Abstract

   Patients with systemic lupus erythematosus (SLE) often have decreased
   fertility. Gene translation is crucial to oocyte meiosis and
   development. However, it remains unclear how SLE affects this process.
   Here, we used single-cell transcriptome and translatome sequencing to
   uncover a notable disruption in protein translation in oocytes from SLE
   mice, associated with the N^4-acetylcytidine (ac^4C) modification.
   Inhibition of ac^4C levels in vitro substantially reduced oocyte
   translation efficiency. Notably, through trace-cell ac^4C-RNA
   immunoprecipitation (acRIP) sequencing, we mapped the ac^4C landscape
   in SLE mouse oocytes and found that deficient ac^4C modification
   substantially impaired the translation of Zygote arrest 1. Furthermore,
   we demonstrated that nicotinamide treatment notably and safely improved
   the quantity and quality of oocytes in SLE mice by enhancing ac^4C
   modification levels. Our findings highlight the essential role of N-
   acetyltransferase 10 (NAT10)-mediated ac^4C modification in abnormal
   oocyte development in SLE and suggest that nicotinamide holds promise
   for improving fertility in patients with SLE.
     __________________________________________________________________

   Nicotinamide supplementation is a potential strategy to improve
   fertility in patients with SLE.

INTRODUCTION

   Systemic lupus erythematosus (SLE) is a common autoimmune disease that
   affects women of childbearing age. With advancements in rheumatology,
   most patients can achieve good disease control and maintain safe
   fertility ([46]1, [47]2). However, SLE can notably affect ovarian
   reserve function ([48]3). On the one hand, many patients use
   cyclophosphamide, a gonadotoxic drug that can cause premature ovarian
   insufficiency (POI) ([49]4); on the other hand, even patients with
   normal menstrual cycles who have not been treated with gonadotoxic
   drugs may experience reduced ovarian reserve, indicating that SLE
   itself can disrupt ovarian function ([50]5). These factors contribute
   to the developmental disorders of oocytes and embryos, making it
   difficult for patients to conceive and achieve successful pregnancy
   outcomes ([51]6). To date, fertility management for patients with SLE
   has been limited to those treated with cyclophosphamide. The guidelines
   suggest that oocytes, embryos, or ovary cryopreservation can be
   considered for fertility preservation before cyclophosphamide
   treatment, and a gonadotropin-releasing hormone agonist can be added
   for fertility protection at the same time ([52]1, [53]7). However, when
   patients with SLE are in a stable condition and ready for pregnancy,
   their oocytes continue to be damaged by SLE, and there is no targeted
   method to help them improve the quality of oocytes and reproductive
   outcomes. A major obstacle to identifying therapeutic targets is the
   lack of a complete understanding of the mechanisms underlying the
   impairment of oocyte development in SLE.

   From the germinal vesicle (GV) stage, oocyte transcription gradually
   silences, and gene expression becomes primarily regulated by
   translation ([54]8). It has been confirmed that the translational
   program is integral to the development of oocytes and embryos ([55]9).
   Because of the limited number and difficulty in obtaining mammalian
   oocytes, microtranslatomics is essential for exploring gene expression
   signatures in oocytes and embryos. Fortunately, multiple research
   teams’ recent breakthroughs in microtranslation omics technology have
   enabled in-depth studies to explore the key mechanisms of oocyte and
   embryo development ([56]10–[57]13). This technology also offers a
   chance to investigate the gene expression characteristics of
   SLE-affected oocytes.

