AKT2 reduces IFNβ1 production to modulate antiviral responses and systemic lupus erythematosus

Abstract Interferon regulatory factor 3 (IRF3)‐induced type I interferon (I‐IFN) production plays key roles in both antiviral and autoimmune responses. IRF3 phosphorylation, dimerization, and nuclear localization are needed for its activation and function, but the precise regulatory mechanisms remain to be explored. Here, we show that the serine/threonine kinase AKT2 interacts with IRF3 and phosphorylates it on Thr207, thereby attenuating IRF3 nuclear translocation in a 14‐3‐3ε‐dependent manner and reducing I‐IFN production. We further find that AKT2 expression is downregulated in viral‐infected macrophages or in monocytes and tissue samples from systemic lupus erythematosus (SLE) patients and mouse models. Akt2‐deficient mice exhibit increased I‐IFN induction and reduced mortality in response to viral infection, but aggravated severity of SLE. Overexpression of AKT2 kinase‐inactive or IRF3‐T207A mutants in zebrafish supports that AKT2 negatively regulates I‐IFN production and antiviral response in a kinase‐dependent manner. This negative role of AKT2 in IRF3‐induced I‐IFN production suggests that AKT2 may be therapeutically targeted to differentially regulate antiviral infection and SLE.

Introduction
I-IFN and IFN-induced transcription of a wide range of IFNstimulated genes (ISGs) are both critical to elicit innate and adaptive immune responses against various infections (Sadler & Williams, 2008;Schneider et al, 2014;Wu & Chen, 2014;Boxx & Cheng, 2016). However, I-IFN is pathogenic during the development of autoimmune disorders, which might be triggered by multiple genetic and environmental factors. Especially, the persistent presence of I-IFN has been proved to accelerate the pathogenesis of SLE (Munz et al, 2009;Bengtsson & Ronnblom, 2017). It is therefore vital to understand the precise mechanism of regulating I-IFN production, which can offer new drug targets to treat infections or SLE.
The induction of I-IFN is elicited by pattern recognition receptors (PRRs) and a cascade of downstream molecules. IRF3 is a master transcription factor for I-IFN production, and IRF3 transcriptional activity and other biological functions are precisely regulated via its phosphorylation (Taniguchi et al, 2001;Saitoh et al, 2006;Chattopadhyay et al, 2016;Huai et al, 2019). Phosphorylated IRF3 undergoes conformational change to form dimers, which then enters the nuclei and associates with interferon transcriptional elements to initiate I-IFN transcription, mainly Ifnb1 (Honda et al, 2006;Sadler & Williams, 2008). Phosphorylation sites of the cluster 1 (Ser385/ Ser386) and the cluster 2 (Ser396, Ser398, Ser402, Thr404, and Ser405) in the C terminus of IRF3 have been demonstrated to indicate IRF3 activation status in antiviral immunity (Lin et al, 1998(Lin et al, , 1999Panne et al, 2007). Further studies suggest that Ser386 is critical, while Ser396 plays a moderate role in IRF3 dimerization and activation (Dalskov et al, 2020;Jing et al, 2020), and PTEN regulates the phosphorylation of IRF3 at Ser97 to prevent IRF3 nuclear import . Despite these findings, it is still unclear whether IRF3 phosphorylation at other sites could affect the intracellular vs. nuclear translocation of IRF3.
The serine/threonine kinase AKT (also named protein kinase B/ PKB) represents a critical kinase family that can phosphorylate different target proteins and control a variety of cellular functions. AKT family contains three highly conserved isoforms in mammals, AKT1/PKBa, AKT2/PKBb, AKT3/PKBc (Fayard et al, 2010). There are growing researches about the distinct role of AKTs in the induction of I-IFN. After herpes simplex virus-1 (HSV-1) infection, AKT1 phosphorylates TBK1 to attenuate STING signaling (Wu et al, 2019); others also suggest that AKT1 phosphorylates cGAS to suppress its enzymatic activity and results in downregulation of IFNb1 production (Seo et al, 2015). In contrast to the negative role of AKT1 on I-IFN production, our group has recently demonstrated that in response to viral infection, AKT3 expression is significantly enhanced in macrophages and promotes IFNb1 induction . It is not fully understood whether and how AKT2 could affect I-IFN production.
In this study, we discovered that Akt2 expression was downregulated in macrophages upon treatment with IFNb1, the Toll-like receptor7/9 (TLR7/9) agonists, and during viral infection or during TMPD (N,N,N 0 , N 0 -tetramethyl-p-phenylenediamine)-induced SLE. Disruption of Akt2 enhanced the production of IFNb1 and protected mice from viral infection. Conversely, Akt2 deficiency led to the much worse SLE for increased IFNb1 production. Mechanismly, AKT2 could directly bind and phosphorylate IRF3 at Thr207, which worked together with 14-3-3e to restrain the nuclear translocation of IRF3. In addition, AKT2 but not AKT2 kinase-dead mutant overexpression aggravated viral infection in zebrafish larvae, while overexpression of the IRF3-T207A promoted IFNb1 production and reduced viral infection which was not reversed by AKT2 in zebrafish larvae. These data have demonstrated that AKT2 phosphorylates IRF3-T207 to reduce IRF3 nuclear localization and IFNb1 induction, providing either protective or pathogenic role in SLE and viral infections.

