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. 2026 Sep 25;5(6):100369. doi: 10.1016/j.cellin.2026.100369

PLIN2 links lipid droplets to GRP78-mediated GPX4 stabilization to suppress neuronal ferroptosis

Qianruo Wang a,b,1, Baokun Sui a,d,1, Jiaxin Zheng a, Chuanyin Yao a, Jianqing Zhao a, Zhen F Fu a, Ling Zhao a,c,d,⁎, Ming Zhou a,d,⁎⁎
PMCID: PMC13734872  PMID: 42990892

Abstract

Neuronal ferroptosis is increasingly recognized as a major driver of neuroinflammation, yet how lipid droplets coordinate neuronal antioxidant defense remains largely unknown. Here, we identify a previously unrecognized PLIN2–GRP78 pathway that links lipid droplets to the ferroptosis regulator GPX4. Mechanistically, the lipid droplet protein PLIN2 directly interacts with GRP78, promoting its expression and translocation from the endoplasmic reticulum (ER) to the cytosol. Cytosolic GRP78 subsequently associates with GPX4, preventing its K48-linked ubiquitination and proteasomal degradation, thereby preserving GPX4 stability and limiting lipid peroxidation, reactive oxygen species accumulation, and ferroptotic cell death. Furthermore, overexpression of PLIN2 failed to restore GPX4 expression or alleviate ferroptosis in GRP78-deficient neurons, indicating that GRP78 is required for the anti-ferroptotic function of PLIN2 and that PLIN2 regulates ferroptosis through the GRP78–GPX4 axis. Importantly, neuron-specific deletion of GRP78 markedly exacerbates ferroptosis, microglial activation, inflammatory cytokine production, and mortality in both lipopolysaccharide (LPS)-induced and virus-induced neuroinflammation, whereas pharmacological inhibition of ferroptosis largely reverses these pathological changes. Together, our findings establish a mechanistic link between lipid droplets and the GPX4 antioxidant system through PLIN2-dependent regulation of GRP78, revealing an unreported pathway controlling neuronal ferroptosis and neuroinflammation and highlighting GRP78-mediated GPX4 stabilization as a potential therapeutic strategy for neuroinflammatory diseases.

Keywords: PLIN2-GRP78 axis, GPX4 K48-ubiquitination, Neuronal ferroptosis, Neuroinflammation, Lipid droplet signaling

Graphical abstract

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Highlights

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    PLIN2-GRP78 stabilizes GPX4 via K48-deubiquitination.

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    LDs govern ferroptosis through protein-protein interaction.

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    Ferroptosis checkpoint links lipid signaling to neuroinflammation.

1. Introduction

During encephalitis, intense neuroinflammation profoundly reshapes neuronal lipid metabolism, disturbing lipid turnover and intracellular metabolic balance, thereby promoting neuronal dysfunction and cellular injury (Hasel et al., 2021). In this pathological context, lipid droplets (LDs) are increasingly recognized as active regulators of cellular metabolism rather than inert lipid storage structures, with emerging roles in controlling oxidative stress and cell fate through mechanisms that remain incompletely understood (Mathiowetz & Olzmann, 2024; Olzmann & Carvalho, 2019). At the structural level, LDs contain a neutral lipid-rich core enclosed by a phospholipid monolayer, on which perilipin 2 (PLIN2), a predominant surface protein, regulates droplet integrity and facilitates the association of functional proteins (Carr & Ahima, 2016; Roberts et al., 2023; Wu et al., 2022). While PLIN2-dependent LD biogenesis is known to influence cellular stress responses, its role in neuronal survival remains elusive.

Ferroptosis is characterized by iron-dependent accumulation of lipid peroxides and has been implicated as an important contributor to neuronal injury during encephalitis and neurodegenerative disorders (Calhoon et al., 2025; Jiang et al., 2021). Glutathione peroxidase 4 (GPX4), the gatekeeper of ferroptosis, directly neutralizes lipid peroxides to maintain membrane integrity (Bersuker et al., 2019; Zhou et al., 2025). Intriguingly, LDs have been proposed to counter ferroptosis by sequestering free iron (Lee et al., 2024; Sassano et al., 2025) and buffering oxidants (Tan et al., 2023), while their lipid composition and surface proteins dynamically regulate GPX4 activity (Chen et al., 2021; Ma et al., 2021). However, the molecular mechanisms linking LD biology to GPX4-dependent ferroptosis suppression in neurons, a cell type exquisitely vulnerable to lipid peroxidation, remain undefined.

GRP78 (HSPA5/BiP), an endoplasmic reticulum (ER)-resident molecular chaperone, orchestrates protein folding and ER stress responses (Akinyemi et al., 2023; Graewert et al., 2024). Strikingly, recent studies reveal GRP78 translocates to LD surfaces, where it interacts with LD membrane proteins to modulate LD morphology and lipid homeostasis (Tian et al., 2023). LDs regulate GRP78 expression through PLIN2-mediated mechanisms, thereby linking LD homeostasis with ER stress-associated ferroptosis resistance in neurons (Zhang et al., 2024). Beyond ER function, GRP78 deficiency exacerbates ferroptosis by disrupting intracellular iron homeostasis (Chen et al., 2019; Li et al., 2023), yet whether GRP78 bridges LDs to GPX4-dependent ferroptosis suppression in neurons remains unknown.

Here, we identify a PLIN2-GRP78-GPX4 signaling axis that enables LDs to suppress neuronal ferroptosis by stabilizing GPX4. This pathway not only unravels a novel mechanism of LD-mediated neuroprotection but also highlights potential therapeutic targets for ferroptosis-driven neuroinflammatory diseases.

