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. 2025 Jun 12;301(7):110367. doi: 10.1016/j.jbc.2025.110367

Regulation of NRF2 by stably associated phosphoinositides and small heat shock proteins in response to stress

Changliang Chen 1,2,‡, Noah D Carrillo 1,2,‡, Mo Chen 2, Tianmu Wen 2, Poorwa Awasthi 2, Richard A Anderson 2,§,∗, Vincent L Cryns 1,2,§,∗
PMCID: PMC12273567  PMID: 40516875

Abstract

Reactive oxygen species are generated by aerobic metabolism, and their deleterious effects are buffered by the cellular antioxidant response, which prevents oxidative stress. The nuclear factor erythroid 2-related factor 2 (NRF2) is a master transcriptional regulator of the antioxidant response. Basal levels of NRF2 are kept low by ubiquitin-dependent degradation of NRF2 by E3 ligases, including the Kelch-like ECH-associated protein 1 (KEAP1). Here, we show that the stability and function of NRF2 is regulated by the type I phosphatidylinositol phosphate kinase γ (PIPKIγ), which binds NRF2 and is required to stably couple phosphatidylinositol 4,5-bisphosphate to NRF2 in response to oxidative stress. Stress also induces the interaction of the small heat shock protein HSP27 and NRF2, and this interaction is enhanced by phosphatidylinositol 4,5-bisphosphate. Silencing PIPKIγ or HSP27 destabilizes NRF2, reduces expression of its target gene heme oxygenase-1 (HO-1), and sensitizes cells to oxidative stress. These data demonstrate an unexpected collaboration between phosphoinositides, which are stably coupled to NRF2, and HSP27, which is recruited to NRF2 by a phosphoinositide-dependent mechanism to regulate NRF2 stability and function. These findings also point to PIPKIγ and HSP27 as drug targets to destabilize NRF2 in cancer.

Keywords: NRF2; phosphoinositide; PtdIns(4,5)P2; small heat shock proteins: oxidative stress; nucleus

Oxidative stress is characterized by elevated levels of reactive oxygen species (ROS), which damage cellular molecules and contribute to the pathogenesis of diverse diseases, including cancer (1, 2). Oxidative stress is a metabolic hallmark of cancer that activates the cellular antioxidant response to enable tumor cells to survive in the setting of high ROS levels (3). The nuclear factor erythroid 2–related factor 2 (NRF2) is a master transcriptional regulator of the cellular antioxidant response that accumulates in the nucleus in response to oxidative stress (4). NRF2 controls the expression of a broad array of genes that contain antioxidant response elements, such as heme oxygenase-1 (HO-1), which act to restore redox homeostasis (3, 5, 6). In the absence of stress, NRF2 is negatively regulated by ubiquitin-dependent proteasomal degradation mediated predominantly by the Kelch-like ECH-associated protein 1 (KEAP1), an adaptor subunit of the Cullin 3-based E3 ubiquitin ligase, which results in low basal levels of NRF2 (4). ROS oxidizes several cysteine residues in KEAP1 to disrupt its interaction with the Neh2 domain of NRF2, resulting in NRF2 protein stabilization and activation (7). Notably, elevated NRF2 expression is evident across diverse tumor types and has been associated with drug resistance and cancer progression (5). While mutations in KEAP1 and NRF2 have been implicated in stabilizing NRF2 within a subset of cancers (4, 8, 9), it is likely that additional regulators contribute to the physiological control and aberrant activation of NRF2 in cancer.

The phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) associates with many cellular proteins to support a broad spectrum of cellular functions, whereas its dysregulated activity contributes to diverse diseases (10, 11). Cellular PtdIns(4,5)P2 is predominantly generated by type I phosphatidylinositol 4-phosphate (PIP) 5-kinases (PIPKIs), which include three isoforms (α, β, and γ) (12). PtdIns(4,5)P2-generating enzymes are physically associated with PtdIns(4,5)P2 protein effectors (13, 14, 15). Among these, several nuclear PtdIns(4,5)P2 effectors, including nuclear speckle–targeted PIPKIα regulated-poly(A) polymerase (Star-PAP), p53, brain acid soluble protein 1 (BASP1), and BRG1, have been reported (14, 15, 16, 17). PIPKIα binds stress-activated wildtype p53 and mutant p53 in the nucleus, facilitating the transfer of PtdIns(4,5)P2 to the C-terminal domain of p53. This interaction leads to the recruitment of the small heat shock proteins (sHSPs), HSP27 and αB-crystallin, to the nuclear p53–PtdIns(4,5)P2 complex (14). Notably, PtdIns(4,5)P2 binding to p53 significantly augments the interaction between sHSPs and p53, and these sHSPs are crucial for stabilizing the nuclear p53–PtdIns(4,5)P2 complex (14).

Here, we report that NRF2 is a novel nuclear phosphoinositide effector. Specifically, we demonstrate that the γ isoform of PIPKI (PIPKIγ) binds NRF2 in response to oxidative stress and is required to stably couple PtdIns(4,5)P2 to NRF2 to generate an NRF2–PtdIns(4,5)P2 complex that is analogous to the p53–phosphoinositide complex (14). Stress induces the recruitment of sHSPs to the nuclear NRF2–PtdIns(4,5)P2 complex, with HSP27 playing a pivotal role in stabilizing NRF2. These findings demonstrate a previously unrecognized role of nuclear phosphoinositides and sHSPs in regulating NRF2, suggesting a broader role of nuclear phosphoinositides in cell signaling via their stress-regulated linkage to a specific subset of nuclear proteins.