Akt2 expression is negatively correlated with IFNb1 production
We firstly examined the distribution of AKT members in different organs. In contrast to the dominant expression of Akt3 in brain, Akt1 was widely expressed and Akt2 was mainly expressed in heart, liver, and kidney ( Fig EV1A). The expression levels of Akt1 and Akt2 were downregulated, while Akt3 levels were upregulated in livers from vesicular stomatitis virus (VSV)-infected mice ( Fig EV1B). We also confirmed the decreased Akt2 expression in spleen and lung from VSV-infected mice (Fig EV1C), as well as from hepatitis B virus (HBV)-infected hepatocellular carcinoma (HCC) patients ( Fig EV1D) that is consistent with the reduced AKT2 expression in livers from HBV-infected patients provided by the GEO database (Fig 1A, left panel). Akt2 mRNA levels were also substantially downregulated in brain or lung from Japanese encephalitis virus (JEV)-infected or the influenza virus (H7N9, H1N1/PR8)-infected mice, respectively ( Fig 1B). Moreover, the reduced Akt2 mRNA expression was detected in bronchial epithelial cells upon H1N1 infection (Fig 1A, right panel), and in murine peritoneal elucidated macrophages (PEMs) after treated with lipopoly(I:C) to activate the RIG-1/MAVS pathway or treated with lipopoly(A:T), lipo-ISD and HSV-1 to activate the cGAS/STING pathway ( Fig 1C). Then, we also detected the decreased expression of total AKT2 and the phosphorylated AKT2-Ser474 at protein levels in VSV-infected PEMs ( Fig EV1E).
Notably, we found that the mRNA levels of Akt2 were negatively correlated with Ifnb1 mRNA levels in VSV-infected PEMs, or in Listeria monocytogenes (LM) infected bone marrow-derived macrophages (BMDMs) (Fig 1D). Therefore, to investigate whether the IFNb1/IFNAR signaling was responsible for Akt2 expression, we obtained PEMs from the I-IFN receptor-deficient (Ifnar1 KO) mice which hardly produced Ifnb1 and Cxcl10 in response to VSV infection or lipo-ISD treatment (Fig EV1F), ensuring PEMs from Ifnar1 KO mice were defective in response to IFNb1 stimulation. Interestingly, Akt2 expressions were no longer downregulated in VSVinfected or lipo-ISD-treated PEMs from the Ifnar1 KO mice ( Fig 1E).
The IFNb1/IFNAR signaling activates several key downstream effectors, and which one could participate in the suppression of Akt2 expression? To answer this, WT PEMs were pretreated with various inhibitors targeting IFNb1/IFNAR downstream proteins, including signal transducer and activator of transcriptions (STATs), p38, activator protein-1 (AP-1), nuclear transcription factor kappa-B (NF-jB), and only the inhibitors targeting STAT3 and STAT6 (not STAT5) could prevent the downregulation of Akt2 expression ( Fig 1F). Furthermore, in the cytokine-activated Janus kinase (JAK)/STAT signaling cascade, the suppressors of cytokine signaling (SOCS) family members, especially SOCS1 and SOCS3, are induced by I-IFN stimulation, which then inhibit STAT activity to form a negative feedback loop (Morris et al, 2018). Knockdown of Socs1 or Socs3 expression by siRNAs further reduced Akt2 mRNA levels and enhanced I-IFN production after VSV treatment ( Fig EV1G). Those results indicate that during viral infection, the IFNb1/IFNAR signal decreases Akt2 expression via STAT3/STAT6, and knockdown of SOCS1/SOCS3 could release their inhibition on STAT3/6 and further reduce Akt2 expression.
To elucidate whether AKT2 regulated Ifnb1 production, PEMs were treated with the AKT2 selective inhibitor CCT128930 (Fig 1G) or Akt2 was knocked down by siRNA (Figs 1H and EV1H). After stimulated with lipo-poly(I:C), lipo-poly(A:T), lipo-ISD or infected with VSV and HSV-1, inhibition of AKT2 or knockdown of Akt2 both enhanced the mRNA levels of Ifnb1 in PEMs (Fig 1G and H). These data suggest that Akt2 expression is reduced upon stimuli or viral infection, and Akt2 negatively regulates IFNb1 production. activate the TLR3 pathway or treated with LPS to activate the TLR4 pathway ( Fig 2C). Following I-IFN production in Akt2 KO macrophages, we further asked whether this affected bystander cells. WT or Akt2 KO PEMs were stimulated with lipo-poly(I:C) for 3 h and washed several times to remove any residual lipo-poly(I:C). Fresh medium was added to WT or Akt2 KO PEMs for another 9 h to collect cell medium, which was used to stimulate MEFs or PEMs. The culture medium from Akt2 KO PEMs indeed induced higher Ifnb1, Ifna4 production in MEFs or Cxcl10, Ifit1, Mx1 production in PEMs ( Fig EV2B). This suggests that Akt2 KO macrophages not only enhance I-IFN production but also enhance the ISG expression in bystander cells such as PEMs and MEFs, therefore better preventing the viral infection and propagation. Consistent with Akt2 KO PEMs (Fig 2A-C), Akt2 KO primary-MEFs also enhanced Ifnb1 mRNA level and reduced the amount of GFP-fused VSV infection shown by immunofluorescence microscopy or FACS assays (Figs 2D and EV2C). Moreover, after VSV infection, overexpression of AKT2 in MEFs and HEK293T cells restrained Ifnb1 expression (Fig 2E), and IFNb1-reporter luciferase assay confirmed that AKT2 inhibited IFNb1 transcription in a dose-dependent manner ( Fig EV2D). Because AKT2 kinase activity is pivotal for its biological function in various cell types, we next overexpressed the kinase-dead mutant AKT2-T309A/S474A in HEK293T cells to explore whether AKT2 kinase activity was functioned in suppressing IFNb1 production. When co-expressed with TBK1 or infected with VSV, AKT2 reduced the IFNb1 luciferase readings, while the AKT2-T309A/S474A failed to inhibit this (Figs 2F and EV2E).
To verify the in vivo antiviral function of AKT2 or its kinase-dead mutant AKT2-T309A/S474A, we employed the zebrafish model as previously described (Meng et al, 2016;Guerra-Varela et al, 2018). Zebrafish can be generated in short term to express the interested proteins and is responsive to VSV infection. We used mpeg1-mCherry transgenic zebrafish larvae in which macrophage-lineage cells express mCherry and could be monitored in vivo. AKT2 mRNA was transcripted in vitro and injected into zebrafish embryos for 48 h to ensure AKT2 expression at protein levels ( Fig EV2F), followed by VSV infection. We observed the colocalization of GFPfused VSV (green) with mpeg1-mCherry macrophages (red) in the dorsal muscle, brain, and abdominal vessels of zebrafish larvae (Figs 2G and EV2G, indicated by arrows). Zebrafish larvae overexpressing AKT2 reduced ifn1 mRNA levels, displayed severer tissue damages in eye and trunk muscle, and decreased the survival rates ( Fig 2H). In contrast, zebrafish larvae overexpressing the AKT2-T309A/S474A no longer aggravate viral infection than PBS control ( Fig 2H). Together, we have demonstrated that AKT2 inhibits Ifnb1 expression and this is dependent on AKT2 kinase activity.
AKT2 did not affect IRF3 transcription and translation (Fig EV3D,. To explore whether AKT2 interacted with IRF3 endogenously, cell lysates of WT or Akt2 KO PEMs were prepared for immunoprecipitation using anti-AKT2 antibody, followed by immunoblotting using anti-IRF3 antibody (Fig 3D, left panel). Alternatively, cell lysates were prepared for immunoprecipitation using anti-IRF3 antibody, followed by immunoblotting using anti-AKT2 antibody ( Fig 3D, right panel). Both sets of data confirmed the endogenous association between AKT2 and IRF3. To map their interaction, HA-tagged AKT2 and Flag-tagged IRF3 or their truncations were co-expressed in HEK293T cells as indicated ( Fig EV3E). Immunoprecipitation with anti-HA or anti-Flag antibodies was performed, which showed that the N terminus of AKT2 interacted with the IAD-domain of IRF3 ( Fig EV3E). Next, GST-tagged AKT2 and His-tagged IRF3 were purified from E.coli to verify their direct interaction via the pull-down assay ( Fig 3E). As a master transcription factor, IRF3 is phosphorylated and undergoes dimerization to ◀ Figure 1. Akt2 expression is negatively correlated with IFNb1 production.

AKT2 phosphorylates IRF3 at Thr207 and blocks IRF3 activation
Our above findings indicated that inhibition of IFNb1 expression was dependent on kinase activity of AKT2; therefore, we speculated that AKT2 kinase activity was critical to inhibit IRF3 translocation into the nuclei. Next, Flag-tagged IRF3 was overexpressed in HEK293T-IRF3 KO cells together with HA-tagged AKT2 or the kinase-dead mutant AKT2-T309A/S474A. As expected, the AKT2 kinase-dead mutant failed to decrease the amount of IRF3 in the nuclei ( Fig 4A). Using the Phos-tag gel, we detected the enhanced IRF3 phosphorylation levels in Akt2 KO macrophages upon VSV challenge ( Fig EV4A). To confirm that AKT2 regulated IRF3 phosphorylation in the in vitro kinase assay, HA-tagged AKT2 were enriched from HEK293T cells via immunoprecipitation and incubated with the purified His-tagged IRF3 protein from E. Coli. In the presence of AKT2, IRF3 was indeed phosphorylated that was shown as the shifted bands in the Phos-tag gel (Fig 4B,red arrow). To further identify the potential phosphorylation sites in IRF3, Flagtagged IRF3 was overexpressed into HEK293T cells with AKT2 or the GFP control, next, anti-Flag immunoprecipitation was performed for the mass spectrometry (MS) analysis. Compared to the control, AKT2 overexpression induced IRF3 phosphorylations at Ser14, Ser173, S175, Thr180, Thr207 ( Fig EV4B). We next mutated these residues into alanine (S14A, S173A/S175A, S180A, and T207A) to prevent phosphorylation. Only when the IRF3-T207A mutant was co-expressed, AKT2 no longer reduced the IFNb1 luciferase activity (Figs 4C and EV4C). Moreover, the nuclear location and phosphorylation of IRF3-T207A were not changed by AKT2 (Fig 4D and E), and overexpression of IRF3-T207A did not change IRF3 Ser396phosphorylation and dimerization after VSV stimulation ( Fig EV4D). To verify the important function of IRF3-T207, IRF3-T207D (D, aspartic acid) was generated to mimic the phosphorylation status. Overexpression of IRF3-T207D inhibited IFNb1 production, and AKT2 co-expression showed no further inhibition ( Fig EV4E).
We were intrigued to assess the in vivo function of IRF3-T207A mutant, and then, zebrafish embryos were overexpressed with IRF3 or IRF3-T207A followed by VSV infection (Fig EV4F). Compared to the controls, zebrafish larvae expressing IRF3 enhanced the survival rates, and expressing IRF3-T207A displayed higher survival rates ◀ Figure 2. AKT2 kinase activity is indispensable to attenuate I-IFN production in macrophages and in zebrafish larvae. and the reduced amount of VSV copies (Fig 4G and EV4G). Importantly, AKT2 overexpression could aggravate VSV copies and reduce the survival rates in zebrafish larvae expressing IRF3, but not expressing the IRF3-T207A mutant (Figs 4G and H,and EV4G). The kinase-dead mutant AKT2-T309A/S474A was included as the control, which showed less VSV copies in zebrafish larvae when compared to those expressing the WT AKT2 ( Fig 4G). Collectively, these results have demonstrated that AKT2 depends on its kinase activity to phosphorylate IRF3 at Thr207 for preventing I-IFN production and antiviral effects.

