference gene. All fold changes were expressed normalized to the untreated control.

Detection of proteins or cytoplasmic DNA by immunofluorescence

Proteins or cytoplasmic DNA was probed according to a previously described protocol (8). Briefly, cells on coverslips were fixed for 20 minutes at room temperature (RT) in 4% paraformaldehyde (PFA) diluted in PBS. Cells were then permeabilized with 0.1% Tween 20 and 0.01% Triton X-100 for an additional 20 min, followed by blocking with 1% BSA, 22.52 mg/ml glycine in TBST (0.1% Tween 20 in TBS) for 30 min at RT. Subsequently, fixed and permeabilized cells were incubated with primary anti-PNKP (Cat. #: PA5-82263, Invitrogen), TFAM (1 in 500 dilution, Cat. #: PA5-29571, Invitrogen), or anti-dsDNA (Cat. #: ab27156; 1 in 1000 dilution; Abcam) overnight at 4°C or 2–3 h at RT. After three washes for 5 min each, coverslips were incubated with species-specific Alexa Fluor-conjugated secondary antibody (1 in 250 dilution) for 1 h at RT. After three consecutive 5-min washes, coverslips were mounted with an anti-fade mounting medium containing DAPI (Cat #: C069; GeneCopoeia). Z-stack images were obtained using a laser scanning confocal microscope (Zeiss LSM710, Oberkochen, Germany) with a 40 × 1.3 objective.

Click-iT plus TUNEL labelling of cytoplasmic DNA

2.5 × 105 MCF7 cells were reverse transfected with 20 nM final concentration of siRNA against PNKP or the non-targeted control. 24 h post-transfection, transfection medium was changed to the appropriate fresh medium and left for an additional 66 h. To detect for the presence of strand breaks, a slightly modified terminal deoxynucleotidyl transferase dUTP nick end labelling (TUNEL) assay protocol was adapted. Briefly, cells were fixed for 10 minutes at RT in 4% PFA in PBS. Cells were then permeabilized with 0.1% Tween 20 and 0.01% Triton X-100 for an additional 10 min. To detect cytoplasmic DNA fragments, fixed and permeabilized cells were labelled with the Click-iT Plus TUNEL assay kit (Cat. #: C10618; Invitrogen) following the manufacturer's instructions. Following labelling of DNA fragments in the cytosol, slides were either imaged or subjected to further dsDNA antibody staining using standard procedures. After three 5-min washes, slides were mounted on a coverslip with an anti-fade mounting medium containing DAPI (Cat #: C069; GeneCopoeia). Z-stack images were obtained using laser scanning confocal microscopy (Zeiss LSM710) with a 40× 1.3 objective.

Cell lysis and western blotting

Western blotting was performed as described (35,36) with some minor modifications. Briefly, cells were harvested at the indicated time points and lysed in RIPA buffer (Cat. #: 89900, Thermo Scientific) supplemented with protease and phosphatase (PhosSTOP, Roche) inhibitor cocktails for 30 min on ice with 10 min intermittent vortexing. Homogenized lysates were centrifuged at 16 000×g for 10 min to remove cellular debris. The Bradford assay was used to determine protein concentration, after which 20–30 μg of protein was denatured by heating at 70°C for 10 min in Laemmli buffer supplemented with 10% β-mercaptoethanol. Denatured proteins were separated by SDS-PAGE electrophoresis and transferred onto a nitrocellulose membrane using a Trans-Blot Turbo Transfer System (Bio-Rad). Membranes were blocked with 5% nonfat dry skim milk in TBST (20 mM Tris base, 500 mM NaCl and 0.1% Tween-20, pH 7.5) for 1 hour at RT. Blocked membranes were cut at the appropriate marker size to allow for the probing of several proteins with different antibodies on the same membrane before incubation with the indicated primary antibodies diluted in the appropriate dilution buffer (TBST or TBST milk) at 4°C overnight or at RT for 3 h. Subsequently, membranes were washed three times in TBST, incubated with species-specific horseradish peroxidase-conjugated (or IRDye-conjugated) secondary antibody (Li-Cor) diluted in TBST (1 in 5000) for 1 h at RT. After three washes in TBST, the signals were detected with an Odyssey imaging system (Li-Cor) using an ECL chemiluminescence kit (Cat. #: PI34577, Thermo Scientific, if using horseradish peroxidase-conjugated antibody). The relative protein levels were analyzed using Image Studio software. In all western blot experiments, β-actin, tubulin or GAPDH served as the loading control.

Radiation treatment

For radiation treatment, cells transiently transfected with the indicated siRNA were exposed to the indicated dose of radiation at least 40–42 h post transfection using a 60Co Gammacell (Atomic Energy of Canada Limited, Ottawa, ON), or left untreated as controls, and maintained in the same media under standard cell culture conditions for different time points up to 24–120 h post treatment. Subsequently, cells were lysed for protein or DNA/RNA extraction or other downstream experiments.

Pharmacological inhibition of proteins

To assess the involvement of a particular protein in the pathway that mediates the T1IFN response, the protein of interest was pharmacologically inhibited using its potent inhibitor. Briefly, 41–66 h post siRNA transfection, cells were treated with inhibitors at the indicated concentrations (or with DMSO as the vehicle control) and left in the media containing the inhibitors before harvesting at the indicated time point.

Mitochondrial DNA depletion

To assess for the contribution of mtDNA to STAT1 activation following downregulation of PNKP, mtDNA was depleted as described (37,38). Briefly, MCF7, PANC-1 or T47D cells cultured under standard culture conditions were incubated with 100–150 ng/ml ethidium bromide (EtBr) for 6–20 days. Media was changed to fresh media containing 100–150 ng/ml EtBr every 2–4 days. In another experiment, MCF7 or T47D cells cultured under standard culture conditions were incubated with 10 μM ddC (supplemented with 50 μg/ml uridine and 1 mM sodium pyruvate) for 6 days as previously described (39,40). Culture media was replenished with fresh media containing 10 μM ddC every 2 days. On day 6 or 20, MCF7, PANC-1 or T47D cells cultured in EtBr or ddC and the untreated control group were reverse transfected with siRNA as described above and left for up to 114 h before DNA content, protein and gene expression analyses by western blot, RT-qPCR and qPCR, respectively.

Subcellular fractionation for immunoblotting

To assess the subcellular localization of proteins, cells were harvested, and proteins extracted from the whole cell, cytosolic, mitochondrial, and nuclear fractions. Briefly, trypsinized and pelleted cells were incubated end-over-end for 10–15 min at 4°C in a cytoplasmic extraction buffer (CEB, 10 mM HEPES pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM DDT, 0.1% triton-X100 and 5 μg/ml digitonin) supplemented with freshly prepared 1× Complete EDTA-free protease inhibitor cocktail and 1x phosphatase inhibitor (PhosSTOP, Roche) for selective plasma membrane permeabilization. Lysed cells were then centrifuged at 1500×g for 10 min at 4°C, generating a supernatant as the crude cytosolic fraction and a bulk pellet as the nuclear fraction. The crude supernatant was collected and further subjected to centrifugation at 16 000×g for 15–20 min. The resulting cleared supernatant was kept as the cytosolic fraction, whereas the small pellet was considered the mitochondrial fraction. Later, the mitochondrial fraction was washed/rinsed once in CEB above, centrifuged at 1500×g for 5 min, and the pellet kept on ice. The bulk nuclear pellet from above was resuspended in CEB (without 0.1% Triton X-100), vortexed repeatedly to lyse any unlysed cells, kept on ice for 5–10 min before centrifugation at 1500×g for 5 min to remove any contaminating cytosolic contents. The resulting supernatant was discarded, and the pellet kept on ice for further lysis. Subsequently, both the nuclear and mitochondrial fraction pellets were incubated in RIPA buffer and allowed to lyse for 30 min on ice with intermittent vortexing every 10 min and then centrifuged at 16 000×g for 10 min at 4°C. The supernatants were collected as the nuclear and mitochondrial fractions, respectively. Protein concentration of the different fraction supernatants was determined using the Bradford assay.

Detection of cytosolic DNA by qPCR

90 h post transfection, MCF7 cells were harvested for subcellular fractionation using cytosolic extraction buffer (CEB: 10 mM HEPES pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.34 M sucrose, 10% glycerol, 1 mM DDT, 0.1% Triton-X100 and 5 μg/ml digitonin) supplemented with freshly prepared 1 × Complete EDTA-free protease inhibitor cocktail and 1x phosphatase inhibitor (PhosSTOP, Roche; if samples were to be used for western blotting). Cells resuspended with the CEB were incubated end-over-end for 10 min at 4°C to allow for selective plasma membrane permeabilization then centrifuged at 21 000×g for 20 min to pellet intact nuclei and other cell debris. The supernatant was collected and kept as the cytosolic fraction, while the crude pellet was resuspended in CEB (this time, CEB was supplemented with 0.5% of Triton X-100), left on ice for 30 min with 10 min intermittent vortexing. All subcellular fractions (400 μl) were pretreated with proteinase K (Qiagen) and incubated at 56°C for 30 min, then with 500 μg/ml RNase A at 37°C for 1 h before downstream DNA purification by traditional phenol/chloroform extraction. Purified cytosolic DNA was dissolved in 30–60 μl of RNase and DNase-free water whereas DNA from the crude pellet fraction was resuspended in 200 μl of RNase and DNase-free water. DNA concentration was determined spectrophotometrically prior to quantitative PCR (qPCR). All purified DNA was initially diluted 1:100 before downstream qPCR. qPCR was then performed on purified DNA from both cytosolic and crude pellet fractions using both nuclear (Alu DNA sequence, 5.8s, 18s and 28s rDNA) and mtDNA (mtDNA1 and 2, Homo-ND1 and 2) primers. The cycle threshold (CT) values obtained for Alu DNA sequence abundance from the nuclear fractions served as the normalization control for both the nuclear and mtDNA values obtained from the cytosolic fractions. This allowed effective standardization among samples and controlled for any variations in the total amount of mtDNA in control and PNKP-deficient samples.

ELISA for secreted human IFNβ1 in culture media

Cell culture supernatants were collected, cleared of debris by centrifugation and then passaged through a 0.22 μm syringe filter before taking out 50 μl in triplicate for measurement of secreted IFNβ1 using the IFN beta Human ELISA Kit (Cat. #: 414101; Invitrogen) in accordance with the manufacturer's protocol.

2′3′- cGAMP ELISA

To detect presence of 2′3′- cGAMP following loss of PNKP in MCF7 cells, the 2′3′-cGAMP ELISA Kit (catalog no. 501700; Cayman Chemical) was used according to the manufacturer's instructions and the protocol described in (41). Briefly, 30 μg of cell lysates prepared in RIPA buffer (50 μl) from PNKP-depleted and control cells were used for the reaction with each sample assayed in duplicate. Following overnight antibody incubation and development, the plate was read using a FLUOStar Omega microplate reader (MBG LabTech) at a wavelength of 450 nm. Data analysis was done using the Arigobio ELISA calculator (https://www.arigobio.com/ELISA-calculator). The concentration of 2′3′-cGAMP was extrapolated from the standard curve generated. The amount of 2′3′-cGAMP in PNKP-depleted MCF7 cells was normalized to that obtained in the control cells.

Determination of mtDNA integrity by long PCR amplification assay

Mitochondrial and nuclear genome-specific semiquantitative PCR assays of long DNA fragments for measuring DNA integrity were performed as described previously (42,43). Purified cellular DNA was subjected to mtDNA long PCR amplification using specific primers to amplify the entire 16.6-kb mtDNA sequence. Here, 40 ng of template DNA, 0.4 μM of primers of both forward and reverse primers, 12.5 μl of LongAmp Taq 2X Master Mix (NEB; Cat #: M0270S) that contains sufficient dNTPs and Taq DNA polymerase were mixed in a 25 μl PCR reaction tube. Cycling parameters were as follows: initial denaturation at 94°C for 2 min followed by 30 cycles of 94°C for 30 s, 60°C for 30 s and 65°C for 15 min (50 s per kb), and a final elongation step of 65°C for 10 min and held at 4°C. Because small DNA target regions are unlikely to experience any significant damage, primers were also designed to amplify a 211-bp region of mtDNA, which was used as an indicator of relative copy number (and to normalize the long amplicon copy number) and a measure of the PCR quality of the genomic DNA extracts. PCR products were quantified using the Qubit dsDNA broad range assay kit (Cat. #: Q32850, Invitrogen). The frequency of DNA lesions was calculated by normalizing the amplification of DNA samples from PNKP-depleted cells to the amplification of the DNA samples from the non-targeted siControl cells, which was set at 100%.