   In recent years, the mechanism of epigenetic regulation of gene
   translation in oocytes has garnered notable research attention. At
   present, there are two main types of epigenetic regulation of gene
   translation efficiency (TE) in oocytes, namely, N^6-methyladenosine
   (m^6A) and N^4-acetylcytidine (ac^4C). The translation of m^6A-modified
   transcripts, facilitated by the m^6A reader Ythdf3, is crucial for
   oocyte development ([58]14). As shown in our previous studies, Ythdf3
   levels are significantly reduced in aged oocytes, and interventions
   targeting Ythdf3 can disrupt oocyte translation and inhibit the TE of
   aging-related maternal factors such as helicase, lymphoid specific
   (Hells), leading to oocyte senescence ([59]15). Furthermore, we
   identified N-acetyltransferase 10 (NAT10)-mediated ac^4C modification
   as an important regulatory factor in the maturation of oocytes in vitro
   ([60]16). Bao and colleagues ([61]17) confirmed that the loss of NAT10
   resulted in global translational inhibition in metaphase II (MII)-stage
   oocytes, which subsequently led to disorders in oocyte meiosis. RNA
   immunoprecipitation (IP) sequencing (RIP-seq), which combines RNA IP
   and high-throughput sequencing, can simultaneously detect thousands of
   bound transcripts (mRNAs, noncoding RNAs, or viral RNAs) in a single
   experiment ([62]18). Therefore, it can be a powerful tool to understand
   the epigenetic regulatory network of gene translation. At present, the
   description of RIP landscape in mammalian oocytes has been limited to
   healthy samples ([63]17, [64]19), and there have been no reports on
   disease-derived samples, especially SLE oocytes. Consequently, it is
   worth mapping the RIP-seq landscape, so as to reveal the role of
   translational regulation of epigenetic modifications in SLE oocyte
   damage.

   In this study, we used medical record data from patients with SLE and a
   mouse model to confirm the effect of SLE on oocyte quantity and
   quality. We next investigated the precise transcriptional and
   translational landscapes by analyzing the trace-cell translatome and
   transcriptome in oocytes from SLE and normal female mice and determined
   the expression, function, and regulation of gene translation of
   NAT10-mediated ac^4C in SLE oocytes. In addition, we used trace-cell
   ac^4C-RNA immunoprecipitation and sequencing (acRIP-seq) to reveal the
   downstream mechanism of ac^4C-mediated translational regulation. We
   further evaluated the efficacy and safety of nicotinamide in improving
   ac^4C modification in oocytes of SLE mice.

RESULTS

Decline in ovarian function in women with SLE leads to poor birth outcomes

   First, we retrospectively analyzed 50 women with a history of SLE who
   had undergone in vitro fertilization with or without intracytoplasmic
   sperm injection (IVF/ICSI) treatment at our reproductive center,
   resulting in a total of 102 IVF/ICSI treatment cycles. To minimize
   imbalances in baseline data, we matched the patients based on age, body
   mass index (BMI), type and duration of infertility, history of
   polycystic ovary syndrome, and history of ovarian endometriosis at a
   1:4 ratio using propensity score matching (PSM). The control group
   included 200 control women who had undergone 309 IVF/ICSI treatment
   cycles (fig. S1). Both groups were well matched and had similar
   baseline characteristics (table S1).

   In terms of ovarian reserve, the SLE group showed significantly lower
   serum anti-Müllerian hormone (AMH) levels and fewer antral follicle
   counts (AFCs) on the third day of the menstrual cycle than the control
   group ([65]Fig. 1, A and B). As for ovarian reactivity, the number of
   oocytes retrieved after controlled ovarian stimulation was
   significantly lower in the SLE group ([66]Fig. 1C). IVF laboratory
   results indicated no significant difference in the normal fertilization
   rate between the SLE group and the control group ([67]Fig. 1D).
   However, the SLE group had a subsequent embryo development disorder,
   with a significantly lower rate of available embryo and oocyte
   utilization ([68]Fig. 1, E and F). In terms of clinical outcomes, the
   SLE group experienced a significantly higher early miscarriage rate and
   a lower live birth rate ([69]Fig. 1, G and H) than the control group.

Fig. 1. Decreased ovarian function and oocyte quality in women with SLE.

