A UFD1 variant encoding a microprotein modulates UFD1f and IPMK ubiquitination to play pivotal roles in anti-stress responses

Li, Xiuzhi; Wang, Xiaolin; Liu, Xu; Shan, Ge; Chen, Liang

doi:10.1038/s41467-025-62073-6

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Published: 21 July 2025

A UFD1 variant encoding a microprotein modulates UFD1f and IPMK ubiquitination to play pivotal roles in anti-stress responses

Nature Communications volume 16, Article number: 6695 (2025) Cite this article

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Abstract

Eukaryotic cells make multiple efforts to cope with internal and external stresses; such mechanisms include metabolic responses and the generation of stress-responsive mRNA isoforms (SR-mRNAisos), such as the classical XBP1s. Here, we identified a mammalian conserved SR-mRNAiso, UFD1s, which encodes a microprotein with anti-stress functions. UFD1s decreased the K63-linked ubiquitination levels of UFD1 full-length protein (UFD1f) via competitive binding to the E3 ubiquitin ligase MARCH7, and therefore regulated the dynamics of protein ubiquitination. Inositol polyphosphate multikinase (IPMK) was identified as the most significantly UFD1s-regulated target in terms of changes in K48- and K11-ubiquitination. UFD1s promoted autophagy and fatty acid oxidation, and IPMK was consistently destabilized. Ufd1s-deficient male mice exhibited metabolic disorders and accelerated NASH progression. Plasmid or circRNA expressing UFD1s alleviated NASH in mice, indicating that UFD1s has therapeutic value. Our findings revealed a mammalian conserved microprotein that plays crucial roles in anti-stress regulation through the modulation of ubiquitination and metabolism.

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Introduction

Eukaryotic cells employ adaptive and conserved mechanisms to maintain homeostasis in response to cellular stresses such as endoplasmic reticulum (ER) stress, oxidative stress, heat shock, and nutrient stress, including glucose or amino acid deprivation^1,2,3. Prolonged exposure to cellular stresses contributes to multiple diseases, such as metabolic disorders and human cancers^1,4,5. Taking the example of ER stress or the unfolded protein response (UPR), eukaryotic cells utilize a key stress sensor, IRE1α, which is activated to selectively splice XBP1 (HAC1 in yeast) mRNA^6,7,8. Spliced XBP1 (XBP1s) encodes a potent transcription factor that regulates the transcription of target genes involved in protein homeostasis^6,7,8,9. The XBP1s protein is also induced under oxidative stress or other types of stress, and is involved in lipid regulation and gluconeogenesis^10,11.

One shared feature of the cellular response to stress is the global dynamic regulation of protein ubiquitination, which is required to degrade a large portion of unwanted proteins, stabilize specific proteins, or regulate cell signaling^2,3. For the ubiquitin system, distinct polyubiquitin chain linkages play different roles^12,13. Individual ubiquitin moieties can be conjugated via one of seven lysine (K) residues, K6, K11, K27, K29, K33, K48, and K63, or the N-terminal methionine residue (M1) in ubiquitin to form different types of polyubiquitin^12,13. The three most abundant types of polyubiquitin chains in cells are K48-, K11-, and K63-linked^12,14. K48-linked polyubiquitin chains, the most abundant and most studied linkage, typically lead to proteasome-mediated protein degradation^13,14. K11- and K63-linked polyubiquitin are less studied^12,13. K11-polyubiquitin conjugates are considered degradation signals in cell cycle regulation^13,15, but have also been shown to stabilize some proteins^16,17,18. K63-polyubiquitin linkages have nonproteolytic functions, and regulate the recruitment, trafficking, and activity of specific proteins^12,19,20. For example, K63-linked ubiquitination of G3BP1 (the core protein of stress granules), which is catalyzed by the TRIM21 E3 ubiquitin ligase, effectively inhibits the formation of stress granules under oxidative stress²⁰. Not much is known about the other types of polyubiquitin^12,13.

Ubiquitin recognition factor in ER-associated degradation 1 (UFD1) is known as a ubiquitin-recognition protein with putative binding sites for monoubiquitin and polyubiquitin that acts as an adapter protein with nuclear protein localization protein 4 (NPL4) to form the eukaryotic conserved adapter complex in the endoplasmic reticulum (ER)-associated degradation (ERAD) pathway, which exports misfolded proteins from the ER to the cytoplasm for UPS-dependent degradation^{21,22,23,24,25}. K48-ubiquitinated proteins are generally recognized by the adapter complex and then subjected to threading and unfolding by the AAA+ ATPase p97 (called Cdc48 in yeasts) for further degradation in the 26S proteasome^26,27,28. Additionally, UFD1, together with NPL4 and p97, promotes the reformation of the nucleus and regulates spindle disassembly at the end of mitosis in Xenopus laevis egg extracts^29,30. On the other hand, UFD1 is reported to play an NPL4- and p97-independent role in at least one case, in which UFD1 recruits the deubiquitinating enzyme USP13 to stabilize SKP2 in response to ER stress³¹.

Dynamics of metabolic processes, such as remodeling of lipid metabolism and energy metabolism always accompany cellular stress responses^32,33,34. Inositol pyrophosphate metabolites are broadly characterized metabolic regulators in a range of cellular stress responses across eukaryotes^35,36. Inositol polyphosphate multikinase (IPMK) is a highly conserved enzyme in the biosynthesis of lipid-derived inositol polyphosphates, including IP4 and IP5^35,37, and catalysis of the production of phosphatidylinositol-3,4,5-trisphosphate (PIP3) lipids^38,39. IPMK also functions via AMPK and ULK1 signaling or by modulating TFEB to regulate autophagy^40,41,42,43, another major proteolytic system responsible for the removal of an array of intracellular materials, including accumulated unfolded proteins under cellular stress^44,45. Autophagy, as a catabolic process, can recycle cellular components, including lipid droplets (LDs) and damaged organelles, to reduce sources of reactive oxygen species (ROS) in response to diverse types of cellular stress^46,47. Autophagy is also a key cellular event associated with lipid metabolism disorders^46,48.

Cellular stresses are linked to a series of human diseases^4,5,49. Taking the most severe form of nonalcoholic fatty liver disease, nonalcoholic steatohepatitis (NASH), as an example, oxidative stress from the overgeneration of reactive oxygen species (ROS) is a major contributor to this disease^50,51. These ROS molecules result from mitochondrial dysfunction, ER stress, and NADPH oxidase activation in hepatocytes in NASH^51,52. Despite progress in understanding the pathology of NASH, effective treatments for this disease are lacking^50,53.

To further elucidate the anti-stress mechanisms, we aimed to isolate stress-responsive transcripts with regulatory effects in mammalian cells. This effort has led to the identification of a stress-responsive mRNA isoform (SR-mRNAiso), UFD1s, encodes a mammalian conserved microprotein UFD1s. An array of molecular, cellular, and mouse experiments have been carried out to evaluate the anti-stress roles of UFD1s and to reveal the corresponding functional mechanisms. We have provided lines of evidence showing that insufficient levels of cellular UFD1s contribute to increased total ubiquitination levels by affecting the activity of the UFD1 full-length protein (UFD1f). Via the destabilization of a key factor, IPMK, by affecting its K48- and K11-linked polyubiquitin, UFD1s promotes autophagy and fatty acid oxidation. Ufd1s-deficient mice exhibit metabolic disorders, stress-related phenotypes in the liver, and accelerated NASH progression. UFD1s has potential biomedical value, as the overexpression of this protein with a plasmid or circular RNA (circRNA) relieves NASH symptoms in mice.

Results

UFD1s is identified as a stress-responsive UFD1 mRNA isoform

To identify the dynamics of alternative splicing events in mammalian cells under stress, we performed ribosomal RNA (rRNA)-depleted RNA sequencing (RNA-seq) on human HEK293T and mouse N2a cells subjected to glucose deprivation (GD, metabolic oxidative stress) or thapsigargin (TG, an ER stress inducer) (Fig. 1a, b). A total of 2529 and 3052 differentially expressed splicing events (DESEs) (fold change >2 or <0.5, P value < 0.05) in HEK293T cells upon GD and TG treatments compared with the control, respectively, were identified; 650 of these events were shared by the two stress conditions (Fig. 1b and Supplementary Data 1). In N2a cells, 3992 and 3840 DESEs were identified upon GD and TG treatment, respectively; 550 of them overlapped between the two stresses (Fig. 1b and Supplementary Data 1). We further performed sequence conservation analysis for these 650 and 550 SR-mRNAisos in human and mouse cells, respectively. Twenty-seven splicing variants with conservation scores over 50% were identified between human and mouse SR-mRNAisos, and XBP1s and a UFD1 splicing variant, UFD1s, were the two with the highest conservation (Fig. 1b and Supplementary Data 1). XBP1s is 100% conserved, and UFD1s is 97.5% conserved (Fig. 1b and Supplementary Data 1).

**Fig. 1: Stress-responsive *UFD1* variant (*UFD1s*) encodes a microprotein in mammals.**

XBP1s, a well-known SR-mRNAiso, was identified, supporting the strategy of our analyses. The UFD1 gene is highly conserved from yeast to mammals, and consists of 12 exons in both humans and mice (Supplementary Fig. 1a, b). The mammalian conserved SR-mRNAiso UFD1s is a previously unrecognized isoform lacking a 26-nucleotide (nt) region present in the UFD1f mRNA (Supplementary Fig. 1b). This 26-nt difference between UFD1s and UFD1f allowed isoform-specific sequence-based examinations such as PCR and fluorescence in situ hybridization (FISH) (Supplementary Fig. 1c). The genomic structure of UFD1 in human and mouse indicated that an alternative 5′ splice site (A5SS) located within exon 2 led to the formation of UFD1s (Fig. 1c). UFD1s levels in both human and mouse cells were unaltered when NMDI14, a nonsense-mediated RNA decay (NMD) inhibitor⁵⁴, was used, indicating that UFD1s is not a substrate for NMD (Supplementary Fig. 1d). We then explored the potential RNA-binding proteins (RBPs) involved in UFD1s formation. Integrative analysis of published RBP eCLIP-seq data (103 RBPs in HepG2 cells and 120 RBPs in K562 cells from ENCODE; see “Methods” for details) revealed that AQR, DDX3X, NCBP2, TIA1, and U2AF65 presented significant binding signals on exon 2b along with the 100-nt flanking sequences (Supplementary Fig. 1e). Knockdown of the splicing factors AQR or U2AF65 but not the other three RBPs significantly reduced UFD1s levels^55,56,57, suggesting that alternative splicing of the UFD1 gene regulated by AQR and U2AF65 generated the SR-mRNAiso UFD1s (Supplementary Fig. 1e, f). Unlike XBP1s, UFD1s was not produced by IRE1α-mediated unconventional splicing, as either knocking down IRE1α with siRNA or attenuating the endonuclease activity of IRE1α with the small-molecule inhibitor 4μ8C^6,7,58 resulted in decreased XBP1s levels but unaltered UFD1s levels in HEK293T cells under normal or stress conditions (Supplementary Fig. 1g, h). IRE1α RNA immunoprecipitation (RIP) also demonstrated that IRE1α bound to unspliced XBP1 but not UFD1f mRNA (Supplementary Fig. 1i). Furthermore, 5′ and 3′ rapid amplification of cDNA ends (RACE) demonstrated that the full length of human UFD1s mRNA was 979 nt (Supplementary Fig. 1j), while the mouse Ufd1s transcript has been annotated (NR_028403.1).

PCR and single-molecule FISH (smFISH) revealed that UFD1s expression was significantly increased after TG or GD treatments from low baseline levels under normal, unstressed conditions in human HEK293T and mouse N2a cells (Fig. 1e, f and Supplementary Fig. 2a). UFD1f levels were unaltered or slightly but significantly increased in N2a cells (Fig. 1e, f and Supplementary Fig. 2a). Both UFD1s and UFD1f mRNAs were predominantly localized in the cytoplasm (Fig. 1f). UFD1s also exhibited significantly increased expression levels in human HeLa and mouse Hepa1-6 cells under various stresses, including TG, GD, Hank′s balanced salt solution (HBSS)-responsive nutrient starvation, and glutamine deprivation (Supplementary Fig. 2b). The levels of UFD1f either did not significantly change or increased to a lesser degree than did the levels of UFD1s in either HeLa or Hepa1-6 cells under these stresses (Supplementary Fig. 2b). Saccharomyces cerevisiae (S. cerevisiae) HAC1 generated a stress-induced isoform, whereas a corresponding stress-induced UFD1s was not detected in the yeasts S. cerevisiae or Schizosaccharomyces pombe (S. pombe) under treatment with tunicamycin (Tm, an ER stress inducer) (Supplementary Fig. 2c). UFD1 is a single-exon gene in S. cerevisiae, and the corresponding exon of the UFD1 gene in S. pombe lacks an A5SS (Supplementary Fig. 2d). These results demonstrated that UFD1s is an upregulated SR-mRNAiso under cellular stresses that is conserved at least in mammalian cells.