2. Results

2.1. PLIN2-dependent lipid droplet biogenesis suppresses neuronal ferroptosis

Our prior work demonstrated that neurotropic viruses (e.g., RABV, JEV, EMCV) exploit LD-mediated suppression of neuronal ferroptosis to enhance replication (Wang et al., 2024). However, whether LDs intrinsically regulate neuronal ferroptosis (independent of viral infection) and thereby influence neuroinflammation remains unclear. To address this, N2a cells were treated with 100 μM oleic acid (OA; 4 h) to promote LD formation (DMSO as control), followed by treatment with 1 μM RSL3 (2 h). Electron microscopy revealed RSL3-induced mitochondrial damage (membrane rupture/shrinkage, red arrows), which was mitigated by OA (Fig. 1(A)). Consistently, OA reversed RSL3-induced reductions in cell viability (CCK-8 assay; Fig. 1(B)) and suppressed ferroptosis markers: lipid peroxidation, MDA, and ROS (Fig. 1(C–E)). These data confirm that OA-induced LD formation inhibits neuronal ferroptosis.

Fig. 1.

Fig. 1

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Lipid droplet protein PLIN2 mediates ferroptosis resistance in neuronal cells. (A) Transmission electron microscopy (TEM) of N2a cells treated with DMSO (control), 1 μM RSL3 (2 h), or RSL3 + 100 μM oleic acid (OA, 4 h). Red arrows indicate mitochondrial damage (scale bar: 500 nm). (B) Cell viability (CCK-8 assay), (C) lipid peroxidation (C11-BODIPY), (D) MDA levels, and (E) ROS (DCFH-DA) in N2a cells after treatments in (A). (F) Western blot of PLIN2-His and Actin in N2a cells transfected with PLIN2-His or empty vector (EV). (G–H) BODIPY 493/503 staining (green) and DAPI (blue) of OA-treated N2a cells overexpressing PLIN2-His or EV (G, scale bar: 10 μm); quantification of LD fluorescence intensity (H). (I) Western blot of PLIN2 and Actin in N2a cells transfected with PLIN2 siRNAs (siPLIN2-1/2/3) or siNC. (J–K) BODIPY 493/503 staining of OA-treated N2a cells transfected with siPLIN2 or siNC (J, scale bar: 10 μm); quantification of LD fluorescence intensity (K). (L–O) Cell viability (L), lipid peroxidation (M), MDA (N), and ROS (O) in RSL3-treated N2a cells transfected with siPLIN2 or siNC. Data are mean ± SEM (n = 5). ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.000 1.

PLIN2, a key LD surface protein, regulates LD biogenesis and stability by preventing fusion/degradation (Wu et al., 2022). Our previous work identified PLIN2 as a mediator of LD formation in virus-infected N2a cells (Wang et al., 2024), prompting us to hypothesize that PLIN2 links LD formation to ferroptosis suppression. To test this, we generated His-tagged PLIN2-overexpressing N2a cells (validated by Western blot; Fig. 1(F)). PLIN2 overexpression significantly enhanced OA-induced LD formation compared to empty vector controls (Fig. 1(G–H)) and reduced ferroptosis in RSL3-treated cells, as evidenced by increased viability and decreased lipid peroxidation, MDA, and ROS (Fig. S1). Conversely, PLIN2 knockdown (three siRNAs, validated by Western blot; Fig. 1(I)) reduced OA-induced LD formation (Fig. 1(J–K)), decreased viability (Fig. 1(L)), and elevated ferroptosis markers (lipid peroxidation, MDA, ROS) in RSL3-treated cells (Fig. 1(M–O)). Together, these results demonstrate that PLIN2-dependent LD formation suppresses neuronal ferroptosis.

2.2. PLIN2 directly binds GRP78 via its C-terminal domain to promote GRP78 expression and ER-to-cytosol translocation

PLIN2 has been shown to promote LD biogenesis and suppress neuronal ferroptosis, yet the mechanistic link between PLIN2 and ferroptosis inhibition remains undefined. To address this, we performed RNA-seq analysis of N2a cells overexpressing PLIN2-His (vs empty vector) and identified GRP78 (HSPA5) as the most significantly upregulated gene (Fig. 2(A)). qRT-PCR validated elevated GRP78 mRNA levels in PLIN2-overexpressing N2a cells (Fig. S2A), with conserved upregulation of human GRP78 (hGRP78) observed in SK-N-SH cells overexpressing human PLIN2 (hPLIN2-His; Fig. S2B). Western blot further confirmed PLIN2-dependent upregulation of GRP78 protein in both murine N2a and human SK-N-SH cells (Fig. S3A–B).

Fig. 2.

Fig. 2

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PLIN2 interacts with GRP78 via C-terminal domain to promote GRP78 expression and ER-to-Cytosol translocation. (A) RNA-seq heatmap of differentially expressed genes in N2a cells transfected with PLIN2-His vs EV (n = 4; adjusted P < 0.05). (B) Semi-endogenous Co-IP: N2a cells transfected with PLIN2-His or GRP78-Flag were immunoprecipitated with anti-His or anti-Flag beads, followed by immunoblotting for PLIN2, GRP78, His, Flag, and Actin. (C) Cytosolic and ER fractions were isolated, and GRP78 levels were analyzed by Western blot. (D) Coomassie blue staining of purified GST-GRP78 protein. (E) GST pull-down: N2a cell lysates (PLIN2-His transfected) incubated with GST-GRP78 or GST beads, immunoblotted for His, GST, and Actin. (F) Schematic diagrams of full-length PLIN2 and its truncation constructs: full-length PLIN2 (1–426 aa) and truncation constructs (1–393 aa, 1–135 aa, 136–209 aa, and 210–393 aa). (G) Co-IP of PLIN2 truncations: N2a cells transfected with His-tagged PLIN2 truncations were immunoprecipitated with anti-His beads, immunoblotted for GRP78, His, and GAPDH. (H) Schematic of GRP78 truncation constructs (NBD: nucleotide-binding domain; NSD: negative regulatory domain; SBD: substrate-binding domain). (I) Co-IP of GRP78 truncations: N2a cells transfected with Flag-tagged GRP78 truncations were immunoprecipitated with anti-Flag beads, immunoblotted for PLIN2, Flag, and GAPDH.