Results

PtdIns(4,5)P2 interacts with NRF2 in the nucleus

Phosphoinositide signaling, particularly within the nucleus, is activated in response to cellular stress (10). Nuclear PtdIns(4,5)P2 levels are dramatically increased by cellular stress, promoting complex formation with proteins such as p53 in the membrane-free nucleoplasm to stabilize p53 and mitigate stress-induced apoptosis (14, 15). In parallel, NRF2 is a central orchestrator of the oxidative stress response, with its nuclear accumulation being a characteristic response to stress (5). Moreover, NRF2 is unstable under basal conditions (4, 5), leading us to hypothesize that phosphoinositides bind and stabilize NRF2 in response to stress analogous to p53 (14, 15). To test this hypothesis, we immunoprecipitated (IPed) endogenous NRF2 and simultaneously assayed for NRF2–PtdIns(4,5)P2 complexes using a dual approach of double fluorescent immunoblotting (IB) combined with metabolic [3H]myo-inositol radiolabeling. Using these approaches, PtdIns(4,5)P2 co-IPed with endogenous NRF2, and the PtdIns(4,5)P2 signal overlapped with NRF2 after SDS-PAGE (Fig. 1A). The PtdIns(4,5)P2 antibody detection after SDS-PAGE was validated using [3H]myo-inositol, which is specifically incorporated into phosphoinositides (18, 19). NRF2 was IPed from labeled cells and resolved via SDS-PAGE showing peak [3H] activity in the gel section containing the NRF2–PtdIns(4,5)P2 complex, which was increased by diethyl maleate (DEM) treatment (Fig. 1B and Extended Data Fig. S1A). This approach indicates that NRF2 is a phosphoinositide effector capable of forming a stable complex with PtdIns(4,5)P2 similar to p53 (14, 15). Moreover, ectopically expressed FLAG-tagged NRF2 was pulled down by PtdIns(4,5)P2-conjugated beads (Fig. 1C), confirming an interaction between NRF2 and PtdIns(4,5)P2. To determine whether PtdIns(4,5)P2 directly binds NRF2, microscale thermophoresis (MST) was performed with purified recombinant NRF2 (20). Ptdlns(4,5)P2 bound NRF2 with greater affinity (Kd = 109 ± 10 nM) than other phosphoinositides (Table 1 and Extended Data Fig. S1B). These data indicate that PtdIns(4,5)P2 directly binds NRF2 with high affinity in vitro. This interaction is distinct from the PtdIns(4,5)P2 stably coupled to NRF2 that is resistant to denaturation and SDS-PAGE.

Figure 1.

Figure 1

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PtdIns(4,5)P2 interacts with NRF2 in the nucleus.A, endogenous NRF2 was IPed from MDA-MB-231 cells, and PtdIns(4,5)P2 was detected by fluorescent IB. IgG was used as a negative control. B, MDA-MB-231 cells were cultured from low confluency in media containing [3H]myo-inositol or unlabeled myo-inositol. After 72 h, cells were treated with 200 μM DEM or vehicle for 4 h before being processed for IP against NRF2. NRF2 and PtdIns(4,5)P2 were confirmed and quantified (Extended Data Fig. S1A) by fluorescent WB and ImageJ before samples were resolved by SDS-PAGE and the gel sectioned corresponding to the NRF2 immunoblot. Gel sections from the indicated lanes were then dissolved and analyzed by liquid scintillation counting (LSC). Graph is mean ± SD C, HEK293FT cells were transiently transfected with FLAG-tagged NRF2 and processed for pulldown of PtdIns(4,5)P2 48 h later. D and E, MDA-MB-231 cells were treated with vehicle or 200 μM DEM for 4 h before being processed for IF against PtdIns(4,5)P2 and NRF2. DAPI was used to stain the nuclei. The images (D) were taken with a Leica SP8 confocal microscope and processed by ImageJ. The graph (E) is shown as mean ± SD of three independent experiments with n = 10 cells scored in each independent experiment. F and G, MDA-MB-231 cells were treated with 200 μM DEM for 4 h before being processed for PLA between PtdIns(4,5)P2 and NRF2. DAPI was used to stain nuclei. The images (F) were taken with a Leica SP8 confocal microscope. LASX (Leica) was used to quantify the PLA signal, and the graph (G) is shown as mean ± SD of three independent experiments with n = 15 cells scored in each independent experiment. For all panels, n = 3 independent experiments; scale bar represents 5 μm. DAPI, 4',6-diamidino-2-phenylindole; DEM, diethyl maleate; HEK293FT, human embryonic kidney 293FT cell line; IB, immunoblotting; IF, immunofluorescence; IgG, immunoglobulin G; IP, immunoprecipitation; IPed, immunoprecipitated; NRF2, nuclear factor erythroid 2–related factor 2; PLA, proximity ligation assay; PtdIns(4,5)P2, phosphatidylinositol 4,5-bisphosphate; WB, Western blot.

Table 1.

MST analysis of NRF2 and PtdIns isomers

Binding partners Binding affinity with NRF2 (Kd)
PtdIns No binding
PtdIns4P 1200 ± 20 nM
PtdIns(4,5)P2 109 ± 10 nM
PtdIns(3,4,5)P3 7400 ± 150 nM
PtdIns5P 2600 ± 1.7 nM
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The interaction of recombinant fluorescently labeled NRF2 and nonlabeled PtdIns isomers was quantitated by MST assay. A constant concentration of fluorescently labeled NRF2 was incubated with increasing concentrations of nonlabeled ligands to calculate the binding affinity. The binding affinities determined by MST are shown as indicated Kd values. Data are presented as the mean ± SD; n = 3.

NRF2 is activated and translocates to the nucleus in response to cellular stress, especially oxidative stress (21, 22, 23, 24). Consistent with these reports, NRF2 expression was substantially increased by genotoxic stress (cisplatin), NRF2 induction (tertiary butylhydroquinone and sulforaphane), oxidative stress (DEM), and the SOD1 inhibitor LCS-1 (Extended Data Fig. S1C). DEM rapidly induced NRF2 expression in a time-dependent manner (Extended Data Fig. S1D). NRF2 and PtdIns(4,5)P2 colocalized in the nucleus as determined by immunofluorescence (IF), and their colocalization was increased by DEM (Fig. 1, D and E). Their stress responsive interaction was confirmed in situ using proximity ligation assay (PLA) (Fig. 1, F and G), an assay used to detect associated proteins and post-translational modifications on target proteins (25, 26). These findings demonstrate that NRF2–PtdIns(4,5)P2 complexes are present in the nucleus and that oxidative stress enhances complex formation in the nucleus, similar to p53 (14, 15).