UFD1s encodes a microprotein

UFD1s contains a potential open reading frame (ORF) of 150- and 174-nt to encode a deduced 50- and 58-amino acid (aa) polypeptide in humans and mice, respectively (Fig. 1g and Supplementary Fig. 2e). Ribosome profiling revealed that polyribosomes bound to UFD1s RNA (Supplementary Fig. 2f, g), indicating that UFD1s might function as a template for protein translation. Flag-tagged constructs were then generated, in which the tag was fused to the deduced C-terminus of the UFD1s ORF sequences (Supplementary Fig. 2h). The UFD1s-Flag fusion protein, which was examined using an anti-FLAG antibody, was expressed in human and mouse cells (Supplementary Fig. 2h). Mutation of the UFD1s-ORF start codon from ATG to ATT in the constructs abolished UFD1s-Flag protein expression (Supplementary Fig. 2h). We produced specific antibodies against the unique 14-aa and 22-aa C-terminal regions of the human and mouse UFD1s proteins, respectively (Fig. 1g). The ability of the anti-UFD1s antibody to detect the UFD1s protein via western blotting was verified with the overexpressed UFD1s-Flag fusion, as both the anti-UFD1s and anti-FLAG antibodies detected the fusion protein (Supplementary Fig. 2i). To exclude the possibility that UFD1s might be derived from the breakdown of UFD1f, as UFD1s shares 36 N-terminal amino acids with UFD1f, N-terminal and C-terminal Flag-tagged UFD1f plasmids were constructed. The western blotting results revealed no shorter proteins in either the Flag-UFD1f or the UFD1f-Flag fusion proteins (Supplementary Fig. 2j). UFD1s may not be a shorter broken-down form of the UFD1f protein derived from the breakdown of UFD1f in cells, while it is an independent protein isoform encoded by a separate mRNA (Supplementary Figs. 1j and 2j).

Immunoprecipitation (IP) experiments with the anti-UFD1s antibody in HEK293T and N2a cells revealed that it could pull down the UFD1s protein, and this result was confirmed by mass spectrometry (MS) (Supplementary Fig. 2k). The endogenous UFD1s protein levels were significantly increased upon TG and GD treatments in HEK293T and N2a cells (Fig. 1h). Immunofluorescence (IF) staining revealed that the UFD1s protein was localized predominantly in the cytoplasm and that its levels were increased upon TG treatment (Fig. 1i). We noted that the UFD1f protein levels did not significantly change under stress conditions in either HEK293T or N2a cells (Fig. 1h, i). Overall, we concluded that UFD1s encodes a microprotein whose expression is significantly increased in stressed cells.

The UFD1s protein has anti-stress effects in mammalian cells

To investigate the cellular roles of UFD1s, the CRISPR/Cas9 technique was used to change the A5SS of UFD1 to reduce the generation of UFD1s while keeping the amino acid sequence of UFD1f unchanged (Fig. 2a). Compared with wild-type (WT) cells, UFD1s-deficient (MUT) HEK293T or N2a cells presented lower UFD1s mRNA and UFD1s protein levels, whereas UFD1f mRNA and UFD1f protein levels remained unchanged (Fig. 2b and Supplementary Fig. 3a, b). When examined via IF staining, HEK293T or N2a WT cells, but not MUT cells, presented increased UFD1s levels upon TG treatment (Supplementary Fig. 3c, d). Alteration of the canonical AG/GT site to CG/CA did not completely abolish the expression of UFD1s, and A5SS has been shown to demonstrate some flexibility in previous studies^59,60.

**Fig. 2: UFD1s protein exhibits anti-stress effects and regulates autophagy and lipid metabolism.**

Under normal culture conditions, the growth rate of UFD1s MUT cells was lower than that of WT cells, and the overexpression of UFD1s promoted cell growth (Supplementary Fig. 3e, f). As a readout of oxygen metabolism, ROS were also measured⁶¹, and we found that relative ROS levels were increased in MUT cells compared with WT cells when all cells were cultured in regular medium without stress treatment (Supplementary Fig. 4a). The overexpression of UFD1s in WT cells decreased ROS levels (Supplementary Fig. 4b). These results indicated that baseline UFD1s expression under normal conditions plays a cellular role in eliminating intrinsic metabolic stress. Cellular stresses are frequently accompanied by abnormal levels of ROS and ATP, which are essential for cell survival^61,62. In UFD1s MUT cells, the increase in ROS levels was more evident in those challenged with GD, and UFD1s-overexpressing cells subjected to GD treatment presented lower ROS levels (Supplementary Fig. 4a, b). The overexpression of UFD1s in MUT cells rescued the UFD1s deficient effects on ROS levels (Fig. 2c). In UFD1s MUT cells, ATP levels were slightly but statistically significantly lower in regular medium than those of WT cells, whereas when the cells were challenged with GD, the ATP levels were dramatically lower in MUT cells than in WT cells (Supplementary Fig. 4c). When UFD1s was overexpressed in WT cells under regular culture conditions, ATP levels did not significantly change; under GD treatment, UFD1s-overexpressing cells presented significantly increased ATP levels (Supplementary Fig. 4d). Moreover, the overexpression of UFD1s in MUT cells reversed the UFD1s deficient effects with increased ATP levels (Fig. 2d). The overall protective effects of UFD1s on cell survival were evaluated, and compared with WT cells, UFD1s MUT cells presented significantly lower survival rates upon GD treatment, and the overexpression of UFD1s in MUT cells rescued the cell survival phenotype (Fig. 2e). The effects of UFD1s were also tested with siRNA-mediated knockdown in two other human cell lines, AGS and SGC7901 (Supplementary Fig. 4e). The siRNA was specific for knocking down UFD1s but not UFD1f mRNA (Supplementary Fig. 4f). In both cell lines with UFD1s knockdown, a significantly lower survival rate was observed when the cells were subjected to GD treatment (Supplementary Fig. 4g).

To further explore the cellular phenotypes of UFD1s MUT cells with respect to energy metabolism, the oxygen consumption rate (OCR) of the cells was measured. The OCR can be used to represent four parameters of mitochondrial respiration: basal, ATP production-linked, maximal respiration, and proton leakage⁶³. We found that UFD1s MUT HEK293T and N2a cells presented reduced basal, ATP production-linked, and proton leakage respiration (Fig. 2f and Supplementary Fig. 4h). The maximal respiration rate was also decreased in MUT N2a cells but not in MUT HEK293T cells (Fig. 2f and Supplementary Fig. 4h). Consistent with these changes in MUT cells, the overexpression of UFD1s in WT cells increased all these parameters of mitochondrial respiration (Fig. 2g and Supplementary Fig. 4i). Basal respiration shows energetic demand under baseline conditions, and ATP production-linked respiration represents the portion of basal respiration that drives ATP production. Proton leak respiration is the remaining basal respiration not coupled to ATP production and is a sign of mitochondrial damage or a regulatory mechanism of mitochondrial ATP production.

These results demonstrated that UFD1s has protective effects on and a regulatory role in mitochondrial function in response to intrinsic and external stresses in mammalian cells.

UFD1s regulates autophagy and lipid metabolism

To further evaluate the physiological function of UFD1s, we performed RNA-seq of WT and UFD1s MUT HEK293T cells under normal or GD conditions. A total of 219 (119 upregulated, 100 downregulated) differentially expressed (DE) mRNAs were identified between MUT and WT cells under normal culture conditions, and supposedly these DE mRNAs were related to the effects of UFD1s against internal stresses (Supplementary Fig. 4j, k and Supplementary Data 2). A total of 261 (146 upregulated, 115 downregulated) DE mRNAs were identified between MUT and WT cells under GD treatment, and these DE mRNAs were associated with the effects of UFD1s against external stresses (Supplementary Fig. 4j, k and Supplementary Data 2). Thirty downregulated and 49 upregulated mRNAs were shared by both culture conditions, and these 79 DE mRNAs would be linked to the functions of UFD1s in both intrinsic and external stresses (Fig. 2h, Supplementary Fig. 4k, and Supplementary Data 2). Gene Ontology (GO) analysis of these 79 DE mRNAs revealed that autophagy and lipid-associated biological processes were the most significantly enriched (Fig. 2i); therefore, we focused on the roles of UFD1s in autophagy and lipid metabolism processes.

Transmission electron microscopy (TEM) revealed that, under normal conditions, fewer and smaller autophagic vesicles (AVs) were present in HEK293T or N2a UFD1s MUT cells than in WT cells (Fig. 2j and Supplementary Fig. 5a). Chloroquine (CQ) treatment inhibits the late events of autophagy by blocking the binding of autophagosomes to lysosomes and facilitates the observation of AVs⁴³, and MUT cells also displayed fewer and smaller AVs than WT cells did (Fig. 2j and Supplementary Fig. 5a). HBSS medium induces autophagy under nutrient starvation conditions⁶⁴. Under HBSS starvation, the MUT cells again presented fewer and smaller AVs than the WT cells did (Fig. 2j and Supplementary Fig. 5a). Microtubule-associated protein 1 light chain 3 (LC3)-phosphatidylethanolamine conjugate (LC3-II) is a marker for monitoring autophagy⁴³. LC3-II levels observed by western blotting were significantly decreased in UFD1s MUT cells (Supplementary Fig. 5b). The mCherry-EGFP-LC3 tandem reporter, whose EGFP signal is quenched while the mCherry signal is stable in acidic compartments, was used to monitor autophagic flux^43,65. In MUT cells, red puncta, which represent acidified mature autophagosomes and autolysosomes, were significantly decrease under both normal and starvation conditions (Fig. 2k and Supplementary Fig. 5c). Autophagy is generally considered a protective mechanism against stress⁶⁶, and the results demonstrated that UFD1s promoted autophagy.

We then examined the regulatory roles of UFD1s in lipid and energy homeostasis. Nile red staining revealed that, compared with WT cells, UFD1s MUT cells had significantly more lipid droplets (LDs) (Fig. 2l and Supplementary Fig. 6a). UFD1s overexpression reduced the number of LDs in WT cells (Supplementary Fig. 5b). LDs store neutral lipids mainly in the form of triglycerides and a small portion of cholesterol, and triglyceride levels were also elevated in UFD1s MUT cells (Supplementary Fig. 6c). We next evaluated fatty acid oxidation (FAO) via OCR assays. UFD1s MUT cells presented a decreased OCR, and UFD1s overexpression in WT cells increased the OCR (Fig. 2m and Supplementary Fig. 6d-f). The levels of acetyl-CoA carboxylase (ACC), which is the rate-limiting enzyme of fatty acid biosynthesis, were examined to evaluate the potential regulatory effect of UFD1s on de novo lipid synthesis⁶⁷. Compared with those in the corresponding controls, neither total nor phosphorylated ACC levels were changed in UFD1s-MUT or UFD1s-overexpressing cells (Supplementary Fig. 6g, h). These results demonstrated that UFD1s might not regulate lipid synthesis, but reduce the accumulation of lipids by accelerating the FAO rate. Overall, we concluded that UFD1s regulate autophagy and FAO in mammalian cells and that both are intervening responses to intrinsic and extrinsic stresses.

UFD1s regulates the K63-linked polyubiquitination of UFD1f

Dynamic regulation of protein ubiquitination is a critical part of the cellular response to stress, and the UFD1f protein is a critical regulator of this process^2,25. Interestingly, the UFD1f protein displayed increased total ubiquitination levels in UFD1s MUT cells (Fig. 3a, b). We then examined UFD1f for the four most abundant types of polyubiquitin chains, i.e., K48-, K11-, K6-, and K63-linked chains¹⁴. K6 polyubiquitination, which is implicated in mitochondrial quality control, is the fourth most abundant type among all polyubiquitin chains^14,68. UFD1s MUT resulted in significantly increased levels of K63-linked but not K48-, K11- or K6-linked polyubiquitin chains on UFD1f in human and mouse cells (Fig. 3a, b). Conversely, the overexpression of UFD1s significantly reduced the total ubiquitination and K63 polyubiquitination levels of UFD1f in WT cells (Fig. 3c, d). K63 polyubiquitination is often linked with regulatory effects rather than serving as a proteolytic signal like K48 polyubiquitination^13,19,20. Consistent with these results, UFD1s MUT or overexpression had no significant effect on total UFD1f protein levels (Figs. 2b, 3a–d and Supplementary Fig. 3b).