To dissect how PLIN2 regulates GRP78, semi-endogenous co-immunoprecipitation (Co-IP) assays revealed a physical interaction between PLIN2 and endogenous GRP78, with PLIN2 overexpression correlating with increased GRP78 protein levels (Fig. 2(B) and S4). To investigate the regulatory mechanism of PLIN2 on GRP78, cytosolic and ER fractions were isolated, followed by assessment of GRP78 protein levels. The results showed that overexpression of PLIN2 promoted the translocation of GRP78 from the ER to the cytosol (Fig. 2(C)). Additionally, to further validate the alteration of GRP78 localization induced by PLIN2, we performed immunofluorescence (IF) staining combined with confocal microscopy analysis. The results demonstrated that, under basal conditions, GRP78 was predominantly colocalized with the ER marker, indicating its canonical ER localization. However, following PLIN2 overexpression, GRP78 exhibited reduced colocalization with the ER marker (Fig. S5).

To confirm direct binding between PLIN2 and GRP78, we performed GST pull-down assays using purified GST-GRP78 protein (Fig. 2(D) and (E)). The results confirmed a direct interaction between PLIN2 and GRP78, ruling out the involvement of intermediate proteins. Next, we mapped the critical interaction domains using truncated constructs. PLIN2 truncations (1–393 aa, 210–393 aa, 1–135 aa, 136–209 aa; Fig. 2(F) and (G)) were subjected to semi-endogenous Co-IP, revealing that only the C-terminal 210–393 aa region of PLIN2 interacts with endogenous GRP78 and upregulates its expression (Fig. 2(G)). Conversely, GRP78 was divided into functional domains (NBD, NSD, SBD; Fig. 2(H)), and Co-IP assays identified the NBD domain as the sole region mediating interaction with endogenous PLIN2 (Fig. 2(I)).

Given that GRP78 is a canonical target of the unfolded protein response (UPR), we next investigated whether PLIN2-mediated GRP78 upregulation was associated with ER stress signaling. We therefore examined the activation status of the PERK–eIF2α–ATF4 pathway in PLIN2-overexpressing N2a cells. PLIN2 overexpression significantly increased the phosphorylation of PERK and eIF2α and elevated ATF4 protein levels (Fig. S6), indicating activation of the PERK–eIF2α–ATF4 branch of the UPR. These findings suggest that PLIN2-induced GRP78 upregulation is associated with activation of ER stress signaling and may occur, at least in part, through the PERK–eIF2α–ATF4-mediated UPR pathway. Thus, rather than directly regulating Grp78 transcription, PLIN2 may promote GRP78 expression indirectly through modulation of ER homeostasis.

Collectively, these data demonstrate that PLIN2 directly interacts with GRP78 via its 210–393 aa region and GRP78's NBD domain, leading to increased GRP78 expression and ER-to-cytosol translocation.

2.3. The PLIN2-GRP78-GPX4 axis suppresses neuronal ferroptosis via GRP78-SBD (137-178 aa)-mediated GPX4 upregulation

We previously demonstrated that PLIN2 inhibits neuronal ferroptosis and directly interacts with GRP78 to upregulate its expression. However, whether PLIN2's anti-ferroptotic effect is intrinsic or mediated through GRP78 remains unclear. Glutathione peroxidase 4 (GPX4), a master regulator of ferroptosis, controls lipid peroxidation via glutathione-dependent antioxidant activity (Liang et al., 2023). Notably, GRP78 indirectly modulates GPX4 expression/function through signaling pathways, thereby regulating oxidative stress responses (Li et al., 2023). We thus hypothesized that GRP78 interacts with GPX4 to suppress neuronal ferroptosis.

To validate this, we transfected N2a cells with GRP78-Flag or GPX4-Myc (empty vector as control) and performed Co-IP assays. GRP78-Flag co-immunoprecipitated with endogenous GPX4 and increased GPX4 expression, whereas GPX4-Myc interacted with endogenous GRP78 without affecting GRP78 levels (Fig. 3(A)).

Fig. 3.

Fig. 3

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GRP78 stabilizes GPX4 via SBD (137–178 aa) to inhibit neuronal ferroptosis. (A) Semi-endogenous Co-IP: N2a cells transfected with GRP78-Flag or GPX4-Myc were immunoprecipitated with anti-Flag or anti-Myc beads, immunoblotted for GRP78, GPX4, Flag, Myc, and Actin. (B) Co-IP of GRP78 domain truncations: N2a cells transfected with Flag-tagged GRP78 (FL/NBD/NSD/SBD) were immunoprecipitated with anti-Flag beads, immunoblotted for GPX4, Flag, and Actin. (C) AlphaFold3 prediction of GRP78-SBD (137–218 aa) and GPX4 interaction interface. (D) Co-IP of GRP78-SBD subdomains: N2a cells transfected with Flag-tagged GRP78 (137–178 aa or 179–218 aa) were immunoprecipitated with anti-Flag beads, immunoblotted for GPX4, Flag, and Actin. (E) Coomassie blue staining of purified GST-PLIN2 protein. (F) GST pull-down: N2a cell lysates (GPX4-Myc transfected) incubated with GST-PLIN2 or GST beads, immunoblotted for Myc, GST, and Actin. (G–J) N2a cells were transfected with EV, GRP78-Flag, or GRP78-SBD (137–178)-Flag then treatment with RSL3 (1 μM, 2 h). (G) Cell viability, (H) lipid peroxidation, (I) MDA levels, and (J) ROS production were measured 48 h post-transfection. Data are mean ± SEM (n = 5). ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.000 1.

Next, we mapped the interaction domains using GRP78 truncations (NBD/NSD/SBD-Flag). Semi-endogenous Co-IP showed that only the SBD domain of GRP78 interacted with endogenous GPX4 and upregulated its expression (Fig. 3(B)). AlphaFold3 modeling predicted GPX4 binds the 137–218 aa region of GRP78-SBD (Fig. 3(C)). We generated SBD subfragments (137–178 aa and 179–218 aa) and confirmed via Co-IP that the 137–178 aa segment mediates GPX4 interaction and upregulation (Fig. 3(D)).