Generation of ectopically overexpressed GFP-tagged wild type PNKP T47D cell line

To generate T47D cell line ectopically overexpressing GFP-tagged wild type PNKP, wild type T47D cells were grown and allowed to attach for at least 24 h prior to transfection. The next day, cells were transfected with 500 ng of the PNKP plasmid (pRP[Exp]-CMV > EGFPBamHI/HindIII/{hPNKP[NM_007254.4]) obtained from Vector Builder (Chicago, IL) using lipofectamine 2000 (Invitrogen) according to manufacturer's instructions. 72 h after transfection, cells were passaged with fresh growth media and allowed to attach overnight before incubation with 0.5 μg/ml puromycin to select for cells that were efficiently transfected and are antibiotic-resistant. Cells were maintained in selective media for up to 7 days such that only cells that retained the plasmid were left growing. Furthermore, cells were left in culture without puromycin for an additional 2–3 weeks until visible colonies that were GFP positive were formed. These GFP-positive colonies were then picked and expanded before performing western blotting to select clones that had GFP-tagged wild type PNKP overexpressed. These clones were then later expanded and used for downstream experiments.

Polymerase chain reaction (PCR) analysis of mtDNA-depleted cells

For analysis of mtDNA copy number following ddC treatment, PCR was performed with primers designed against regions of both mitochondrial and nuclear DNA. DNA was extracted from cells using the DNeasy Blood and Tissue kit (Cat #: 69504, Qiagen) following the manufacturer's instructions. DNA concentration was determined using the Qubit dsDNA broad range assay kit (ThermoFisher, Cat #: Q32850). Here, 40 ng of template DNA, 0.4 μM of primers targeting regions of genes encoding mitochondrial cytochrome c oxidase subunits 1 and 2 and nuclear GAPDH or β-globin, 12.5 μl of LongAmp Taq 2X Master Mix (NEB; Cat #: M0270S) that contains sufficient dNTPs and Taq DNA polymerase were mixed in a 25 μl PCR reaction tube. Cycling parameters were as follows: initial denaturation at 94°C for 2 min followed by 30 cycles of 94°C for 30 s, 60°C for 30 s and 65°C for 50 s, and a final elongation step at 65°C for 10 min and held at 4°C. PCR products were electrophoresed on a 2% agarose gel, stained with RedSafe nucleic acid staining solution (Froggabio Scientific Solutions, Ontario, Canada) and photographed under UV light to confirm that the correct lengths of amplicon were generated. Lastly, PCR products were quantified using the Qubit dsDNA broad range assay kit as above.

Statistical analysis

All statistical analyses and generation of bar graphs were performed using GraphPad Prism software, whereas some numerical data tabulation was done in Microsoft Excel. All data are presented as the mean ± standard error of the mean of at least three independent experiments. Results were considered statistically significant if the statistical difference between two groups was less than 0.05. In general, *P < 0.05; ** P < 0.01; *** P < 0.001; and ****P < 0.0001. Comparison between two groups was conducted using Student's two-tailed t-test. Further statistical information can be found in the figure legends.

Results

SiRNA-mediated downregulation of PNKP activates T1IFN response independent of ionizing radiation

Initially, we treated MCF10A cells (an immortalized human mammary epithelial cell line) with short interfering RNA (siRNA) targeting PNKP to determine if downregulation of PNKP activated the T1IFN response using STAT1 phosphorylation (pSTAT1) as a recognized indicator of the response. We observed a time-dependent significant increase in pSTAT1 levels over 90 h of exposure to the siRNA (Figure 1A and B). We also observed a modest increase in total STAT1, an observation that is consistent with previous reports that persistent stimulation of the T1IFN response can significantly increase the protein levels of STAT1 (11,44). Similar results were obtained with the MCF7 and T47D breast cancer cell lines (Figure 1C and D). The western blot also revealed a robust and significant phosphorylation of IRF3 (another indication of the T1IFN response) in PNKP-depleted MCF7 and T47D cells relative to the non-targeted control cells (Supplementary Figure S1A). This was accompanied by an increase in the protein levels of ISG15, one of the ISGs that become induced following STAT1 activation and subsequent nuclear translocation. However, not all cell lines responded in the same manner as shown by analysis of two other breast cancer cell lines, MDA-MB-231 and MDA-MB-436 (Figure 1C). It is worth noting that while MDA-MB-436 cells showed a significant increase in STAT1 phosphorylation in response to PNKP loss, MDA-MB-231 cells did not, which agrees with reports that not all cell lines are adept at activating the T1IFN response (45–47). To ensure that the observed STAT1 phosphorylation was not due to off-target effects of the pooled siRNA targeting PNKP, we tested additional siRNAs against different sequences of PNKP (with one targeting its 3′-untranslated region (3′-UTR)) and included additional non-targeted siControls. Consistently, we observed an increase in STAT1 phosphorylation only in PNKP-depleted cells (Figure 1E and F), demonstrating that the siRNAs were specifically targeting PNKP. Additionally, we performed immunofluorescence and provided further evidence that the siRNA effectively downregulated PNKP (Supplementary Figure S1B). We further corroborated our claim that PNKP loss indeed promotes T1IFN response by ectopically overexpressing GFP-tagged wild type PNKP. We then used the siRNA targeted against the 3′-UTR of PNKP to selectively deplete the GFP-tagged PNKP-overexpressing T47D cells of its endogenous PNKP. As a proof of concept, the GFP-tagged PNKP-overexpressing T47D cells failed to induce sufficient level of pSTAT1 relative to the wild type T47D cells that had the endogenous PNKP depleted (Figure 1G). The preceding result demonstrates that the T1IFN response that we observed was indeed due to the silencing effect of the siRNAs against PNKP as overexpressing PNKP reverses this observation.

Figure 1. siRNA-mediated downregulation of PNKP activates the type I IFN response. (A and B) Downregulation of PNKP in MCF10A cells upregulated the T1IFN response as shown by the increase in activated STAT1 (phosphorylation of tyrosine 701) in a time-dependent fashion post-transfection of PNKP siRNA. + sign indicates the use of siRNA targeting PNKP and – sign indicates transfection with control siRNA. (B) The densitometric analysis of the pSTAT1/STAT1 blots normalized to siControl. (C and D) PNKP depletion revealing differential response in four human breast cancer cell lines. While MCF7, T47D cells displayed a robust and significant activation of the type-I IFN response as determined by elevated levels of ISG15 and phosphorylated IRF3 and STAT1 with MDA-MB-436 also significantly increasing its pSTAT1 level, MDA-MB-231 cells appear unresponsive. (D) The densitometric analysis of the pSTAT1/STAT1 blots normalized to siControl. (E and F) Validation that loss of PNKP mediates the type-I IFN response. siRNAs designed against other sequences of PNKP mRNA also effectively depleted MCF7 cells of PNKP and activated type-I IFN response. (F) The densitometric analysis of the pSTAT1/STAT1 blots normalized to siControl, indicating that loss of PNKP using different siRNA sequences consistently and significantly increases the levels of pSTAT1 in MCF7 cells. All data are presented as standard error of the mean of n = 3 independent experiments. (ns = not significant, *P< 0.05, **P< 0.01; ***P< 0.001 and ****P< 0.0001). Statistical significance was determined using unpaired Student's t-test. (G) Western blot showing that PNKP overexpression reverses the observed T1IFN response following depletion of the endogenous PNKP. Following downregulation of the endogenous PNKP, T47D cells overexpressing PNKP failed to robustly phosphorylate STAT1 and IRF3 compared to the control T47D cells that robustly increased the levels the phosphorylated STAT1 and IRF3 following downregulation of the endogenous PNKP. CNTRL = Control T47D cell line; OE = overexpression. For the western blots data shown in (A–G) whole cell extracts were used to determine protein expression.

Since PNKP plays an important role in both single and double DNA strand break repair pathways and its depletion has been shown to sensitize cancer cells to radiation due to accumulation of DNA strand breaks (25,26,48), we hypothesized that loss of PNKP together with radiation may lead to an additional increase in the T1IFN response. However, the combination of siRNA-mediated depletion of PNKP and irradiation failed to elicit a greater response in MCF7 cells than depletion of PNKP alone, even with a dose as high as 30 Gy (Figure 2A). Similar results were observed with T47D and the PANC-1 cells (Figure 2B and C). Next, we performed RT-qPCR to measure changes in genes known to be upregulated following induction of the T1IFN response (Figure 2D). Depletion of PNKP alone in MCF7 cells led to a significant upregulation of the transcript levels of ISG15 and IFNβ1 as well as the dendritic cell attractant CCL5 (49). Of these, only CCL5 transcript levels showed a modest increase when PNKP-depleted cells were irradiated. Noticeably, irradiation of the cells transfected with the control siRNA did not stimulate increased expression of these genes. Similar results were obtained with PANC-1 cells (Supplementary Figure S1C).

Figure 2. siRNA-mediated downregulation of PNKP activates type I IFN response independent of ionizing radiation (IR). (A) IR dose-independent activation of the T1IFN response in PNKP-depleted MCF7 cells. Following transfection, MCF7 cells were exposed to increasing doses of ionizing radiation, and protein expression was assessed 72 h later to determine if radiation contributes to the T1IFN response in MCF7 cells following downregulation of PNKP. (B, C) IR-independent activation of the T1IFN response in T47D and PANC-1 cells, respectively. For the western blots data shown in (A–C), whole cell extracts were used to determine protein expression. (D) siRNA-mediated knockdown of PNKP upregulates the expression of ISGs, including CCL5, ISG15 and IFNβ1 independent of radiation 3 days post-IR in MCF7 cells. Data are presented as standard error of the mean of n = 3 independent experiments. (*P< 0.05, **P< 0.01; ***P< 0.001 and ****P< 0.0001). Statistical significance was determined using unpaired Student's t-test.

PNKP-depleted cells secrete IFNβ1 that promotes STAT1 phosphorylation and induction of pro-inflammatory genes

To be fully activated, STAT1 in complex with its partner STAT2 requires cytokine/interferon secretion and subsequent paracrine or autocrine binding of interferons to cognate interferon alpha and beta receptor (IFNAR) on the cell surface (50,51). Once bound, type 1 interferons (interferon α or β) promote JAK/TYK2 tyrosine phosphorylation, which is required for the downstream phosphorylation of STAT1 on tyrosine 701 (52). Phosphorylated STAT1 in turn forms a dimer with STAT2 and IRF9 (the ISGF3 complex) enabling STAT1 to translocate to the nucleus to turn on a number of genes collectively called the interferon-stimulated genes (ISGs) (53,54). To confirm whether the observed STAT1 phosphorylation was driven by secreted cytokines and/or interferons and not due to PNKP loss directly activating STAT1, we incubated naïve MCF7 cells with conditioned media (CM) derived from unirradiated and irradiated MCF7 cells with or without PNKP depletion. Our results indicate that the CM from unirradiated PNKP-depleted cells was more potent at activating STAT1 compared to CM from cells exposed to an IR dose of 8 Gy (Figure 3A), further validating the observation above that loss of PNKP alone is sufficient to induce a significant T1IFN response independent of IR. We also tested the ability of other cell lines to induce STAT1 phosphorylation following incubation with CM from PNKP-depleted MCF7 cells. Indeed, we showed that in addition to naïve MCF7 cells, other cell lines responded to the interferons present in PNKP-depleted MCF7 cells by increasing STAT1 phosphorylation (Figure 3B).

Figure 3. Loss of PNKP promotes IFNβ1 secretion that mediates downstream STAT1 activation. (A) Immunoblots showing activation of the T1IFN response as determined by elevated IRF3 and STAT1 phosphorylation following depletion of PNKP in MCF7 cells (first 4 lanes). Secreted interferons/cytokines present in conditioned media (CM) from PNKP-depleted MCF7 cells activate STAT1 independent of radiation (last four lanes). Naïve MCF7 cells were incubated with CM from the corresponding first four lanes) for 24 h prior to cell harvesting. (B) CM from PNKP-depleted MCF7 cells stimulate the T1IFN response in other naïve cancer cells. Cells were allowed to attach before 24-hour incubation with CM from PNKP-depleted MCF7 cells. (C) ELISA assay to determine levels of secreted IFNβ1 in media supernatants in PNKP-depleted MCF7, T47D and PANC-1 cells relative to the siControl cells 90 h post transfection with siRNA (replicates: n = 5 for MCF7, and n = 4 for both T47D and PANC-1). Data are presented as standard error of the mean of n number of experiments (*P< 0.05, **P< 0.01 and ***P< 0.001). Statistical significance was determined using unpaired Student's t-test. (D) Western blot analysis showing result of neutralizing antibodies (NA) against IFNβ and IFNAR2 in PNKP-depleted MCF7 cells. (E) Western blot analysis showing result of neutralizing antibodies against IFNβ, IFNα and IFNAR2 in PNKP-depleted MCF7 cells; time of incubation = 24 h. For the western blots data shown in (A–E) whole cell extracts were used to determine protein expression. (F) Subcellular fractionation of MCF7 cells reveals the localization of proteins that mediate T1IFN response following downregulation of PNKP. Fractionation purity was assessed using lamin A/C and nucleolin as nuclear markers, and tubulin and GAPDH as cytoplasmic markers. Whole cell, cytosolic and nuclear extracts were used to determine protein expression.