PIPKIγ controls NRF2 stability

Ptdlns(4,5)P2 and the enzymes responsible for its generation, specifically the PIP kinases, PIPKIα, PIPKIγ, PIPKIIα, and PIPKIIβ, are all present in the nucleus (27). We postulated that one or more of these PIP kinases interact with NRF2 to generate Ptdlns(4,5)P2 on NRF2, thereby stabilizing the protein. KO of these PIP kinases by CRISPR–Cas9 revealed that only PIPKIγ regulated NRF2 protein levels (Fig. 2, A and B). Transient knockdown (KD) of PIPKIγ reduced basal protein levels of NRF2, whereas mRNA levels were unaffected (Extended Data Fig. S2, A–C). Furthermore, PIPKIγ KD with different siRNAs or PIPKIγ KO inhibited NRF2 induction by tHBQ as well as basal NRF2 levels (Fig. 2, C and D). Notably, the destabilization of NRF2 by PIPKIγ KD was attenuated by the proteasome inhibitor MG132 (Fig. 2E and Extended Data Fig. S2D), indicating that the ubiquitin–proteasome pathway contributes to the observed effects. Collectively, these results indicate that PIPKIγ regulates NRF2 protein stability under basal conditions and in response to oxidative stress.

Figure 2.

Figure 2

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PIPKIγ controls NRF2 stability.A and B, the expression of NRF2 was analyzed by IB in MDA-MB-231 cells with deletion of PIPKIα, PIPKIγ, PIPKIIα, or PIPKIIβ (A). The intensity of the immunoblots was quantified, and the graph is shown (B) as mean ± SD. C, KD of PIPKIγ using two shRNAs in HCT116 cells. Cells were treated with vehicle or 100 μM tBHQ for 4 h. The expression of NRF2 was analyzed by IB, and the graph is shown as mean ± SD. D, MDA-MB-231 cells (parental or with PIPKIγ deletion) were treated with 100 μM tBHQ for 4 h. NRF2 protein expression was analyzed by IB. E, MDA-MB-231 cells transfected with control siRNAs or siRNAs targeting PIPKIγ were treated with or without 10 μM MG132 for the indicated time. NRF2 protein levels were determined by IB and quantified (Extended Data Fig. S2B) by ImageJ. For all panels, n = 3 independent experiments. KD, knockdown; IB, immunoblotting; NRF2, nuclear factor erythroid 2–related factor 2; PIPKIγ, phosphatidylinositol phosphate kinase γ; tBHQ, tertiary butylhydroquinone.

PIPKIγ generates the NRF2–PtdIns(4,5)P2 complex

Having demonstrated that PIPKIγ regulates NRF2 stability, we next examined whether PIPKIγ binds NRF2. NRF2 co-IPed with PIPKIγ, and the association between PIPKIγ and NRF2 was increased by oxidative stress (Fig. 3, A and B). IF revealed that NRF2 and PIPKIγ colocalized in the nucleus, and their colocalization was significantly enhanced by oxidative stress similar to the NRF2 and PtdIns(4,5)P2 observations (Fig. 3, C and D). The association of PIPKIγ and NRF2 was also examined by PLA, which revealed that oxidative stress increased NRF2–PIPKIγ foci in the nucleus (Fig. 3, E and F). Functionally, PIPKIγ KD reduced NRF2–PtdIns(4,5)P2 complexes as detected and quantified by PLA, indicating that a stress responsive interaction between NRF2 and PIPKIγ mediates NRF2–PtdIns(4,5)P2 production (Fig. 3, G and H). Taken together, these data demonstrate that PIPKIγ associates with NRF2 in the nucleus and is necessary for generating the nuclear NRF2–Ptdlns(4,5)P2 complex.

Figure 3.

Figure 3

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PIPKIγbinds nuclear NRF2. A and B, MDA-MB-231 (A) or HCT116 (B) cells were treated with vehicle or 200 μM DEM for 4 h. Endogenous PIPKIγ was IPed, and NRF2 was analyzed by IB and quantified by ImageJ. C and D, MDA-MB-231 cells were treated with 200 μM DEM for 4 h before being processed for IF against PIPKIγ and NRF2. DAPI was used to stain nuclei. The images (C) were taken with a Leica SP8 confocal microscope and processed by ImageJ. The graph (D) is shown as mean ± SD of three independent experiments with n = 10 cells scored in each independent experiment. E and F, MDA-MB-231 cells were treated with 200 μM DEM for 4 h before being processed for PLA between PIPKIγ and NRF2. DAPI was used to stain nucleic acids. The images (E) were taken with a Leica SP8 confocal microscope. The red PLA signal was quantified by LASX (Leica), and the graph (F) is shown as mean ± SD of three independent experiments with n = 15 cells scored in each independent experiment. G and H, MDA-MB-231 cells with or without PIPKIγ knockdown (KD) were treated with vehicle or 200 μM DEM for 4 h before being processed for PLA between PtdIns(4,5)P2 and NRF2 (G). DAPI was used to stain nucleic acids. The red PLA signal was quantified by LASX (Leica). The experiments were repeated three times. The graph is shown as mean ± SD of n = 15 cells from one representative experiment (H). For all panels, n = 3 independent experiments; scale bar represents 5 μm. DAPI, 4',6-diamidino-2-phenylindole; DEM, diethyl maleate; IB, immunoblotting; IF, immunofluorescence; IPed, immunoprecipitated; NRF2, nuclear factor erythroid 2-related factor 2; PIPKIγ, phosphatidylinositol phosphate kinase γ; PLA, proximity ligation assay.