Given PLIN2-GRP78 binding, we investigated whether PLIN2 directly regulates GPX4. Semi-endogenous Co-IP initially suggested PLIN2 interacts with GPX4 (Fig. S7), but GST pull-down with purified GST-PLIN2 showed no direct binding (Fig. 3(E) and (F)), indicating the PLIN2-GPX4 interaction is GRP78-dependent.

To confirm GRP78-GPX4 binding suppresses ferroptosis, we transfected full-length GRP78 or GRP78-SBD (137–178 aa) into RSL3-treated N2a cells. Both constructs increased cell viability and reduced lipid peroxidation, MDA, and ROS levels (Fig. 3(G–J)).

Collectively, these data reveal a hierarchical pathway: PLIN2 binds GRP78 to enhance its interaction with GPX4 via GRP78-SBD (137–178 aa), thereby upregulating GPX4 and suppressing neuronal ferroptosis.

2.4. GRP78 stabilizes GPX4 by inhibiting K48-linked ubiquitination at lysine 47 to suppress neuronal ferroptosis

To investigate how GRP78 upregulates GPX4 protein levels, we first measured GPX4 mRNA abundance following GRP78 overexpression. No significant increase was observed (Fig. S8A), indicating GRP78 does not transcriptionally regulate GPX4. This was confirmed in primary neurons from neuron-specific GRP78 knockout C57BL/6J mice, where GPX4 mRNA levels remained unchanged (Fig. S8B). CHX chase assays revealed that GRP78 overexpression significantly prolonged GPX4 half-life in N2a cells (Fig. 4(A)), suggesting post-translational stabilization via reduced degradation.

Fig. 4.

Fig. 4

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GRP78 inhibits K48-linked Ubiquitination of GPX4 at Lysine 47 to prevent ferroptosis. (A) Cycloheximide (CHX) chase assay: Western blot of GPX4, Flag, and Actin in N2a cells transfected with GRP78-Flag or EV (CHX: 100 μg/mL, 0–24 h). (B) Western blot of GPX4 in GRP78-knockdown (siGRP78) N2a cells treated with MG132 (10 μM, proteasome inhibitor), chloroquine (CQ, 10 μM, lysosome inhibitor), or DMSO. (C–E) Ubiquitination assay: N2a cells co-transfected with GPX4-Myc, HA-Ub (C), HA-Ub (K48) (D), or HA-Ub(K63) (E), plus GRP78-Flag, GRP78-SBD (137–178)-Flag, or EV. Co-IP with anti-Myc beads followed by immunoblotting for HA, Myc, Flag, and Actin (MG132: 10 μM, 4 h). (F) Ubiquitination assay of GPX4 mutants: N2a cells co-transfected with HA-Ub(K48), GPX4-Myc (WT/K47R/K58R), and GRP78-Flag. Co-IP with anti-Myc beads and immunoblotting as in (C). (G–H) Western blot of GPX4, PLIN2-His, GRP78-Flag, and Actin in N2a cells transfected with (G) siGRP78 + PLIN2-His or (H) siPLIN2 + GRP78-Flag. (I–L) Cell viability (I), lipid peroxidation (J), MDA (K), and ROS (L) in RSL3-treated N2a cells transfected with siGRP78, siGRP78+PLIN2-His, siPLIN2, or siPLIN2+GRP78-SBD (137-178aa)-Flag. Data are mean ± SEM (n = 5). ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.000 1.

To define the degradation pathway, GRP78 was knocked down in N2a cells using three specific siRNAs, followed by treatment with the proteasome inhibitor MG132 or lysosomal inhibitor chloroquine (CQ). Western blot showed GPX4 levels were fully restored by MG132 (but not CQ) in GRP78-knockdown cells (Fig. 4(B)), confirming GRP78 stabilizes GPX4 through the proteasomal pathway.

Since ubiquitination is the primary proteasomal degradation signal, we assessed whether GRP78 modulates GPX4 ubiquitination. Co-IP assays in N2a cells co-transfected with GPX4-Myc and full-length GRP78-Flag or GRP78-SBD (137–178 aa)-Flag (plus MG132) demonstrated that both GRP78 constructs reduced GPX4 ubiquitination (Fig. 4(C)). Further analysis revealed GRP78 specifically suppressed K48-linked (but not K63-linked) polyubiquitination of GPX4 (Fig. 4(D–E)).

To map the critical ubiquitination site, we generated GPX4 mutants (K47R and K58R) and performed Co-IP with K48-ubiquitin plasmids. GRP78 failed to reduce K48-ubiquitination of GPX4-K47R, whereas K58R had no effect (Fig. 4(F)), identifying GPX4-K47 as the target lysine residue.

To clarify the functional hierarchy between PLIN2 and GRP78, we conducted rescue experiments. Firstly, the GRP78 knockdown: PLIN2 overexpression did not restore GPX4 levels (Fig. 4(G) and (H)) or reverse ferroptosis markers (reduced viability, elevated lipid peroxidation/MDA/ROS) induced by GRP78 depletion (Fig. 4(I–L)). Secondly, the PLIN2 knockdown: GRP78 overexpression rescued GPX4 expression (Fig. 4(H)) and ferroptosis phenotypes (Fig. 4(I–L)), with GRP78-SBD (137–178 aa) alone sufficient to reverse PLIN2 depletion-induced ferroptosis.

Collectively, these data demonstrate that GRP78 directly inhibits K48-linked ubiquitination of GPX4 at lysine 47, stabilizing GPX4 protein to suppress neuronal ferroptosis, with PLIN2 acting upstream to promote GRP78 function rather than directly regulating GPX4.