Targeting AKT2 enhances antiviral defense in mice
Akt2 KO mice showed normal percentages of macrophages (CD11b + F4/80 + ) in bone marrow (BM) and spleen, or normal macrophage development ex vivo derived from BM measured by FACS or quantitative reverse transcription-PCR (qRT-PCR) for the mRNA levels of adhesion G protein-coupled receptor E1 (Adgre1, also known as F4/ 80) and Mer tyrosine kinase (MerTK) (Fig 5A and Appendix Fig  S1A). We have shown that Akt2 KO PEMs enhanced IRF3 activity, and previous work suggests that the host could induce RIPA (RLRinduced IRF3-mediated pathway of apoptosis) to eliminate viral infection and pathogenesis (Chattopadhyay et al, 2016;Chattopadhyay & Sen, 2017). Therefore, we examined and found that in the absence or presence of viral infection, Akt2 deficiency in PEMs did not obviously affect cell apoptosis (Appendix Fig S1B). This excludes the possibility that AKT2 might influence macrophage apoptosis to contribute to its antiviral function. Akt2 KO mice also showed normal percentages of CD4 + T cells, CD8 + T cells, or B220 + B cells in thymus, spleen, or BM (Appendix Fig S1C). We next used VSV to infect WT and Akt2 KO mice and evaluate the in vivo function of Akt2 on antiviral defense. After VSV challenge, Akt2 KO mice enhanced the survival rates ( Fig 5B) and elevated Ifnb1 mRNA expression in livers, spleens, or lungs ( Fig 5C) as well as serum IFNb1 concentrations (Fig 5D). Moreover, the amount of VSV in the liver of Akt2 KO mice was reduced than that of WT mice (Fig 5E).
To verify the role of Akt2-deficient immune cells, we adoptively transferred WT and Akt2 KO BMs respectively to reconstitute the irradiated CD45.1 recipient mice, followed by VSV challenge (Fig EV5A). The reconstituted CD45.1 Akt2 KO mice showed higher survival rates with the increased concentrations of serum IFNb1 (Fig 5F), which also reduced the copy numbers or titers of VSV from livers, spleens, or lungs (Fig 5G and H). To further explore the in vivo function of Akt2-deficient macrophages, we followed a previous study to effectively delete macrophages in vivo by clodronate liposome treatment (Jordan et al, 2003). Then, these mice were intravenous injected with WT and Akt2 KO BMDMs followed by VSV challenge (as indicated in Fig EV5B). The mice reconstituted with Akt2 KO-macrophages also enhanced serum IFNb1 concentrations ( Fig EV5C).
What's more, to ascertain if AKT2 might be a potential target to enhance antiviral effect in the host, the AKT2 selective inhibitor CCT128930 was used to treat mice, which indeed enhanced the survival rates in response to VSV infection ( Fig 5I). These data together demonstrated that deletion or blocking AKT2 could be an effective strategy to enhance I-IFN production and facilitate the host against viral infection.
Akt2 is associated with the pathology of SLE Given that heightened I-IFN is a prominently hallmark in SLE patients and correlated with disease severity (Bengtsson & Ronnblom, 2017), we next assessed how Akt2 functioned in TMPDinduced murine SLE model. The expression of Akt2 in different organs or tissue from TMPD-induced mice was analyzed at 2 weeks (the early stage) or 12 weeks (the late stage). Akt2 expression was reduced in peritoneal infiltrating cells (PICs) at both the early and late stages, as well as in liver and lung at the late stage (Fig EV5D  and E). Consistently, the expression of Akt2 was decreased in the PEMs after treated with R848 or CpG to activate the TLR7 and TLR9 ◀ Figure 3. AKT2 restrains IRF3 nuclear translocation via 14-3-3e.
A, B IFNb1 luciferase assays (upper panels) in HEK293T (A) and HEK293T-IRF3 KO (B) cells transfected with the indicated plasmids or infected with VSV. Immunoblot analysis showed the indicated constructs in HEK293T cells (bottom panels). n = 3, respectively. C The mRNA levels of Ifnb1 were measured by qRT-PCR in WT (n = 3) and Akt2 KO (n = 3) PEMs with Irf3 knockdown for 48 h and VSV treatment for 6 h. D WT and Akt2 KO PEMs were infected with or without VSV for 6 h. Then, cell lysates of WT or Akt2 KO PEMs were prepared for immunoprecipitation using anti-AKT2 antibody, followed by immunoblotting using anti-IRF3 antibody (left panel). Alternatively, cell lysates were prepared for immunoprecipitation using anti-IRF3 antibody, followed by immunoblotting using anti-AKT2 antibody (right panel) to detect the endogenous interaction between AKT2 and IRF3. E Immunoblot analysis of GST-AKT2 and His-IRF3 interaction in a GST pull-down assay. F Immunoassay of IRF3 in dimer or monomer form by native-gel and p-IRF3 (Ser396), p-TBK1 (Ser172), AKT2 and ACTIN by SDS-gel in WT and Akt2 KO PEMs with VSV stimulation for 6 h. G Immunofluorescent microscopic imaging (left panel) and statistics analysis (right panel) for IRF3 nuclear translocation in WT and Akt2 KO PEMs at 6 h post-VSV infection. IRF3 (red), Nuclei (Hoechst, green). The white arrows indicate the nuclei with IRF3 translocation. Mock, n = 2; VSV, n = 3. Bar, 20 lm. H Immunoassay of IRF3, AKT2, LaminB1, and Tubulin in the nuclear and cytoplasmic fractions from the WT and Akt2 KO PEMs after VSV treatment for 6 h. Tubulin and LaminB1 were used as cytoplasmic and nucleic protein loading control, respectively. I Immunoassay of IRF3, SP1, and Tubulin in the nuclear and cytoplasmic fractions of the HEK293T cells overexpressed with indicated constructs for 24 h and stimulated with VSV for 6 h. Tubulin and SP1 served as cytoplasmic and nucleic protein loading control, respectively. J IFNb1 luciferases assays were performed in HEK293T cells transfected with AKT2 and 14-3-3e followed by VSV treatment. n = 3, respectively. K qRT-PCR analysis for the Ifnb1 mRNA levels in VSV-stimulated WT and Akt2 KO PEMs with 14-3-3e siRNA pretreatment. n = 3, respectively.
Data information: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 and ns, not significant (P > 0.05); using a one-way ANOVA pathways (Fig EV5F), and both pathways were previously demonstrated to aggravate SLE (Celhar et al, 2012). According to others' studies, the diffuse pulmonary hemorrhage (DPH) can occur in some SLE patients with the increased mortality (Aguilera-Pickens & Abud-Mendoza, 2018). We found that TMPDtreated Akt2 KO mice developed more serious DPH and increased leukocyte recruitment than that in WT mice at 2 weeks ( Fig 6A, left  panel). In parallel, Akt2 KO mice exhibited splenomegaly and disrupted splenic architecture (Fig 6A, right panel) with the elevated percentages of CD11b + cells (Fig EV5G). Not only the expression of Ifnb1 and Ifna4 increased in PIC and peripheral blood mononuclear cells (PBMCs) (Fig 6B) but also the amount of nuclear IRF3 enhanced in PICs (Fig 6C) from TMPD-induced Akt2 KO mice.
Next, CD45.1 mice were also adoptively transferred with CD45.2 WT or Akt2 KO BMs and treated with TMPD for 12 weeks to induce SLE ( Fig EV5H). CD45.1 Akt2 KO mice showed severer DPH and splenomegaly ( Fig EV5I). TMPD injection could induce strong immune responses and the formation of lipogranuloma which is adhering to the mesothelial surface of mice (Reeves et al, 2009). Additionally, TMPD-induced SLE is accompanied with glomerulonephritis with the elevated anti-dsDNA IgG level in serum. In CD45.1 Akt2 KO mice, we observed the exacerbated lipogranulomas (Fig 6D), glomerulonephritis, and the enhanced anti-IgG staining in kidney after TMPD treatment for 12 weeks (Fig 6E). In agreement with this, CD45.1 Akt2 KO mice developed higher amount of anti-dsDNA IgG in serum (Fig 6F), increased Ifnb1 and Ifna4 expression in PICs (Fig 6G), and enhanced percentages of B220 + and CD19 + MHCII + cells in PBMCs (Fig 6H).
We then confirmed the reduced expression of AKT2 in PBMCs isolated from SLE patients, when compared that from healthy donors (Fig 6I). Also, the expression of AKT2 was negatively correlated with I-IFN in CD14 + monocytes from SLE patients (Fig 6J). Interestingly, in resting PBMCs from SLE patients, IRF3 was located in the cytoplasm and AKT2 was expressed at a relative higher level, which was contrast to the decreased AKT2 expression and the increased amount of IRF3 in the nucleus once PBMCs were at activated state ( Fig 6K). Together, we demonstrated that AKT2 plays a protective role in the pathology of SLE.