NRF2 binds sHSPs in the nucleus

PtdIns(4,5)P2 binding to p53 was recently reported to recruit sHSPs to the p53–PtdIns(4,5)P2 complex and stabilize p53 (14). Intriguingly, sHSPs have also been implicated in resistance to oxidative stress (28, 29, 30). Hence, we postulated that sHSPs bind and stabilize NRF2 to inhibit oxidative stress. Consistent with this idea, the sHSPs HSP27 and αB-crystallin co-IPed with NRF2 under basal conditions, whereas HSP40, HSP60, and HSP90, which are highly expressed, interacted minimally with NRF2 (Fig. 4, A and B). Incubation of recombinant NRF2 with increasing amounts of recombinant HSP27 or αB-crystallin revealed direct and saturable binding (Extended Data Fig. S3, A and B). These interactions were confirmed by MST, which revealed high affinity binding of HSP27 (Kd = 213 ± 30 nM) and αB-crystallin (Kd = 859 ± 70.1 nM) to NRF2, respectively (Extended Data Fig. S3, C and D). Oxidative stress enhanced the interaction of NRF2 with each sHSP as determined by coimmunoprecipitation (co-IP) (Fig. 4, C–E and Extended Data Fig. S3E). Moreover, the nuclear signal of sHSPs was increased upon oxidative stress in agreement with prior reports (31, 32), and the colocalization of HSP27 with NRF2 in the nucleus was increased in response to DEM treatment as determined by IF (Fig. 4, F and G). Strikingly, the NRF2–HSP27 and NRF2–αB-crystallin interactions detected by PLA were dramatically increased by DEM treatment (Fig. 4, H and I and Extended Data Fig. S3F). These results indicate that nuclear NRF2 associates with sHSPs and that these interactions are enhanced by oxidative stress.

Figure 4.

Figure 4

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HSP27 interacts with NRF2. A and B, endogenous NRF2 was IPed from MDA-MB-231 (A) or MCF7 (B) cells, and HSP27, αB-crystallin, HSP40, HSP60, and HSP90 were analyzed by IB. IgG was used as a negative control. C, MDA-MB-231 cells were treated with vehicle or 200 μM DEM for 4 h. Endogenous NRF2 was IPed, and HSP27 was analyzed by IB. D, MDA-MB-231 cells were treated with vehicle or 200 μM DEM for 4 h. Endogenous HSP27 was IPed, and NRF2 was analyzed by IB. E, quantification using ImageJ of co-IPed protein levels (C and D). F and G, MDA-MB-231 cells were treated with vehicle or 200 μM DEM for 4 h before being processed for IF staining against HSP27 and NRF2 (F). DAPI was used to stain nuclei. The images (F) were taken with a Leica SP8 confocal microscope and processed by ImageJ. The experiments were repeated three times, and the graph (G) is shown as mean ± SD of three independent experiments with n = 10 cells scored in each independent experiment. H and I, MDA-MB-231 cells were treated with vehicle or 200 μM DEM for 4 h before being processed for PLA between HSP27 and NRF2. DAPI was used to stain nuclei. The images (H) were taken with a Leica SP8 confocal microscope. The red PLA signal was quantified by LASX (Leica), and the graph (I) is shown as mean ± SD of three independent experiments with n = 15 cells scored in each independent experiment. For all panels, n = 3 independent experiments; scale bar represents 5 μm. DAPI, 4',6-diamidino-2-phenylindole; DEM, diethyl maleate; HSP27, heat shock protein 27; IB, immunoblotting; IF, immunofluorescence; IgG, immunoglobulin G; IPed, immunoprecipitated; NRF2, nuclear factor erythroid 2-related factor 2; PLA, proximity ligation assay.

The Neh2 domain of NRF2 binds sHSPs and their interaction is regulated by PtdIns(4,5)P2

We used a series of NRF2 truncation mutants (Fig. 5A) to map its binding domain for sHSPs and PtdIns(4,5)P2. Recombinant His-Tag NRF2 truncation mutants were incubated with HSP27, αB-crystallin, or PtdIns(4,5)P2 beads, and the NRF2 constructs that were pulled down by those beads were analyzed by IB. Our results demonstrate that deleting the Neh2 domain (amino acid residues 2–86) of NRF2 eliminates the interaction of NRF2 with HSP27, αB-crystallin, and PtdIns(4,5)P2 (Fig. 5, B–E), indicating a crucial role of the Neh2 domain in these interactions. However, we cannot exclude a potential contribution of the Neh6/7 domains (amino acid residues 205–400) in NRF2 as deletion of these domains partially attenuates the interaction with αB-crystallin. Notably, we also observed that PtdIns(4,5)P2 enhanced NRF2 binding to both HSP27 and αB-crystallin (Fig. 5, F and G), suggesting that PIPKIγ stabilizes NRF2 by enhancing the PtdIns(4,5)P2-dependent recruitment of HSP27 and αB-crystallin to the Neh2 domain of NRF2.

Figure 5.

Figure 5

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PtdIns(4,5)P2 binding to NRF2 promotes its interaction with sHSPs.A, schematic representation of NRF2 truncation mutants used in the study. B, recombinant His-Tag NRF2 truncation mutants were immobilized on HSP27 or αB-crystallin beads, and HSP27 or αB-crystallin bound to each mutant was analyzed by IB and the bound NRF2 mutants were quantified for HSP27 (C) and αB-crystallin (D) using ImageJ. E, recombinant His-Tag NRF2 truncation mutants were pulled down by PtdIns(4,5)P2 beads, and PtdIns(4,5)P2 bound to each NRF2 mutant was analyzed by IB. F and G, 6 nM recombinant His-tagged NRF2 and 200 nM HSP27 or 120 nM αB-crystallin were incubated with the indicated concentrations of PtdIns(4,5)P2. NRF2 was pulled down, and the bound HSP27 (F) or αB-crystallin (G) was analyzed by IB. For all panels, n = 3 independent experiments. HSP27, heat shock protein 27; PtdIns(4,5)P2, phosphatidylinositol 4,5-bisphosphate; sHSP, small heat shock protein.

HSP27 stabilizes NRF2

To specifically investigate the role of sHSPs in NRF2 stability, we used transient KD of αB-crystallin and HSP27. HSP27 KD reduced basal and oxidative stress–induced NRF2 protein expression in multiple human cancer cell lines (Fig. 6, A–C). NRF2 mRNA (NFE2L2) levels were not affected by HSP27 KD (Fig. 6D). Moreover, NRF2 destabilization by HSP27 KD was attenuated by MG132, consistent with a role for the ubiquitin–proteasome pathway as a mechanism by which NRF2 levels are reduced in the absence of HSP27 (Fig. 6E). In contrast, αB-crystallin KD did not reduce NRF2 protein levels, whereas HSP27 KD in the same cell line decreased NRF2 levels (Extended Data Fig. S4, A and B). HSP27 KD also decreased NRF2 expression in a KEAP1 mutant cancer cell line (33) suggesting that sHSP stabilization of NRF2 may be downstream of KEAP1 regulation (Fig. 6F). Because both HSP27 and KEAP1 bind to the Neh2 domain of NRF2 (Fig. 5B) (34), we investigated whether HSP27 disrupts the KEAP1–NRF2 interaction. However, recombinant HSP27 did not inhibit the interaction between KEAP1 and NRF2 as determined by NRF2 IP in a cell-free system (Fig. 6G). Taken together, these findings demonstrate that HSP27 stabilizes NRF2 by a KEAP1-independent mechanism.