**Fig. 3: UFD1s modulates K63-polyubiquitination of UFD1f via competitively binding to MARCH7.**

To investigate how UFD1s regulates the K63-polyubiquitination of UFD1f, we employed IP followed by MS (IP‒MS) of UFD1s and UFD1f to identify their common partners in HEK293T cells (Fig. 3e). We speculated that UFD1s may regulate UFD1f by competing with or regulating shared interacting partners. One E3 ubiquitin ligase, membrane-associated RING-CH-type finger 7 (MARCH7), was identified as the only shared E3 ligase for both UFD1s and UFD1f (Fig. 3e and Supplementary Data 3). MARCH7, a RING domain-containing E3 ubiquitin ligase, can catalyze the K63 polyubiquitination of Mdm2, which leads to the stabilization of Mdm2 and the degradation of p53⁶⁹. The UFD1f-MARCH7 and UFD1s-MARCH7 interactions were further validated by co-IP assays in HEK293T and N2a cells (Supplementary Fig. 7a, b). UFD1s MUT resulted in increased binding of UFD1f to MARCH7 in HEK293T and N2a cells, whereas UFD1s overexpression decreased the interaction between UFD1f and MARCH7 (Fig. 3f). Short hairpin RNA (shRNA)-mediated MARCH7 knockdown led to substantial reductions in the total ubiquitination and K63 polyubiquitination levels of UFD1f in human and mouse cells (Fig. 3g). These results demonstrated that UFD1s competitively binds to MARCH7 to regulate the K63 polyubiquitination of UFD1f.

To identify the K63-ubiquitinated lysine sites of UFD1f, we performed a two-step IP assay (Re-IP) followed by MS in HEK293T cells (see “Methods” for details) (Fig. 3h and Supplementary Data 4). Two K63-ubiquitinated K residues, K240 and K280, were identified from two independent replicates, and both are conserved in human and mouse UFD1f (Supplementary Data 4). Three mutants of Flag-tagged UFD1f, including the K240R mutant, K280R mutant, and K240R/K280R double mutant, were then generated, and all three mutants presented significantly decreased K63-linked polyubiquitination levels (Fig. 3i). The K240R/K280R double mutant presented very low levels of K63-linked polyubiquitination (Fig. 3i). Furthermore, the K63 polyubiquitination levels of the K240R/K280R double mutant were not sensitive to UFD1s deficiency in HEK293T MUT cells (Supplementary Fig. 7c). Taken together, these results suggest that through competitive interactions with MARCH7, UFD1s modulates the K63-polyubiquitination of UFD1f, primarily at the K240 and K280 sites.

UFD1s regulates the ubiquitination dynamics of proteins, including IPMK

We next aimed to examine the global changes in protein ubiquitination levels in UFD1s MUT and WT cells. Compared with those in WT HEK293T or N2a cells, total ubiquitination levels were significantly increased in UFD1s MUT cells (Supplementary Fig. 7d). On the other hand, overexpression of UFD1s reduced the total ubiquitination levels in HEK293T or N2a cells (Supplementary Fig. 7e). UFD1s knockdown with siRNA also led to remarkable increases in global ubiquitination in HEK293T and N2a cells (Supplementary Fig. 7f).

To further explore changes in UFD1s-modulated ubiquitination, we conducted ubiquitin (label-free)-modified proteome profiling in UFD1s MUT and WT HEK293T cells (see “Methods” for details) (Fig. 4a)⁷⁰. A total of 178 proteins were significantly upregulated, and 93 proteins had significantly downregulated ubiquitination levels in UFD1s MUT cells (Fig. 4b and Supplementary Data 4). We focused on proteins with higher ubiquitination levels because global protein ubiquitination levels were also increased in MUT cells. GO analysis of the 178 proteins revealed several biological processes associated with responses to an array of internal or external cellular stresses, protein polyubiquitination, ribonucleoprotein complex assembly, the cell cycle, protein localization to organelle, autophagy, and lipid metabolic process (Fig. 4c). IPMK, which belongs to the GO term metabolic process, was identified as the most significantly UFD1s-regulated target in terms of changes in ubiquitination (Fig. 4b, c).

**Fig. 4: UFD1s regulates ubiquitination dynamics of proteins, including IPMK and IPMK is regulated by UFD1f and MARCH7.**

We observed that IPMK protein levels were markedly increased in UFD1s MUT HEK293T and N2a cells without affecting IPMK protein translation (Fig. 4d). The IPMK mRNA levels and the binding of the polyribosome to IPMK mRNA were unchanged in UFD1s MUT cells (Supplementary Fig. 7g, h). Western blotting revealed that the total ubiquitination level of IPMK was significantly greater in UFD1s MUT cells than in WT cells (Fig. 4e, f), whereas the overexpression of UFD1s prominently reduced the total ubiquitination level of IPMK (Supplementary Fig. 7i). K48-, K11-, and K63-linked polyubiquitin chains of IPMK were then examined, and the levels of the K11-linked type were significantly increased, those of the K63-linked type displayed no significant change, and those of the K48-linked type were lower in UFD1s MUT cells (Fig. 4e, f). Consistently, overexpression of UFD1s in WT cells caused reduced K11 polyubiquitination and increased K48 polyubiquitination in mammalian cells (Supplementary Fig. 7i). K48-polyubiquitin chains are the canonical signal for proteasomal degradation^12,13. K11-polyubiquitin conjugates are believed to stabilize proteins^16,17; for example, K11-polyubiquitin stabilizes β-catenin and activates the Wnt/β-catenin signaling pathway¹⁶. Taken together, these results demonstrated that UFD1s affects the ubiquitination of many proteins involved in stress responses and metabolic processes, among which IPMK is regulated mostly by K11-polyubiquitin.

MARCH7-intermediated ubiquitination of UFD1f regulates IPMK ubiquitination

To test whether the regulation of IPMK by UFD1s was intermediated by UFD1f, we assessed the interaction between UFD1f and IPMK. Co-IP assays demonstrated that UFD1f could co-pull down IPMK (Fig. 4g). Furthermore, compared with the K240R/K280R double mutant, the overexpression of WT UFD1f resulted in higher total IPMK protein levels, and compared with the K240R/K280R double mutant, the overexpression of WT UFD1f resulted in lower K48-linked and higher K11-linked ubiquitination levels of IPMK (Fig. 4h, i). Given that MARCH7 intermediated UFD1f K63-ubiquitination, we then investigated whether dysregulation of MARCH7 affects the stability and ubiquitination of IPMK. ShRNA-mediated knockdown of MARCH7 led to a substantial reduction in IPMK protein levels, with increased K48- and decreased K11-polyubiquitination levels of IPMK in human and mouse cells (Fig. 4j, k). These results indicate that UFD1s modulates dynamic changes in protein ubiquitination. IPMK was the most sensitive target of the changes in ubiquitination under UFD1s insufficiency, and UFD1s-modified UFD1f K63 ubiquitination regulated the ratio of K48- and K11-linked ubiquitination of IPMK.

IPMK contributes significantly to UFD1s phenotypes

We next sought to examine the involvement of IPMK in the effects of UFD1s on autophagy and lipid metabolism. LC3-positive puncta, which are markers of autophagy, were also examined⁴³, and UFD1s MUT resulted in a reduction in the number of LC3-positive puncta (Fig. 5a, b). The knockdown of IPMK with shRNA rescued the repressive effects of LC3-positive puncta in UFD1s MUT cells (Fig. 5a, b). Nile red staining demonstrated that IPMK knockdown reduced the number of LDs in WT cells and abolished the increased LD phenotype in UFD1s MUT cells (Fig. 5c, d). Furthermore, IPMK knockdown partially but significantly restored the decreased FAO rate shown in human and mouse UFD1s MUT cells (Fig. 5e, f). To elucidate the molecular mechanism by which IPMK regulates energy metabolism, we attempted to identify the key downstream effectors of IPMK. AMP-activated protein kinase (AMPK) is known as a highly conserved central sensor of energy metabolism under stress conditions, and is implicated in multiple featured mechanisms of autophagy and mitochondrial function⁷¹. We observed that IPMK knockdown led to increased levels of active phosphorylated AMPK in human and mouse cells (Fig. 5g). AMPK can promote peroxisome proliferator-activated receptor α (PPARα) and peroxisome proliferator-activated receptor γ coactivator 1α (PGC-1α) to regulate mitochondrial function, including oxidative phosphorylation (OXPHOS) and fatty acid/lipid metabolism^71,72. Upregulated levels of the PPARα and PGC-1α proteins were observed after AMPK activation in IPMK-knockdown cells (Fig. 5g, h). These data indicated that IPMK is a critical UFD1s functional target that suppresses autophagy, stimulates lipid accumulation by regulating FAO, and functions through the inhibition of AMPK activation.

Metabolic phenotypes of Ufd1s^−/− mice

To explore the physiological roles of UFD1s in animals, we applied the CRISPR/Cas9 technique to generate Ufd1s-deficient (Ufd1s^−/−) mice, in which the A5SS of Ufd1 was mutated and the amino acid sequence of the Ufd1f protein remained unchanged (Fig. 6a). The Ufd1s mRNA and protein levels were significantly decreased in Ufd1s^−/− mice, whereas the Ufd1f mRNA and Ufd1f protein levels were not altered (Supplementary Fig. 8a, b). Under normal conditions, Ufd1s^−/− mice presented lower energy expenditure, less oxygen consumption, and a higher respiratory exchange rate (RER) than did wild-type (WT) mice (Supplementary Fig. 8c). Carbon dioxide production, food uptake, water uptake, and spontaneous movement were unchanged (Supplementary Fig. 8d).

**Fig. 6: Ufd1s-deficiency leads to liver defects and exacerbates NASH progression.**

The liver, an organ with a central role in metabolic processes⁷³, was then examined. The Ipmk protein levels were significantly increased in the liver of Ufd1s^−/− mice (Fig. 6b). On the other hand, starvation led to reduced Ipmk protein levels in WT mouse livers (Fig. 6c). Ufd1s deficiency led to abnormal liver morphology, including mild watery degeneration, swelling, and cytosolic looseness of hepatocytes, as shown by hematoxylin‒eosin (H&E) staining, which was used to measure the area of the nucleus relative to that of the cytoplasm, and abnormalities were more severe after starvation for 24 h (Fig. 6d). Immunohistochemical (IHC) staining of the autophagy marker LC3 revealed significantly decreased signals in Ufd1s^−/− livers under both normal and starvation conditions (Fig. 6d). Starvation is well known to induce autophagy⁷⁴. Moreover, compared with WT mouse livers, Ufd1s^−/− mouse livers presented more LDs via Oil red O staining under a normal food supply, and the increase in the number of LDs became more dramatic after starvation (Fig. 6d). The total and phosphorylated ACC levels were unchanged in Ufd1s^−/− mouse livers (Supplementary Fig. 8e), similar to the results in Ufd1s MUT cells (Supplementary Fig. 6g). To further reveal the anti-stress function of Ufd1s in hepatocytes, we examined the ROS and ATP levels in primary hepatocytes from Ufd1s^−/− mice under GD conditions. Compared with WT cells, primary hepatocytes from Ufd1s^−/− mice presented higher ROS and lower ATP levels (Supplementary Fig. 8f). OCR assays assessing FAO in primary hepatocytes revealed that Ufd1s^−/− cells exhibited significantly lower OCR than WT cells (Supplementary Fig. 8g). These results demonstrated that Ufd1s also has functions in anti-stress responses and the regulation of lipid metabolism in primary hepatocytes. MARCH7-mediated ubiquitination of UFD1f regulated the stability of IPMK; thus, we explored whether MARCH7 contributes to lipid accumulation in the liver. Cell cultures under hepatic steatosis conditions via free fatty acid induction were established^75,76, and MARCH7 knockdown alleviated the accumulation of LDs in human HepG2 and mouse Hepa1-6 cell cultures under free fatty acid induction (Supplementary Fig. 8h).