Next, we asked whether the secreted interferons contained IFNβ1 using an enzyme-linked immunosorbent assay (ELISA) to quantify the amount of IFNβ1 present in the media of PNKP-depleted MCF7 cells relative to the siControl cells. Our result indicated that media from PNKP-depleted MCF7 cells had >5-fold increase in secreted IFNβ1 relative to the siControl cells (Figure 3C). A similar result was obtained with T47D and PANC-1 cells (Figure 3C). This suggests that the increased level of phosphorylated STAT1 in PNKP-depleted cells arises in response to the elevated levels of secreted IFNβ1. Secreted IFNβ1 binds to its cognate cell surface receptors, composed of IFNAR1 and IFNAR2, which in turn leads to downstream JAK/TYK2 and STAT1 phosphorylation (51). To determine whether the cell surface receptor was mediating downstream STAT1 phosphorylation through autocrine/paracrine binding of IFNβ1 or the different subunits of IFNα, we treated PNKP-depleted MCF7 cells with IFNAR2 blocking/neutralization antibody. We observed a marked abrogation of STAT1 phosphorylation with a corresponding reduction in the protein and mRNA levels of STAT1-regulated ISG15, and the cytokine CXCL10, respectively (Figure 3D and Supplementary Figure S2A), which agrees with the data by Zhang et al. (55) of reduced STAT1 phosphorylation following neutralisation of IFNAR2 in PANC-1 cells. A similar result was obtained with PNKP-depleted T47D and PANC-1 cells (Supplementary Figure S2B and C).

IFNAR co-ordinates the downstream signalling effects of IFNα as well as IFNβ1 (56). Since we observed a sharp decrease in STAT1 phosphorylation following neutralization of IFNAR2, we next performed antibody neutralization experiments to specifically identify which of the type 1 interferons was playing a significant role in the autocrine/paracrine T1IFN signalling following loss of PNKP in MCF7 cells. Western blot and RT-qPCR identified IFNβ1 as the key driver of STAT1 phosphorylation, as incubation with an IFNβ1 neutralizing antibody robustly impaired STAT1 phosphorylation with a corresponding reduction in downstream ISGs and the cytokine CXCL10 (Figure 3E and Supplementary Figure S2A). A similar result was obtained with T47D cells incubated with neutralizing antibody against IFNβ1 (Supplementary Figure S2B). Although there was a slight reduction, neutralizing antibodies against IFNβ1 and IFNAR2 did not have a significant effect on CCL5, suggesting that CCL5 may be regulated by other pathways. Moreover, Supplementary Figure S2A shows that the treatment with neutralizing antibodies against IFNβ1 or IFNAR2 did not affect the transcript levels of IFNβ1, suggesting that IFNβ1 was not being further induced by STAT1 activation. Neutralizing antibodies against IFNα, however, failed to abrogate STAT1 phosphorylation in MCF7 cells (Figure 3E). A similar result was obtained with PANC-1 cells, except that western blot data suggests that IFNα might also be mediating STAT1 activation in PANC-1 cells (Supplementary Figure S2C) unlike in MCF7 cells where IFNβ1 appears to be the key cytokine.

STAT1 is a transcription factor and has been shown to translocate to the nucleus upon phosphorylation through dimerization with its partner STAT2 and IRF9 (53,54). We, therefore, sought to determine the localisation of key proteins, including phosphorylated STAT1 and IRF3 following loss of PNKP by performing subcellular fractionation followed by immunoblotting. We first confirmed the efficiency of the fractionation protocol using markers specific for proteins in the cytosol (beta tubulin and GAPDH) and nucleus (Lamin A/C and nucleolin) (Figure 3F). Subsequently, we showed that both phosphorylated STAT1 and IRF3 localised to the nucleus upon activation due to loss of PNKP (Figure 3F), indicating that both STAT1 and IRF3 undergo their canonical transcriptional activities that mediate the T1IFN response in MCF7 cells. Taken together, the preceding results suggest that loss of PNKP induces the T1IFN response by increasing the levels of secreted IFNβ1 in MCF7 and T47D cells, and both IFNβ1 and IFNα in PANC-1 cells. These cytokines subsequently bind to their cognate cell surface IFNAR receptor and induce downstream factors, including STAT1 to translocate to the nucleus to promote the secretion of pro-inflammatory cytokines and chemokines.

Involvement of the cGAS-STING signalling pathway following PNKP depletion

Defects in DDR proteins and/or exposure to IR have been demonstrated to cause DNA damage that consequently activates the cGAS-STING pathway (11,57). Activation of this pathway is dependent on the sensing of mitochondrial or nuclear-sourced cytosolic DNA fragments by the DNA sensor cGAS, and the downstream activation of proteins such as STING and TBK1 followed by the nuclear translocation of phosphorylated IRF3. However, non-canonical, cGAS-independent activation of the T1IFN response pathway has also been described (55,58). Since we consistently observed a persistent accumulation of cytosolic DNA (described in detail below) in PNKP-depleted cells, we sought to determine if the observed T1IFN response in PNKP-depleted cells was mediated by the DNA sensor cGAS and its canonical downstream protein partners. To test the involvement of cGAS, we employed siRNA to simultaneously deplete MCF7 cells of PNKP and cGAS. As expected, combined downregulation of PNKP and cGAS abrogated both STAT1 and IRF3 phosphorylation in MCF7 cells (Figure 4A), suggesting that the observed T1IFN response following PNKP loss is mediated by the canonical cGAS-STING signalling pathway in MCF7 cells. We next assessed changes in gene expression and showed by RT-qPCR that simultaneous depletion of PNKP and cGAS led to a significant reduction in the transcript levels of ISG15, ZBP1, CCL5, CXCL10 and IFNβ1 (Supplementary Figure S3A). We confirmed the involvement of cGAS by pharmacological inhibition using a recently described selective human cGAS inhibitor (cGASi) called G140 (59). Treatment of PNKP-depleted MCF7 cells with either 5 or 10 μM G140 for 48 h significantly reduced the levels of phosphorylated STAT1 and ISG15 (Figure 4B and C). It should be noted that cGAS has been previously identified as the DNA sensor in MCF7 cells responsible for the induction of STAT1 phosphorylation (60). Additional evidence for the involvement of cGAS was provided by measuring the amount of 2′3′-cGAMP, the product of cGAS, produced in PNKP-depleted MCF7 cells relative to the control cells (Figure 4D). Pharmacological inhibition and/or downregulation by siRNA of proteins that act downstream of cGAS, i.e. STING, TBK1 and JAK1/2, led to reduced levels of phosphorylated STAT1 and IRF3 as well as ISG15 in MCF7, PANC-1 and PC-3 cells (Figure 5A and Supplementary Figure S3B-D), suggesting that the canonical cGAS-STING pathway may be required for T1IFN response in PNKP-depleted cells. However, an important exception to this observation is in T47D cells, which activates STAT1 despite lacking cGAS (61) (Figure 1C and Supplementary Figure S3E).

Figure 4. Involvement of the cGAS-STING signalling pathway in the T1IFN response following PNKP depletion. (A) Effect of RNAi-mediated double downregulation of PNKP and cGAS or ZBP1 on the T1IFN response in MCF7 cells. (B) Effect of cGAS inhibition by G140 on the T1IFN response in PNKP-depleted MCF cells. For the western blots shown in (A) and (B), whole cell extracts were used to determine protein expression. (C) Densitometric analysis of the of the western blot signal for pSTAT1/STAT1 ratio in (B) above. (D) Measurement of cGAS activity in PNKP-depleted MCF7 cells relative to the control cells. 2′3′-cGAMP was determined from cell lysates using an ELISA. All data are presented as standard error of the mean of n = 4 experiments (**P< 0.01, ***P< 0.001). Statistical significance was determined using unpaired Student's t-test.

Figure 5. Involvement of STING and ZBP1 in the T1IFN response following PNKP depletion. (A) T1IFN response observed following PNKP depletion in MCF7 cells is STING-dependent. MCF7 cells were double depleted of STING and PNKP before cells were harvested for western blotting using whole cell lysates 90 h post transfection. (B) Effect of RNAi-mediated double downregulation of PNKP and ZBP1 on the T1IFN response in T47D cells. For the western blots shown in (A) and (B), whole cell extracts were used to determine protein expression. (C) RT-qPCR data showing that siRNA-mediated double depletion of PNKP and ZBP1 abrogates T1IFN response in MCF7 cells. All data are presented as standard error of the mean of n = 4 experiments (*P< 0.05, **P< 0.01, ***P< 0.001 and ****P< 0.0001. Statistical significance was determined using unpaired Student's t-test.

ZBP1 synergises with the cGAS-STING signalling pathway to drive sustained STAT1 activation following PNKP depletion

Other than cGAS, cytosolic DNA, particularly of mitochondrial origin, has been reported to be sensed by Z-DNA binding protein 1 (ZBP1) under low-level oxidative conditions (62–65). Moreover, ZBP1 was recently reported to co-operate with cGAS in the sensing of doxorubicin-induced mitochondria-sourced cytosolic DNA, consequently leading to T1IFN response in cardiomyocytes (65). Because PNKP has been reported to maintain mitochondrial genome stability (23,26), we reasoned that its loss could generate oxidized or Z-form mtDNA that are sensed by ZBP1, which, in co-operation with cGAS, may be driving STAT1 phosphorylation as was recently reported (65). We, therefore, performed double depletion of PNKP and ZBP1 in MCF7 and T47D cells using siRNA against either protein. As shown in Figures 4A and 5B, double depletion of PNKP and ZBP1 led to an impairment of STAT1 and IRF3 phosphorylation and a diminished level of ISG15. Similarly, RT-qPCR revealed a significant reduction of ISG15 as well as the cytokines CCL5, CXL10 and IFNβ1 in MCF7 cells (Figure 5C). Notably, ZBP1 is remarkably upregulated following loss of PNKP (Figure 5C), which is consistent with the role of interferons in the induction of ZBP1 (66,67). Although we identified cGAS as a major DNA sensor in PNKP-depleted cells, ZBP1, independent of cGAS, appears to play an important role in the sensing of cytosolic DNA in T47D cells, which has been shown to lack cGAS (61). We confirmed cGAS deficiency in T47D cells and also in MDA-MB-468 (68) and HEK293T (1) cells (Supplementary Figure S3E). We also consistently observed an accumulation of STING at the protein (Figures 3D, 4A and Supplementary Figure S2B), and RNA levels (Supplementary Figure S2A and D) in both MCF7 and T47D cells, and asked whether this was a direct response to PNKP loss or due to the effect of activated STAT1 since STING has itself been previously described as an ISG involved in a T1IFN response positive feedback loop (69). We showed that blocking the STAT1 signalling pathway with neutralizing antibodies against IFNβ1 or IFNAR2 failed to dramatically reduce STING RNA or protein levels in a PNKP-depleted background (Supplementary Figure S2A and B). The preceding results suggest that the T1IFN response following PNKP depletion is orchestrated via a canonical pathway involving ZBP1-cGAS-STING-TBK1 activities.

T1IFN response in PNKP-depleted cells is not mediated via the RIG-I/MAVS RNA sensing pathway

A recent study showed that ATR inhibition and exposure of multiple human and murine cells to IR can generate cytosolic RNA that subsequently potentiates a mitochondrial antiviral-signalling (MAVS)-dependent T1IFN response (11). Moreover, RNA viruses have been demonstrated to induce the T1IFN response (70). We, therefore, asked whether depletion of PNKP led to the generation of cytosolic RNA, which could potentially be sensed by the melanoma differentiation‐associated gene 5 (MDA5)/retinoic acid‐inducible gene I (RIG‐I)/MAVS pathway (71). RIG1 and MDA5 recognise short and long double-stranded RNA, respectively, become activated and converge on MAVS to signal through TBK1 and IRF3 to stimulate T1IFN response (72–76). Double depletion of PNKP and MAVS using siRNA failed to abrogate STAT1 phosphorylation in MCF7 cells (Supplementary Figure S4A), suggesting that the MDA5/RIG1/MAVS pathway is not required for the observed T1IFN response in MCF7 cells following downregulation of PNKP. This result also indicates that the transfected siRNAs used for many of our experiments were not inadvertently activating the cytosolic RNA sensors, which would have suggested a false positive result.