Discussion
Phosphorylation, dimerization, and nuclear accumulation are three hallmarks of IRF3 activation. A great number of studies have elucidated the mechanism of IRF3 phosphorylation, but less about how to regulate IRF3 translocation from cytoplasm to nuclei. Our study has provided a new perspective that cooperating together with 14-3-3e, AKT2 binds and phosphorylates IRF3 at Thr207 to restrain IRF3 nuclear translocation and blunt IFNb1 production. Once macrophage responds to viral infection, IFNb1 is induced to downregulate Akt2 expression, which leads to IRF3 release from the cytoplasm for its nuclear entry and the induction of I-IFN (Appendix Fig S3). Upon viral infection, TBK1 phosphorylates MAVS, TRIF, and STING to recruit IRF3, which then allows TBK1 to phosphorylate IRF3. Once phosphorylated, IRF3 is licensed to dissociate from them, dimerizes and entries into the nuclei. Since IRF3-Thr207 has no impact on IRF3 dimerization, we could speculate that like WT IRF3, IRF3-Thr207 is recruited to and then dissociated from the other adaptor proteins (STING/TRIF/TBK1). In addition, the luciferase reporter assays in Fig 3A and B have shown that IRF3-5D-induced IFNb1 production was not affected by AKT2 overexpression. This suggests that IRF3-5D might form a dimer and is not accessible to be phosphorylated by AKT2 at Thr207. Previous work has suggested that PTEN antagonizes the PI(3)K/AKT pathway (Maehama & Dixon, 1998;Stambolic et al, 1998), and PTEN can positively facilitate IRF3 nuclear localization through de-phosphorylating IRF3 at Ser97 . Those clues imply a possibility about whether PTEN could inactivate AKT2, which results in the reduced phosphorylation levels of IRF3 at Thr207 and allows IRF3 activation and I-IFN production.
Besides phosphorylation, other types of post-translational modifications (PTMs) are also critical for IRF3 activation, including ◀ Figure 4. AKT2 phosphorylates IRF3 at Thr207 and blocks IRF3 activation.
A Immunoblot analysis of IRF3 in the cytoplasmic and nucleic fractions of HEK293T-IRF3 KO cells with overexpression of IRF3 and AKT2 or AKT2-S309A/T474A. B Immunoblot analysis of in vitro kinase assay by the phosphorylation gel. HA-AKT2 was immunoprecipitated from HEK293T cells by anti-HA/IgG antibody and Protein-G beads, and His-IRF3 was purified from E.coli cells by anti-His antibody-conjugated agarose beads. His-IRF3 alone and anti-IgG-beads added with His-IRF3 served as controls. The red arrow indicated the phosphorylated IRF3. C IFNb1 luciferase assays and immunoblot analysis showed the HEK293T-IRF3 KO cells with overexpression of the indicated plasmids for 24 h and VSV infection for 6 h. n = 3, respectively. D Immunofluorescence microscopy (top panel) of AKT2-, Flag-IRF3-, or Flag-IRF3-T207A-overexpressed MEF cells after poly(A:T) stimulation for 6 h. Flag (IRF3, red) and Hoechst (nuclei, blue). Immunoassay of nuclear-cytoplasm extractions (bottom panel) from HEK293T-IRF3 KO cells with overexpression of the indicated plasmids for 24 h followed by VSV infection for 6 h. GAPDH and SP1 were used as cytoplasmic and nucleic protein loading control, respectively. Bar, 20 lm. E HEK293T-IRF3 KO cells were overexpressed with indicated plasmids, and the cell lysates were conducted to immunoblot analysis by phosphorylation gel for the assay of phosphorylated IRF3 (the upper band) by anti-Flag antibody. F Survival rates (left panel) until 72 h and H&E staining (right panel) at 18 h of the zebrafish larvae with indicated protein expression and VSV challenge. The arrows indicated the VSV-infected eye and skeletal muscle in zebrafish larvae. Bars, 100 lm. G qRT-PCR analysis of the VSV copies in overexpressed zebrafish embryos with the VSV challenge at 18 h later. Every dot represents three zebrafish embryos. Horizontal vertical square bracket shows the statistical analysis of comparison with "PBS mock", the rest shows the comparison with "PBS VSV". PBS mock, n = 15; PBS VSV, n = 9; IRF3 VSV, n = 16; IRF3-T207A VSV, n = 11; AKT2 VSV, n = 14; AKT2-T309A/S474A VSV, n = 9; IRF3 + AKT2 VSV, n = 9; IRF3 + AKT2-T309A/S474A VSV, n = 13; IRF3-T207A + AKT2 VSV, n = 10. H The survival rates of over-expressed zebrafish larvae as long as 72 h after the VSV challenge.
Data information: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001 and ns, not significant (P > 0.05); using a one-way ANOVA test (G), or two-way ANOVA test (C), or log-rank (Mantel-Cox) test (F left panel and H). Data are from at least three independent experiments (C), or representative of three independent experiments (A, B, D-H). Error bars (C, mean AE SEM; G, mean AE SD). Source data are available online for this figure.
10 of 18 The EMBO Journal 41: e108016 | 2022 ª 2022 The Authors ubiquitination, methylation, acetylation, SUMOylation, and ADPribosylation. Further studies suggest that one type of PTMs might be related to other types of PTMs to affect IRF3 activation. For example, the prolyl isomerase Pin1 phosphorylates IRF3 at Ser339/ Pro340, leading to IRF3 polyubiquitination and then degradation (Saitoh et al, 2006); lysine methyltransferase nuclear receptorbinding SET domain 3 (NSD3) is associated with IRF3 directly and methylates IRF3 at K366, which can maintain IRF3 phosphorylation to promote I-IFN production (Wang et al, 2017). These studies suggest that different signal pathways are participated and The EMBO Journal 41: e108016 | 2022 cooperated to precisely regulate IRF3 activation. It is interesting to further reveal whether AKT2-mediated IRF3-Thr207 phosphorylation could affect or to be affected by other types of PTMs. Antiviral responses are complicated processes including the early and late stages. We confirmed that Akt2 deficiency could enhance Ifnb1 and ISGs production at the early stage (Fig 2A and B), which could also affect bystander cells for the induction of ISGs ( Fig  EV2B). In addition, vRNA extracted from VSV was used to treat Akt2 KO macrophages to mimic 5'pppRNA stimulation (Goulet et al, 2013), which differentially regulated the expression of several genes at the early or late stages. For example, Akt2 KO PEMs enhanced the mRNA levels of Ifnb1 and Ifna4 at the early stage, but this enhancement was diminished at the late stage; in contrast, the mRNA levels of Ccl2 and Ccl5 were enhanced in Akt2 KO PEMs at the late stage (Appendix Fig S2A). These results suggest that AKT2 might affect the expression of multiple genes directly or indirectly during different phases of antiviral responses. For instance, IRF7 is another IRF family member which could also induce Ifnb1 transcription (Levy et al, 2002;Ning et al, 2011). We found that Ifna was enhanced in livers and lungs of Akt2 KO mice (Appendix Fig S2B), and Ifna transcription is driven mainly by IRF7. IRF7 is also downstream of the IFNb1/IFNAR pathway; therefore, Ifna enhancement might be indirectly induced by the increased IFNb1 in Akt2 KO macrophages. Future studies are needed to explore additional functions of AKT2, including its role in regulating IRF7 activation. In addition, AKT2 activation could induce glucose transporter 1 expression, glucose uptake, and lactate production in neoplastic cells (Moro et al, 2009). Interestingly, lactate was reported to inhibit RLR-mediated IFN production . It is therefore worthy to test whether AKT2 might also affect lactate production to inhibit IFN production.
Meanwhile, the significance of AKTs has been studied extensively since its first discovery 30 years ago (Coffer & Woodgett, 1991). AKTs play critical roles in gene transcription, protein synthesis, proliferation, migration, autophagy, anti-apoptosis, and the maintenance of metabolic homeostasis (Dummler & Hemmings, 2007;Xue et al, 2015). AKTs also regulate immune cell development and immune responses (Zhang et al, 2013;Xue et al, 2015). We have noticed that in viral-infected macrophages, Akt3 expression is upregulated as an ISG (Hubel et al, 2019; Xiao et al, 2020), while Akt2 transcription is suppressed, which are both dependent on the IFNb1/IFNAR pathway. STAT is the key transcription factor to induce ISGs, and STAT3 has been identified to bind to Akt2 promoter in website forecast (QIAGEN) or through ChIP experiment in HCC cells (Xie et al, 2018). In this study, we demonstrated that inhibition of STAT3/STAT6 could prevent the reduced Akt2 transcription in Fig 1F; knockdown of Socs1/Socs3, which are key molecules in the negative feedback loop of the IFNb1/IFNAR-JAK/STAT pathway, could release STAT3/STAT6 to further decrease Akt2 expression in Fig EV1G. Importantly, our study elucidated that the downregulated expression of Akt2 in macrophages might be used as a biomarker in I-IFN-related diseases including SLE. Apart from infections, I-IFN is positively related to the development of SLE, and the JAK/STAT pathway participates in the pathogenesis of SLE (Goropevsek et al, 2019). Therefore, targeting JAK/STAT is a promising strategy to protect SLE patients. Our study provides different perspectives about the underline mechanism: targeting JAK/ STAT could not only block I-IFN signaling, but also maintaining Akt2 expression in macrophages that could reduce IRF3 nuclei entry and inhibit I-IFN production. On the other hand, AKT2 is a critical negative regulator for I-IFN production, we propose a new application of the selective AKT2 inhibitor CCT128930 in antiviral therapy. Together, considering AKT2 expression and function, AKT2 could be used as a biomarker for diagnosis or a drug target in SLE or viral infection.
AKTs also play critical roles in the induction and progression of various tumors, and CCT128930 was previously investigated as an antitumor drug in preclinical studies (Yap et al, 2011). Tumorassociated macrophages (TAMs) are essential to promote tumor development, but I-IFN was previously reported to enhance tumor cell apoptosis (Parker et al, 2016). Interestingly, we found that when macrophages were treated with the genomic DNA from the tumor cell line MC38 and B16, Akt2 was downregulated and negatively related to Ifnb1 expression (Appendix Fig S2C). Moreover, Akt2 was downregulated when macrophages are treated with IL-6 and TNF-a (Appendix Fig S2D), which are critical proinflammatory cytokines in the tumor microenvironments (TME). Could Akt2 deficiency in TAMs enhance I-IFN production, which is able to shape TAM function to inhibit tumor development? In addition, several studies have indicated that AKT2, but not AKT1, could promote macrophage migration and phagocytosis (Shiratsuchi & Basson, 2007;Zhang et al, 2009), and TAM-mediated phagocytosis ◀ Figure 5. Targeting AKT2 enhances antiviral defense in mice.
Data information: *P < 0.05, **P < 0.01; using a paired t-test (C and F right panel), or unpaired t-test ( 12 of 18 The EMBO Journal 41: e108016 | 2022 ª 2022 The Authors facilitate tumor growth (Kitamura et al, 2015;Roy et al, 2018). Therefore, it is exciting to explore whether targeting AKT2 represents "one stone kills two birds" strategy, which not only reduces tumor cell survival but also reverses the function of TAMs to better block tumor development. This potential application should be investigated using the in vivo tumor models in the future. . The related mutations in AKT2 or in IRF3 were constructed by standard PCR cloning strategy. To purify proteins from E. coli, AKT2 was inserted into the pGEX4T-1-GST plasmid (GE Healthcare) with GST-tag in the N terminus and IRF3 was inserted into the pET-28a-His plasmid with 6xHis tag in the N terminus. Human 14-3-3f and 14-3-3e cDNAs were amplified from reverse-transcribed cDNA of HEK293T cells. vRNA was prepared from VSV in TRIzol (Takara).