Together, these results demonstrated that the Ufd1s^−/− mice presented phenotypes associated with energy metabolism and stress responses. These phenotypes are consistent with the abnormalities observed in mammalian cell lines with Ufd1s deficiency. Ufd1s is necessary for the ability of animals to cope with intrinsic stresses.

Ufd1s deficiency accelerates NASH progression

We further examined the role of Ufd1s under pathogenic stress by inducing NASH in mice fed a methionine- and choline-deficient (MCD) diet (Fig. 6e), which led to a deficiency in phosphatidylcholine synthesis, increased oxidative stress, and subsequent liver damage^77,78. Serum triglyceride and total cholesterol levels were elevated in Ufd1s^−/− mice compared with WT mice under normal and NASH conditions (Fig. 6f). The serum levels of aspartate aminotransferase (AST) and alanine aminotransferase (ALT), which are markers of liver injury, were also measured. The serum AST levels of Ufd1s^−/− mice were higher than those of WT mice under normal and NASH conditions, while the ALT levels were not significantly different (Fig. 6g). H&E staining revealed that the Ufd1s^−/− livers exhibited more severe steatosis and a greater number of LDs compared to WT livers (Fig. 6h). Masson staining to assess extracellular matrix accumulation revealed more severe fibrosis in Ufd1s^−/− livers compared to WT controls under NASH conditions (Fig. 6h)⁷⁹. IHC analysis revealed that Ipmk protein levels were markedly higher, while the autophagy marker LC3 levels were significantly lower in Ufd1s^−/− mouse livers than WT mice under both normal and NASH conditions (Fig. 6h). We collected 8 pairs of para-hepatocellular carcinoma tissues from patients, with HE analysis diagnosed with non-NASH or NASH. IHC results showed that the expression of UFD1s was significantly decreased in NASH samples, compared to those in non-NASH samples (Fig. 6i and Supplementary Fig. 8i).

Immune and inflammatory pathways play crucial roles in the pathogenesis of NASH⁸⁰. A broad spectrum of immune cells, including macrophages, natural killer (NK) cells, B cells, and T cells, are found in the liver and contribute to NASH progression^80,81. Multiplex immunofluorescence (mIF) assays revealed that infiltration of macrophages, natural killer (NK) cells, B cells and T cells remained unchanged between WT and Ufd1s^−/− livers under normal conditions, whereas under NASH conditions, the density of NK cells was significantly decreased, and that of B cells was increased in Ufd1s^−/− mouse livers (Supplementary Fig. 9a). Ufd1s deficiency in mice led to increased levels of IL-6 and TNF (as inflammation indicators) in NASH conditions, while these levels were not significantly changed in Ufd1s^−/− livers under normal conditions (Supplementary Fig. 9b). Consistently, the liver of Ufd1s^−/− mice had a greater degree of inflammation in the NASH condition (Supplementary Fig. 9c). Collectively, these results demonstrated that the UFD1s expression was linked to inflammation inhibition in NASH conditions.

Ufd1s possesses medical value in treating NASH

To provide further insights into the therapeutic potential of Ufd1s, we employed two strategies, plasmid injection and circRNA administration, to express the Ufd1s protein in a NASH model (Fig. 7a). For plasmid DNA, the mouse Ufd1s ORF was introduced into the pLIVE vector (pLIVE-Ufd1s), which was driven by a liver-specific promoter and led to liver gene expression in mice⁸². The control plasmid and pLIVE-Ufd1s were injected hydrodynamically into the mice once via the intravenous tail vein (Fig. 7a). A circRNA with the mouse Ufd1s coding sequence along with an internal ribosome entry site (IRES) (circUfd1s) was synthesized in vitro, and a circRNA with the start codon of Ufd1s mutated to ATT (circUfd1s^ATG-ATT) was used as a negative control (Supplementary Fig. 9d). CircRNAs enveloped in Lipofectamine 3000 were delivered into mice twice through tail vein injection (Fig. 7a). IHC staining of mouse livers demonstrated that the Ufd1s levels were significantly higher in the pLIVE-Ufd1s and circUfd1s groups than in the corresponding controls (Fig. 7b). Administration of either pLIVE-Ufd1s or circUfd1s led to reduced serum triglyceride and total cholesterol levels (Fig. 7c). Serum AST and ALT levels were clearly decreased upon pLIVE-Ufd1s injection, whereas AST but not ALT levels were reduced in the circUfd1s-injected group compared with those in the control group (Fig. 7d). Compared with the corresponding controls, both strategies of Ufd1s overexpression in the NASH model resulted in less severe steatosis, fewer LDs, less fibrosis, lower Ipmk levels, higher autophagic LC3 levels, and lower-grade inflammation in the liver (Fig. 7e and Supplementary Fig. 9e). Whereas glucose tolerance and insulin tolerance remained unchanged in the NASH model after Ufd1s overexpression, indicating that the glucose metabolism was not affected (Supplementary Fig. 9f, g). Collectively, Ufd1s has the potential to serve as a therapeutic drug for NASH.

**Fig. 7: Application of Ufd1s-coding plasmid or circRNA alleviates NASH progression.**

Discussion

Accumulating evidence has demonstrated that prolonged exposure to cellular stresses such as ER stress, mitochondrial stress, heat shock, and oxidative stress contributes to human diseases such as metabolic disorders and cancers^1,4,5,83. The underlying mechanisms involved in the response to cellular stresses are crucial to elucidate. Here, we report an anti-stress microprotein, UFD1s, that promotes autophagy and lipid metabolism by modulating the ubiquitination dynamics of key factors in mammals (Fig. 8), expanding our understanding of cellular stress responses and providing a new strategy for developing treatments for human diseases.

The regulation of alternative splicing is one of the core mechanisms in eukaryotic stress responses and has been linked to diseases and therapeutics^59,84,85,86. It has been over two decades since the discovery of the unconventional cytoplasmic splicing mechanism that generates XBP1s under cellular stress, while the discovery of other SR-mRNAisos with key regulatory roles has been lacking. Unlike the formation of XBP1s, the UFD1s variant is generated by alternative splicing from UFD1 pre-mRNA, which is regulated by AQR and U2AF65 (Fig. 1d and Supplementary Fig. 1e). Both AQR and U2AF65 are involved in multiple alternative splicing events^55,56,57. UFD1s may not be eukaryotic conserved as XBP1s is, as we failed to detect the existence of UFD1s in yeasts (Supplementary Fig. 2c, d). Interestingly, UFD1s regulates the ubiquitination of UFD1f through competing for the E3 ligase MARCH7 (Fig. 3e–g). Presumably, the same sequences shared by UFD1s and the N-terminus of UFD1f are the reasons for this interference. UFD1s is a microprotein derived from a short splicing variant, and in plants, some microproteins generated from alternative splicing or gene duplication have been found to interfere with the function of the full-length protein and are therefore termed small interfering peptides⁸⁷. No small interfering peptides have been found in animal cells⁸⁸, and the UFD1s discovered in this study may be recognized as an animal small interfering peptide. In comparison, the role of XBP1s is not directly related to that of XBP1u, the unspliced form of XBP1^6,7. UFD1s is not only induced under external stresses like XBP1s, but also expressed under normal physiological conditions at background levels, and our data show that UFD1s is required for the response to both internal and external stresses (Figs. 2 and 6). UFD1s is subjected to the identification of protein translational modifications (PTMs), and exploring the possible coordination between PTMs and the anti-stress roles of UFD1s would be interesting. Future investigations should also examine whether the other SR-mRNAisos identified in this study that are not as conserved as XBP1s and UFD1s may also play important roles in stress responses (Fig.1b and Supplementary Data 1).

Dynamics in protein ubiquitination constitute another core molecular feature of stress responses^2,4. The classical K48-ubiquitination that promotes protein degradation is relatively well studied in stressed cells to remove unwanted and deleterious proteins^12,13. Our study unexpectedly reveals the roles of less studied K63- and K11-linked ubiquitination in anti-stress responses, and K63-linked ubiquitination is proved to be the major (if not the only) contributor to the regulation of UFD1f total ubiquitination, although the abundance of K63-linked ubiquitination is less than that of K48-linked ubiquitination in cells. It is also reasonable to speculate that the regulation of UFD1f ubiquitination by UFD1s is a complex process that is potentially orchestrated by multiple factors, and in the anti-stress responses, protein ubiquitination plays a larger role than just the degradation of surplus or damaged proteins via K48-linked ubiquitination. UFD1f with or without K63-linked ubiquitination seems to have distinct functions. In the regulation of IPMK ubiquitination, UFD1f with K63-ubiquitination favors K11-ubiquitination and stabilizes IPMK, whereas UFD1f without K63-ubiquitination favors K48-ubiquitination and IPMK degradation (Fig. 4h, i). Changes in the protein levels and ubiquitination status of IPMK upon MARCH7 knockdown also indicated that UFD1s-modified UFD1f K63 ubiquitination potentially regulates the ratio of K48- to K11-linked ubiquitination of IPMK (Fig. 4k), further confirming that the ratio of K48- to K11-linked ubiquitination of IPMK is precisely orchestrated by conditions. The role of UFD1f as a component of the UFD1-NPL4 adapter complex in proteasome-mediated protein degradation has been better studied, but how changes in the K63 ubiquitination of UFD1f modulate the ubiquitination of hundreds of proteins, including IPMK, requires further exploration.

At the cellular level, we demonstrate that UFD1s regulates autophagy and lipid metabolism. Both differentially expressed genes and proteins with significant alteration in ubiquitination in the UFD1s deficient cells are enriched for GO terms related to autophagy and lipid metabolism (Figs. 2i and 4c). As a preserved adaptive response to environmental changes, autophagy is critical for regulating cell survival and death, while lipid metabolism maintains energy and cellular homeostasis, and both of which are essential elements of material and energy homeostasis, and are fine-tuned in anti-stress responses^46,66. Even in normal conditions, UFD1s is essential to ensure balanced autophagy and lipid metabolism in cells against internal stresses (Fig. 2j–m, Supplementary Figs. 5 and 6). In mice under 24-hour starvation, which is a mild nutrient stress, Ufd1s is also essential to keep liver lipid storage and autophagy in check (Fig. 6d). In starvation-stressed cells, lipid droplet formation is known to buffer autophagy-derived lipids, protect mitochondria from lipotoxic damage, and store lipids for future consumption^64,89. In the past decade, significant progress has been made in comprehending the crosstalk between autophagy and lipid metabolism. For instance, autophagy can be induced by inhibiting biosynthesis of fatty acids or by increasing biosynthesis of cell membrane lipids⁹⁰. Conversely, inhibition of autophagy increases lipid storage in both cultured cells and mice⁹¹. We provide some insights about how protein ubiquitination regulates autophagy and lipid metabolism, yet the interplay of ubiquitination, autophagy, and lipid metabolism in response to stresses requires further elucidation.

Stress-induced or accelerated diseases such as NASH have become an increasing health problem, and NASH presents disordered metabolic homeostasis characterized by dysregulation of glucose, lipid, and bile acid metabolism⁹². To date, no effective NASH drug has been approved by the FDA or any other leading agencies^{50,53,93,94,95}. When under stresses that cause NASH, Ufd1s is essential in protecting the liver from fat storage and promoting autophagy (Fig. 6h). Interestingly, a recent study has shown that XBP1 promotes NASH progression, and XBP1 inhibitors have protective effects in NASH mice⁹⁶. Despite the fact, this study examines the XBP1 gene as a whole, and a more specific investigation for the roles of XBP1s in NASH may help to understand this stress-induced isoform. For NASH, a mitochondrial noncoding circRNA, SCAR, has been found to mitigate insulin resistance and cirrhosis, and downregulation of SCAR is associated with disease progression⁵¹. CircRNAs with translation capability also hold potential as protein substitution, including a recent effort in developing circRNA vaccine against COVID-19⁹⁷. UFD1 and its variants have not been reported to be associated with metabolic diseases such as NASH. In this study, Ufd1s expressed with either plasmid DNA or circRNA alleviates NASH progression in mice (Fig. 7). Considering that Ufd1s protects cells from both internal and external stresses, we hope this microprotein can be developed into effective drugs to treat NASH, or furthermore, to protect cells from stresses in other metabolic diseases.