ROS combined with the loss of PNKP causes mtDNA damage that drives the T1IFN response

To further understand the mechanisms behind the increase in STAT1 phosphorylation following loss of PNKP, we next focused attention on the role that ROS may play in the process. We confirmed a role for ROS by incubating PNKP-depleted MCF7 cells with the general ROS scavenger, N-acetyl cysteine (NAC), as well as the superoxide scavenger, tiron. Incubation with either compound significantly reduced the levels of STAT1, and IRF3 phosphorylation in a dose dependent manner (Figure 6A, B and F, Supplementary Figure S5A, B and F). Next, we showed that incubation with the mitochondrial-targeting superoxide dismutase mimetic, mitoTEMPO, impaired STAT1 phosphorylation (Figure 6C and Supplementary Figure S5C), implicating the mitochondria as a source of the ROS in MCF7 cells following PNKP depletion. Treatment of PANC-1 with the different ROS scavengers also yielded a similar result (Supplementary Figure S5D).

Figure 6. Loss of PNKP cooperates with ROS to cause mtDNA damage that drives T1IFN response. (A) Increasing concentrations of the reactive oxygen species scavenger, N-acetyl cysteine (NAC) reduced the levels of phosphorylated STAT1 in PNKP-depleted MCF7 cells. Cells were treated with the indicated concentrations of NAC for up to 72 h prior to immunoblot analysis. (B) Increasing concentrations of the superoxide scavenger, tiron, reduced the T1IFN response in PNKP-depleted MCF7 cells. Cells were treated with increasing concentrations of tiron for 72 h prior to immunoblot analysis. (C) Treatment of PNKP-depleted MCF7 cells with mitoTEMPO abrogates T1IFN response. Following transfection, cells were incubated with the indicated concentrations of mitoTEMPO for 65 h prior to cell harvesting and immunoblotting. (D and E) Long PCR amplification of mitochondrial DNA in control and PNKP-depleted MCF7 (D) and PANC-1 (E) cells. Compared to the siControl cells, PNKP-depleted cells show reduced level of mtDNA integrity, indicating an accumulation of mtDNA damage following loss of PNKP. The relative levels of endogenous DNA damage were calculated by quantitating the long amplicon (16.6-kb for mtDNA) PCR product after normalizing for mtDNA copy number by PCR of a 211-bp mtDNA region. n = 3 and 4 replicates, respectively. Data are presented as standard error of the mean of n number of experiments (*P< 0.05, and ***P< 0.001 and ****P< 0.0001; ns = not significant). Statistical significance was determined using unpaired Student'st-test. (F) PNKP-depleted MCF7 cells incubated with the ROS scavengers, tiron and NAC reduced STAT1 phosphorylation and H2AX phosphorylation, suggesting a reduced level of DNA damage upon treatment with ROS scavengers. PNKP-depleted MCF7 cells were incubated with 3 mM tiron or NAC for 66 h prior to cell harvesting and western blotting. For the western blots shown in (A–F), whole cell extracts were used to determine protein expression.

Since we showed that treatment with the ROS scavengers, tiron, NAC and mitoTEMPO strongly reduced STAT1 activation, it suggested that oxidative damage to mtDNA and subsequent generation of DNA fragments and their eventual leakage into the cytosol may be responsible for the observed increase in STAT1 phosphorylation in PNKP-depleted cells. Therefore, to directly link loss of PNKP to mtDNA damage, we performed a long PCR amplification assay (42,43,77). This assay provides a sensitive method to detect DNA damage and is based on the concept that any alterations – in the form of single or double strand breaks or base modifications – due to oxidative damage to one or both strands of DNA will inhibit DNA polymerisation, thereby reducing the amount of the amplicon (78). The relative levels of endogenous DNA damage were calculated by quantitating the long amplicon (16.6-kb for mtDNA) PCR products after normalizing for mtDNA by amplification of a 211-bp mtDNA region. As shown in Figure 6D and E, downregulation of PNKP led to a significant reduction in the integrity of mtDNA in PNKP-depleted MCF7 and PANC-1 cells relative to the siControl cells, suggesting that loss of PNKP promotes the accumulation of oxidative damage to mtDNA. This agrees with previous reports by Mandal et al. (26) and Tahbaz et al. (23) of a reduction in the amplified mtDNA product following loss of PNKP in HEK293 and A549 cells. Furthermore, we validated the presence of PNKP in MCF7 cell mitochondria as previously identified in HEK293 and A549 cells (Supplementary Figure S5E). We also confirmed that ROS were propagating DNA damage in the absence of PNKP by detecting the presence of phosphorylated H2AX (γH2AX), a known marker of DNA damage (Figure 6F). Compared to the non-targeted control, loss of PNKP increased the levels of γH2AX, whereas incubation of PNKP-depleted MCF7 cells with 3 mM tiron or NAC for 66 h abrogated phosphorylated H2AX protein (Figure 6F), indicating that ROS induces significant levels of persistent DNA damage in the absence of PNKP. Collectively, the above results strongly support the involvement of ROS and their consequent damage to mtDNA to be responsible for the increase in STAT1 phosphorylation and resultant T1IFN response in a PNKP-deficient background. PNKP, therefore, plays an important role in maintaining mtDNA integrity and its loss can synergise with ROS to promote damage to mtDNA, which can potentially propagate induction of pro-inflammatory genes in certain cell lines.

We next sought to confirm that mtDNA serves as a source of the cytosolic DNA that is driving the T1IFN response in PNKP-depleted cells. Compared to nuclear DNA, mtDNA is more sensitive to oxidative damage, requiring longer repair time (78,79). Additionally, due to its bacterial origin, mtDNA is recognized as a pathogen-associated molecular pattern (PAMP) by the innate immune system (80) and is more likely to be resistant to clearance by cytosolic nucleases such as TREX1 than nuclear DNA due to its burden of oxidative damage (81), thereby making it more persistent and immunostimulatory. We, therefore, pre-incubated MCF7 cells for 6 days with 10 μM 2′,3′-dideoxycytidine (ddC), a deoxyribonucleoside analogue that is routinely used to inhibit mtDNA replication and consequently reduce mtDNA copy number (39,60,82), and then downregulated PNKP in both mtDNA-depleted and untreated MCF7 cells. We initially validated mtDNA depletion by ascertaining the relative levels of mtDNA-encoded genes at the genomic, mRNA and protein levels, with nuclear genome-encoded genes serving as controls. Our results showed that following depletion of mtDNA, the mtDNA-encoded genes (cytochrome c oxidase subunits 1 and 2, MTCO1 and 2) reduced significantly (≥50%) at the DNA, mRNA and protein levels relative to the untreated control cells (Supplementary Figure S6A–C). We also observed a marked decrease in the levels of mitochondrial transcription factor A (TFAM). In contrast, the nuclear DNA-encoded cytochrome c oxidase subunit 4 (MTCO4) and GAPDH were unchanged in both the treated and untreated cells (Supplementary Figure S6A and B). We then assessed STAT1 activation in mtDNA- and PNKP-depleted MCF7 cells. As shown in Figure 7A, depletion of mtDNA led to a reduction in pSTAT1 compared to the untreated control cells. A similar result was obtained with PANC-1 and T47D cells (Figure 7B and Supplementary Figure S6D and E). A similar response was also observed following exposure of MCF7 and T47D cells to low dose ethidium bromide (EtBr), another reagent used to deplete mtDNA copy number (37,38) (Figure 7B, C and Supplementary Figure S6D). This agrees with previous reports of a reduced T1IFN response following mtDNA depletion (37,83). Our data also agrees with a report that cytosolic DNA present in MCF7 cells is mainly derived from the mitochondria (60). We further validated these data by conducting RT-qPCR to analyse differential gene expression at the mRNA level. Compared to the untreated control cells, mtDNA-depleted MCF7 cells displayed significantly reduced levels of ISG15, CXCL10 and IFNβ1 (Figure 7D). Collectively, these results identify mtDNA damage following loss of PNKP to play an important role in the elevation of STAT1 phosphorylation and downstream upregulation of the interferon-stimulated genes. It is worth noting that we validated the non-involvement of TREX1 in the T1IFN response induced by PNKP loss by double depletion of PNKP and TREX1 and observed that pSTAT1 level was not dramatically affected relative to the pSTAT1 level obtained in PNKP-depleted cells (Supplementary Figure S4B). Additionally, we observed that loss of PNKP upregulates the levels of TREX1, likely in response to the accumulating cytosolic DNA (Figure 3D and Supplementary Figure S4B), which is in line with a report in the literature of induced TREX1 in response to increasing amount of cytosolic DNA (8). One would have expected the increased TREX1 level to clear the accumulating cytosolic DNA in the PNKP-depleted cells, thereby reducing pSTAT1 levels. However, the increased TREX1 levels in PNKP-depleted cells have no effect on pSTAT1 levels, suggesting that the cytosolic DNA is resistant to degradation by TREX1. This agrees with reports in the literature that TREX1 is unable to degrade oxidized DNA or DNA with a terminal 3′-phosphate (81,84).

Figure 7. Confirmation that leakage of mtDNA fragments into the cytoplasm is required to activate STAT1 in PNKP-depleted MCF7 cells. (A–C) Immunoblots showing that depletion of mtDNA using ddC (A and B) or EtBr (B and C) together with loss of PNKP in MCF7 (A and C), T47D and PANC-1 (B) cells reduced STAT1 activation to a level comparable to that obtained in the siControl cells with intact/depleted mtDNA. This suggests that loss of PNKP leads to mtDNA damage that resulted in leakage of mtDNA into the cytosol to activate STAT1. For the western blots shown in (A–C), whole cell extracts were used to determine protein expression. (D) RT-qPCR data show that mtDNA depletion together with loss of PNKP significantly reduced the expression of IFNβ1 and the ISGs, ISG15 and CXCL10, relative to MCF7 cells with intact mtDNA. Error bars represent standard error of the mean. Statistical difference determined using unpaired two-tailed Student's t-test. n = 3 independent experiments. ns = not significant. (*P< 0.05; **P< 0.01; ***P< 0.001; ****P< 0.0001). Cells were pre-treated with 10 μM ddC for 6 days or 100–150 ng/ml of EtBr for 20 days (in MCF7 cells) or for 6 days (in T47D cells) prior to PNKP depletion, cell harvesting and immunoblotting.

PNKP-depleted cells display increased accumulation of mitochondria-sourced cytosolic DNA

Immunofluorescence using an anti-dsDNA antibody revealed increased accumulation of cytosolic dsDNA in PNKP-depleted MCF7 cells relative to the non-targeted control MCF7 cells (Figure 8A). We further validated this by conducting a modified Terminal deoxynucleotidyl transferase (TdT) dUTP Nick-End Labeling (TUNEL) assay that is sensitive enough to detect the presence of cytosolic DNA. Our result shows colocalization of both positive signal from the anti-dsDNA antibody and TUNEL click chemistry reaction (Figure 8B), indicating the presence of dsDNA in the cytoplasm of PNKP-depleted cells. To determine the source of the cytosolic dsDNA generated following loss of PNKP, we depleted MCF7 cells of mtDNA using ddC over 6 days before knocking down PNKP in these cells, again using TFAM protein levels to confirm depletion of mtDNA copy number. We observed that downregulation of PNKP in control MCF7 cells generated cytoplasmic dsDNA, while this cytoplasmic dsDNA was dramatically reduced in PNKP-depleted MCF7 cells that were initially depleted of mtDNA (Figure 8C). We used the mtDNA stably-associated protein, TFAM (85), as a marker of the mitochondria in these experiments.

Figure 8. Loss of PNKP leads to the accumulation of cytoplasmic DNA sourced from the mitochondria. (A) Immunofluorescence reveals that PNKP-depleted MCF7 cells accumulate more cytoplasmic double stranded DNA (dsDNA), compared to the wild type (or siControl) counterparts. (B) Cells depleted of PNKP accumulate DNA fragments in the cytoplasm as identified by anti-dsDNA antibody and by performing a TUNEL reaction involving click-chemistry to label the cytoplasmic DNA. PNKP-depleted cells stained positive for both anti-dsDNA antibody and TUNEL click chemistry reaction with many regions of co-localisation (shown by white circles). (C) PNKP-depleted MCF7 cells accumulated cytoplasmic DNA, which was reduced following depletion of mtDNA, suggesting that mtDNA damage and consequent leakage into the cytoplasm could be responsible for STAT1 activation following loss of PNKP. In (A–C), fixed and permeabilised cells were stained with anti-dsDNA antibody (green), and DAPI (blue) to counterstain the nucleus. Immunolabelled cells were imaged using confocal microscopy (×40 oil lens). Z-stack images were taken and processed using maximum image projections function. (D and E) Treatment of PNKP-depleted MCF7 (D) and PANC-1 (E) cells with either CSA or VBIT4 led to a reduced level of STAT1 phosphorylation relative to the untreated control cells, indicating that leakage of mtDNA contributes to the observed STAT1 activation following loss of PNKP. For the western blots shown in (D and E), whole cell extracts were used to determine protein expression. (F) PNKP-depleted MCF7 cells accumulate more cytosolic DNA derived from the mitochondria compared to the siControl cells. 90 h post transfection, cells were harvested and subjected to subcellular fractionation prior to DNA purification and subsequent qPCR procedure. Error bars represent standard error of the mean. Statistical difference determined using unpaired two-tailed Student t-test. n = 4 independent experiments. (**P< 0.01; ***P< 0.001; statistical significance was determined using unpaired Student's t-test).