Macrophage preparation and treatment
BMDMs were generated from BM cells followed by culturing in complete 1640 supplemented with 10% (vol/col) FBS, penicillin and streptomycin (100 U/ml), and M-CSF (20 ng/ml) for 5-7 days. PEMs were harvested from mice injected peritoneally with 2 ml 3% Brewer thioglycolate medium for 4 days and cultured with complete DMEM supplemented with 10% (vol/col) FBS and penicillin and streptomycin (100 U/ml). Primary MEFs were trypsin digested from mouse embryo at day13.5 without the fetal liver, expanded and cultured with complete DMEM. To active different pathways in macrophages, poly(I:C), poly(A:T), ISD, or vRNA were transfected via Lipofectamine 2000 (Invitrogen); or poly(I:C), LPS, VSV, or ◀ Figure 6. Akt2 is associated with the pathology of SLE.
A Representative images of lungs and spleens from WT and Akt2 KO mice treated with TMPD (500 ll per mouse) for 2 weeks, and the corresponding H&E staining of the tissues. Bar, 200 lm. B, C qRT-PCR analysis of the relative Ifnb1 and Ifna4 mRNA expression levels in PICs (n ≥ 5) and PBMCs (n ≥ 4) (B), and immunoblot analysis of IRF3, LaminB1, and GAPDH in the PIC nuclear and cytoplasmic fractions (C) from WT and Akt2 KO mice treated with TMPD for 2 weeks. GAPDH and LaminB1 were used as cytoplasmic and nucleic protein loading control, respectively. D, E Representative images of lipogranulomas in peritoneal cavity (D), H&E staining (E left panel) and immunofluorescence of IgG deposits (E right panel) in the kidney sections were collected from WT and Akt2 KO bone marrow-reconstituted CD45.1 mice 12 weeks after TMPD treatment. Bar, 50 lm. F-H Quantitative analysis of serum anti-dsDNA IgG antibody by ELISA (F, n ≥ 4), mRNA expression levels of Ifnb1 and Ifna4 in PIC by qRT-PCR (G, n ≥ 4) and FACS analysis of the percentages of B220 + and CD19 + MHCII + cells in PBMC (H, n ≥ 4) from WT and Akt2 KO bone marrow-reconstituted CD45.1 mice 12 weeks after TMPD treatment. I qRT-PCR analysis of the relative mRNA expression levels of AKT2 in PBMC isolated from SLE patients (n = 9) or healthy donors (n = 9). J qRT-PCR analysis of the relative mRNA expression levels of AKT2 (left panel) and IFNA (middle panel) or IFNb1 (right panel) in CD14-positive monocytes isolated from SLE patients (n = 9) or healthy donors (n = 9). K Immunofluorescence of IRF3 (Red), AKT2 (Green), DAPI (Blue, nucleus) in CD14 + cells from SLE patients followed by analysis of the cells about mean amount of AKT2 in this cell as well as mean amount of IRF3 in the same nucleus with ImageJ software. Cells (n = 39) were collected from 3 patients. Bar, 2 lm.

Animal experiment
8-week-old mice were infected with VSV intravenously (i.v.) at a dose of 10 7 pfu per gram of animal weight to monitor the survival rate. The inhibitor CCT128930 at a dose of 20 mg/kg or the DMSO control was injected intraperitoneally (i.p) twice every other day before VSV infection. Mice were infected with GFP-fused VSV at a dose of 10 6 pfu/g of animal weight and sacrificed at 3 h, 6 h, or 18 h to collect serum and organs. Serum was used to detect IFNb1 concentrations by enzyme-linked immunosorbent assay (ELISA, PBL, #42400), and organs were collected for hematoxylin and eosin (H&E) staining or for RNA extraction followed by qRT-PCR assay. For adoptive transfer, BM cells were isolated from mice without erythrolysis and i.v. injected into the lethally irradiated CD45.1 recipient mice (2 million per mice). Alternatively, clodronate liposome (200 ll/20 g) was i.v. injected twice every other day to delete endogenous macrophages, followed by i.v. transferring of BMDMs (3 million per mice), and mice were then infected with VSV. SLE was induced by i.p. injection with 500 ll of TMPD (Sigma, P2870) per mouse (female, 6-8 weeks old), and PBS was used as the control. 2 or 12 weeks after injection, mice were sacrificed for FACS assay or for the detection of serum anti-dsDNA IgG levels (kexinbiotech) by ELISA and others.

Overexpression and VSV challenge of zebrafish larvae
Ectopic overexpression and VSV challenge of zebrafish larvae were performed as previously described (Meng et al, 2016). Briefly, wildtype or mpeg:mCherry embryos were injected with the 50 pg (< 2 nl/embryo) of the in vitro transcribed mRNA (Thermo Fisher Scientific) in the first 20 min after fertilization. After 48 h, VSV was micro-injected (< 4 nl/larva) into the embryo yolk. After 6 or 18 h, zebrafish larvae were harvested to extract mRNA or for the microscopy assay.

FACS assay and microscopy assay
FACS assay and immunofluorescence assay were performed as described (Liu et al, 2020). Single cell suspension from spleen, BM, thymus, PEMs, and blood were prepared for immunostaining with anti-IRF3, anti-Flag, anti-AKT2, and other appropriate antibodies. Annexin V/7AAD staining or DAPI/Hoechst staining of PEMs was conducted according to the instruction. Data and images were collected with BD Accuri C6 and Fortessa, or Olympus BX51, Zeiss LSM880 Leica SP8 WLL, then analyzed with the respective software.

IFNb1 Luciferase reporter assay
The plasmids expressing the indicated proteins (0.25 lg), the IFNb1 luciferase reporter (0.25 lg), and the internal control TK-renilla (0.01 lg) were transfected with 1.53 ll PEI into HEK293T cells, which were 80-90% confluent in a 24-well plate. After 6-8 h, fresh culture medium was changed to culture the cells for at least another 18 h. If necessary, HEK293T cells were infected with or without VSV for 6 h. Then, cells were harvested to measure the luciferase readings with the guidance of a dual-luciferase assay kit (Promega, #0000456500). Briefly, culture supernatant was discarded, and cells were washed with PBS. Passive Lysis Buffer (diluted with water to 1×, 100 ll/well) was added to lyse cells, and supernatants were collected. Next, 20 ll lytic supernatants of each well were mixed with 20 ll Luciferase Assay Buffer II and measured as Results 1. Then, 20 ll Stop & Glo â Buffer was added for Results 2. Finally, we divided Result 1 by Result 2 to get the relative IFNb1 Luciferase activity, and calculated the fold changes of IFNb1 Luciferase activity by compared to the control groups.
Each experiment was performed in triple wells using the same plasmids synchronously for technical replicate; and three independent experiments were repeated. The data were shown from three independent experiments or from a representative experiment as indicated.