Methods

Human study and approval

All clinical samples (non-NASH controls and NASH liver specimens) were hepatocellular carcinomas adjacent normal tissues embedded in paraffin, and were obtained from the First Affiliated Hospital of USTC. The study was approved by the Institutional Review Board of the First Affiliated Hospital of USTC (2024-RE-491). All tissue samples were collected in compliance with the informed consent policy. Patient information is provided in Supplementary Table 1.

Mice

Ufd1s-deficient (Ufd1s^−/−) mice were generated by the Laboratory Animal Research Center of USTC with the CRISPR/Cas9 system⁹⁸. Briefly, Cas9 mRNA and the corresponding sgRNA (5′-TGCTAGCAGGGCCTAATGAC-3′) were synthesized with mMESSAGE mMACHINE^TM T7 ULTRA Transcription Kit (Invitrogen, AM1345) and MEGAshortscript^TM T7 Transcription Kit (Invitrogen, AM1354), respectively, and purified with MEGAclear^TM Transcription Clean-Up Kit (Invitrogen, AM1908). The single-stranded oligodeoxynucleotide (ssODN) containing the mutated splice sites (AGGTCA to CGCAGC, also a synonymous mutation for Ufd1f) and flanking homologous sequences (119 nucleotides) were synthesized (Sangon). Then, Cas9 mRNA/sgRNA and ssODN were microinjected into zygotes from C57BL/6 mice. All mice were genotyped with PCR amplification at 2 weeks after birth, and the synonymous mutation was confirmed by Sanger sequencing. Specific PCR primers were listed in Supplementary Table 2. All Ufd1s^−/− mice for further analyses were in parallel with gender- and age-matched wild-type littermates as a control group.

All mice were bred and housed in a special pathogen-free (SPF) facility maintained in a 12/12-h light/dark cycle (lights from 8:00 to 20:00) at a temperature of 23–25 °C with 40–60% humidity. Maximal five mice were housed in a cage and had free access to water and food for the duration of the experiment, unless specified in the corresponding situation.

For the 24-h starvation treatment, mice were kept in the environment (23–25 °C with a 12-h light-dark cycle and lights on at 8:00 AM) with free access to water but no food supply at 9:00 PM on the first day followed by continued starvation for 24 h, and fasted again for 12 h before bleeding and sacrifice at 9:00 AM on the third day. All control mice were fasted for 12 h before bleeding and sacrifice at 9:00 AM.

To evaluate metabolic physiology, including total energy expenditure, O₂, CO₂, RER, food uptake, and water uptake, and spontaneous movement, mice were randomly placed into the Promethion Metabolic Cage System (Sable Systems International) and acclimated to individual cage for a week prior to data collection with normal feeding and living conditions. The Cage System fitted with indirect open circuit calorimetry and food consumption and activity monitors to measure activity, food uptake, and energy expenditure. O₂, CO_2, and water uptake were measured with the GA-3 Series (GA3mX) gas analyzer. Respiratory exchange rate (RER) was calculated as VCO₂/VO₂. And then data from a continuous 24-h period were used for analysis.

All animal procedures were approved by the Animal Care and Use Committee of University of Science and Technology of China, and performed in accordance with the approved guidelines (USTCACUC202101048). Male mice (9-week-old) were used and randomly assigned to experimental groups in this study, due to the fact that males are more prone to develop metabolic disorders in response to nutritional challenges⁹⁹.

NASH model

For inducing NASH mice model, methionine and choline-deficient diet (MCD diet, purchased from Jiangsu Synergy Pharmaceutical Biological Engineering Co., Ltd) was used. Mice (9-week-old) were starved overnight followed by feeding MCD diet for 4 weeks. All mice were kept in a controlled environment (23–25 °C with a 12-h light-dark cycle and lights on at 8:00 AM) with free access to water.

Yeast strains

The yeast strains Saccharomyces cerevisiae BJ2168 (MATa his3∆1 leu2∆0 met15∆0 ura3∆0) and Schizosaccharomyces pombe PR109 (h- leu1-32 ura4-D18) were kindly provided by Dr. Gang Cai (USTC) and were cultured with the YPD and YES liquid mediums in the shaker at 30 °C.

Cell culture

HEK293T, HeLa, N2a, Hepa1-6, and HepG2 cells were purchased from the American Type Culture Collection (ATCC). SGC-7901 and AGS cells were kindly provided by Dr. Wenchu Lin (The Second Affiliated Hospital, The Chinese University of Hong Kong). All above cells were cultured under standard conditions with DMEM containing 10% FBS, 1% penicillin/streptomycin at 37 °C under 5% CO₂. Cells were checked with a PCR-based method and DAPI staining to ensure no contamination of mycoplasma. All cell lines were authenticated by short-tandem-repeat (STR) profiling.

Library preparation for RNA-seq

Total RNAs from HEK293T and N2a cells were extracted using TRIzol reagent (Invitrogen) according to the manufacturer′s instructions. The ribo-minus libraries were performed with the TruSeq Ribo Profile Library Prep Kit (Illumina) according to the manufacturer′s instructions and then subjected to 150-nt paired-end sequencing generating ~100 million read pairs with an Illumina Nextseq 500 system (Novogene).

Stress responsive mRNA isoform (SR-mRNAiso) identification

Identification of SR-mRNAisos was performed as previously described with several modifications⁵⁷. Briefly, the adapters were trimmed with Cutadapt (-e 0.1 -O 5) to obtain clean reads. Reads continuously aligned to the reference genomes (human: hg19; mouse: mm10) were filtered out with Bowtie (-v 2 -m 20 -best). The remaining reads were aligned to the reference genomes using Blat with at least a 5-nt overhang and obtained the correspondent split junctions. The split junctions were further annotated to genomic locus and the junction reads per million (RPM) were calculated for SR-mRNAisos. Then, the differentially expressed (DE) SR-mRNAisos were determined by DEseq2 with the cutoff (Fold change >2 or <0.5, P value < 0.05). For the conservation analysis of SR-mRNAisos, exon sequences (20-nt upstream and 20-nt downstream) flanking the split junction were aligned between human and mouse, and the identity were defined as conservation scores.

Differentially expressed (DE) genes analysis

For data processing, clean reads were obtained after trimming the adapters with Cutadapt (-e 0.1 -O 5), and were subsequently aligned to the human reference genome (hg19) with Bowtie2 (-v 1). For mRNAs, the corresponding reads were counted with BEDtools and read per million (RPM) was used to calculate mRNA levels. The DE genes were determined by DEseq2 with the cutoff (RPM > 1, fold change >2 or <0.5 and P value < 0.01).

Gene Ontology (GO) analysis

GO analysis was performed by GOrilla web-server with default parameters¹⁰⁰, and was visualized by the ggplot2 package in R software.

eCLIP-seq data analysis

Published eCLIP-seq data of 103 RBPs (HepG2) and 120 RBPs (K562) were obtained from ENCODE. 26-nt spliced sites in UFD1s versus UFD1f (GTCAGATGTGGAGAAAGGAGGGAAGA) along with 100-nt genomic flanking sequences were used for searching eCLIP binding signals of RBPs. RBPs with binding signals (RPM ≥ 2) for the 226-nt sites in both cell lines were considered as candidates to regulate UFD1s formation.

Plasmid construction and cell transfection

All plasmids were constructed with restriction enzyme digestion and ligation, or with a recombinant method (Vazyme). The p3×FLAG-Myc-CMV24 backbone vector (Sigma, E9283) was used for overexpressing UFD1s, IRE1α, UFD1f, UFD1f mutants, and IPMK. For human UFD1s and mouse Ufd1s, the FLAG tag was constructed to the C terminus of the corresponding UFD1s open reading frame (ORF); for human Flag-UFD1f and UFD1f-Flag plasmids, the FLAG tag was constructed to the N terminus and C terminus of UFD1f ORF, respectively. The coding sequence of human IRE1α was constructed into the backbone between HindIII and BamHI double-digested sites. Human and mouse IPMK were inserted into the C terminus of Myc tag to generate Myc- IPMK fusion proteins. Mouse March7 were constructed into the pKH3 backbone (Sigma, 12555) vector between BamHI and EcoRI. Mouse Ufd1s ORF with a FLAG tag was inserted into the liver-specific expressed pLIVE vector (Mirus Bio, MIR 5420) between NheI and SacI. All constructions were sequenced for confirmation. The shRNA plasmids for knockdown of human AQR (shAQR, TRCN0000074872), human DDX3X (shDDX3X, TRCN0000000003), human NCBP2 (shNCBP2, TRCN0000059995), human TIA1 (shTIA1, TRCN0000074464), human U2AF65 (shU2AF65, TRCN0000001162), human MARCH7 (shMARCH7, TRCN0000073068), human IPMK (shIPMK, TRCN0000195137), mouse March7 (shMarch7, TRCN0000182847), mouse Ipmk (shIpmk, TRCN0000024840) with negative-control shRNA (shCOO2, MFCD07785395) was obtained from the MISSION shRNA Library (Sigma). Further information about plasmids is available upon request. Transfection of plasmids, siRNAs, and shRNAs was performed with Lipofectamine 2000 (Invitrogen, 11668019) for HEK293T, N2a, SGC-7901 and AGS cells or Lipofectamine 3000 (Invitrogen, L3000015) for HepG2 and Hepa1-6 cells according to the manufacturer′s protocol. Oligonucleotide sequences for primers used in plasmid construction, shRNAs and siRNAs are included in Supplementary Table 2.

Single molecule fluorescence in situ hybridization (smFISH)

The smFISH of UFD1f and UFD1s RNAs in HEK293T cells was performed¹⁰¹. Briefly, cells were fixed in 4% PFA for 10 min at room temperature, washed with DPBS buffer twice, and permeabilized with ice-cold 70% ethanol overnight at −20 °C. For hybridization, cells were washed with 2×SSC twice, pre-hybridized at 37 °C for 30 min in hybridization buffer (30% formamide, 5×SSC, 9 mM citric acid, pH 6.0, 0.1% Tween 20, 50 μg/mL heparin, 1×Denhardt′s solution and 10% dextran sulfate), and incubated with 4 nM denatured probes overnight at 37 °C in hybridization buffer. Then, cells were washed with the wash buffer (30% formamide, 5×SSC, 9 mM citric acid, pH 6.0, 0.1% Tween 20, 50 μg/mL heparin) four times at 37 °C and 5×SSCT (5×SSC, 0.1% Tween 20) twice at room temperature. For amplification, cells were pre-amplified at room temperature for 30 min in the amplification buffer (5×SSC, 0.1% Tween 20, 10% dextran sulfate). 3 μM individual hairpin was heated at 95°C for 90 s and cooled to room temperature for 30 min, protected from light. Cells were incubated in the amplification buffer with prepared hairpins overnight (12–16 h) at room temperature followed by three 5 min and two 30 min washes in 5×SSCT. Finally, nuclei were stained with DAPI (Sigma, F6057). The images were captured using the LSM 980 confocal microscope (Zeiss) and further processed with Image J, and the ImageJ Plot Profile tool was used to calculate signal intensity. All probe sequences are included in Supplementary Table 2.

Cellular stress, cycloheximide (CHX), and NMDI14 treatments

To induce ER stress, human and mouse cells were incubated with 300 nM TG (Sigma, T9033) for 6 h and 12 h for RNA examination and protein analysis, respectively. For yeasts, ER stress was induced with 5 μg/ml Tm (Sigma, T7765) in the corresponding liquid mediums for the indicated time. For glucose deprivation (GD), cells were cultured in the DMEM medium without glucose (Gibco, 11966025) at 37 °C for 24 h. For total ROS and ATP levels detection, HEK293T, N2a, and primary hepatocytes were treated with GD for 6 h. For cell survival analysis, HEK293T, N2a, AGS, and SGC-7901 cells were treated with GD for 12 h. For glutamine deprivation or HBSS-induced nutrient starvation, cells were incubated with the DMEM medium without glutamine (Gibco, 10313021) or the Hank′s Balanced Salt Solution (HBSS) (Gibco, 14025092) at 37 °C for 24 h, respectively. For autophagy, cells were treated with 50 μM chloroquine (CQ) (Sigma, C6628) for 10 h. For the cycloheximide (CHX) chase experiment, cells were treated with 20 μg/mL CHX (Sigma, C7698) for 4 h. For inhibiting the nonsense mediated RNA decay (NMD) pathway, cells were treated with 50 μM NMDI14 (MCE, HY-111374) for 6 h.