Under certain physiological conditions, such as oxidative stress or mitochondrial damage, the mitochondrial permeability transition pore (mPTP) is triggered to open, allowing leakage of mitochondrial contents including mtDNA into the cytosol to mediate the T1IFN response (86–89). We corroborated our findings by pharmacologically inhibiting mPTP using a specific and potent inhibitor, cyclosporin A (CSA) (88,90–94). Interestingly, treatment of PNKP-depleted MCF7 and PANC-1 cells with 5 μM CSA for 48 h reduced the levels of STAT1 phosphorylation compared to the untreated control group (Figure 8D and E). This is consistent with previous studies showing reduced STAT1 activation following pharmacological inhibition of mPTP with CSA (62,86,95,96). We validated this result by incubating PNKP-depleted MCF7 or PANC-1 cells with VBIT-4, an inhibitor of outer mitochondrial membrane voltage-dependent anion channel (VDAC), which blocks the oligomerization of mitochondrial pores, thereby preventing the release of mtDNA fragments into the cytosol (97,98). VBIT-4 treatment reduced STAT1 phosphorylation compared to the untreated control group in both MCF7 and PANC-1 cells (Figure 8D and E), strongly indicating that loss of PNKP causes leakage of mtDNA into the cytosol. To further corroborate our findings, we conducted qPCR on DNA extracted from the cytosolic and crude nuclear fractions after first confirming the purity of the different fractions (Supplementary Figure S7A). Relative to the siControl MCF7 cells, PNKP-depleted cells displayed up to 50% increase in the levels of cytosolic mtDNA accumulation (Figure 8F), which agrees with previous findings of increased mtDNA accumulation following DNA damage (40,65). Although we found that mtDNA leakage plays a significant role in the activation of the T1IFN response following PNKP depletion, nuclear DNA damage, and subsequent leakage of some DNA fragments into the cytosol could also be contributing to the increased STAT1 and IRF3 phosphorylation. This is because PNKP-depleted T47D or MCF7 cells incubated with the CDK1 specific inhibitor (RO-3306) – which induces cell cycle arrest at the G2/M phase thereby blocking mitotic progression – displayed a reduced level of STAT1 phosphorylation (Supplementary Figure S7B and C), which agrees with reports that prolonged cell cycle arrest prior to mitosis suppresses inflammatory signalling (99,100). In addition, to identify whether both mitochondrial- and nuclear-sourced DNA were co-operating to drive STAT1 phosphorylation following PNKP depletion, we incubated mtDNA-depleted T47D cells that already had PNKP downregulated with 10 μM CDK1 inhibitor and assessed for pSTAT1 levels. We observed that while mtDNA depletion reduced pSTAT1 levels relative to the NT control, CDK1 inhibitor treatment of the mtDNA-depleted cells modestly reduced pSTAT1 levels but did not completely abrogate pSTAT1 levels (Supplementary Figure S6F). Similarly, growth limitation by serum starvation in PNKP-depleted T47D reduced STAT1 phosphorylation, indicating that cell cycle progression was required for the induction of T1IFN response in the cell lines that are T1IFN-responsive to PNKP loss (Supplementary Figure S7D). Furthermore, incubation of PNKP-depleted T47D cells with the nuclear export (CRM1) inhibitor, leptomycin B (LMB), reduced STAT1 phosphorylation relative to the untreated control cells (Supplementary Figure S7C and E). Nonetheless, our data indicate that mtDNA makes significant contribution to the T1IFN response as dual inhibition of nuclear (using LMB) and mitochondrial (using VBIT4) export of genomic contents into the cytosol further reduced STAT1 phosphorylation compared to single treatment (Supplementary Figure S7E).

Discussion

Mutations in PNKP or its loss have been associated with different neurodegenerative and neurodevelopmental disorders (28–30), increased mtDNA damage (26) and elevated sensitivity of cancer cells to hydrogen peroxide and radiation (25). More recently, it was reported that radiation together with PNKP inhibition increases DNA damage that eventually activates the cGAS-STING pathway in non-small cell lung cancer (101). Our study is the first to directly link PNKP loss to activation of the innate immunity independent of radiation or exposure to other genotoxic agents. Our results agree with reports in the literature implicating DNA repair proteins as a limiting factor in the priming and activation of innate immunity in tumours (reviewed in (102,103)).

In this study, we have demonstrated that PNKP regulates T1IFN response in a variety of cell lines and that its loss significantly enhances ZBP1/cGAS/STING/TBK1/IRF3-dependent STAT1 phosphorylation via induction of mtDNA damage induced by ROS (see model Figure 9). Notably, this increase in T1IFN response following loss of PNKP was not further potentiated by radiation, suggesting that loss of PNKP is sufficient to activate this pathway at least in some cell lines. It is possible that the time point of 72 h post-irradiation used in this study is not sufficient to activate STAT1 as a report suggests that STAT1 activation is time (between 3 and 6 days) and cell cycle dependent following irradiation (99). Although loss of PNKP robustly activated STAT1 in most of the cancer cell lines we tested, some cell lines, including MDA-MB-231, were non-responsive. It is possible that some of these non-responsive cell lines are defective in (or express very low levels of) key proteins such as STING that are involved in the T1IFN response (45). Moreover, it has been demonstrated that genotoxic stress failed to activate STAT1 or upregulate ISGs in MDA-MB-231 (104). In addition, not all cancer cell lines are adept at activating the cGAS-STING-TBK1-IRF3 signalling pathway, either due to harbouring a defective downstream protein or an inability to respond to secreted interferons in a paracrine or autocrine fashion. This differential response of the different cancer cell lines to loss of PNKP warrants further study to understand the underlying mechanisms.

Figure 9. Schematic diagram illustrating the underlying mechanism of how loss of PNKP activates the type 1 interferon response. Left panel: in the presence of PNKP, ROS-induced DNA damage fails to activate the T1IFN response as the strand break termini are constantly repaired by PNKP. However, in the absence of PNKP, the strand breaks induced by ROS are persistent, leading to the generation of smaller DNA fragments that are bound by ZBP1 and/or cGAS. Middle panel: activated cGAS synthesizes the second messenger molecule 2′3′-cGAMP, which binds to and activates STING leading to the downstream phosphorylation and activation of TBK1, IRF3 and subsequent synthesis and secretion of type 1 interferons such as IFNβ. Likewise, ZBP1, bound by oxidized DNA, is activated, and in complex with cGAS augments the activity of cGAS and STING. Right panel: the secreted IFNβ binds to its cognate receptor thereby promoting downstream phosphorylation events involving sequential JAK/TYK1 activation, STAT1/STAT2 activation and complex formation with IRF9. This ISGF3 complex translocates to the nucleus where it turns on interferon-stimulated genes. (Figure created with BioRender.com).

Our results identify IFNβ1 as the key cytokine responsible for the activation of STAT1 following loss of PNKP. We show that the secreted IFNβ1 was present in the conditioned media and that it binds to its cognate receptor to promote STAT1 phosphorylation as determined by neutralization, immunoblotting, and gene expression experiments with anti-IFNAR2 blocking antibody. Our findings above agree with the canonical role of IFNβ1 in the activation of STAT1 via binding to cognate cell surface receptors and consequent downstream STAT1 phosphorylation facilitated by increased JAK1/2 autophosphorylation (105). The consistent induction of the chemo-attractants, CXCL10 and CCL5 in PNKP-depleted cells strongly indicates that pharmacological inhibition of PNKP could potentially drive the recruitment of dendritic cells to the tumour microenvironment, and in turn convert a rather immunologically silent tumour to an immune-competent tumour.

We also mapped the upstream proteins responsible for the increased STAT1 phosphorylation following loss of PNKP, and identified cGAS, STING, TBK1, IRF3 and JAK1/2 as important mediators of innate immunity in cancer cells, suggesting that the observed T1IFN response following loss of PNKP follows the canonical cGAS-STING pathway, except in T47D cells where we observed that ZBP1 was the major driver of STAT1 activation independent of cGAS. Recently, ZBP1 was shown to stabilize the Z-form of mtDNA in a complex involving cGAS to promote the T1IFN response (65). Moreover, ZBP1 has been associated with low-level oxidative stress-induced release of mtDNA that mediates ZBP1 activation and a subsequent induction of pro-inflammatory genes (62,63,65,106). Additionally, one study reported direct in vitro interaction of ZBP1 with mtDNA (107), suggesting that ZBP1 can potentially bind mtDNA. Furthermore, it was found that depletion of either ZBP1 or STING suppresses IFNβ1 or IFNα induction in necrotic HT29 cells (64). Similarly, we showed that ZBP1 is required to activate the T1IFN response as double depletion of PNKP and ZBP1 abrogates STAT1 phosphorylation in both T47D and MCF7 cells. It is plausible that loss of PNKP generates Z-form DNA that is sensed and stabilized by ZBP1, and in concert with cGAS leads to the activation of the T1IFN response. Our study, therefore, identifies a unique pathway involving ZBP1-cGAS-STING-TBK1-IRF3 signalling pathway to be responsible for STAT1 phosphorylation in MCF7 and partly in T47D cells following loss of PNKP. Recently, it was reported that ZBP1 regulates STING activity as its loss led to reduced STING dimerization and phosphorylation of TBK1 following irradiation (108). It is also possible that, once activated, ZBP1 directly regulates STING activity in MCF7 and T47D cells, hence its loss dramatically impairs STAT1 activation.

Although STING has been shown to be an ISG (69), inhibition of IFNβ1 and IFNAR2 failed to significantly reduce its protein levels, which is in contrast to ISG15, which was reduced both at the protein and transcript levels following incubation with the neutralizing antibodies against IFNβ1 or IFNAR2. The disparity between the two ISGs in response to IFNβ1 or IFNAR2 inhibition could stem from the effect of activated NF-κB on STING degradation. Zhang et al. recently showed that upon activation, NF-κB prevents STING degradation by altering microtubule-based STING transport from the Golgi apparatus to the lysosome, thereby increasing STING signalling (109). It is, therefore, possible that STING is initially upregulated by IFNβ1 stimulation, but later becomes stabilised by activated NF-κB, hence inhibition of IFNβ1 or IFNAR2 no longer has an effect on STING compared to ISG15. The above assumption requires further investigation in the context of PNKP inhibition. Irrespective of the mechanisms involved in the accumulation of STING following loss of PNKP, the observed increase in STING protein levels in PNKP-depleted cells is very promising as high STING expression in tumours has been associated with overall immune cell infiltration and modulation of the anti-cancer immune response (110–113). High STING expression in PNKP-depleted cancer cells further highlights the need to develop inhibitors against PNKP to improve the efficacy of immune checkpoint inhibitors in combination with radiotherapy against tumours.

Our data obtained with ROS scavengers and mtDNA depletion indicate that loss of PNKP combines with ROS to augment the level of endogenous mtDNA damage. Low-level oxidative stress has been shown to increase mtDNA damage (62). The elevated STAT1 phosphorylation levels in PNKP-depleted cells could be further driven by ROS, since STAT1-induced ROS production (and vice versa) has been described previously in MCF7 cells (114). Our results also identify the mitochondria as a source of ROS production as treatment with the mitochondria-targeted antioxidant, mitoTEMPO, impaired STAT1 phosphorylation. The loss of mtDNA integrity due to ROS-induced damage translates into leakage of mtDNA into the cytosol. This cytosolic mtDNA is bound and stabilised by ZBP1, which in complex with cGAS drives downstream STAT1 activation (65). This was corroborated by demonstrating that depletion of mtDNA impaired STAT1 phosphorylation and induction of pro-inflammatory genes, and that treatment with CSA and VBIT-4, both of which block release of mitochondrial content into the cytosol (90,97,98), also led to reduced phosphorylated STAT1. These results are congruent with the already established role that PNKP plays in the maintenance of mtDNA within the base excision repair pathway (23,26) and highlight the need for cells to possess an arsenal of DNA repair proteins to prevent mtDNA-induced autoimmunity. While we have demonstrated that much of the cytosolic DNA is sourced from the mitochondria, there is also the possibility that nuclear DNA may contribute to the observed STAT1 phosphorylation. First, the observed increase in γH2AX levels is suggestive of nuclear DNA double strand breaks due to loss of PNKP. Second, treatment of PNKP-depleted MCF7 and T47D cells with the CDK1 inhibitor reduces STAT1 and IRF3 phosphorylation, implying that leakage of nuclear DNA fragments, which is dependent on cell cycle progression (99,100), contributes to the observed STAT1 phosphorylation. Lastly, incubation of PNKP-depleted cells with the nuclear export inhibitor, LMB, dramatically reduces STAT1 phosphorylation, indicating that export of materials (including proteins and DNA fragments) into the cytosol is a prerequisite for T1IFN response following loss of PNKP. It could be inferred from the latter that LMB treatment of PNKP-depleted cells prevents nuclear export of cGAS to the cytosol, hence reduced STAT1 phosphorylation. Buttressing this assertion is a recent finding that a nuclear export signal is required for cGAS to sense cytosolic DNA, and that LMB blocks cGAS accumulation in the cytosol (115).