Figure 6.

Figure 6

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HSP27 stabilizes NRF2.A, MDA-MB-231 cells were transiently transfected with control siRNAs or siRNAs targeting HSP27, and NRF2 expression was analyzed by IB 72 h later. The graph is shown as mean ± SD. B, RT–PCR analysis of NFE2L2 mRNA in MDA-MB-231 cells. NFE2L2 mRNA levels were normalized to GAPDH mRNA. The graph is shown as mean ± SD of n = 4 independent experiments. C, MDA-MB-231 cells transfected with control siRNAs or siRNAs targeting HSP27 were treated with or without 10 μM MG132 for the indicated time. NRF2 protein levels were determined by IB. D and E, HCT116 (D) or HS578 (E) cells were transiently transfected with control siRNAs or siRNAs targeting HSP27. About 68 h later, transfected cells were treated with vehicle or 100 μM tBHQ for an additional 4 h, and NRF2 expression was then analyzed by IB. The graph is shown as mean ± SD. F, A549 cells were transiently transfected with control siRNAs or siRNAs targeting HSP27, and NRF2 expression was analyzed by IB 72 h later. The graph is shown as mean ± SD. G, 0.5 μg of recombinant His-tagged NRF2 and 9 μg KEAP1 were incubated with the indicated concentrations of HSP27. NRF2 was pulled down, and the bound KEAP1 and HSP27 were analyzed by IB. For all panels, n = 3 independent experiments unless otherwise indicated. HSP27, heat shock protein 27; IB, immunoblotting; KEAP1, Kelch-like ECH-associated protein 1; NRF2, nuclear factor erythroid 2–related factor 2; tBHQ, tertiary butylhydroquinone.

PIPKIγ and HSP27 regulate HO-1 expression, ROS levels, and oxidative stress–induced cell death

These stepwise observations prompted us to investigate PIPKIγ-dependent HSP27 stabilization on NRF2 function. NRF2 regulates the expression of HO-1, a crucial player in cellular adaptation to oxidative stress (35). KD of PIPKIγ or HSP27 individually reduced basal and oxidative stress–induced HO-1 protein levels (Fig. 7, A and B and Extended Data Fig. S4C). In parallel, KD of PIPKIγ or HSP27 increased ROS levels in response to DEM treatment in multiple cell lines (Fig. 7, C and D). Furthermore, PIPKIγ and HSP27 KD enhanced oxidative stress–induced cell death, underscoring the therapeutic relevance of the NRF2–PtdIns(4,5)P2 pathway (Fig. 7, E and F). Collectively, our data reveal a previously unrecognized role of the NRF2–PtdIns(4,5)P2–sHSP complex in regulating NRF2 stability and function (Fig. 7G).

Figure 7.

Figure 7

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PIPKIγ and HSP27 regulate HO-1 expression, ROS levels, and oxidative stress–induced cell death.A, HCT116 cells expressing control shRNA or shRNAs targeting PIPKIγ were treated with 200 μM DEM for 4 h. The expression of PIPKIγ, HO-1, and NRF2 was analyzed by IB and quantified by ImageJ (Extended Data Fig. S4C). B, MDA-MB-231 cells were transiently transfected with control siRNAs or siRNAs targeting HSP27 for 72 h. Cells were then treated with vehicle or 200 μM DEM for an additional 4 h, and HO-1, NRF2, and HSP27 expression was analyzed by IB. C and D, ROS levels were determined in MDA-MB-231 and HCT116 cells transiently transfected with control siRNAs or siRNAs targeting HSP27 (C) or PIPKIγ (D) for 72 h. Cells were treated with vehicle or 200 μM DEM for the final 4 h before analyzing ROS levels. The graph is shown as mean ± SD. E and F, cell viability was determined by MTT assay in MDA-MB-231 and HCT116 cells transiently transfected with control siRNAs or siRNAs targeting HSP27 (E) or PIPKIγ (F) for 72 h. Cells were treated with vehicle or 200 μM DEM for the final 4 h before analyzing cell viability. The graph is shown as mean ± SD. G, a schematic model of NRF2 regulation by PIPKIγ, PtdIns(4,5)P2, and sHSPs. For all panels, n = 3 independent experiments. DEM, diethyl maleate; HO-1, heme oxygenase-1; HSP27, heat shock protein 27; IB, immunoblotting; MTT, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; NRF2, nuclear factor erythroid 2–related factor 2; PIPKIγ, phosphatidylinositol phosphate kinase γ; PtdIns(4,5)P2, phosphatidylinositol 4,5-bisphosphate; ROS, reactive oxygen species; sHSP, small heat shock protein.

Discussion

Numerous studies have demonstrated that NRF2 is destabilized under basal conditions via the activities of three E3 ubiquitin ligase complexes, KEAP1/Cullin-3, β-TrCP/SCF, and HRD1, which interact with different domains in NRF2 to trigger its destruction by the ubiquitin–proteasome system (6, 36, 37). In contrast, NRF2 is commonly hyperactivated in cancer by mutations in KEAP1 or the Neh2 domain in NRF2 that disrupt the KEAP1–NRF2 interaction and prevent negative regulation (5). Oncogenic K-Ras, B-Raf, and Myc also increase NRF2 gene expression and promote oxidative stress resistance and tumor growth (38, 39). In addition, hyperactivated PI3K–Akt signaling stabilizes NRF2 by Akt-dependent p21 induction, which disrupts KEAP1 binding to NRF2, and GSK-3 inhibition, which negatively regulates β-TrCP-mediated NRF2 degradation (5, 36, 40, 41).