5′ and 3′ rapid amplification of cDNA ends (RACE)

3′ RACE and 5′ RACE of human UFD1s were performed with the SMARTer RACE cDNA Amplification kit (Takara, 634858) following the manufacturer′s instructions. 5′ RACE of the SMARTer RACE cDNA Amplification kit relies on the terminal transferase activity of the RT enzyme, which adds 3–5 residues to the 3′ end of the first-strand cDNA, thus ensuring amplification of the full 5′ end of the RNA. 3′ RACE with the SMARTer RACE cDNA Amplification kit relies on the 3′ poly(A) tail of the RNA. The primers were listed in Supplementary Table 2.

PCR reactions

RNA was extracted using TRIzol reagent according to the manufacturer′s protocol. 500 ng RNA was reverse-transcribed into complementary DNA (cDNA) with GoScript Reverse Transcription System (Promega, A5000) according to the protocol. For semi-quantitative RT-PCR gels, 25-30 cycles were always carried out with 2×Taq PCR mix (TransGen, AS122). Real-time quantitative PCR (RT-qPCR) was performed with GoTaq SYBR Green qPCR Master Mix (Promega, A6001) according to standard procedures, normalized to ACTB mRNA. All PCR products were sequenced for confirmation. All used primer information was included in Supplementary Table 2.

Western blotting

For sample preparation, cells or tissues were homogenized in RIPA buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 5 mM EDTA, 1 mM DTT, 1% NP-40, 0.1% SDS, and 1×protease inhibitor cocktail (Roche, 05892970001)). For western blotting, the lysates were separated on SDS-PAGE gels and transferred onto nitrocellulose membranes (PALL). Membranes were processed according to the ECL Western blotting protocol (GE Healthcare). Images were taken using the ImageQuant LAS4000 Biomolecular Imager (GE Healthcare). The gray-scale statistics of western blotting were performed by Image J. The following antibodies were used in this study: anti-FLAG (Sigma, F1804, 1:2000), anti-β-ACTIN (TransGen, HC201, 1:2000), anti-UFD1s (human) (this study, 1:500), anti-Ufd1s (mouse) (this study, 1:500), anti-UFD1f (Proteintech, 10615-1-AP, 1:1000), anti-total Ub (CST, 3936), anti-K48 (CST, 8081, 1:1000), anti-K63 (CST, 5621, 1:1000), anti-K11(Invitrogen, PA5-120621, 1:1000), anti-K6 (Abclonal, A18106, 1:500), anti-MARCH7 (Santa Cruz, sc-166945, 1:500), anti-IPMK (Immunoway, YT2384, 1:1000), anti-HA (TransGen, HT301, 1:2000); anti-Myc (Proteintech, 60003-2-Ig, 1:2000), anti-LC3 (Proteintech, 14600-1-AP, 1:1000), anti-P-AMPK (CST, 2535, 1:1000), anti-AMPK (CST, 5832, 1:1000), anti-PGC-1α (Abclonal, A20995, 1:1000), anti-PPARα (Immunoway, YM8234, 1:1000), anti-p-ACC (CST, 3661, 1:1000), anti-ACC (CST, 3662, 1:1000), anti-GAPDH (EnoGene, E12-052, 1:2000).

Anti-UFD1s antibody preparation

Human and mouse UFD1s peptides synthesis and anti-UFD1s specific antibodies preparation were carried out by GL Biochem (Shanghai). In brief, KLH-coupled peptides Cys-SNYATLGPGPTQPT (human) and Cys-SNYATLSPRSTQPAQHYLSY (mouse) were synthesized, respectively. Polyclonal antibodies against the UFD1s peptide were obtained from two inoculated rabbits. Antibodies were purified using affinity chromatography on columns containing the corresponding peptides and antibodies titer was detected by ELISA.

Immunofluorescence (IF) staining

Cells were fixed with 4% PFA at room temperature for 15 min and permeabilized with PBS, 1% Triton X-100 on ice for 20 min. After washing with PBST (PBS, 0.4% Tween 20) twice, they were blocked with blocking buffer (PBST, 1% BSA) at room temperature for 30 min and incubated with the diluted antibodies at room temperature for 4 h. Subsequently, cells were incubated with the corresponding Alexa Fluor 488- or 546-conjugated secondary IgG antibodies in the dark for 1 h at room temperature after washing with PBST twice. Nuclei were stained with DAPI and the images were acquired with Zeiss LSM 980 confocal microscope. Antibodies used in the IF staining are as follows: anti-UFD1s (human) (this study, 1:200), anti-Ufd1s (mouse) (this study, 1:200), anti-UFD1f (Proteintech, 10615-1-AP, 1:500), anti-LC3 (Proteintech, 14600-1-AP, 1:500), anti-PGC-1α (Abclonal, A20995, 1:500), anti-PPARα (Immunoway, YM8234, 1:500), Goat anti-Rabbit IgG Secondary Antibody, Alexa Fluor 488 (Invitrogen, A-11008, 1:500), Goat anti-Rabbit IgG Secondary Antibody, Alexa Fluor 546 (Invitrogen, A-11010, 1:500), Goat anti-Mouse IgG Secondary Antibody, Alexa Fluor 488 (Invitrogen, A-11001, 1:500).

CRISPR/Cas9-mediated UFD1s MUT cells

For UFD1s-deficient (MUT) HEK293T and N2a cells, a single-guide RNA (sgRNA) targeting human and mouse UFD1s (5′-TGCTAGCAGGGCCTAATGAC-3′) was designed and cloned into the backbone px330-mCherry (Addgene, 98750). The mutated splice sites (AGGTCA to CGCAGC) for UFD1s and flanking homologous sequences were inserted into the vector pcDNA3. Firstly, 500,000 cells were seeded into 6-well plates and transfected with 1 μg px330-mCherry expressing Cas9 mRNA and sgRNA, and 1 μg linearized pcDNA3 (Invitrogen, V79020) with homologous sequences. Forty-eight hours later, hundreds of individual mCherry-positive colonies were generated into 96-well plates with the BD FACSAria™ III cell sorting system (Beckman). Finally, we performed a PCR-mediated genotype and confirmed the corresponding products by Sanger sequencing. The primer sequences are presented in Supplementary Table 2.

MTT assay and cell survival assay

MTT assay was performed with MTT cell proliferation and Cytotoxicity Detection Kit (KeyGEN, KGA311) according to the manufacturer′s instructions. The 490 nm absorbance was measured by Multiskan Go microplate spectrophotometer (Thermo). Cell survival was detected via propidium iodide (PI) (Beyotime, ST511) and Hoechst 33342 (Sigma, B2261) double staining for 20 min at 37 °C. Cells images were taken by the Olympus IX81 inverted microscope.

Mass spectrometry (MS) to identify proteins

Protein identification by mass spectrometry (MS) was performed at Shanghai Applied Protein Technology Co., Ltd. Specific silver-stained bands, coomassie blue-stained gels or mixture solutions were digested, extracted, purified and identified. For sample preparation, gel samples were destained with destaining solution (30% ACN/100 mM NH₄HCO₃) until transparent, followed by reduction (100% ACN with 10 mM DTT, 37 °C for 1 h) and alkylation (60 mM IAA, room temperature in dark for 1 h). After washing with 100 mM NH₄HCO₃ (room temperature in dark, 2 × 15 min), samples were lyophilized for 10 min and then digested with 2.5–10 ng/μL trypsin or chymotrypsin at 37 °C for 20 h (chymotrypsin for UFD1s identification; trypsin for remaining all). Peptides were extracted with 60% CAN/0.1% TFA (15 min sonication), lyophilized and redissolved with 2% FA. While solution samples were reduced with 100 mM DTT at 95 °C for 5 min, buffer-exchanged using UA buffer (8 M Urea, 150 mM Tris‐HCl, pH 8.5) in 10 kDa centrifugal filter units, and alkylated with 100 mM IAA (room temperature in dark for 30 min). Trypsin digestion (4 μg in 25 mM NH₄HCO₃) proceeded at 37 °C for 16–18 h. The digested peptide mixtures were separated by nanoElute system (Bruker) on a column (Thermo EASY column, 25 cm, 3 μm, C18), loaded with 0.1% FA/water (A), and separated with 0.1% FA/ACN (B) using a gradient time of 60 min at a flow rate of 300 nL/min: 0–60% B (50 min), 60–90% B (4 min), 90% B (6 min). LC‐MS/MS analysis was performed on a timsTOF Pro mass spectrometry (Bruker) and operated in PASEF mode (positive ion, 1.5 kV) with m/z 100–1700 range, using 1 MS and 8 MS/MS PASEF scans per 0.95 s cycle; ion charge: 0–5; active exclusion was enabled with a release time of 24 s; automatic gain control (AGC) target was set to 1e6. For data processing, raw data analyzed with MaxQuant1.6.14 against the UniProt database (Homo sapiens) or sequences of UFD1s proteins with following parameters, enzyme: trypsin (cleavage sites: lysine (K) and arginine (R)) or chymotrypsin (cleavage sites: phenylalanie (F), leucine (L), tryptophan (W) and tyrosine (Y)); max missed cleavages: 2; fixed modification: carbamidomethyl (C); variable modifications: oxidation (M), GlyGly (K); peptide mass tolerance: 20 PPM; fragment mass tolerance: 0.1 Da; database pattern: reverse; peptide/protein/site FDR ≤ 0.01. For the identification of UFD1s peptides, co-immunoprecipitation of UFD1s and UFD1f, no biological replicates were used.

Immunoprecipitation of human and mouse UFD1s peptides for MS

For the identification of UFD1s peptide fragments, whole HEK293T or N2a cell lysates were lysed with IP buffer (50 mM Tris-HCl, pH 8.0, 150 mM NaCl, 1.5 mM MgCl₂, 1 mM EDTA, 0.5% NP-40, 10% glycerol, 4 mM DTT and 1×protease inhibitor cocktail) on ice for 20 min. The lysis was collected and sonicated for 5 min with the Sonics Vibra-Cell (3 s on, 6 s off, 30% amplitude). After centrifugation at 13,000 rpm for 10 min at 4 °C, the supernatant was incubated with 5 μg UFD1s antibodies or normal rabbit IgG (CST, 2729) coupled to protein G magnetic beads (Invitrogen) overnight at 4 °C in a rotating wheel. After four 5-min washes with the IP buffer, the purified beads were boiled with SDS loading buffer for 10 min. Then the mixture was separated and the gel was performed by silver staining with Protein Stains K kit (Sangon, C500021). The specific silver-stained bands were then cut for mass spectrometry and identified peptide sequences were searched against the sequence of human and mouse UFD1s proteins, respectively.

Co-immunoprecipitation (Co-IP)

HEK293T or N2a cells were harvested in ice-cold IP buffer for 30 min on ice. The supernatant was collected after centrifugation at 1000 × g for 5 min at 4 °C, and was sonicated for 5 min with the Sonics Vibra-Cell (3 s on, 6 s off, 30% amplitude) on ice. After centrifugation at 13,000 rpm for 10 min at 4 °C, the supernatant was incubated with 3 μg specific antibodies or normal IgG coupled to protein G magnetic beads overnight at 4 °C in a rotating wheel. After two 5-min washes with IP buffer and another two 5-min washes with high-salt IP buffer (50 mM Tris-HCl, pH 8.0, 500 mM NaCl, 1.5 mM MgCl₂, 1 mM EDTA, 0.5% NP-40, 10% glycerol, 4 mM DTT and 1×protease inhibitor cocktail), the purified beads were boiled with SDS loading buffer for 10 min. Then the mixture was analyzed by western blotting or subjected to MS.

Two-step immunoprecipitation assay (Re-IP)

The Re-IP assay was performed to identify K63-ubiquitinated lysine (K) sites of UFD1f¹⁰². In brief, HEK293T cells were co-expressing Flag-tagged UFD1f and HA-tagged K63Ub for 48 h. After two cold PBS washes, cells were harvested in the IP buffer and subjected to sonication for 5 min with the Sonics Vibra-Cell (3 s on, 6 s off, 30% amplitude) on ice. After centrifugation at 13,000 rpm for 10 min at 4 °C, the supernatant of lysate was incubated with the anti-HA antibody (3 μg) coupled to protein G magnetic beads overnight at 4 °C in a rotating wheel. After two 5-min washes with IP buffer and one 5-min washes with high-salt IP buffer (500 mM NaCl), the purified complex was denatured by boiling in the IP buffer containing 1% SDS. Subsequently, the supernatant was diluted 1:10 with IP buffer and then performed the secondary UFD1f IP with the anti-UFD1f antibody (3 μg) coupled to protein G magnetic beads. After three 5-min washes with the IP buffer, the purified complex was boiled with SDS loading buffer for 10 min, and separated on the SDS-PAGE gels. Coomassie blue stained gels with size more than 35 kDa were subjected to MS, followed by identifying K63-ubiquitinated lysine (K) sites of UFD1f with two biological replicates.