Immunoprecipitation, immunoblot, native PAGE
Immunoprecipitation and Mass-spec assay were performed as described previously (Xue et al, 2016;Xiao et al, 2020). In native PAGE, cells were collected in PBS without residuary FBS and lysed with cold lysis buffer. 8% acrylamide gel without SDS was pre-run at 40 mA for 30 min on ice, with 25 mM Tris-HCl (pH 8.4) in a cathode added 0.5% deoxycholate, and 192 mM glycine in anode chamber. Before immunoblotting, the cell lysates in the native loading buffer (Beyotime) were loaded on the gel and electrophoresed for 60 min at 200V on ice.
GST pull-down and the in vitro kinase assay pGEX-4T-1-GST-hAKT2 and pET-28a-6*His-hIRF3 were transformed into Escherichia coli BL21 (DE3) (TranGen Biotech). Protein expression was induced via 0.25 mM IPTG treatment for 24 h at 16°C. Recombinant proteins were harvested as described previously . The purified proteins at the same molar quantity including 6*His-hIRF3 and GST-hAKT2 or GST were incubated for 2 h followed by incubation with GST-beads for another 1 h at 4°C in 400 ll pull-down buffer (20 mM Tris-HCl, 100 mM NaCl, 5 mM MgCl 2 , 1 mM EDTA, 1 mM DTT, 0.5% (v/v) NP-40 and 10 mg/ml BSA, pH 7.5), and washed four times with the same buffer. Samples ª 2022 The Authors The EMBO Journal 41: e108016 | 2022 were treated with the addition of SDS loading buffer and the supernatants were subjected to 10% SDS-PAGE.
To ensure AKT2 kinase activity, HA-AKT2 was immunoprecipitated from HEK293T cells using anti-HA antibody, and anti-IgG antibody was used as a control. Samples were then incubated with 6*His-hIRF3 at 30°C for 30 min with gentle shaking in the kinase reaction buffer (Cell Signaling Technology). This kinase reaction was stopped by adding SDS loading buffer, and the supernatants were subjected to a phosphate gel for immunoblotting to measure the in vitro phosphorylation reaction. Phosphate gel was 8% acrylamide gel with the addition of 5 mM Phos-tag TM (#034-93521(AAL-107)) and 10 mM Mn 2+ in the resolving gel. Before transferring, the gel was washed 3 times every 10 min in the transfer buffer containing EDTA (10 mM), which is required to increase the transfer efficiency. Phos-tag TM and Mn 2+ cooperates to bind a phosphorylated protein; when the phosphorylation levels are increased, the migration velocity of this protein is slower. Therefore, nonphosphorylated and phosphorylated proteins could be separated in the gel.

RNA isolation and qRT-PCR
Total RNA was extracted from cells or grated tissue in TRIzol (Takara), reverse transcribed with random hexamers (Sangon) and M-MLV transcriptase (Takara). qRT-PCR was performed to detect the mRNA levels of the indicated genes on a CFX-96 real-time PCR system (Bio-Rad) or LightCycler-480-384 (Roche) with SYBR Green Master Mix (Yeasen). qRT-PCR primer sequences are listed in Table EV2. b-Actin in human and mouse samples, or GAPDH in zebrafish samples were used to normalize gene expression. The "relative" mRNA levels were calculated using the 2 ÀDCt method with the normalized gene. The "fold" changes were calculated using the 2 ÀDDCt method with the normalized gene and compared to the control groups including PBS, mock, WT mock, DMSO mock, siNC, Vector, PBS mock, WT siNC mock.

Statistics
Adequate power was ensured when choosing the sample sizes. Data are represented as the means of at least three experiments or representative of three independent experiments. Statistical analyses are performed using GraphPad Prism software, version 8. Statistical analysis was performed using unpaired or paired t-test or correlation or one-way ANOVA or two-way ANOVA test. The log-rank (Mantel-Cox) test is used for survival comparisons. ns, not significant (P > 0.05); *P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001.

Data availability
This study includes no data deposited in external repositories.
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Improving Anticancer Activity of Chrysin using Tumor Microenvironment pH-Responsive and Self-Assembled Nanoparticles

Chrysin is a natural bioactive compound with potential biological activities. However, unfavorable physicochemical properties of native chrysin make it difficult to achieve good therapeutic efficacies. In this study, poly(ethylene) glycol (PEG4000)-conjugated chrysin nanoparticles were prepared. The PEG4000 was conjugated to chrysin through cis-aconityl and succinoyl linkers to achieve tumor microenvironment-specific drug release from PEGylated nanoparticles. The conjugation of PEG and chrysin via succinoyl (PCNP-1) and cis-aconityl (PCNP-2) linkers was confirmed by the 1H NMR and FTIR analysis. The nanoparticles were characterized by DLS, TEM, XRD, and DSC analysis. Comparatively, PCNP-2 showed a better drug release profile and higher anticancer activity against human breast cancer cells than chrysin or PCNP-1. The apoptosis studies and colony formation inhibition assay revealed that the PCNP-2 induced more apoptosis and more greatly controlled the growth of human breast cancer cells than pure chrysin. Thus, the use of PCNPs may help to overcome the issues of chrysin and could be a better therapeutic approach.