2.5. Neuronal GRP78 deficiency exacerbates ferroptosis and neuronal injury in LPS-challenged and virus-infected mice

We previously demonstrated that LPS induces ferroptosis in mouse neuronal cells (Wang et al., 2025). To determine whether the PLIN2–GRP78 pathway is endogenously regulated under neuroinflammatory conditions, we examined the dynamic expression profiles of PLIN2 and GRP78 in vivo. Wild-type C57BL/6J mice were intraperitoneally challenged with LPS, and brain tissues were collected at 0, 2, 6, 12, 24, and 48 h after treatment for detecting PLIN2 and GRP78 expression by Western blot analysis. Notably, the protein levels of PLIN2 and GRP78 progressively increased and reached their highest levels at 12 h post treatment, followed by a relative decline at 24 and 48 h post treatment (Fig. S9A). These results indicate that both PLIN2 and GRP78 are dynamically induced during acute neuroinflammatory responses. We further examined the expression profiles of PLIN2 and GRP78 in virus-induced encephalitis models. Wild-type C57BL/6J mice were infected with HSV-1 or EV71, and brain tissues were collected at the indicated time points after infection for detecting PLIN2 and GRP78 expression by Western blot analysis. In contrast to the transient induction observed following LPS challenge, both HSV-1 and EV71 infection induced a progressive increase in PLIN2 and GRP78 protein levels. The expression of both proteins became apparent at 24 h post-infection and reached the highest levels at 48 h post-infection (Fig. S9B and Fig. S9C). Together, these results demonstrate that the PLIN2–GRP78 pathway is endogenously and dynamically activated under both inflammatory and viral encephalitic conditions, supporting its physiological relevance during neuroinflammatory stress.

To directly assess GRP78's in vivo function, the neuronal GRP78 knockout mice were introduced, and the efficiency and specificity of neuronal GRP78 knockout were initially determined, and the knockout efficiency reached 80%-85% in neurons, with no knockout in microglia (Fig. S10A and B). We next studied GRP78 function in ferroptosis in vivo. Neuron-specific GRP78 knockout (Grp78−/−) markedly reduced brain GRP78 protein (Fig. S10C). In LPS-challenged mice, Grp78−/− brains showed lower GPX4 protein than wild-type (Fig. S10D), similar to Grp78−/− mice infected with HSV-1 or EV71 (Fig. S10E–F). These data indicated that neuronal GRP78 regulates GPX4 protein in vivo, prompting us to explore its role in ferroptosis.

To investigate the role of neuronal GRP78 in neuroinflammation, we intraperitoneally injected LPS (5 mg/kg) into Grp78−/− mice pre-treated with Fer-1 (10 mg/kg, ferroptosis inhibitor) or DMSO 1 h prior. Wt mice receiving DMSO + LPS served as controls. Brain tissues (cerebral cortex/cerebellum) collected at 12 h post-injection showed that Grp78−/− mice had significantly elevated ROS and 4-hydroxynonenal (4-HNE) levels compared with Wt, which were reversed by Fer-1 (Fig. 5(A–E)). Neuronal injury was further evaluated by Nissl staining. Compared with mock-injected Wt mice, LPS challenge induced evident neuronal morphological damage in the cerebral cortex and cerebellum, characterized by neuronal shrinkage, irregular cellular morphology, and loss or dispersion of Nissl substance. Notably, these pathological alterations were markedly exacerbated in Grp78−/− mice, whereas Fer-1 treatment substantially alleviated neuronal morphological damage (Fig. 5(F) and (G)). Quantitative analysis further confirmed that the percentage of morphologically damaged neurons was significantly increased following LPS challenge and further elevated by Grp78 deficiency, while Fer-1 treatment markedly reduced neuronal damage. To validate viral infection models, Wt/Grp78−/− mice were infected with HSV-1 or EV71. Infected Wt brains showed elevated lipid peroxidation and ROS, which were further increased in Grp78−/− mice. Fer-1 reversed these ferroptosis markers to near-Wt levels (Fig. S11).

Fig. 5.

Fig. 5

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Neuronal GRP78 deficiency exacerbates ferroptosis and neuronal injury in LPS-induced encephalitis. (A) Schematic of in vivo treatments: Wt mice (saline/LPS) and neuron-specific Grp78 knockout (Grp78−/−) mice (LPS + DMSO/LPS + Fer-1, 10 mg/kg i.p.). (B–E) Brain sections from mice stained for (B) ROS (DCFH-DA, red) and DAPI (blue, scale bar: 20 μm); (D) 4-HNE (green) and DAPI (blue, scale bar: 20 μm); quantification of (C) ROS and (E) 4-HNE fluorescence intensity. (F) Nissl staining of cortical neurons (F, scale bar: 20 μm). (G) Representative images of Nissl staining and quantification of morphologically damaged neurons. Morphologically damaged neurons were identified based on neuronal shrinkage, irregular cellular morphology, nuclear condensation, and loss or dispersion of Nissl substance. Data are presented as the percentage of damaged neurons among the total neurons counted. Data are mean ± SEM (n = 5). ∗∗∗P < 0.001, ∗∗∗∗P < 0.000 1.

Collectively, these findings demonstrate that neuronal GRP78 deficiency exacerbates ferroptosis and neuronal injury in vivo, regardless of pathogenic trigger (LPS or neurotropic viruses), with Fer-1 rescue confirming ferroptosis as the primary cell death mechanism.

2.6. PLIN2-GRP78 axis suppresses neuronal ferroptosis by stabilizing GPX4, attenuating microglial activation and neuroinflammation to Improve Survival in LPS/virus-challenged mice

Neuronal cell death—including ferroptosis—releases cellular components and molecular signals that trigger and exacerbate neuroinflammation (Sarkar et al., 2014). To investigate how GRP78 deficiency impacts this process, we analyzed neuroinflammation in LPS-challenged wild-type (Wt) and neuron-specific Grp78 knockout (Grp78−/−) mice with/without Fer-1 (ferroptosis inhibitor) pretreatment. H&E staining revealed LPS induced perivascular cuffing and glial accumulation (hallmarks of neuroinflammation (Varadkar et al., 2014)) in the cerebral cortex and cerebellum, which were exacerbated in Grp78−/− mice and alleviated by Fer-1 (Fig. 6(A)). A similar pattern was observed in HSV-1-infected mice (Fig. S12). The qRT-PCR analysis confirmed LPS upregulated pro-inflammatory cytokines (TNF-α, IL-1β, IL-6) in Wt mice. Grp78−/− mice showed further elevation of IL-1β/TNF-α/IL-6, effects fully reversed by Fer-1 (Fig. 6(B–D)). Given that ferroptosis activates microglia to release pro-inflammatory cytokines (Castro-Gomez & Heneka, 2024), we assessed cytokine distribution via immunofluorescence (IBA-1 for microglia, NEUN for neurons (Morizane et al., 2013)). LPS increased neuronal release of IL-1β/TNF-α (primarily extracellular), with Grp78−/− mice showing exacerbated release and Fer-1 treatment reversing this effect (Fig. 6(E–G)).