Examination of protein ubiquitination

For examining the overall levels of protein ubiquitination, cells were homogenized in RIPA buffer and further analyzed by western blotting using the α-total ubiquitin antibody. For examining different polyubiquitin chains of specific protein, after 6 h MG132 (25 μM) treatment, cells were lysed by boiling in denaturing buffer (150 mM Tris-HCl pH 8.0, 1% SDS and 30% glycerol) for 10 min. Cell lysates were then diluted 10 times with buffer A (50 mM Tris-HCl pH 8.0, 150 mM NaCl, 0.5% NP-40 and protease inhibitor cocktail). After centrifugation at 13,000 rpm for 10 min at 4 °C, the supernatant was incubated with the antibodies against the specific protein under investigation. The antibodies (UFD1f, Myc tag, or IPMK antibodies) were coupled to Protein G magnetic beads. After overnight incubation at 4 °C in a rotating wheel, beads were subjected to four 5-min washes with the buffer A, and the purified beads were boiled with SDS loading buffer for 10 min and subjected to western blotting. Proteins immunoprecipitated were then analyzed by immunoblotting using α-total, K48-, K11-, K63-, or K6-linked polyubiquitin antibodies, respectively.

RNA immunoprecipitation (RIP)

To perform the IRE1α RIP assay¹⁰³, 10⁷ HEK293T cells expressing Flag-tagged IRE1α were crosslinked in a UV-cross-linker (120 mJ/cm², 2 min), and then harvested in 1 mL ice-cold RIPA buffer with 0.1 U/mL RNase inhibitor (Promega, N2511), 2 mM DTT, 1×protease inhibitor cocktail, and 1×phosphatase inhibitor cocktail (TransGen, DI201) on ice for 30 min. Cells were sonicated for 5 min with the Sonics Vibra-Cell (3 s on, 6 s off, 30% amplitude). After centrifugation at 13,000 rpm for 10 min at 4 °C, the supernatant was collected and incubated with 3 μg FLAG antibody or normal mouse IgG (Proteintech, B900620) at 4 °C for 4 h. Protein G magnetic beads were blocked with 500 ng/µL yeast total RNA (Invitrogen, AM7120G) and 1 mg/mL BSA (Sigma, A7030) for 1 h at room temperature, washed with RIPA buffer twice, and then incubated with the above mixture for 2 h at 4 °C. The protein-RNA complexes were washed with RIPA buffer twice and another two high-salt RIPA buffer (500 mM NaCl) washes. One-fifth of protein-RNA complexes was boiled for 10 min in SDS-loading buffer and examined by western blotting. The remaining one was digested with proteinase K at 55 °C for 20 min, followed by RNA extraction with TRIzol reagent. The purified RNA was analyzed by RT-qPCR.

Ubiquitin (label-free)-modified proteome

The ubiquitin (label-free)-modified proteome was carried out at Shanghai Applied Protein Technology Co., Ltd. For sample preparation, 10⁸ WT or UFD1s MUT HEK293T cells with three biological replicates were treated with 25 μM MG132 (APExBIO, A2585) for 6 h and then washed twice by cold PBS. Cells were immediately lysed in UA buffer (8 M Urea, 100 mM Tris-HCl, pH 8.5) on ice for 10 min. Cells were harvested, frozen in liquid nitrogen and stored at −80 °C for further ubiquitin (label-free)-modified proteome¹⁰⁴. Briefly, 3 mg total proteins were digested with trypsin (1:50) overnight at 37 °C and mixed with 0.1% trifluoroacetic acid (TFA). Then, the digest peptides were desalted on C18 Cartridges (Sigma, bed I.D. 7 mm, volume 3 ml) and lyophilized according to the manufacturer′s protocol. Further immunoaffinity purification (IAP) of K-ε-GG-modified peptides was performed using a PTMScan Ubiquitin Remnant Motif (K-ε-GG) Kit (CST, 5562). LC-MS/MS assays were conducted on a timsTOF Pro mass spectrometer (Bruker) that was coupled to Nanoelute (Bruker Daltonics) for 60 min. The peptides were loaded on a C18-reversed phase analytical column in buffer A (0.1% formic acid) and separated with a linear gradient of buffer B (84% acetonitrile and 0.1% formic acid) at a flow rate of 300 nL/min. The mass spectrometer was operated in positive ion mode with data dependent acquisition (DDA). The mass spectrometer collected ion mobility MS spectra over a mass range of m/z 100–1700 and 1/k0 of 0.6–1.6, and then performed 10 cycles of PASEF MS/MS with a target intensity of 1.5k and a threshold of 2500. The MS data were analyzed using the MaxQuant software and searched against the UniProKB Homo sapiens database with related parameters, max missed cleavages: 2; peptide mass tolerance: 20 PPM; fixed modification: carbamidomethyl (C); variable modifications: oxidation (M), GlyGly (K); database pattern: reverse; peptide/protein/site FDR ≤ 0.01. The differential ubiquitin-modified proteins were determined with the cutoff (Fold change >2 or <0.5 and P value < 0.05). P values were calculated with two-tailed unpaired Student′s t-tests.

Transmission electron microscopy (TEM)

For TEM, cells were firstly fixed with 2% PFA and 3% glutaraldehyde in PBS for 2 h at room temperature followed by three 0.1 M cacodylate buffer (pH 7.4) washes. After post-fixation in 1.5% potassium ferrocyanide and 1% osmium tetroxide in 0.1 M cacodylate buffer for 40 min, cells were washed with 0.1 M cacodylate buffer three times. Then, cells were further fixed with 1% osmium tetroxide in 0.1 M cacodylate buffer for 40 min. After one 0.1 M cacodylate buffer and three ddH₂O washes, cells were incubated with 2% uranyl acetate in ddH₂O for 1 h. After washing with ddH₂O three times, treated cells were dehydrated with a graded series of ethanol solutions (30, 50, 70, 80%, 90, 100%) and 100% acetone, and were embedded in epoxy resin. Ultrathin sections were stained with uranyl acetate and lead citrate, and examined using the 120 kV electron microscope (FEI Tecnai T12). Images were captured at room temperature with an Eagle CCD camera using Tecnai User Interface (TUI) software.

Nile red staining

Cells were washed with PBS and fixed in 4% PFA for 10 min at room temperature. Then, they were stained with 0.05 μg/mL Nile red (Sigma, 72485) for 10 min after washing with PBS twice. Nuclei were stained with Hoechst 33342 and images were acquired by laser confocal microscopy LSM 980 (Zeiss). The Nile red signal is defined as the mean gray value per cell quantified by Image J.

ROS and ATP measurements

Cellular ROS and ATP levels were detected using the ROS Assay Kit (Beyotime, S0033) and ATP assay Kit (Beyotime, S0026), respectively, according to the manufacturer′s instructions.

Measurements of triglyceride, total cholesterol, AST, and ALT

Cellular or serum triglyceride and total cholesterol levels were measured with Triglyceride Assay Kit (NJJC, F001-1-1) and Total cholesterol Assay Kit (NJJC, F002-1-1), respectively, according to the manufacturer′s instructions. For cellular triglyceride, cells were collected and lysed with RIPA buffer for 30 min, an equal volume of cell lysate was used to measure triglyceride. The triglyceride levels were normalized to total proteins detected by Bradford Protein Assay Kit (Thermo, 23200). The serum ALT and AST levels of mice were detected with Glutamic-pyruvic Transaminase (GPT) Activity Assay Kit (Solarbio, BC1555) and Micro Glutamic-oxalacetic Transaminase (GOT) Assay Kit (Solarbio, BC1565), respectively. All mice were kept in a controlled environment (23–25 °C with a 12-h light-dark cycle and lights on at 8:00 AM) with free access to water and food, and were fasted for 12 h before bleeding and sacrifice at 9:00 AM.

Ribosome profiling

For ribosome profiling assays⁸⁴, 10⁷ indicated cells were cultured with DMEM containing 100 mg/mL CHX for 1 h, followed by lysis in ribosome lysis buffer (100 mM KCl, 2 mM MgCl₂, 10% Glycerol, 50 mM HEPES, 0.1% Triton X-100, 1 mM DTT, 1×protease inhibitor cocktail, 20 U/mL RNase inhibitor, 100 μg/mL CHX) on ice for 30 min. Ribosome components were separated on a 20–50% linear sucrose gradient containing 100 mM NaCl, 5 mM MgCl₂, 10 mM Tris, pH 7.5 and 1 mM DTT after centrifuging at 38,000 rpm for 4 h at 4 °C. The gradients were fractionated with optical scanning at 254 nm using a Gradient Fractionator (BioComp). GAPDH mRNA is a positive control, and H19 lncRNA, a negative control, is known to be noncoding.

Oxygen consumption rate (OCR) and fatty acid oxidation (FAO) measurements

All assays were performed using the Seahorse XFe96 analyzer (Seahorse Bioscience, Agilent) according to the manufacturer′s instructions. Briefly, 10⁴ cells were seeded in a 96-well XF cell culture microplate in the DMEM medium for 24 h. OCR was measured with the XF96 analyzer in Seahorse XF DMEM medium (Agilent, 103575-100) containing 1 mM pyruvate (Sangon, A600884), 2 mM glutamine (Gibco, 25030081) and 10 mM glucose (Sigma, G7021) using Cell Mito Stress Test Kit (Agilent, 103015-100). For the FAO rate analysis, after seeding cells in a 96-well XF cell culture microplate for 24 h, the cell culture medium was replaced with substrate-limited DMEM (Gibco, A1443001) supplemented with 0.5 mM glucose, 1 mM glutamine, 0.5 mM carnitine (MCE, HY-B0399), 1% FBS for 12 h. Before starting the XF assay, substrate-limited DMEM was replaced with FAO assay medium (KHB buffer: 111 mM NaCl, 4.7 mM KCl, 1.25 mM CaCl₂, 2 mM MgSO₄, 1.2 mM NaH₂PO₄ supplemented with 2.5 mM glucose, 0.5 mM carnitine, 5 mM HEPES, adjusted to pH 7.4 in 37 °C) in a non-CO₂ incubator for 45 min. Finally, just prior to starting the assay, added BSA or BSA-conjugated palmitate (50 μM) (Agilent, 102720-100). FAO assays were carried out using Cell Mito Stress Test Kit. All data were obtained using Wave software and graphs were obtained by GraphPad Prism 8.

Hydrodynamic injection of plasmid DNA

To obtain liver-specific expression of Ufd1s, 15 μg pLIVE empty vector or pLIVE-Ufd1s plasmids diluted in a volume of 2 mL saline (0.9% NaCl), about equivalent to 10% of the bodyweight, were injected into the mouse tail vein within 5 s with a 2.5-mL syringe and a 27-gauge needle⁸². And no mortality was observed after hydrodynamic injection.

In vitro RNA circularization

Linear RNA precursors were synthesized using a TranscriptAid T7 High Yield Transcription Kit (Thermo, K0441). In brief, 1 μg template, 2 μL T7 RNA polymerase, 0.5 mM NTPs, and 2 mM GMP were mixed and incubated at 37 °C for 4 h. After DNase I (Thermo, EN0521) treatment, additional 2 mM GTP was added along with the circularization buffer containing magnesium (15 mM MgCl₂, 50 mM Tris-HCl, pH 7.5, 1 mM DTT), and the reaction was incubated at 55°C for 20 min. After circularization, RNA was digested with RNase R (Epicenter, RNR07250) to enrich circRNA. Circularized RNA was separated on 5% Urea PAGE gel, cut in the corresponding band, eluted overnight in elution buffer (20 mM Tris-HCl, pH 7.5, 250 mM NaOAc, 1 mM EDTA, and 0.25% SDS) and purified using phenol/chloroform (pH 4.5)¹⁰⁵.