INTRODUCTION
In the 21st century, phytoconstituents have emerged as the most prominent source of many anticancer agents and represent an alternative to synthetic chemotherapeutic agents for cancer treatment. 1 A wide range of phytoconstituents, particularly bioflavonoids, have shown potential anticancer activities against various cancers. Chrysin (5,7-dihydroxy-2phenyl-4H-chromen-4-one) is one such natural, plant-based bioflavonoid with broad-spectrum anticancer activity. 2 This molecule is extracted from Passiflora caerulae, Passiflora incarnata, honey, propolis, and honeycombs. 3 The preclinical studies of chrysin have shown its potential anticancer effects through selective cell signaling mechanisms, which are connected to cell proliferation, inflammation, angiogenesis, and cancer metathesis. 4−6 Chrysin was found to be effective against various cancers including breast, lung, bladder, thyroid, prostate, colon, etc. 2,5,7−9 Despite the potential efficacy of chrysin in cancer treatment, some major challenges hinder the clinical use of chrysin. These challenges are low aqueous solubility (3 μg/mL), poor cellular uptake, physiochemical instability, rapid metabolism, and low oral bioavailability. 10,11 Therefore, in the past few years, many attempts have been made to improve the physicochemical properties and delivery of chrysin. Researchers have used micronization, 12 nanoemulsion, nanomicelles, nanosuspension, host−guest complexes, different derivatization, and salt formation approaches. 13−17 However, these systems still have a few drawbacks and cannot deliver the drug site-specifically. Therefore, inspired by the specific tumor microenvironment, we have developed self-assembled nanoparticles of chrysin by conjugating it with PEG 4000 via acid-sensitive linkers. Further, it is reported that the drug−polymer conjugated system shows high drug loading and enhanced therapeutic efficacy in comparison to the physical encapsulation of the drug-loaded system. 18 In a study, chrysin-loaded PLGA−PEG nanoparticles were also prepared and used against T47D (IC 50 = 31.28 μM) and MCF7 (IC 50 = 52.54 μM) breast cancer cells 19 and gastric cancer AGS cell (IC 50 = 36.8 μM) lines. 20 In another interesting study, PEG 1K/2K -chrysin conjugate was developed for the delivery of doxorubicin where the directly conjugated chrysin did not show any sign of toxicity against the HepG2 cells. 21 PEG is an FDA-approved, hydrophilic polymer and is generally used as an excipient in different pharmaceutical formulations. 22−25 High biocompatibility and hydrophilic nature make PEG an attractive polymer for formulation development. 26 PEGylation of hydrophobic drugs improves their aqueous solubility, improves cellular uptake, prolongs circulation in the blood and therefore in vivo performance, and improves overall therapeutic efficacy. 27−31 The development of a pH-responsive system helps in the site-specific delivery of therapeutic agents, particularly in cancer chemotherapy. The pH of the tumor microenvironment (pH 6.5−6.8) is slightly acidic as compared with physiological pH 7.4. This pH difference provides an opportunity for selective delivery of drugs using pH-sensitive drug carriers. 32−34 Herein, we have conjugated chrysin to PEG using two pH-sensitive linkers, i.e., succinoyl and cis-aconityl, and evaluated its anticancer activity. The synthesized system was able to self-assemble at the nanoscale, which was characterized by different analytical techniques.
2.2. Synthesis of pH-Sensitive PEGylated Chrysin Conjugate. The PEGylated chrysin conjugates were synthesized by a two-step reaction. First, the hydroxyl group of PEG 4000 was modified into succinoyl and cis-aconityl functionals using succinic anhydride and cis-aconitic anhydride, respectively. 35 In the next step, after 6 h of stirring, a solution of modified PEG 4000 (mPEG, 0.500 g, 0.12 mmol), EDC (0.285 g, 0.18 mmol), and NHS (0.07 g, 0.06 mmol) in DMSO was dropwise added to the solution of chrysin (0.30 g, 0.12 mmol) in DMSO. The mixture was stirred for 24 h under nitrogen conditions. The resulting solution was dialyzed (MWCO 2 kDa) against ultrapure Milli-Q water, with the water changed every 6 h cycle, up to 3 days, and the product was collected by lyophilization.
2.3. Characterization of PEGylated Chrysin Conjugates. The PEGylated chrysin conjugates were characterized by proton nuclear magnetic resonance (Bruker 500-MHz Ultra shield plus NMR instrument) and Fourier transform infrared (PerkinElmer Spectrum 65 series, USA) spectroscopic techniques. For the NMR analysis, the PEGylated chrysin conjugates were dissolved in CDCl 3 , and pure chrysin was dissolved in DMSO-d 6 solvent. Then both were scanned using a 1 H NMR (500 MHz) instrument. For the FTIR analysis, samples were mixed with KBr, pelletized, and scanned from 4000 to 400 cm −1 . The critical aggregation concentrations of the conjugates were measured using the pyrene fluorescent method, as previously reported. 36 A 25 μL aliquot of pyrene solution (6 × 10 7 M) was incubated with 5−500 μg/mL of PEGylated chrysin conjugates in the dark and at room temperature for 3 h. After that, samples were used to measure fluorescence intensities at a 339 nm excitation and 383 and 373 nm emission intensities. The fluorescence intensities were measured using a microplate reader (Synergy H1 Hybrid Reader, Biotek; Winooski, VT, USA). The intensity ratio and concentration of PEGylated chrysin conjugates were plotted to determine the critical micelle concentration (CMC) value.
2.4. Preparation of PEGylated Chrysin Nanoparticles (PCNP). To obtain PEGylated chrysin nanoparticles, i.e., PCNP-1 (PEG 4000 −chrysin via succinoyl linker) and PCNP-2 (PEG 4000 −chrysin via cis-aconityl linker), the conjugates were dissolved in acetone at a concentration of 10 mg/mL. Then, 3 mL of this solution was dropwise added to 15 mL of water under stirring at 1000 rpm. After that, the mixtures were kept stirring (1000 rpm, RT) for 3 h to evaporate acetone. The chrysin content in nanoparticles was quantified by a UV− visible spectrophotometer at 266 nm, and % drug content was calculated by the following equation.
amount of chrysin in PCNPs total weight of PEG chrysin conjugate 100 4000 2.5. Characterizations of PCNP. The size distribution of PCNP-1 and PCNP-2 was determined using DLS technique (Malvern Instruments, UK). TEM instrument (JEOL JEM-2100 TEM, Tokyo, Japan) was used to confirm the morphology of nanoparticles. Powder XRD (D8 Advance, Bruker, Germany) and DSC (DSC 4000, PerkinElmer, USA) instruments were used to confirm the physical state of the prepared nanoparticles. 35 2.6. In Vitro Drug Release Study. The release of chrysin from PCNP-1 and PCNP-2 was performed using the dialysis method. 37 PCNP-1 and PCNP-2 nanoparticles were separately placed in a dialysis bag (MWCO 2 kDa). The formulationcontaining dialysis bag was immersed in 100 mL of sodium acetate buffer (pH 5.0), phosphate buffer (pH 6.5), or phosphate buffer saline (pH 7.4). The experiment was performed at 37°C and with 100 rpm stirring speed. At different intervals, aliquots from the released media were collected for drug analysis and replenished with an equivalent volume of fresh media.
2.7. Stability Study. The stability study of prepared PCNPs was performed by measuring particle size (colloidal stability) and % drug content (drug stability) over a period of 96 h at refrigeration temperature. The change in particle size was measured by DLS method, and % chrysin content was measured using a UV−visible spectroscopic method.
2.8. Cell Culture. Human breast cancer (MDA-MB-231) and MCF-7 cells were purchased from NCCS (Pune, India) and cultured in Dulbecco's Modified Eagle's Medium-high Glucose (DMEM-HG) and MEM, respectively, with 10% fetal bovine serum (FBS) in a fully humidified atmosphere of 5% CO 2 at 37°C. After reaching 80% to 90% confluency, cells were trypsinized for experiments. 38 2.8.1. In Vitro Cytotoxicity Assay. To evaluate the anticancer activity of prepared nanoparticles, an in vitro cytotoxicity assay was performed. About 7.0 × 10 3 cells/well were seeded in 96-well plates and incubated for 24 h in a CO 2 incubator. After that, cells were treated with pure chrysin, PCNP-1, and PCNP-2 at different concentrations (5, 10, 20, 40, 60, 80, and 100 μg/mL). After the completion of the incubation period (24 and 48 h), media from the wells was discarded, and fresh media containing 0.5 mg/mL of MTT reagent was added to each well. After 4 h of incubation, MTT solution was removed, and crystal formazan was dissolved by adding 100 μL of DMSO to each well. After 10 min of incubation, absorbance was measured at 570 nm using a multimode plate reader (Synergy H1, Biotek, USA). Untreated cells were considered as a control, and % cell viability was determined with respect to the viability of the control cells.
2.8.2. Apoptotic Assay. To examine the cellular changes after treatment with PCNP, an apoptotic assay was performed by acridine orange/ethidium bromide (AO/EtBr) staining, as previously reported. 38 About 4.0 × 10 4 cells/well were seeded in 12-well plates. After 24 h, the cells were treated with pure chrysin, PCNP-1 and PCNP-2, equivalent to 10 μg/mL of chrysin. After the completion of the incubation period (24 and 48 h), the cells were trypsinized and centrifuged at 1800 rpm for 5 min. The resultant cell pellets were dispersed in 1× PBS, and 10 μL of suspension was taken for further examination. The cell suspension was mixed with 5 μL of AO dye (1 mg/ mL) and EtBr dye (1 mg/mL) and mounted on a clean glass slide. Then, cells were observed using a fluorescence microscope (Zeiss, Germany). The cells were counted using ImageJ software and plotted as % cells in a stacked column graph by using Graphpad Prism 6.
2.8.3. Colony Formation Assay. To assess the colony formation potential of MDA-MB-231 cells after the treatment with PCNP, a colony formation assay was performed. 39 For that, about 300 cells/well were seeded in 6-well plates. After 24 h, cells were treated with a concentration of 10 μg/mL of pure chrysin, PCNP-1, and PCNP-2. The plates were kept in a CO 2 incubator for 7 days to allow the cells to form colonies. Further, the media was discarded and washed twice with 1× PBS. The cells were fixed with cell-fixing solution (95:5, ethanol and glacial acetic acid) for 15 min. The cell-fixing solution was removed, and the cells were washed with PBS. Finally, the colonies were stained by crystal violet staining solution (5 mg/mL crystal violet in 2% ethanol) for 15 min at room temperature and washed with PBS. The number of colonies were counted under microscope (Carl Zeiss, Germany), and a graph was plotted by using GraphPad Prism software (GraphPad 6.0). Representative images were taken by a digital camera (Nikon 1000D camera, Taiwan).
2.9. Statistical Analysis. All the experimental measurements were performed in triplicate, and the values are expressed as the mean ± standard deviation (SD). A p-value less than 0.05 was considered statistically significant.