Fig. 6.

Fig. 6

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GRP78 suppresses neuroinflammation and improves survival in LPS- and virus-induced encephalitis. (A) H&E staining of brain sections from Wt and Grp78−/− mice (LPS ± Fer-1) showing perivascular cuffing and glial infiltration (scale bar: 20 μm). (B–D) qRT-PCR analysis of IL-1β (B), TNF-α (C), and IL-6 (D) mRNA in brain tissues from mice. (E–G) Immunofluorescence staining of brain sections for neurons (NEUN, cyan), microglia (IBA-1, red), TNF-α (green), and IL-1β (yellow) (E, scale bar: 10 μm); quantification of (F) TNF-α and (G) IL-1β fluorescence intensity. (H) Survival curves of Wt mice without LPS treatment, Wt mice with LPS treatment, Grp78⁻/⁻ mice with LPS treatment, and Grp78⁻/⁻ mice with LPS and Fer-1 treatment (n = 10 mice/group). Data are mean ± SEM ((B–F), (H–I), n = 5). ∗∗P < 0.01, ∗∗∗P < 0.001, ∗∗∗∗P < 0.000 1.

Survival studies revealed that Grp78−/− mice succumbed 2 days earlier to LPS-induced lethal encephalitis (0% survival) compared with Wt mice, while Fer-1 rescued survival to 20%. Similarly, Grp78−/− mice died 3 days earlier from HSV-1 infection (0% survival), with Fer-1 improving survival to 25% (Fig. 6(H) and S13). These data demonstrate that neuronal GRP78 deficiency accelerates mortality by promoting ferroptosis-driven microglial activation and pro-inflammatory cytokine amplification.

In summary, the LD protein PLIN2 interacts with GRP78 to stabilize GPX4 by inhibiting K48-linked ubiquitination at GPX4-K47. This suppresses lipid peroxidation and ROS, thereby restricting neuronal ferroptosis and attenuating neuroinflammation progression.

3. Discussion

Neuronal ferroptosis has emerged as a key driver of neuroinflammatory and neurodegenerative pathology; however, how lipid metabolic organelles regulate ferroptotic cell fate in neurons remains incompletely understood (Becher et al., 2024; Gu et al., 2024; Wang et al., 2023). LDs, traditionally viewed as inert lipid storage compartments, are increasingly recognized as dynamic organelles involved in cellular stress adaptation (Farías et al., 2022; Gu et al., 2019; Wang et al., 2022). In this study, we identify a PLIN2-centered regulatory axis that links lipid droplet biology to ferroptosis control through ER-associated proteostasis mechanisms.

A central finding of this work is that PLIN2 functions as an upstream regulator of the ER chaperone GRP78, thereby connecting LDs to ferroptosis signaling. While previous studies have primarily focused on LDs as metabolic buffers or lipid detoxification sites (Mehlem et al., 2013; Scorletti & Carr, 2022; Yang et al., 2022), our results support a broader role for LD-associated proteins in coordinating inter-organelle stress communication. Specifically, PLIN2 promotes the expression and cytoplasmic redistribution of GRP78, suggesting that LDs actively engage ER proteostasis networks rather than serving as passive lipid reservoirs.

Mechanistically, we demonstrate that GRP78 acts as a key post-translational regulator of the ferroptosis effector GPX4. GRP78 directly interacts with GPX4 and suppresses its K48-linked ubiquitination, thereby preventing proteasomal degradation and maintaining GPX4 protein stability. This regulation occurs independently of transcriptional control, highlighting protein stability as a critical checkpoint in ferroptosis sensitivity. Importantly, these findings extend the functional scope of GRP78 beyond its canonical role as an ER-resident chaperone involved in unfolded protein response signaling, revealing a broader capacity to regulate cytosolic protein homeostasis linked to oxidative cell death.

Neuron-specific deletion of GRP78 in mice markedly exacerbates lipid peroxidation, neuroinflammation, and disease severity in both LPS- and virus-induced encephalitis models. These pathological phenotypes are significantly alleviated by pharmacological inhibition of ferroptosis, indicating that ferroptosis is a major downstream effector of GRP78 deficiency in neuroinflammatory injury. Importantly, GRP78 was selected as the in vivo genetic target because it represents the critical downstream effector node within the PLIN2–GRP78–GPX4 axis. Previous studies have demonstrated that systemic PLIN2 deficiency leads to substantial alterations in LD homeostasis and metabolic phenotypes, including reduced hepatic lipid accumulation and protection against diet-induced obesity, adipose inflammation, and fatty liver disease (Tsai et al., 2017). Moreover, tissue-specific deletion of PLIN2 has been reported to alleviate hepatic lipid accumulation, inflammation, and fibrosis under metabolic stress conditions (Wang et al., 2024). These findings support the role of PLIN2 as an important upstream regulator of lipid droplet homeostasis and cellular stress responses in vivo. At the same time, the broad metabolic and tissue-specific phenotypes associated with PLIN2 deficiency highlight its pleiotropic functions and indicate that PLIN2 deletion may affect multiple biological processes beyond the PLIN2–GRP78–GPX4 pathway. Given that PLIN2 operates upstream of GRP78 to regulate GPX4 stability in our experimental system, disruption of GRP78 provides a functionally focused strategy to interrogate the physiological relevance of this pathway in vivo while minimizing the broader pleiotropic consequences associated with PLIN2 deficiency. Thus, while PLIN2 initiates the regulatory cascade, GRP78 serves as a critical downstream execution checkpoint linking LD signaling to ferroptotic cell death in the brain.