Here, we report the discovery of a previously unknown regulator of NRF2, PtdIns(4,5)P2. PIPKIγ binds NRF2 directly, and the complex is detected in the nucleus. Then, PIPKIγ generates PtdIns(4,5)P2 that is stably coupled to NRF2 and is detectable by IB following SDS-PAGE similar to p53 (14, 15). Moreover, this approach is validated by [3H]myo-inositol labeling, which shows incorporation of the radiolabel into the NRF2 protein. The Neh2 domain of NRF2 also contains a binding site for PtdIns(4,5)P2. Of note, this noncovalent binding between NRF2 and PtdIns(4,5)P2 is distinct from the PIPKIγ-dependent stably associated PtdIns(4,5)P2 that is resistant to denaturing conditions. The generation of NRF2–PtdIns(4,5)P2 complexes is enhanced by oxidative stress, and the nuclear NRF2–PtdIns(4,5)P2 complex then recruits the sHSPs HSP27 and αB-crystallin, but not other molecular chaperones such as HSP40/60/90, indicating specificity. Although these sHSPs play critical roles in drug resistance, metastasis, and redox homeostasis (28, 29, 30, 42, 43, 44), the underlying mechanisms by which they act are poorly understood. Interestingly, HSP27, but not αB-crystallin, regulates the stability of NRF2 by a ubiquitin–proteasome pathway–dependent mechanism as well as the activity of NRF2, increasing expression of its transcriptional target HO-1 and counteracting oxidative stress. Like KEAP1 (34), HSP27 binds to the Neh2 domain of NRF2, but HSP27 has not been shown to disrupt the interaction between KEAP1 and NRF2. Moreover, HSP27 KD destabilizes NRF2 in KEAP1 mutant cancer cells, indicating that HSP27 stabilizes NRF2 by a KEAP1-independent mechanism.

The observed stabilization of NRF2 by a nuclear phosphoinositide complex is reminiscent of the recently reported regulation of nuclear p53 by phosphoinositides (14, 15). In both cases, a type I phosphoinositide 5-kinase binds to a nuclear protein and generates a PtdIns(4,5)P2 complex on the nuclear protein. Moreover, PtdIns(4,5)P2 binding to p53 and NRF2 acts as a signal to recruit sHSPs to the nuclear complex (14). These conserved features suggest that phosphoinositides may play a broad role in nuclear signaling by stress-regulated linkage to a subset of nuclear proteins, which serves as a signal to recruit sHSPs to phosphoinositide-bound proteins to regulate their stability and/or activity. However, there are some notable differences. p53 and NRF2 interact with different PIP kinase isoforms: p53 with PIPKIα (14) and NRF2 with PIPKIγ. In addition, while both sHSPs HSP27 and αB-crystallin stabilize nuclear p53, only HSP27 stabilizes NRF2. These differences indicate that the nuclear phosphoinositide pathway is modified for distinct protein targets or different cell types. Although both wildtype and mutant p53 bind NRF2 and regulate its transcriptional activity (45), it remains to be seen whether phosphoinositide linkage to either protein regulates their interaction. Moreover, it is unknown whether the NRF2–PtdIns(4,5)P2 complex is sequentially modified by multiple PIP kinases and phosphatases like p53 (15).

The discovery of this dual regulation of NRF2 by phosphoinositides and HSP27 and its role in regulating oxidative stress/cell death has profound therapeutic implications for cancer and other diseases characterized by aberrant NRF2 activation. These findings provide new mechanistic insights into the reported oncogenic and prometastatic actions of PIPKIγ (45, 46, 47, 48, 49, 50) and HSP27 (29, 44), neither of which has been previously linked to NRF2. These data also point to new strategies to destabilize NRF2 in cancer by inhibiting PIPKIγ, HSP27, or their respective interactions with NRF2. Moreover, NRF2 can be added to a growing list of PtdIns(4,5)P2 effectors, including p53 (14, 15) and Star-PAP (16), which contain stably associated phosphoinositides that regulate their activity, a fundamentally new signaling mechanism distinct from the canonical membrane–localized phosphoinositide second messenger pathway (17).

Experimental procedures

Antibodies and reagents

Antibodies against NRF2 (ab137550) were purchased from Abcam; β-Actin (#4967), HSP40 (#4868), HSP60 (#12165), HSP90 (#4877), Keap1 (#8047), HSP27 (D6W5V) (#95357), PIPKIα (#9693), PIPKIγ (#3296), PIPKIIα (#5527), PIPKIIβ (#9694), and DYKDDDDK Tag (#14793) were purchased from Cell Signaling Technology; HSP27 (ADI-SPA-801) and αB-crystallin (clone 1B6.1-3G4) from Enzo Lifesciences; HSP27 (sc-13132) and His-tag(sc-8036) from Santa Cruz Biotechnology; PtdIns(4,5)P2 (IgM) from Echelon Biosciences; and HO-1 (A303-662A) from Fortis Life Sciences. Goat anti-rabbit immunoglobulin G (IgG) (H + L) (code: 111-035-144) and donkey anti-mouse IgG (H + L) (code: 715-035-151) antibodies were purchased from Jackson ImmunoResearch Labs. Polyclonal and monoclonal antibodies against PIPKIα and PIPKIγ were produced as described previously (50). Diethyl maleate (D97703), tert-butylhydroquinone (#112941), LCS-1 (SML0466), and sulforaphane (#574215) and MG132 (#M7449) were obtained from Sigma–Aldrich. Cisplatin was purchased from Selleckchem and dissolved in water (0.3 mM working solution). DCFDA/H2DCFDA-Cellular ROS Assay Kit (ab113851) was purchased from Abcam; NRF2-FLAG plasmid (#36971) was purchased from Addgene; and PtdIns(4,5)P2 beads (#P-B045) were purchased from Echelon Biosciences. For recombinant protein production, His-tagged NRF2, HSP27, and αB-crystallin complementary DNAs were purchased from GenScript, and the recombinant proteins were purified as previously described (15). The full-length NRF2 construct (#OHU26812) and each of the NRF2 truncation mutant constructs Δ2–86 amino acids (#U617HEL120-4), Δ87 to 204 amino acids (#U617HEL120-5), Δ205 to 400 amino acids (#U617HEL120-6), and Δ401 to 605 amino acids (#U617HEL120-7) were purchased from GenScript. For the KD experiments, ON-TARGETplus siRNA SMARTpools with four siRNAs in combination against human PIPKIγ (L-004782-00-0050), HSP27 (L-005269-0050), and αB-crystallin (L-009743-00-0020) were purchased from Dharmacon. Nontargeting siRNA (#D-001810-01; Dharmacon) was used as a control. siRNAs were delivered to cells by RNAiMAX reagent (#13778150; Thermo Fisher Scientific), and the KD efficiency was determined by IB.