   [70]Fig. 1.
   [71]Open in a new tab

   (A) Serum AMH levels for the SLE and control groups (n = 50/200). (B)
   AFC on the third day of the menstrual cycle for the SLE and control
   groups (n = 50/200). (C) Number of retrieved oocytes for the SLE and
   control groups (n = 99/287). (D) Normal fertilization rate for the SLE
   and control groups (n = 416/1735). (E) Available embryo rate for the
   SLE and control groups (n = 243/1104). (F) Oocyte utilization rate for
   the SLE and control groups (n = 416/1735). (G) Early miscarriage rate
   for the SLE and control groups (n = 33/175). (H) Live birth rate for
   the SLE and control groups (n = 74/333). Data are shown as the
   mean ± SD.

   These results suggested that women with SLE had decreased ovarian
   function and reduced oocyte quality, leading to poor reproductive
   outcomes. Understanding the effects of SLE on oocytes and elucidating
   the underlying mechanisms are essential for developing effective
   treatment strategies to improve fertility in women with SLE.

Oocyte damage in mice with SLE

   To rule out the effects of gonadotoxic drugs such as cyclophosphamide,
   we used a mouse model of SLE to study the direct effects of SLE on
   oocytes. Given the difficulty in obtaining mammalian oocytes and their
   rapid decline with age, we selected a Toll-like receptor 7
   (TLR7)–induced SLE mouse model with a younger age of onset. Consistent
   with previous reports ([72]20–[73]22), topical application of the TLR-7
   agonist imiquimod promoted an SLE phenotype in the mice, as evidenced
   by chronic nephritis with hypercellular glomeruli and increased
   mesangial matrix (fig. S2, A and B). The mice also exhibited systemic
   inflammation, including lung inflammation (fig. S2C), megakaryocyte
   hyperplasia (fig. S2D), splenomegaly (fig. S2, E and F), liver
   inflammatory infiltration (fig. S2G), and hepatomegaly (fig. S2H).
   These findings indicated that a substantial SLE-like inflammatory
   response was induced by imiquimod treatment, successfully establishing
   the SLE mouse model.

   We hormonally induced ovulation in both SLE and wild-type (WT) mice to
   compare the quantity and quality of ovulated oocytes between the two
   groups. The number of GV-stage oocytes in the SLE group was
   significantly lower than that in the WT group ([74]Fig. 2A).
   Tetramethylrhodamine, methyl ester (TMRM) staining showed that the
   mitochondrial membrane potential of GV-stage oocytes from the SLE mice
   was significantly reduced ([75]Fig. 2, B and C). In vivo maturation of
   oocytes to the MII stage was induced by human chorionic gonadotrophin
   (hCG) trigger treatment. The number of mature MII-stage oocytes
   retrieved from the SLE mice was also significantly decreased ([76]Fig.
   2D). In addition, the mitochondrial membrane potential of MII-stage
   oocytes in the SLE group was significantly lower than that in the WT
   group ([77]Fig. 2, E and F). Spindle morphology is another important
   indicator of oocyte quality. MII-stage oocytes should be in the
   metaphase of the second meiotic division, with spindle fibers arranged
   in a spindle-like pattern, pulling chromosomal centromeres to line the
   equatorial plate uniformly. Immunofluorescence staining revealed that
   the proportion of MII-stage oocytes with normal spindle morphology was
   significantly lower in the SLE group than in the WT group ([78]Fig. 2,
   G and H). We further performed IVF experiments with MII-stage oocytes
   from both groups and found that oocytes from the SLE mice exhibited
   embryonic development disorders. To be precise, at all stages after the
   four-cell stage, the embryo maturation rate of the SLE mice was lower
   than that of the WT mice ([79]Fig. 2, I and J). These results indicated
   that SLE significantly affected the development of mouse oocytes,
   reducing both the quantity and quality of the oocytes.

Fig. 2. SLE damages mouse oocytes.