CircRNA administration in mice

The delivery of circRNA via tail vein injection mediated by Lipofectamine 3000 was performed in mice¹⁰⁶. The circUfd1s or circRNA control were mixed with Lipofectamine 3000 according to the manufacturer′s protocol. Briefly, a dose of 10 μg circRNA with 20 μL Lipofectamine 3000 in PBS (100 μL total volume) was prepared. The circRNA-Lipofectamine 3000 mixture were injected via the tail vein within 30 s, using a 27-gauge needle and 1 mL syringe.

Histological staining

Liver tissues were fixed with 4% PFA for 24 h, dehydrated in ethanol, and embedded in paraffin. The 5 μm sections of livers were prepared and stained with hematoxylin and eosin (H&E) dye (Servicebio, G1003) and Masson dye solution (Servicebio, G1006). The images were captured using the Olympus IX81 Inverted Fluorescence Microscope (Olympus). The steatosis grade (scale of 0–3) determined by the percentage of lipid droplets and lobular inflammation grade (scale of 0–3) determined by all inflammation foci, were assessed from H&E-stained liver sections by a pathologist (Wuhan Servicebio Technology, Wuhan, China). The percentage of Masson-positive area is calculated by Image J.

Immunohistochemistry (IHC)

For IHC, paraffin sections of mouse or human livers were incubated with anti-IPMK (Immunoway, YT2384, 1:200), anti-LC3 (Proteintech, 14600-1-AP, 1:200), and anti-UFD1s (Human, 1:200; Mouse, 1:200) antibodies overnight at 4 °C followed by incubating with the correspondent HRP-conjugated secondary antibodies (ServiceBio, GB23303 and GB23301) for 1 h at room temperature. Then the sections were visualized with DAB (Servicebio, G1212) chromogenic reaction and nucleus counterstaining with hematoxylin solution. Finally, the sections were dehydrated (75% ethanol, 85% ethanol, 100% ethanol, N-butanol, xylene for 5 min each) and sealed with sealant (Servicebio, G1404). All images were acquired with the Olympus IX81 Inverted Fluorescence Microscope. The average optical density (AOD) is calculated by Image J to indicate the IHC signal.

Oil Red O staining

Frozen sections of mouse livers were fixed with 4% PFA for 15 min, stained with freshly prepared 3 μg/mL Oil Red O solutions (Servicebio, G1015) for 10 min, and saturated Oil Red O dye with 60% isopropanol. After one 5 min hematoxylin-incubation and one ddH₂O wash, sections were checked using the Olympus IX81 microscope. The percentage of Oil Red O positive area was calculated by Image-Pro plus.

Isolation of primary hepatocytes

For isolation of primary hepatocytes from wild-type and Ufd1s^−/− mice¹⁰⁷, mice were first anesthetized by intramuscular injection of isoflurane (Sigma, 792632), and mouse livers were perfused with LPM buffer (Gibco, 17701038) via the inferior vena cava (IVC) for 3–5 min. Then, preheated LDM buffer (GBSS (Sigma, G9779) containing 40 μg/mL Liberase TL (Roche, 05401020001)) was used to perfuse upon the liver color changed from red to beige or light brown. When the liver was monitored with the cracking (always within 2 min), perfusion was stopped immediately and the liver was excised out into the ice-cold liver digestion medium (high-glucose DMEM, 1% penicillin and streptomycin, 15 mM HEPES). The perfused hepatocytes were filtered with a 100 μm cell strainer and washed with liver digestion medium for three times after centrifugation at 50 × g for 5 min at 4 °C. The hepatocytes were resuspended in Williams Medium (Gibco, A1217601) (containing 10% FBS, 1% penicillin and streptomycin, 5 mM HEPES, and 200 mM glutamine) and cultured on collagen (Solarbio, C8062)-coated plates.

Live-cell LC3 imaging

Cells were cultured on glass-bottom dishes (NEST, 801001) and transfected with mCherry-EGFP-LC3 plasmids. After 48 h transfection, cells were starved with HBSS medium for 4 h and then imaged at 37 °C by LSM 980 confocal microscope (Zeiss). Images were analyzed with Image J software.

Free fatty acid induction of cultured cells

To model hepatic steatosis in cellular level, hepatocellular carcinoma cells (human HepG2 and mouse Hepa1-6) were treated with cell culture medium containing the indicated concentrations of oleic acid (OA) (0.5 mM) and palmitic acid (PA) (0.25 mM) (Kunchuan, KC006) for 3 days, then those cells were stained with the Oil Red O staining kit (Solarbio, G1262) specifically for cells, respectively. Cells were imaged with Olympus IX81 microscope and the percentage of Oil Red O positive area was calculated by Image-Pro plus.

Intraperitoneal glucose tolerance test (IPGTT) and insulin tolerance test (ITT)

For IPGTT, mice were fasted overnight for 16 h and then followed by the intraperitoneal injection of D (+)-glucose (Sigma, G7021) solution in saline (2 g/kg body weight). Blood glucose levels were measured from the tail vein by glucometer (Contour care) at 0, 15, 30, 60, 90, and 120 min after glucose injection. For ITT, after 4 h fasting (9:00 am–13:00 pm), mice were intraperitoneally injected with 0.75 U/kg of insulin (Novorapid, Insulin Aspart Injection). Blood glucose levels were similarly measured at 0, 15, 30, 60, 90, 120, 150, and 180 min.

Multiplex immunofluorescence (mIF)

Paraffin slides of mouse livers were first placed on a hot container to melt the paraffin, then hydrated in in xylene for 15 min, followed by immersions in 100, 85, and 75% ethanol for 5 min each and finally in the ddH₂O bath. After hydration, slides were placed in the microwave oven and treated with citrate buffer antigen solutions pH 6.0 (G1202, Servicebio) at a medium–low heat to allow the solution to remain boiling for 15 min. During this process, the buffer should avoid drying out. After the heat treatment, BSA was added to block for 30 min. The first antibody was applied and incubated overnight at 4 °C in a humidified environment. After the first antibody incubation and three 5-min washes with PBS, the correspondent HRP-conjugated second antibody incubation was conducted for 1 h and the TSA fluorescent dye (Recordbio, RC0086-45) was added for 10 min. All steps are repeated for each subsequent round of staining with other antibodies and other chosen fluorescent dyes, including F4/80 (Servicebio, GB113373, 1:500), NK1.1 (CST, 39197, 1:100), CD19 (Abcam, AB245235, 1:800) and CD3 (Abcam, AB16669, 1:200) antibodies. Finally, nuclei were stained with DAPI. Images were captured using the LSM 980 confocal microscope (Zeiss) and further processed with Image J.

Image processing

All images of the Nile Red staining, smFISH, IF, live-cell LC3 imaging, and mIF were taken by confocal microscope LSM980 (Zeiss). All images of PI/Hoechst, H&E, Masson, and Oil red O staining, and IHC were taken with an inverted fluorescence microscope IX81 (Olympus). Image processing and quantification were performed by Image J or Image-Pro plus (as stated specifically in the corresponding method).

Statistical analysis

For the RNA and protein detection of UFD1s and UFD1f expression, the fluorescence intensity from 15 randomly selected cells was quantified. To analyze autophagy activity, the LC3 puncta fluorescence intensity from randomly selected 25 or 30 cells each sample was measured. The number of autophagic vacuoles (AVs) from 5 cells and the size of 10 AVs were measured. Lipid droplets (LDs) of 30 cells were counted to examine lipid metabolism.

All cellular and biochemical experiments were repeated for at least three times. Error bars in the figures represent standard error of the mean (SEM). N indicated times of an experiment, numbers of quantified cells, mice or clinical specimens employed for the experiment. All image processing and quantification were performed by Image J or Image-Pro plus (as stated specifically in the corresponding method). Statistical analyses were performed using GraphPad Prism 8. Statistical significance was calculated using two-tailed unpaired Student′s t-test when two independent groups were compared. The differences were considered statistically significant when P values below 0.05. For RNA-seq analysis, DE SR-mRNAisos and genes were determined by DEseq2. The differences were considered statistically significant when P values < 0.05 for DE SR-mRNAisos and P values < 0.01 for DE genes. For ubiquitin-modified proteome, statistical significance was calculated using two-tailed unpaired Student′s t-tests, and P values < 0.05 were considered statistically significant.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

Raw SR-mRNAiso RNA-seq and DE analysis RNA-seq data are deposited in the NCBI Gene Expression Omnibus (https://www.ncbi.nlm.nih.gov/geo/) with accession GSE249158. All raw ubiquitin proteomics data have been deposited to ProteomeXchange Consortium (http://proteomecentral.proteomexchange.org) via the iProX partner repository with the dataset identifier PXD038683. The detailed results of DE splicing events derived from SR-mRNAiso RNA-seq data are presented in Supplementary Data 1; the detailed results of DE mRNAs from DE analysis RNA-seq data are presented in Supplementary Data 2; The detailed results of UFD1s and UFD1f co-IP MS data are presented in Supplementary Data 3; The K63-ubiquitinated lysine sites of UFD1f and the significantly changed protein candidates from ubiquitin-modified proteome data are presented in Supplementary Data 4. Patient information is provided in Supplementary Table 1. All the sequences of oligos used in this study are presented in Supplementary Table 2. Source data are provided with this paper.

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Acknowledgements

We thank Dr. Wenchu Lin (The Second Affiliated Hospital, The Chinese University of Hong Kong) for offering AGS and SGC7901 cell lines. We also thank Dr. Gang Cai (USTC) for gifting S. pombe and S. cerevisiae strains, Dr. Yide Mei (USTC) for providing plasmids (HA-Ub, HA-K63Ub, and MARCH7), Dr. Chunlei Cang (USTC) for providing the mCherry-EGFP-LC3 plasmid, Dr. Xiaohu Zheng (USTC) for the constructive comments on immunity, and Dr. Fanzheng Meng (USTC) for providing the clinical samples. We thank the Laboratory Animal Research Center of USTC and Experiment Center for Life Science of USTC for technique and experiment supports, respectively; and also thank the Bioinformatics Center of USTC, School of Life Sciences, for providing supercomputing resources. This study was supported by the National Natural Science Foundation of China (32200431, 32470596, 32000438, U23A20164, and 32270590), Anhui Universities Outstanding Youth Research Project (2024AH020017), Fundamental Research Funds for the Central Universities (WK9100000050), Research Funds of Center for Advanced Interdisciplinary Science and Biomedicine of IHM (QYZD20230012), USTC Research Funds of the Double First-Class Initiative (YD9100002049), and Open Project of State Key Laboratory of Systems Medicine for Cancer (KF2402-93).

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These authors contributed equally: Xiuzhi Li, Xiaolin Wang.
These authors jointly supervised this work: Xiaolin Wang, Ge Shan, Liang Chen.

Authors and Affiliations

Department of Cardiology, Department of Obstetrics and Gynecology, The First Affiliated Hospital of USTC, The RNA Institute, School of Basic Medical Sciences, Division of Life Science and Medicine, University of Science and Technology of China (USTC), Hefei, China
Xiuzhi Li, Xiaolin Wang, Xu Liu, Ge Shan & Liang Chen

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Xiuzhi Li
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Xiaolin Wang
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Xu Liu
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Ge Shan
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Liang Chen
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Contributions

G.S. conceived and designed this project. X.Li, X.W., and X.Liu performed experiments. X.W. performed bioinformatics analyses of RNA-seq data. G.S., X.W., X.Li, and L.C. analyzed the data and wrote the manuscript. All authors have discussed the results and made comments on the manuscript. All authors approved the final manuscript.

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Correspondence to Xiaolin Wang, Ge Shan or Liang Chen.

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The authors declare the following patent application related to this work. Patent applicants: Institute of Health and Medicine, Hefei Comprehensive National Science Center; University of Science and Technology of China. Inventors: G.S., X.Li., X.W. (co-authors of this article); Application number: CN 2024114540916; Status: Pending; The patent claims cover the application of small protein UFD1s for the prevention, treatment, and diagnosis of diseases, including NASH. The remaining authors declare no competing interests.

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Li, X., Wang, X., Liu, X. et al. A UFD1 variant encoding a microprotein modulates UFD1f and IPMK ubiquitination to play pivotal roles in anti-stress responses. Nat Commun 16, 6695 (2025). https://doi.org/10.1038/s41467-025-62073-6

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Received: 12 July 2024
Accepted: 10 July 2025
Published: 21 July 2025
DOI: https://doi.org/10.1038/s41467-025-62073-6