Cell culture, transfection, and lentivirus infection

Human embryonic kidney HEK293FT cells (#CRL-3216) and human cancer cell lines A549 (#CCL-185), BT-549 (#HTB-122), HCT116 (#CCL-247), HS578T (#HTB-126), MDA-MB-231 (#HTB-26), and MDA-MB-468 (#HTB-132) were purchased from American Type Culture Collection. MDA-MB-231 cells with deletion of PIPKIα, PIPKIγ, PIPKIIα, or PIPKIIβ were previously described (14). Cells were maintained in Dulbecco's modified Eagle's medium (#10-013-CV; Corning) supplemented with 10% fetal bovine serum (#SH30910.03; Hyclone) and 1% penicillin/streptomycin (#15140-122; Gibco) at 37 °C in a humidified 5% CO2 incubator. Cells were periodically tested for mycoplasma contamination, and mycoplasma-negative cells were used for all analyses. NRF2 plasmid (##36971) from Addgene was transiently transfected into HEK293FT cells using Lipofectamine 2000 (#11668019) from ThermoFisher. The shRNA sequences for PIPKIγ (RHS3979-201766531, RHS3979-201766535) and the shRNA control (pLKO.1, RHS4080) were obtained from Dharmacon. A lentiviral expression system from ThermoFisher was used to generate target virus supernatants that were used to infect MDA-MB-231 and HCT116 cells. After 72 h, puromycin was used to select resistant cells.

IP and IB

IP and IB were performed, and bands were quantitated as previously described (14).

Fluorescent IP–WB

Fluorescent IP–WB was performed as previously described (15) with some modifications. Briefly, cells were lysed in a buffer containing 1% Brij58, 150 mM NaCl, 20 mM Hepes, pH 7.4, 2 mM MgCl2, 2 mM CaCl2, 1 mM Na3VO4, 1 mM Na2MoO4, and protease inhibitors. For endogenous NRF2 IP, 0.5 to 1 mg of cell lysate was incubated with 20 μl anti-NRF2 (#sc-365949 AC; Santa Cruz Biotechnology). Normal IgG-conjugated agarose was used as a negative control (#sc-2343; Santa Cruz Biotechnology). After washing five times with lysis buffer, the protein complex was eluted with SDS sample buffer. The sample was then heated at 98 °C for 5 min. For double fluorescent IP–WB to detect NRF2–PI4,5P2 complexes, anti-pNRF2 rabbit monoclonal IgG at 1:2000 dilution and anti-PI4,5P2 mouse monoclonal IgM antibody (#Z-P04; Echelon Biosciences at 1:2000 dilution) was mixed in blocking buffer and incubated with the membrane at 4 °C overnight. For the secondary antibody incubation, goat anti-rabbit IgG antibody conjugated with IRDye 800CW fluorophore (#926-32211; LI-COR) and goat anti-mouse IgM antibody conjugated with IRDye 680RD fluorophore (#926-68180; LI-COR) at 1:10,000 dilution was mixed in blocking buffer with 0.01% SDS and 0.1% Tween-20 and incubated with the membrane at room temperature for 2 h. Next, the membrane was washed with PBS with Tween-20 (PBST) three times (10 min for each wash). Images were captured on the Odyssey Fc Imaging System (LI-COR Biosciences) using the 700 and 800 nm wavelength channels simultaneously. The NRF2-associated PI4,5P2 complexes were visualized by overlapping the 700 and 800 nm channels.

In vitro binding assay

His-tagged NRF2, HSP27, αB-crystallin proteins were expressed in Escherichia coli and purified with Ni–NTA–agarose as described (14). Recombinant KEAP1 protein (#11981-H20B) was purchased from Sino Biological. The binding assay was performed in PBST by incubating a constant amount of His-tagged NRF2 with an increasing amount of HSP27 or αB-crystallin in the presence of 20 μl anti-NRF2 antibody–conjugated agarose (#sc-365949 AC; Santa Cruz) at 4 °C overnight. The unbound proteins were then removed by washing with PBST three times, and the protein complex was subsequently eluted using SDS sample buffer. The sample was then heated to 98 °C for 5 min and analyzed by IB. Images were captured using the Chemi channel on the Odyssey Fc Imaging System (LI-COR Biosciences).

MST assay

MST was performed as described previously (15). The synthetic proteinase inhibitors for MST, including PtdIns diC16 (#P-0016), PtdIns(4,5)P2 diC16 (#P-4516), PtdIns(3,4,5)P3 diC16 (#P-3916), PtdIns4P diC16 (#P-4016), and PtdIns5P diC16 (#P-5016), were purchased from Echelon Biosciences and prepared as described (15).

Immunofluorescence

For IF studies, cells were grown on coverslips coated with 0.2% gelatin (#G9391; MilliporeSigma), fixed with 4% paraformaldehyde (#sc-281692; Santa Cruz Biotechnology) for 30 min at room temperature, permeabilized with 0.1% Triton X-100 for 10 min, and blocked with 3% bovine serum albumin in PBS for 1 h at room temperature. Next, cells were incubated with primary antibody at 4 °C overnight, then washed three times with PBS, and incubated with fluorescent-conjugated secondary antibodies for 1 h at room temperature. Cells were then washed three times with PBS and mounted in Prolong Glass Antifade Mountant with NucBlue Stain (#P36985; Thermo Fisher Scientific). Images were acquired using the 100× objective lens (numerical aperture 1.4 oil) of a Leica SP8 confocal microscope controlled by LASX software (Leica Microsystems). The mean fluorescent intensity of channels in each cell measured by LASX was used for quantification. Colocalization was determined by ImageJ (NIH), and Pearson’s coefficient was used to quantify the degree of colocalization.