There are several potential clinical implications of this study. First, insights from this study offer further support for the development of inhibitors against PNKP. Since radiation and ROS can induce strand breaks with 3′-phosphate and 5′-hydroxyl termini that must be processed by PNKP prior to re-joining by DNA polymerase and ligases (116,117), a PNKP inhibitor could potentially be a good sensitizer to induce further DNA damage. Second, it further highlights the potential to target DNA repair proteins to increase the immunogenicity of some rather immunologically silent tumours, of which breast cancers are a good candidate (reviewed in (118,119). Moreover, this study adds PNKP to a growing list of candidate DNA repair proteins, including ATM (10), ATR (11) and PARP1 (12) whose dysregulation has been demonstrated to enhance T1IFN-mediated innate immunity in cancer cells. Another potential consequence of activation of the T1IFN response as a result of PNKP depletion relates to the neurological disorders associated with PNKP mutations. These include the neurodevelopmental disorder microcephaly with seizures (MCSZ) and the neurodegenerational disorders ataxia with ocular apraxia type 4 (AOA4) and a variant of Charcot Marie Tooth disease (28,29). Given the growing evidence that STING activation can lead to neuroinflammation and CNS injury (120–123), it will be important to ascertain if any of the PNKP-related disorders stem from an aberrant innate immune response.

In conclusion, our study emphasizes the importance of PNKP in regulating the TI1FN-mediated innate immune response and shows that loss of PNKP cooperates with ROS to cause mtDNA damage that eventually results in leakage and accumulation of mtDNA fragments in the cytosol and the resultant downstream STAT1 activation and induction of the ISGs (see model Figure 9). In addition, this study suggests a strong link between PNKP inhibition and modulation of the tumour microenvironment since PNKP inhibition can potentially increase IFNβ1 secretion in vivo to drive dendritic cell recruitment and maturation, and subsequent priming and activation of T cells for tumour cell recognition and eradication. Lastly, this study provides strong support for the development of inhibitors against PNKP to increase the immunogenicity of some cancers as well as improve the outcome of immunotherapy.

Supplementary Material

gkae654_Supplemental_File

Acknowledgements

We are indebted to Drs Xuejun Sun and Guobin Sun for their help with cell imaging, and Drs Kristi Baker, Mark Glover, and Gordon Chan (University of Alberta) for their useful discussion and advice. The authors are also thankful to ThermoFisher Scientific for the provision of free sample antibodies and reagents through their Aspire program. Graphical abstract and Figure 9 created with BioRender.com.

Author contributions: Conceptualization, W.D.K., M.W.; Methodology, W.D.K., M.F.; Writing – Original Draft, W.D.K; Writing – Review and Editing, M.W.; Funding Acquisition, M.W., W.D.K.; Supervision M.W.

Data availability

The data underlying this article will be shared on reasonable request to the corresponding author.

Supplementary data

Supplementary Data are available at NAR Online.

Funding

Cancer Research Institute (CLIP) [CRI3714]; Canadian Institutes of Health Research [PJT168869]; W.D.K. received scholarships through the following initiatives: the Terry Fox Foundation – Strategic Training in Transdisciplinary Radiation Science for the 21st Century (STARS21) program; Dr Herbert Meltzer Memorial Fellowship through the Alberta Cancer Foundation, Marathon of Hope Graduate Studentships in Breast Cancer or Glioblastoma Research through the Cancer Research Institute of Northern Alberta (CRINA), University of Alberta (Doctoral Recruitment Scholarship, Faculty of Medicine and Dentistry 75th Anniversary and Antoine Noujaim Graduate Entrance Scholarship). Funding for open access charge: Canadian Institutes of Health Research held at the University of Alberta.

Conflict of interest statement. None declared.

Notes

Present address: Michael Weinfeld, Department of Oncology, University of Alberta, Edmonton, Alberta T6G 2R3, Canada.
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References