More broadly, our findings suggest that ferroptosis is not merely a passive consequence of neuroinflammation but a tightly regulated cell death program governed by proteostasis networks. The convergence of lipid metabolic dysfunction, ER stress, and oxidative injury observed in neurological disorders implies that the PLIN2–GRP78–GPX4 axis may represent a shared regulatory node of neuronal vulnerability across diverse pathological conditions. In this context, LDs emerge as active signaling platforms that integrate metabolic and proteotoxic stress to shape neuronal fate decisions (Ackerman et al., 2018; Angius et al., 2015; Bezawork-Geleta et al., 2022).

Several important questions remain to be addressed. The precise mechanism by which PLIN2 regulates GRP78 abundance and subcellular distribution remains unclear, including whether this involves direct protein–protein interaction, modulation of lipid droplet–ER contact sites, or indirect effects on ER stress signaling. Although emerging evidence suggests that ER stress and LD metabolism can influence each other under various pathological conditions, whether ER stress directly feeds back to regulate LD formation during neuroinflammation remains unclear. Therefore, the potential reciprocal regulation between LD dynamics and ER homeostasis warrants further investigation. In addition, whether other LD-associated proteins contribute to ferroptosis regulation, or whether PLIN2 confers functional specificity in neurons, warrants further investigation. Finally, the upstream signals that drive PLIN2 activation during neuroinflammatory conditions remain to be defined and may represent an early regulatory checkpoint in disease progression.

In summary, this study establishes PLIN2 as an upstream regulator of an LD–ER–ferroptosis intracellular signaling axis that governs neuronal survival under inflammatory stress. By defining a PLIN2-dependent mechanism that engages GRP78 to stabilize GPX4, we provide a conceptual framework linking lipid droplet biology to protein quality control and ferroptotic cell death. These findings expand the functional landscape of lipid droplets and identify PLIN2 as a key regulator of neuronal stress responses with potential relevance to neuroinflammatory disease.

4. Materials and methods

4.1. Cells, viruses, antibodies, inhibitors and drugs

N2a (ATCC CCL-131) and SK-N-SH (ATCC HTB-11) cells were obtained from the American Type Culture Collection and authenticated by STR profiling. Cells were cultured in DMEM supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin at 37 °C under 5% CO2. Primary cortical neurons were isolated from embryonic day 16–18 C57BL/6J mouse embryos as previously described (Zhao et al., 2012). Neurons were seeded on poly-L-lysine-coated plates and maintained in Neurobasal medium supplemented with 2% B-27 and 1% penicillin–streptomycin.

Enterovirus 71 (EV71, BrCr strain) and herpes simplex virus type 1 (HSV-1, F strain) were propagated in Vero cells. The following primary antibodies were used: PLIN2 (A20843), GRP78 (A0241), GPX4 (A21440), Flag (AE601), Myc (AE010), His (AE003), and 4-HNE (A24456) from ABclonal; Actin (23660-1-AP) from Proteintech. Secondary antibodies included HRP-conjugated goat anti-mouse (BA1051) and goat anti-rabbit (BA1055) from Boster, as well as DyLight 594-labeled goat anti-mouse IgG (H + L, 35511; Invitrogen).

The following reagents were used: BODIPY 493/503 (GC42959; GLPBIO, CA, USA) for lipid droplet staining; oleic acid (OA, HY-N1446), RSL3 (HY-100218A), Ferrostatin-1 (Fer-1, HY-100579), and lipopolysaccharide (LPS, HY-D1056) were purchased from MedChemExpress.

4.2. Mouse experiment

Neuron-specific GRP78-deficient C57BL/6J mice were obtained from Cyagen Biosciences and housed under standard conditions at Huazhong Agricultural University with free access to food and water. The neuron-specific conditional GRP78 knockout mice used in this study were generated by Cyagen Biosciences using the Cre/loxP system combined with tamoxifen-inducible Cre recombinase to achieve spatiotemporally controlled gene deletion. The specific mouse strain information is as follows: the Cre driver line is the Thy1-CreERT2 transgenic line, which expresses tamoxifen-inducible Cre recombinase specifically in neurons; the floxed allele is Grp78^(flox/flox), in which loxP sites are inserted flanking exon 5∼7 of the Grp78 gene. The genetic background is C57BL/6J. Experimental mice were Thy1-CreERT2; Grp78^(flox/flox), while control mice included Thy1-CreERT2; Grp78^(flox/wt) and littermate Grp78^(flox/flox) negative mice. All mice used in this study were female and 8 weeks of age at the time of tamoxifen induction. Tamoxifen was administered intraperitoneally at a dose of 20 mg/kg body weight for 5 consecutive days. After tamoxifen induction, mice were allowed a 5-day recovery period before subsequent experiments. For the inflammatory model, mice were intraperitoneally injected with LPS (5 mg/kg) or saline, with or without Ferrostatin-1 (10 mg/kg) administered 1 h before LPS treatment. Brain tissues were collected 12 h later for histological analysis and measurement of ROS and 4-hydroxynonenal (4-HNE) levels. For viral infection, 8-week-old female mice were intraperitoneally injected with EV71 (104 PFU/mouse), HSV-1 (102 PFU/mouse), or DMEM as mock control, with or without Ferrostatin-1 pretreatment (10 mg/kg, 1 h before infection).

4.3. Western blot

Total proteins were extracted using RIPA buffer, separated by SDS-PAGE, and transferred onto PVDF membranes. After blocking with 5% skim milk, membranes were incubated with primary antibodies overnight at 4 °C and corresponding secondary antibodies for 1 h at room temperature. Signals were visualized using an Amersham Imager 600.

4.4. Semi-endogenous Co-IP

N2a cells were transfected with the indicated plasmids for 24 h, lysed in RIPA buffer, and subjected to immunoprecipitation using His-tag or Flag-tag beads (Beyotime Biotechnology) at 4 °C overnight. After six washes with lysis buffer, bound proteins were eluted in SDS sample buffer at 98 °C for 10 min, separated by SDS-PAGE, and analyzed by immunoblotting with the indicated antibodies.