RESULTS AND DISCUSSION
3.1. Synthesis and Characterizations of pH-Sensitive PEGylated Chrysin Conjugate. The synthesis procedure of PEGylated chrysin conjugate is schematically shown in Figure  S1. These amphiphilic conjugates of chrysin were prepared by a two-step reaction. In the first step, the PEG 4000 was reacted to succinoyl and cis-aconityl linkers via ring-opening mechanism. In the second step, chrysin was conjugated to succinoyl− PEG 4000 or cis-aconityl−PEG 4000 conjugates through esterification. The formation of the PEGylated chrysin conjugates was confirmed by 1 H NMR and FTIR techniques. Figure S2 shows the 1 H NMR spectra of PEG 4000 , PEG 4000 −SA, and PEG 4000 − CAA where PEG 4000 shows the characteristic chemical shift values at 3.65−3.68 and 4.26 ppm due to CH 2 −CH 2 and −OH groups, respectively. 41 44 The succinoyl linkage containing PEGylated chrysin was confirmed by observed  Figure 2b shows the predictive model representing a possible arrangement of chrysin, linkers, and PEG 4000 molecules in self-assembled PCNP formed by PEG 4000 −chrysin conjugates. The observed chrysin content was found to be 4.05% and 4.11% in PCNP-1 and PCNP-2, respectively ( Figure S6). The TEM images and particle size distribution of PCNP-1 (Figure 2c,d) and PCNP-2 ( Figure  2e,f) show the formation of spherical nanoparticles with a particle size of about 60 nm. The hydrodynamic diameter and surface charge of nanoparticles were measured by DLS technique ( Figure S7). The particle size, polydispersity index, and zeta potential values were 77.3 nm, 0.173 ± 0.003, and −4.25 mV for PCNP-1 and 75.5 nm, 0.185 ± 0.040, and −3.47 mV for PCNP-2, respectively. These results suggested the formation of nanosized and monodisperse particles.
The thermal behavior and physical state of chrysin after formation of the PCNPs were confirmed by DSC and XRD analysis. Figure 2g shows the DSC scans of chrysin, PEG 4000 , PCNP-1, and PCNP-2. The pure chrysin shows a sharp endothermic peak at 289°C 45 while pure PEG 4000 shows an endothermic peak at 64°C, corresponding to their melting points. The PCNP-1 and PCNP-2 show endothermic peaks at 55 and 59°C, respectively, because of the presence of PEG 4000 . However, the typical endothermic peak of pure chrysin was not observed in the scans of both PCNP-1 and PCNP-2, which specifies the phase conversion of chrysin. This change in the physical state of chrysin was further confirmed by PXRD analysis (Figure 2h). The XRD pattern of pure chrysin shows sharp peaks at 2θ values 7.2°, 12.2°, 13.2°, 14.7°, 17.5°, 20.3°, 22.3°, 24.6°, and 27.4°because of the crystalline nature of chrysin. 45 The pure PEG 4000 shows three peaks at 2θ values 13.8°, 19.2°, and 23.3°. The PCNP-1 and PCNP-2 show the 2θ values at 18.8°and 23.05°, and 18.4°and 22.7°, respectively. So, because no sharp peaks of pure chrysin were observed in the XRD spectra of PCNP-1 and PCNP-2, the formation of amorphous nanoparticles was confirmed. 35 3.3. PCNP Releases Chrysin in a pH-Dependent Manner. To evaluate the effect of succinoyl and cis-aconityl linkers on the release of chrysin from PCNPs, PCNP-1 and PCNP-2 were put into dialysis bags and dialyzed against SAB (pH 5.0), PB (pH 6.5), and PBS (pH 7.4). As shown in Figure  3a, the amount of chrysin released from PCNP-1 and PCNP-2 was 23.18 and 20.18%, respectively, after 24 h at pH 7.4. The chrysin release was 42.13% and 46.98% from PCNP-1 and PCNP-2, respectively, at pH 6.5. Hence, at the slightly acidic pH of the tumor microenvironment, the amount of chrysin released was higher than at the physiological pH 7.4. However, the chrysin release was about 60% in the first 3 h at pH 5.0 from both PCNP-1 and PCNP-2. Further, more than 90% of chrysin was released from PCNP-2 at pH 5.0 in 24 h, while 72.9% of chrysin was released from PCNP-1. As expected, a higher release of chrysin was observed at the lower pH 5.0 while a slow chrysin release was observed at pH 7.4. However, at pH 5.0 the accelerated chrysin release was observed because of the rapid hydrolysis of cis-aconityl linker. Hence, the observed results from this study suggest the effect of the pHresponsive (cis-aconityl) linker on the release of chrysin from the prepared PCNP-2 nanoparticles. Importantly, no burst release of drug was observed at pH 6.5 and pH 7.4 from either PCNP-1 or PCNP-2. Therefore, this pH-sensitive response of PCNP-2 may help in the site-specific release and delivery of chrysin.
3.4. Stability of PCNP. In order to check the stability of PCNP, particle size and drug content were measured at freeze temperature. Figure S8 shows the stability results of PCNP-1 and PCNP-2. Initially the particle sizes were observed as 77 and 75 nm for PCNP-1 and PCNP-2, respectively. After a period of 96 h, the particle sizes were observed as 91 and 78 nm for PCNP-1 and PCNP-2, respectively. Hence, no significant particle size changes were observed for either PCNP, which confers physical stability of the nanoparticles. Similarly, % chrysin content was observed as 91 and 98% over the period of 96 h. Hence, the observed physical as well as chemical stability results clearly confers the stability of the prepared PEGylated nanoparticles.
3.5. PCNP Causes Concentration-and Time-Dependent Cytotoxicities for Human Breast Cancer Cells. The cytotoxicity of prepared PCNP was evaluated against MCF-7 and MDA-MB-231 human breast cancer cells. The results of cell viability after treatment with chrysin, PCNP-1, and PCNP-2 at concentrations of 5, 10, 20, 40, 60, 80, and 100 μg/mL are presented in Figure 3b. It was observed that the PCNP-1 and PCNP-2 showed a significant decrease in the viability of breast cancer cells compared to chrysin in both dose-and timedependent manner. In addition, PCNP-2 showed a higher inhibitory effect (p value < 0.05) as compared with PCNP-1 or chrysin after 24 and 48 h. The half-maximal inhibitory concentration (IC 50 ) of the pure chrysin, PCNP-1, and PCNP-2 after 24 and 48 h incubation time are presented in Table 1. The IC 50 values (at 24 h) were found to be 52.2, 49.6, and 34.8 μg/mL for pure chrysin, PCNP-1, and PCNP-2, respectively. After 48 h, the IC 50 values were decreased to 19.4, 15.2, and 6.2 μg/mL for pure chrysin, PCNP-1 and PCNP-2, respectively. In the case of MCF-7 cells, the IC 50 values were 89.2, 48.3, and 35.7 μg/mL after 24 h of incubation with chrysin, PCNP-1, and PCNP-2, respectively ( Figure 3c). The IC 50 values were decreased after 48 h of incubation and observed to be 68.4, 44, and 26.1 μg/mL for chrysin, PCNP-1, and PCNP-2, respectively. The results of the cytotoxicity studies depict the potent and significantly enhanced anticancer activity of chrysin-conjugated nanoparticles in comparison to pure chrysin. The values of IC 50 also suggest that PCNP-2 had comparatively higher cytotoxicity than PCNP-1.
3.6. Induction of Apoptosis by PCNP. Cytotoxicity of prepared PCNP was further confirmed by an apoptosis induction assay using AO/EtBr staining. This assay helps to check the changes at the cellular level and to differentiate between apoptotic, necrotic, and live cells after treatment with pure chrysin, PCNP-1, and PCNP-2. The AO dye gives green fluorescence after staining the nuclei of the live cells that have an intact cellular membrane, whereas EtBr emits red fluorescence after staining the nuclei of dead cells that have a compromised/damaged cellular membrane. Figure 4 represents the fluorescent images of MDA-MB-231 cells treated with 10 μg/mL concentration of pure chrysin, PCNP-1, and PCNP-2. After 24 h of incubation, MDA-MB-231 cells treated with pure chrysin did not show any significant signs of apoptosis and necrosis.
After 48 h, the cells started to emit red fluorescence that implies chrysin exhibited cell death after 48 h. However, cells treated with PCNP-1 and PCNP-2 showed apoptosis and necrosis after 24 h, which were further increased after 48 h of incubation. The number of cell deaths was higher for PCNP-2 than PCNP-1 or pure chrysin at 10 μg/mL after the 24 and 48 h periods of time. The viable, apoptotic, and necrotic cells were also counted for each group represented in Figure S9. More than 95% of the control cells produced green fluorescent, implying healthy cells.    revealed that PCNP-1 and PCNP-2 can induce apoptosis in a time-dependent manner in MDA-MB-231 cells, which also corroborates the results of the in vitro cytotoxicity assay. Therefore, these findings suggest that prepared PCNP have higher anticancer activity and can induce more apoptosis in MDA-MB-231 cells as compared with pure chrysin. 3.7. Colony Formation Assay. To further validate the cytotoxicity and growth inhibitory effects of chrysin and PCNP, the clonogenic assay was performed. As shown in Figure 5, with a 10 μg/mL concentration of chrysin, PCNP-1, and PCNP-2, the number of colony formations by MDA-MB-231 cells was decreased as compared with untreated control cells. Quantitatively, the colony formation activity of cells was observed to be 31.4%, 7.3%, and 2.9% after incubation with chrysin, PCNP-1, and PCNP-2, respectively. Further, the cells treated with PCNP-1 and PCNP-2 showed fewer colonies in comparison with the cells treated with pure chrysin. Following the patterns observed in the cytotoxicity assay and apoptosis study, PCNP-2 also showed more inhibition than PCNP-1 of colony formation by MDA-MB-231 cells.

CONCLUSION
In the present work, pH-sensitive PEGylated chrysin nanoparticles (PCNPs) were prepared and evaluated for their anticancer activity against MDA-MB-231 human breast cancer cells. The conjugation of chrysin with PEG via different pHsensitive groups was confirmed by FTIR and 1 H NMR analysis. The prepared conjugates were self-assembled into nanoparticles with a particle size of about 60 nm. Both the PCNPs showed a pH-dependent drug release, that is, a faster and more complete drug release at acidic pH than physiological pH. The PCNPs with cis-aconityl linkage showed higher toxicity toward MDA-MB-231 cells as compared with pure chrysin or succinoyl linked PCNPs. Further, these PCNPs caused induction of apoptosis and a decrease in the colony formation ability of breast cancer cells. Therefore, this study highlights the development and significance of PEGylated chrysin as an anticancer agent.
Reaction schemes representing the PEGylation of chrysin; 1 H NMR, FTIR, and UV−visible spectra of chrysin and PEGylated chrysin conjugates; chemical structure of chrysin; particle size distribution of PCNP-1 and PCNP-2; stability results of the PEGylated chrysin nanoparticles; and percentage of viable, early apoptotic, late apoptotic, and necrotic cells after the treatment with chrysin, PCNP-1, and PCNP-2 for 24 and 48 h (PDF)