1. Sun L. , WuJ., DuF., ChenX., ChenZ.J. Cyclic GMP-AMP synthase is a cytosolic DNA sensor that activates the type I interferon pathway. Science. 2013; 339 :786–791.23258413
2. Li X. , ShuC., YiG., ChatonC.T., SheltonC.L., DiaoJ., ZuoX., KaoC.C., HerrA.B., LiP. Cyclic GMP-AMP synthase is activated by double-stranded DNA-induced oligomerization. Immunity. 2013; 39 :1019–1031.24332030
3. Li X.D. , WuJ., GaoD., WangH., SunL., ChenZ.J. Pivotal roles of cGAS-cGAMP signaling in antiviral defense and immune adjuvant effects. Science. 2013; 341 :1390–1394.23989956
4. Samanta M. , IwakiriD., KandaT., ImaizumiT., TakadaK. EB virus-encoded RNAs are recognized by RIG-I and activate signaling to induce type I IFN. EMBO J. 2006; 25 :4207–4214.16946700
5. Lund J.M. , AlexopoulouL., SatoA., KarowM., AdamsN.C., GaleN.W., IwasakiA., FlavellR.A. Recognition of single-stranded RNA viruses by toll-like receptor 7. Proc. Nat. Acad. Sci. U.S.A. 2004; 101 :5598–5603.
6. Dobbs N. , BurnaevskiyN., ChenD., GonuguntaV.K., AltoN.M., YanN. STING activation by translocation from the ER is associated with infection and autoinflammatory disease. Cell Host Microbe. 2015; 18 :157–168.26235147
7. Bai J. , LiuF. The cGAS-cGAMP-STING pathway: a molecular link between immunity and metabolism. Diabetes. 2019; 68 :1099–1108.31109939
8. Vanpouille-Box C. , AlardA., AryankalayilM.J., SarfrazY., DiamondJ.M., SchneiderR.J., InghiramiG., ColemanC.N., FormentiS.C., DemariaS. DNA exonuclease Trex1 regulates radiotherapy-induced tumour immunogenicity. Nat. Commun. 2017; 8 :15618.28598415
9. Erdal E. , HaiderS., RehwinkelJ., HarrisA.L., McHughP.J. A prosurvival DNA damage-induced cytoplasmic interferon response is mediated by end resection factors and is limited by Trex1. Genes Dev. 2017; 31 :353–369.28279982
10. Hu M. , ZhouM., BaoX., PanD., JiaoM., LiuX., LiF., LiC.-Y. ATM inhibition enhances cancer immunotherapy by promoting mtDNA leakage/cGAS-STING activation. J. Clin. Invest. 2020; 131 :e139333.
11. Feng X. , TubbsA., ZhangC., TangM., SridharanS., WangC., JiangD., SuD., ZhangH., ChenZ.et al . ATR inhibition potentiates ionizing radiation-induced interferon response via cytosolic nucleic acid-sensing pathways. EMBO J. 2020; 39 :e104036.32484965
12. Shen J. , ZhaoW., JuZ., WangL., PengY., LabrieM., YapT., MillsG.B., PengG. PARPi triggers STING-dependent immune response and enhances therapeutic efficacy of immune checkpoint blockade independent of BRCAness. Cancer Res. 2018; 318980.
13. Andrade B. , Jara-GutiérrezC., Paz-AraosM., VázquezM.C., DíazP., MurgasP. The relationship between reactive oxygen species and the cGAS/STING signaling pathway in the inflammaging process. Int. J. Mol. Sci. 2022; 23 :15182.36499506
14. Shekhova E. Mitochondrial reactive oxygen species as major effectors of antimicrobial immunity. PLoS Pathog. 2020; 16 :e1008470.32463825
15. Commoner B. , TownsendJ., PakeG.E. Free radicals in biological materials. Nature. 1954; 174 :689–691.13213980
16. Dizdaroglu M. Formation of an 8-hydroxyguanine moiety in deoxyribonucleic acid on gamma-irradiation in aqueous solution. Biochemistry. 1985; 24 :4476–4481.4052410
17. Haghdoost S. , CzeneS., NäslundI., SkogS., Harms-RingdahlM. Extracellular 8-oxo-dG as a sensitive parameter for oxidative stress in vivo and in vitro. Free Radic. Res. 2005; 39 :153–162.15763963
18. Dianov G.L. , ParsonsJ.L. Co-ordination of DNA single strand break repair. DNA Repair (Amst.). 2007; 6 :454–460.17123872
19. Caldecott K.W. Single-strand break repair and genetic disease. Nat. Rev. Genet. 2008; 9 :619–631.18626472
20. Fisher L.A. , SamsonL., BesshoT. Removal of reactive oxygen species-induced 3′-blocked ends by XPF-ERCC1. Chem. Res. Toxicol. 2011; 24 :1876–1881.22007867
21. Freschauf G.K. , Karimi-BusheriF., Ulaczyk-LesankoA., MereniukT.R., AhrensA., KoshyJ.M., Rasouli-NiaA., PasarjP., HolmesC.F.B., RininslandF.et al . Identification of a small molecule inhibitor of the Human DNA repair enzyme polynucleotide kinase/phosphatase. Cancer Res. 2009; 69 :7739–7746.19773431
22. Karimi-Busheri F. , DalyG., RobinsP., CanasB., PappinD.J.C., SgourosJ., MillerG.G., FakhraiH., DavisE.M., Le BeauM.M.et al . Molecular characterization of a Human DNA kinase. J. Biol. Chem. 1999; 274 :24187–24194.10446193
23. Tahbaz N. , SubediS., WeinfeldM. Role of polynucleotide kinase/phosphatase in mitochondrial DNA repair. Nucleic Acids Res. 2012; 40 :3484–3495.22210862
24. Weinfeld M. , ManiR.S., AbdouI., AceytunoR.D., GloverJ.N. Tidying up loose ends: the role of polynucleotide kinase/phosphatase in DNA strand break repair. Trends Biochem. Sci. 2011; 36 :262–271.21353781
25. Rasouli-Nia A. , Karimi-BusheriF., WeinfeldM. Stable down-regulation of human polynucleotide kinase enhances spontaneous mutation frequency and sensitizes cells to genotoxic agents. Proc. Nat. Acad. Sci. U.S.A. 2004; 101 :6905–6910.
26. Mandal S.M. , HegdeM.L., ChatterjeeA., HegdeP.M., SzczesnyB., BanerjeeD., BoldoghI., GaoR., FalkenbergM., GustafssonC.M.et al . Role of Human DNA glycosylase nei-like 2 (NEIL2) and single strand break repair protein polynucleotide kinase 3′-phosphatase in maintenance of mitochondrial genome. J. Biol. Chem. 2012; 287 :2819–2829.22130663
27. Shin W. , AlpaughW., HallihanL.J., SinhaS., CrowtherE., MartinG.R., Scheidl-YeeT., YangX., YoonG., GoldsmithT.et al . PNKP is required for maintaining the integrity of progenitor cell populations in adult mice. Life Sci. Allian. 2021; 4 :e202000790.
28. Bras J. , AlonsoI., BarbotC., CostaM.M., DarwentL., OrmeT., SequeirosJ., HardyJ., CoutinhoP., GuerreiroR. Mutations in PNKP cause recessive ataxia with oculomotor apraxia type 4. Am. J. Hum. Genet. 2015; 96 :474–479.25728773
29. Taniguchi-Ikeda M. , MorisadaN., InagakiH., OuchiY., TakamiY., TachikawaM., SatakeW., KobayashiK., TsuneishiS., TakadaS.et al . Two patients with PNKP mutations presenting with microcephaly, seizure, and oculomotor apraxia. Clin. Genet. 2018; 93 :931–933.29243230
30. Leal A. , Bogantes-LedezmaS., EkiciA.B., UebeS., ThielC.T., StichtH., BerghoffM., BerghoffC., MoreraB., MeisterernstM.et al . The polynucleotide kinase 3′-phosphatase gene (PNKP) is involved in Charcot-Marie-Tooth disease (CMT2B2) previously related to MED25. Neurogenetics. 2018; 19 :215–225.30039206
31. Gao R. , ChakrabortyA., GeaterC., PradhanS., GordonK.L., SnowdenJ., YuanS., DickeyA.S., ChoudharyS., AshizawaT.et al . Mutant huntingtin impairs PNKP and ATXN3, disrupting DNA repair and transcription. eLife. 2019; 8 :e42988.30994454
32. Quek H. , LuffJ., CheungK., KozlovS., GateiM., LeeC.S., BellinghamM.C., NoakesP.G., LimY.C., BarnettN.L.et al . Rats with a missense mutation in Atm display neuroinflammation and neurodegeneration subsequent to accumulation of cytosolic DNA following unrepaired DNA damage. J. Leukocyte Biol. 2017; 101 :927–947.27895165
33. Crow Y.J. , StetsonD.B. The type I interferonopathies: 10 years on. Nat. Rev. Immunol. 2022; 22 :471–483.34671122
34. Tsukada K. , MatsumotoY., ShimadaM. Linker region is required for efficient nuclear localization of polynucleotide kinase phosphatase. PLoS One. 2020; 15 :e0239404.32970693
35. Banerjee D. , LangbergK., AbbasS., OdermattE., YerramothuP., VolaricM., ReidenbachM.A., KrentzK.J., RubinsteinC.D., BrautiganD.L.et al . A non-canonical, interferon-independent signaling activity of cGAMP triggers DNA damage response signaling. Nat. Commun. 2021; 12 :6207.34707113
36. Zheng X. , WangK., PanL., HaoW., XueY., BacsiA., VlahopoulosS.A., RadakZ., HazraT.K., BrasierA.R.et al . Innate immune responses to RSV infection facilitated by OGG1, an enzyme repairing oxidatively modified DNA base lesions. J. Innate Immun. 2022; 14 :593–614.35512649
37. White M.J. , McArthurK., MetcalfD., LaneR.M., CambierJ.C., HeroldM.J., van DelftM.F., BedouiS., LesseneG., RitchieM.Eet al . Apoptotic caspases suppress mtDNA-induced STING-mediated type I IFN production. Cell. 2014; 159 :1549–1562.25525874
38. Hashiguchi K. , Zhang-AkiyamaQ.M. Establishment of human cell lines lacking mitochondrial DNA. Methods Mol. Biol. 2009; 554 :383–391.19513686
39. Rongvaux A. , JacksonR., HarmanC.C., LiT., WestA.P., de ZoeteM.R., WuY., YordyB., LakhaniS.A., KuanC.Y.et al . Apoptotic caspases prevent the induction of type I interferons by mitochondrial DNA. Cell. 2014; 159 :1563–1577.25525875
40. West A.P. , Khoury-HanoldW., StaronM., TalM.C., PinedaC.M., LangS.M., BestwickM., DuguayB.A., RaimundoN., MacDuffD.A.et al . Mitochondrial DNA stress primes the antiviral innate immune response. Nature. 2015; 520 :553–557.25642965
41. Uggenti C. , LepelleyA., DeppM., BadrockA.P., RoderoM.P., El-DaherM.-T., RiceG.I., DhirS., WheelerA.P., DhirA.et al . 2020) cGAS-mediated induction of type I interferon due to inborn errors of histone pre-mRNA processing. Nat. Genet. 52 :1364–1372.33230297
42. Moro L. , ArbiniA.A., YaoJ.L., di Sant’AgneseP.A., MarraE., GrecoM. Mitochondrial DNA depletion in prostate epithelial cells promotes anoikis resistance and invasion through activation of PI3K/Akt2. Cell Death Differ. 2009; 16 :571–583.19079138
43. Santos J.H. , MeyerJ.N., MandavilliB.S., Van HoutenB. Quantitative PCR-based measurement of nuclear and mitochondrial DNA damage and repair in mammalian cells. Methods Mol. Biol. 2006; 314 :183–199.16673882
44. Cheon H. , Holvey-BatesE.G., SchogginsJ.W., ForsterS., HertzogP., ImanakaN., RiceC.M., JacksonM.W., JunkD.J., StarkG.R. IFNβ-dependent increases in STAT1, STAT2, and IRF9 mediate resistance to viruses and DNA damage. EMBO J. 2013; 32 :2751–2763.24065129
45. Xia T. , KonnoH., AhnJ., BarberG.N. Deregulation of STING signaling in colorectal carcinoma constrains DNA damage responses and correlates with tumorigenesis. Cell Rep. 2016; 14 :282–297.26748708
46. Chen J. , MarkelcB., KaepplerJ., OgundipeV.M.L., CaoY., McKennaW.G., MuschelR.J. STING-dependent interferon-λ1 induction in HT29 cells, a Human colorectal cancer cell line, after gamma-radiation. Int. J. Radiat. Oncol. Biol. Phys. 2018; 101 :97–106.29619982
47. Suter M.A. , TanN.Y., ThiamC.H., KhatooM., MacAryP.A., AngeliV., GasserS., ZhangY.L. cGAS–STING cytosolic DNA sensing pathway is suppressed by JAK2-STAT3 in tumor cells. Sci. Rep. 2021; 11 :7243.33790360
48. Breslin C. , CaldecottK.W. DNA 3′-phosphatase activity is critical for rapid global rates of single-strand break repair following oxidative stress. Mol. Cell. Biol. 2009; 29 :4653–4662.19546231
49. Chabot V. , ReverdiauP., IochmannS., RicoA., SénécalD., GoupilleC., SizaretP.Y., SensebéL. CCL5-enhanced human immature dendritic cell migration through the basement membrane in vitro depends on matrix metalloproteinase-9. J. Leukocyte Biol. 2006; 79 :767–778.16434695
50. Levy D.E. , DarnellJ.E. STATs: transcriptional control and biological impact. Nat. Rev. Mol. Cell Biol. 2002; 3 :651–662.12209125
51. Platanias L.C. Mechanisms of type-I- and type-II-interferon-mediated signalling. Nat. Rev. Immunol. 2005; 5 :375–386.15864272
52. Krämer O.H. , KnauerS.K., GreinerG., JandtE., ReichardtS., GührsK.-H., StauberR.H., BöhmerF.D., HeinzelT. A phosphorylation-acetylation switch regulates STAT1 signaling. Genes Dev. 2009; 23 :223–235.19171783
53. Fu X.Y. , SchindlerC., ImprotaT., AebersoldR., DarnellJ.E. The proteins of ISGF-3, the interferon alpha-induced transcriptional activator, define a gene family involved in signal transduction. Proc. Nat. Acad. Sci. U.S.A. 1992; 89 :7840–7843.
54. Schindler C. , FuX.Y., ImprotaT., AebersoldR., DarnellJ.E.Jr Proteins of transcription factor ISGF-3: one gene encodes the 91-and 84-kDa ISGF-3 proteins that are activated by interferon alpha. Proc. Nat. Acad. Sci. U.S.A. 1992; 89 :7836–7839.
55. Zhang Q. , GreenM.D., LangX., LazarusJ., ParselsJ.D., WeiS., ParselsL.A., ShiJ., RamnathN., WahlD.R.et al . Inhibition of ATM increases interferon signaling and sensitizes pancreatic cancer to immune checkpoint blockade therapy. Cancer Res. 2019; 79 :3940–3951.31101760
56. Murira A. , LamarreA. Type-I interferon responses: from friend to foe in the battle against chronic viral infection. Front. Immunol. 2016; 7 :609.28066419
57. Deng L. , LiangH., XuM., YangX., BurnetteB., ArinaA., LiX.D., MauceriH., BeckettM., DargaT.et al . STING-dependent cytosolic DNA sensing promotes radiation-induced type I interferon-dependent antitumor immunity in immunogenic tumors. Immunity. 2014; 41 :843–852.25517616
58. Dunphy G. , FlanneryS.M., AlmineJ.F., ConnollyD.J., PaulusC., JønssonK.L., JakobsenM.R., NevelsM.M., BowieA.G., UnterholznerL. Non-canonical activation of the DNA sensing adaptor STING by ATM and IFI16 mediates NF-κb signaling after nuclear DNA damage. Mol. Cell. 2018; 71 :745–760.30193098
59. Lama L. , AduraC., XieW., TomitaD., KameiT., KuryavyiV., GogakosT., SteinbergJ.I., MillerM., Ramos-EspirituL.et al . Development of human cGAS-specific small-molecule inhibitors for repression of dsDNA-triggered interferon expression. Nat. Commun. 2019; 10 :2261.31113940
60. Wu L. , CaoJ., CaiW.L., LangS.M., HortonJ.R., JansenD.J., LiuZ.Z., ChenJ.F., ZhangM., MottB.T.et al . KDM5 histone demethylases repress immune response via suppression of STING. PLoS Biol. 2018; 16 :e2006134.30080846
61. Zierhut C. , YamaguchiN., ParedesM., LuoJ.-D., CarrollT., FunabikiH. The cytoplasmic DNA sensor cGAS promotes mitotic cell death. Cell. 2019; 178 :302–315.31299200