Lipid bead-protein pull-down

Lipid Bead-Protein pull-down was performed according to the manufacturer’s instructions with modifications. Briefly, control beads (P-B000; Echelon Biosciences, Inc) and PtdIns(4,5)P2 beads (#P-B045a; Echelon Biosciences, Inc) were pelleted by centrifugation at 1000g or lower. The supernatant was then removed, and the beads were resuspended in an equal volume of binding buffer. The protein (10 μg) was diluted in binding buffer and added to 50 to 100 μl beads. The protein-bead solution was then incubated overnight at 4 °C with continuous motion. The beads were pelleted, the supernatant removed, and the beads were then washed five times with 10× excess of the wash/binding buffer. Next, the bound proteins were eluted by adding an equal volume of 2× Laemmli sample buffer to the beads and then heating them to 98 °C for 5 min. After heating, the beads were pelleted, and the supernatant was removed and stored at 4 °C until analysis.

Proximity ligation assay

Cells were grown on coverslips coated with 0.2% gelatin. After fixation and permeabilization, cells were blocked before incubation with primary antibodies as described in IF. The cells were subsequently processed using the PLA kit (#DUO92101; MilliporeSigma) following the manufacturer’s instructions. The slides were then mounted with Duolink In Situ Mounting Medium with 4',6-diamidino-2-phenylindole (#DUO82040; MilliporeSigma). Punctate PLA foci were detected using a Leica SP8 confocal microscope and quantified by ImageJ analysis.

3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide assay

Around 5 × 104 cells/well in 96-well plates were transfected with control siRNAs or siRNAs targeting PIPKIγ or HSP27 for 48 h. Next, the cells were treated with vehicle or 200 μM DEM for 24 h and then incubated with 100 μl of fresh medium and 10 μl of the 12 mM 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide stock solution from the Vybrant MTT cell proliferation assay kit (#V13154; Thermo Fisher Scientific). After 4 h, all media were removed, 100 μl of dimethyl sulfoxide was added, and the mixture was incubated at 37 °C for 10 min. Absorbance was then measured at 540 nm using a Synergy H1 Hybrid MultiMode Microplate reader (BioTek Instruments).

ROS measurement

In 96-well plates, 4 × 104 cells/well were transfected with control or siRNAs targeting PIPKIγ or HSP27 for 48 h. The cells were then treated with the vehicle or 200 μM DEM for 4 h. The media were removed, and 100 μl/well of 1× buffer was added. After the 1× buffer was removed, the cells were incubated with 100 μl/well of the diluted DCFDA solution for 45 min at 37 °C in the dark. The DCFDA solution was then removed, and 100 μl/well of 1× buffer was added. Fluorescence was measured on a Synergy H1 Hybrid Multimode Microplate reader (BioTek Instruments) at excitation/emission = 485/535 nm in endpoint mode in the presence of a buffer.

[3H]Myo-inositol metabolic labeling

MDA-MB-231 cells were split and cultured in low serum Opti-MEM (#31985070; Thermo) supplemented with 10% dialyzed fetal bovine serum with a 10,000-molecular weight cutoff (#F0392; Sigma) and 1% penicillin–streptomycin (#15140-122; Gibco). Cells were seeded to achieve ∼5% confluency after 24 h after which either 1.322 μM (25 μCi/ml) of [3H]myo-inositol (#NET1156005MC; PerkinElmer) or unlabeled myo-inositol (#J60828.22; Thermo) were added to the media. Cells were incubated with the label for 72 h so the [3H]myo-inositol was incorporated into the cellular metabolism. Cells were treated with control vehicle or 200 μM DEM for another 24 h. Cells were then lysed, and NRF2 was immunoprecipitated before performing SDS-PAGE. SDS-PAGE samples corresponding with the NRF2 immunoblot were excised from the gel and dissolved in 30% H2O2 before liquid scintillation counting (LSC). Dissolved gel samples were added to LSC vials with LSC cocktail (#6013319; PerkinElmer) before being processed by PerkinElmer Tri-Carb 4910 TR liquid scintillation analyzer. Analysis and DPM calculation were automated using QuantaSmart (PerkinElmer) software.

Statistics and reproducibility

Two-sided paired t tests were used to determine pair-wise significance unless indicated otherwise. Although no power calculations were performed, sample sizes were determined based on previously published experiments that observed significant differences between groups. At least three independent experiments or biologically independent samples were used for statistical analysis, and the number of repeats is indicated in the figure legends.

Data availability

All data supporting the findings of this study are available from the corresponding authors on reasonable request.

Supporting information

This article contains supporting information.

Conflict of interest

The authors declare that they have no conflicts of interest with the contents of this article.

Acknowledgments

Author contributions

C. C., M. C., T. W., R. A. A., and V. L. C. conceptualization; C. C., N. D. C., M. C., T. W., P.A., R. A. A., and V. L. C. methodology; C. C., N. D. C., M. C., T. W., and P. A. formal analysis; C. C., M. C., T. W., and P. A. investigation; C. C. and N. D. C. writing–original draft; N. D. C., T. W., P. A., R. A. A., and V. L. C. writing–review & editing; N. D. C. visualization; R. A. A. and V. L. C. supervision; R. A. A. and V. L. C. funding acquisition.

Funding and additional information

We thank Anissa Maravilla and other members of the Cryns and Anderson labs for discussions and comments. This work was supported in part by a National Institutes of Health grant R35GM134955 (to R. A. A.), Department of Defense Breast Cancer Research Program grants W81XWH-17-1-0258 (to R. A. A.), W81XWH-17-1-0259 (to V. L. C.), and W81XWH-21-1-0129 (to V. L. C.), and a grant from the Breast Cancer Research Foundation (to V. L. C.). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Reviewed by members of the JBC Editorial Board. Edited by Alex Toker

Contributor Information

Richard A. Anderson, Email: raanders@wisc.edu.

Vincent L. Cryns, Email: vlcryns@medicine.wisc.edu.

Supporting information

Supportng figures
mmc1.pdf (607.6KB, pdf)

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Associated Data

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Supplementary Materials

Supportng figures
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Data Availability Statement

All data supporting the findings of this study are available from the corresponding authors on reasonable request.

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