4.5. Ubiquitination assay

For the ubiquitination assay, N2a cells were transfected with the indicated plasmid. 48 h post-transfection, cells were treated with MG132 (10 μM) for 4 h before harvest. After lysis of cells in RIPA buffer, they were subjected to ultrasound treatment, as described earlier (Yuan et al., 2024). Cells were lysed and immunoprecipitated with Myc antibody-conjugated magnetic beads to capture total or K48-/K63-linked ubiquitinated proteins. The precipitates were analyzed by immunoblotting using anti-Myc and ubiquitin antibodies.

4.6. GST pull-down

GST-GRP78 or GST-PLIN2 were expressed in BL21(DE3) cells and purified using glutathione-Sepharose beads (Cytiva). For pull-down assays, purified GST-GRP78 or GST-PLIN2 were immobilized on glutathione beads and incubated with lysates from N2a cells transfected with PLIN2-His/GPX4-Myc or empty vector for 2 h at 4 °C. After washing, bound proteins were eluted in Laemmli buffer, separated by SDS-PAGE, and analyzed by immunoblotting with anti-His and anti-GST antibodies.

4.7. Transfection of siRNA

Three pairs of small interfering RNAs (siRNAs) used in this study were designed and synthesized by Tsingke Biotechnology. To verify the transfection efficiency, siRNAs were diluted to a final concentration of 50 nM and transfected into N2a cells according to the manufacturer's instructions. The specific target sequence for siPLIN2-1 was 5′ - AGUGCAUGAU GUUGCUGGACAGGUG - 3′, and siPLIN2-2 sequence was 5′ - CUAAAAGCUCAGAUGCACUU - 3′, and siPLIN2-3 sequence was 5′ - UAGAUAAGCAUCUUCUUAGA - 3′. The specific target sequence for siGRP78-1 was 5′ - GUGAACGACCCCUAACA- 3′, and siGRP78-2 sequence was 5′ - GCUGGAAAGCCACCAGGAUG- 3′, and siGRP78-3 sequence was 5′ - UACAGCUGCUAUUGCA- 3′.

4.8. RNA-seq library construction and sequencing

N2a cells were transfected with either the PLIN2-His expression vector or an empty His-tagged control vector for 48 h. Total RNA was subsequently extracted, and samples with an RNA integrity number (RIN) > 8.0 were selected for downstream analysis. Following quality assessment, mRNA was enriched and subjected to high-throughput sequencing performed by Novogene (Beijing, China). The sequencing workflow included raw data quality control, alignment of clean reads to the reference genome, transcriptome assembly, coding potential prediction, conservation analysis, target gene prediction, gene expression quantification, and differential expression analysis. Gene expression levels were normalized using fragments per kilobase of transcript per million mapped reads (FPKM) to ensure comparability across genes and samples. The RNA-seq dataset generated in this study has been deposited in the Gene Expression Omnibus (GEO) database under accession number GSE327846.

4.9. Ethical approval

C57BL/6J mice were housed in the animal facility at Huazhong Agricultural University following the Regulations on the Administration of Laboratory Animals of Hubei Province, China. All animal experimental procedures were reviewed and approved by the Institutional Animal Care and Use Committee (IACUC) of Huazhong Agricultural University (permit number HZAUMO-2020-0098).

4.10. Statistical analysis

All statistical analyses were performed using GraphPad Prism software. Data are presented as mean ± SEM unless otherwise indicated. For comparisons between two groups, an unpaired two-tailed Student's t-test was used. For experiments involving more than two groups with a single independent variable, one-way analysis of variance (ANOVA) followed by an appropriate multiple-comparison test was performed. Tukey's multiple-comparisons test was used for comparisons among all groups, whereas Dunnett's test was used when multiple experimental groups were compared with a common control group. For experiments involving two independent variables, two-way ANOVA followed by an appropriate multiple-comparison test was performed. The exact statistical test used for each experiment is indicated in the corresponding figure legends.

Unless otherwise indicated, n represents the number of biologically independent samples used for statistical analysis. For cell-based experiments, biological replicates represent independently performed experiments using separate cell cultures on different occasions. For animal experiments, n represents biologically independent animals. For imaging-based experiments, multiple fields of view were acquired from each biological replicate using consistent imaging criteria. These fields were considered technical measurements and were averaged to obtain a single value for each biological replicate before statistical analysis; individual fields of view were not treated as independent samples. All experiments were independently repeated at least three times unless otherwise indicated. A P value < 0.05 was considered statistically significant.

CRediT authorship contribution statement

Qianruo Wang: Writing – original draft, Investigation, Formal analysis, Data curation. Baokun Sui: Writing – original draft, Validation, Investigation, Formal analysis. Jiaxin Zheng: Investigation, Data curation. Chuanyin Yao: Investigation. Jianqing Zhao: Investigation. Zhen F. Fu: Supervision. Ling Zhao: Writing – review & editing, Writing – original draft, Supervision, Project administration. Ming Zhou: Writing – review & editing, Writing – original draft, Supervision, Conceptualization.

Data availability

Data supporting the findings of this work and supplementary information are available from the corresponding author on reasonable request.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgments

This study was supported by the Natural Science Foundation of Heilongjiang Province (QC2026C021 to Q.R.W.), Young Scientists Fund of the National Natural Science Foundation of China (32604642 to Q.R.W.), and the Major Basic Research Project of Hubei Provincial Natural Science Foundation (2024AFE010 to L.Z.). HSV-1-GFP, a gift from Prof. Gang Cao (Huazhong Agricultural University, China), and EV71, a gift from Prof. Min Cui (Huazhong Agricultural University, China), were used in this study.

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.cellin.2026.100369.

Contributor Information

Ling Zhao, Email: zling604@outlook.com, lingzhao@mail.hzau.edu.cn.

Ming Zhou, Email: mikchail@163.com, mingzhou@mail.hzau.edu.cn.

Appendix A. Supplementary data

The following is the supplementary data to this article:

Multimedia component 1
mmc1.docx (2.7MB, docx)

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Supplementary Materials

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Data Availability Statement

Data supporting the findings of this work and supplementary information are available from the corresponding author on reasonable request.

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