62. Szczesny B. , MarcattiM., AhmadA., MontalbanoM., BrunyánszkiA., BibliS.-I., PapapetropoulosA., SzaboC. Mitochondrial DNA damage and subsequent activation of Z-DNA binding protein 1 links oxidative stress to inflammation in epithelial cells. Sci. Rep. 2018; 8 :914.29343810
63. Takaoka A. , WangZ., ChoiM.K., YanaiH., NegishiH., BanT., LuY., MiyagishiM., KodamaT., HondaK.et al . DAI (DLM-1/ZBP1) is a cytosolic DNA sensor and an activator of innate immune response. Nature. 2007; 448 :501–505.17618271
64. Chen D. , TongJ., YangL., WeiL., StolzD.B., YuJ., ZhangJ., ZhangL. PUMA amplifies necroptosis signaling by activating cytosolic DNA sensors. Proc. Nat. Acad. Sci. U.S.A. 2018; 115 :3930–3935.
65. Lei Y. , VanPortflietJ.J., ChenY.-F., BryantJ.D., LiY., FailsD., Torres-OdioS., RaganK.B., DengJ., MohanA.et al . Cooperative sensing of mitochondrial DNA by ZBP1 and cGAS promotes cardiotoxicity. Cell. 2023; 186 :3013–3032.37352855
66. Fu Y. , ComellaN., TognazziK., BrownL.F., DvorakH.F., KocherO. Cloning of DLM-1, a novel gene that is up-regulated in activated macrophages, using RNA differential display. Gene. 1999; 240 :157–163.10564822
67. Yang D. , LiangY., ZhaoS., DingY., ZhuangQ., ShiQ., AiT., WuS.Q., HanJ. ZBP1 mediates interferon-induced necroptosis. Cell Mol Immunol. 2020; 17 :356–368.31076724
68. Hu Y. , ManasrahB.K., McGregorS.M., LeraR.F., NormanR.X., TuckerJ.B., ScribanoC.M., YanR.E., HumayunM., WisinskiK.B.et al . Paclitaxel induces micronucleation and activates pro-inflammatory cGAS-STING signaling in triple-negative breast cancer. Mol. Cancer Ther. 2021; 20 :2553–2567.34583980
69. Ma F. , LiB., YuY., IyerS.S., SunM., ChengG. Positive feedback regulation of type I interferon by the interferon-stimulated gene STING. EMBO Rep. 2015; 16 :202–212.25572843
70. Flür K. , AllamR., ZecherD., KulkarniO.P., LichtnekertJ., SchwarzM., BeutlerB., VielhauerV., AndersH.-J. Viral RNA induces type I interferon-dependent cytokine release and cell death in mesangial cells via melanoma-differentiation-associated gene-5: implications for viral infection-associated glomerulonephritis. Am. J. Pathol. 2009; 175 :2014–2022.19850889
71. Wu J. , ChenZ.J. Innate immune sensing and signaling of cytosolic nucleic acids. Annu. Rev. Immunol. 2014; 32 :461–488.24655297
72. Brisse M. , LyH. Comparative structure and function analysis of the RIG-I-Like receptors: RIG-I and MDA5. Front. Immunol. 2019; 10 :1586–1586.31379819
73. Triantafilou K. , VakakisE., KarS., RicherE., EvansG.L., TriantafilouM. Visualisation of direct interaction of MDA5 and the dsRNA replicative intermediate form of positive strand RNA viruses. J. Cell Sci. 2012; 125 :4761–4769.22797917
74. Kato H. , TakeuchiO., Mikamo-SatohE., HiraiR., KawaiT., MatsushitaK., HiiragiA., DermodyT.S., FujitaT., AkiraS. Length-dependent recognition of double-stranded ribonucleic acids by retinoic acid-inducible gene-I and melanoma differentiation-associated gene 5. J. Exp. Med. 2008; 205 :1601–1610.18591409
75. Anchisi S. , GuerraJ., GarcinD. RIG-I ATPase activity and discrimination of self-RNA versus non-self-RNA. mBio. 2015; 6 :e02349.25736886
76. Hornung V. , EllegastJ., KimS., BrzózkaK., JungA., KatoH., PoeckH., AkiraS., ConzelmannK.K., SchleeM.et al . 5′-Triphosphate RNA is the ligand for RIG-I. Science. 2006; 314 :994–997.17038590
77. Ayala-Torres S. , ChenY., SvobodaT., RosenblattJ., Van HoutenB. Analysis of gene-specific DNA damage and repair using quantitative polymerase chain reaction. Methods. 2000; 22 :135–147.11020328
78. Yakes F.M. , Van HoutenB. Mitochondrial DNA damage is more extensive and persists longer than nuclear DNA damage in human cells following oxidative stress. Proc. Nat. Acad. Sci. U.S.A. 1997; 94 :514–519.
79. Furda A.M. , MarrangoniA.M., LokshinA., Van HoutenB. Oxidants and not alkylating agents induce rapid mtDNA loss and mitochondrial dysfunction. DNA Repair (Amst.). 2012; 11 :684–692.22766155
80. Zhang Q. , RaoofM., ChenY., SumiY., SursalT., JungerW., BrohiK., ItagakiK., HauserC.J. Circulating mitochondrial DAMPs cause inflammatory responses to injury. Nature. 2010; 464 :104–107.20203610
81. Gehrke N. , MertensC., ZillingerT., WenzelJ., BaldT., ZahnS., TütingT., HartmannG., BarchetW. Oxidative damage of DNA confers resistance to cytosolic nuclease TREX1 degradation and potentiates STING-dependent immune sensing. Immunity. 2013; 39 :482–495.23993650
82. Kasashima K. , SumitaniM., EndoH. Human mitochondrial transcription factor A is required for the segregation of mitochondrial DNA in cultured cells. Exp. Cell. Res. 2011; 317 :210–220.20955698
83. Yamazaki T. , KirchmairA., SatoA., BuquéA., RybsteinM., PetroniG., BloyN., FinotelloF., StaffordL., Navarro ManzanoE.et al . Mitochondrial DNA drives abscopal responses to radiation that are inhibited by autophagy. Nat. Immunol. 2020; 21 :1160–1171.32747819
84. Inamdar K.V. , YuY., PovirkL.F. Resistance of 3′-phosphoglycolate DNA ends to digestion by mammalian DNase III. Radiat. Res. 2002; 157 :306–311.11839093
85. Kang D. , KimS.H., HamasakiN. Mitochondrial transcription factor A (TFAM): roles in maintenance of mtDNA and cellular functions. Mitochondrion. 2007; 7 :39–44.17280879
86. Yu C.-H. , DavidsonS., HarapasC.R., HiltonJ.B., MlodzianoskiM.J., LaohamonthonkulP., LouisC., LowR.R.J., MoeckingJ., De NardoD.et al . TDP-43 triggers mitochondrial DNA release via mPTP to activate cGAS/STING in ALS. Cell. 2020; 183 :636–649.33031745
87. Huang L.S. , HongZ., WuW., XiongS., ZhongM., GaoX., RehmanJ., MalikA.B. 2020) mtDNA activates cGAS signaling and suppresses the YAP-mediated endothelial cell proliferation program to promote inflammatory injury. Immunity. 52 :475–486.32164878
88. García N. , GarcíaJ.J., CorreaF., ChávezE. The permeability transition pore as a pathway for the release of mitochondrial DNA. Life Sci. 2005; 76 :2873–2880.15808887
89. Wu C.C. , BrattonS.B. Regulation of the intrinsic apoptosis pathway by reactive oxygen species. Antioxid. Redox. Signal. 2013; 19 :546–558.22978471
90. Briston T. , RobertsM., LewisS., PowneyB., M. StaddonJ., SzabadkaiG., DuchenM.R. Mitochondrial permeability transition pore: sensitivity to opening and mechanistic dependence on substrate availability. Sci. Rep. 2017; 7 :10492.28874733
91. Carroll E.C. , JinL., MoriA., Muñoz-WolfN., OleszyckaE., MoranH.B.T., MansouriS., McEnteeC.P., LambeE., AggerE.M.et al . The vaccine adjuvant Chitosan promotes cellular immunity via DNA sensor cGAS-STING-dependent induction of type I interferons. Immunity. 2016; 44 :597–608.26944200
92. Shimada K. , CrotherT.R., KarlinJ., DagvadorjJ., ChibaN., ChenS., RamanujanV.K., WolfA.J., VergnesL., OjciusD.M.et al . Oxidized mitochondrial DNA activates the NLRP3 inflammasome during apoptosis. Immunity. 2012; 36 :401–414.22342844
93. Patrushev M. , KasymovV., PatrushevaV., UshakovaT., GogvadzeV., GazievA. Mitochondrial permeability transition triggers the release of mtDNA fragments. Cell. Mol. Life Sci. 2004; 61 :3100–3103.15583871
94. Nakahira K. , HaspelJ.A., RathinamV.A., LeeS.J., DolinayT., LamH.C., EnglertJ.A., RabinovitchM., CernadasM., KimH.P.et al . Autophagy proteins regulate innate immune responses by inhibiting the release of mitochondrial DNA mediated by the NALP3 inflammasome. Nat. Immunol. 2011; 12 :222–230.21151103
95. Zhang W. , LiG., LuoR., LeiJ., SongY., WangB., MaL., LiaoZ., KeW., LiuH.et al . Cytosolic escape of mitochondrial DNA triggers cGAS-STING-NLRP3 axis-dependent nucleus pulposus cell pyroptosis. Exp. Mol. Med. 2022; 54 :129–142.35145201
96. Kim B.-R. , KimB.-J., KookY.-H., KimB.-J. Mycobacterium abscessus infection leads to enhanced production of type 1 interferon and NLRP3 inflammasome activation in murine macrophages via mitochondrial oxidative stress. PLoS Pathog. 2020; 16 :e1008294.32210476
97. Kim J. , GuptaR., BlancoL.P., YangS., Shteinfer-KuzmineA., WangK., ZhuJ., YoonH.E., WangX., KerkhofsM.et al . VDAC oligomers form mitochondrial pores to release mtDNA fragments and promote lupus-like disease. Science. 2019; 366 :1531–1536.31857488
98. Xian H. , WatariK., Sanchez-LopezE., OffenbergerJ., OnyuruJ., SampathH., YingW., HoffmanH.M., ShadelG.S., KarinM. Oxidized DNA fragments exit mitochondria via mPTP- and VDAC-dependent channels to activate NLRP3 inflammasome and interferon signaling. Immunity. 2022; 55 :1370–1385.35835107
99. Harding S.M. , BenciJ.L., IriantoJ., DischerD.E., MinnA.J., GreenbergR.A. Mitotic progression following DNA damage enables pattern recognition within micronuclei. Nature. 2017; 548 :466–470.28759889
100. Chen J. , HardingS.M., NatesanR., TianL., BenciJ.L., LiW., MinnA.J., AsanganiI.A., GreenbergR.A. Cell cycle checkpoints cooperate to suppress DNA- and RNA-associated molecular pattern recognition and anti-tumor immune responses. Cell Rep. 2020; 32 :108080.32877684
101. Xue A. , ShangY., JiaoP., ZhangS., ZhuC., HeX., FengG., FanS. Increased activation of cGAS-STING pathway enhances radiosensitivity of non-small cell lung cancer cells. Thoracic Cancer. 2022; 13 :1361–1368.35429143
102. Zhang J. , ShihD.J.H., LinS.-Y. Role of DNA repair defects in predicting immunotherapy response. Biomark. Res. 2020; 8 :23.32612833
103. Bever K.M. , LeD.T. DNA repair defects and implications for immunotherapy. J. Clin. Invest. 2018; 128 :4236–4242.30272580
104. Gaston J. , CheradameL., YvonnetV., DeasO., PouponM.-F., JuddeJ.-G., CairoS., GoffinV. Intracellular STING inactivation sensitizes breast cancer cells to genotoxic agents. Oncotarget. 2016; 7 :77205–77224.27791205
105. Morris R. , KershawN.J., BabonJ.J. The molecular details of cytokine signaling via the JAK/STAT pathway. Protein Sci. 2018; 27 :1984–2009.30267440
106. Saada J. , McAuleyR.J., MarcattiM., TangT.Z., MotamediM., SzczesnyB. Oxidative stress induces Z-DNA-binding protein 1–dependent activation of microglia via mtDNA released from retinal pigment epithelial cells. J. Biol. Chem. 2022; 298 :101523.34953858
107. Baik J.Y. , LiuZ., JiaoD., KwonH.J., YanJ., KadigamuwaC., ChoeM., LakeR., KruhlakM., TandonM.et al . ZBP1 not RIPK1 mediates tumor necroptosis in breast cancer. Nat. Commun. 2021; 12 :2666.33976222
108. Yang Y. , WuM., CaoD., YangC., JinJ., WuL., HongX., LiW., LuL., LiJ.et al . ZBP1-MLKL necroptotic signaling potentiates radiation-induced antitumor immunity via intratumoral STING pathway activation. Sci. Adv. 2021; 7 :eabf6290.34613770
109. Zhang L. , WeiX., WangZ., LiuP., HouY., XuY., SuH., KociM.D., YinH., ZhangC. NF-κb activation enhances STING signaling by altering microtubule-mediated STING trafficking. Cell Rep. 2023; 42 :112185.36857187
110. Sokolowska O. , NowisD. STING signaling in cancer cells: important or not?. Arch. Immunol. Ther. Exp. (Warsz.). 2018; 66 :125–132.28748479
111. Parkes E.E. , WalkerS.M., TaggartL.E., McCabeN., KnightL.A., WilkinsonR., McCloskeyK.D., BuckleyN.E., SavageK.I., Salto-TellezM.et al . Activation of STING-dependent innate immune signaling by S-phase-specific DNA damage in breast cancer. J. Natl. Cancer Inst. 2016; 109 :djw199.27707838
112. Ho S.S. , ZhangW.Y., TanN.Y., KhatooM., SuterM.A., TripathiS., CheungF.S., LimW.K., TanP.H., NgeowJ.et al . The DNA structure-specific endonuclease MUS81 mediates DNA sensor STING-dependent host rejection of prostate cancer cells. Immunity. 2016; 44 :1177–1189.27178469
113. An X. , ZhuY., ZhengT., WangG., ZhangM., LiJ., JiH., LiS., YangS., XuD.et al . An analysis of the expression and association with immune cell infiltration of the cGAS/STING pathway in Pan-cancer. Mol. Ther. Nucleic Acids. 2019; 14 :80–89.30583098
114. Wang Y. , YuX., SongH., FengD., JiangY., WuS., GengJ. The STAT-ROS cycle extends IFN–induced cancer cell apoptosis. Int. J. Oncol. 2018; 52 :305–313.29115415
115. Sun H. , HuangY., MeiS., XuF., LiuX., ZhaoF., YinL., ZhangD., WeiL., WuC.et al . A nuclear export signal is required for cGAS to sense cytosolic DNA. Cell Rep. 2021; 34 :108586.33406424
116. Henner W.D. , RodriguezL.O., HechtS.M., HaseltineW.A. Gamma Ray induced deoxyribonucleic acid strand breaks. 3′ Glycolate termini. J. Biol. Chem. 1983; 258 :711–713.6822504
117. Karimi-Busheri F. , LeeJ., WeinfeldM., TomkinsonA.E. Repair of DNA strand gaps and nicks containing 3′-phosphate and 5′-hydroxyl termini by purified mammalian enzymes. Nucleic Acids Res. 1998; 26 :4395–4400.9742240
118. Clark C.A. , YangE.S. Harnessing DNA repair defects to augment immune-based therapies in triple-negative breast cancer. Front. Oncol. 2021; 11 :703802.34631532
119. Gilmore E. , McCabeN., KennedyR.D., ParkesE.E. DNA repair deficiency in breast cancer: opportunities for immunotherapy. J. Oncol. 2019; 2019 :4325105.31320901
120. Szego E.M. , MalzL., BernhardtN., Rösen-WolffA., FalkenburgerB.H., LukschH. Constitutively active STING causes neuroinflammation and degeneration of dopaminergic neurons in mice. eLife. 2022; 11 :e81943.36314770
121. Fryer A.L. , AbdullahA., TaylorJ.M., CrackP.J. The complexity of the cGAS-STING pathway in CNS pathologies. Front. Neurosci. 2021; 15 :621501.33633536
122. Madsen P.M. , PintoM., PatelS., McCarthyS., GaoH., TaherianM., KarmallyS., PereiraC.V., DvoriantchikovaG., IvanovD.et al . Mitochondrial DNA double-strand breaks in oligodendrocytes cause demyelination, axonal injury, and CNS inflammation. J. Neurosci. 2017; 37 :10185–10199.28931570
123. Hu X. , ZhangH., ZhangQ., YaoX., NiW., ZhouK. Emerging role of STING signalling in CNS injury: inflammation, autophagy, necroptosis, ferroptosis and pyroptosis. J. Neuroinflamm. 2022; 19 :242.