   [80]Fig. 2.
   [81]Open in a new tab

   (A) Comparison of the number of GV oocytes collected from WT and SLE
   mice after superovulation (n = 7/7). (B and C) Immunofluorescence
   images showing mitochondrial membrane potential (TMRM; red) of the WT
   and SLE GV oocytes and the statistical histogram (n = 25/25). Scale
   bars, 10 μm. (D) Comparison of the number of MII oocytes collected from
   WT and SLE mice after superovulation and trigger (n = 5/5). (E and F)
   Immunofluorescence images showing mitochondrial membrane potential
   (TMRM; red) of the WT and SLE MII oocytes and the statistical histogram
   (n = 12/13). Scale bars, 10 μm. (G and H) Immunofluorescence images
   showing chromosome morphology [4′,6-diamidino-2-phenylindole (DAPI);
   blue] and microtubules (α-tubulin; red) of the WT and SLE MII oocytes
   and the statistical histogram (n = 45/52). Scale bars, 10 μm. (I)
   Representative images of embryo development after IVF of MII oocytes
   harvested from SLE and WT mice. Scale bars, 50 μm. (J) Statistical
   analysis of the rate of embryo formation at each stage in SLE and WT
   mice: embryo number/oocyte number per mouse (n = 5/5). Data are shown
   as the mean ± SD.

Abnormal gene expression in SLE mouse oocytes primarily occurs in the
translation process at the GV stage

   Previous studies on oocyte development have suggested that the growth
   process of GV-stage oocytes is primarily dominated by DNA replication
   and transcription of the genome ([82]23). As oocytes mature toward the
   MII stage with the resumption of meiosis, posttranscriptional
   regulation and active translation become the major events in this
   process ([83]24). To better understand the gene expression
   characteristics of SLE-affected oocytes, we collected GV- and MII-stage
   oocytes from both SLE and WT mice and performed transcriptome and
   translatome sequencing (T&T-seq) ([84]Fig. 3A). Three-dimensional
   principal components analysis (3D PCA) revealed the greatest
   distinction between the SLE and WT oocytes in the GV stage ([85]Fig. 3,
   B to E). The GV-stage translatomics also exhibited the most
   differentially expressed genes (DEGs). In the SLE group, 2065 genes
   were down-regulated, while 442 genes were up-regulated, with the number
   of genetic variations far exceeding those observed in the MII stage or
   the transcriptome data of both stages ([86]Fig. 3, F to I). These
   findings highlighted notable differences in gene expression and
   regulation at the GV stage in SLE-affected oocytes, suggesting a
   critical impact of SLE on the early stages of oocyte development.

Fig. 3. Distinct transcriptomics and translatomics patterns of GV and MII
oocytes from SLE and WT mice.

   [87]Fig. 3.
   [88]Open in a new tab

   (A) Schematic diagram depicting major procedures and principles of
   T&T-seq. (B to E) 3D PCA plot of the translatomics and transcriptomics
   of GV and MII oocytes from SLE and WT mice. (F to I) Volcano plots
   showing DEGs of SLE and WT oocytes identified by T&T-seq. Nodiff, no
   significant differences. P < 0.05, Wald test, log[2] fold change (FC)
   > 1.5.

   To determine whether differential basal transcriptional activity
   between SLE and WT oocytes could account for the observed impairment,
   we performed a comprehensive analysis of the overall transcriptional
   status in both groups. The cumulative distribution analysis of gene
   expression profiles demonstrated comparable total RNA levels between
   the SLE and WT oocytes (fig. S3, A and D). Furthermore, the expression
   levels of reference genes glyceraldehyde-3-phosphate dehydrogenase and
   Actb showed no significant differences between the SLE and control
   groups at both GV and MII stages (fig. S3, B, C, E, and F). However, we
   analyzed the gene TE of mouse oocytes and found that the overall TE of
   genes in GV-stage SLE mouse oocytes was significantly lower than that
   in WT mouse oocytes ([89]Fig. 4A). Specifically, only 1601 genes in the
   GV-stage SLE mice had higher TE than those in the WT mice, while 6557
   genes had significantly lower TE ([90]Fig. 4B). If the translation
   process in SLE mouse oocytes is more impaired than that in WT oocytes,
   then more genes are expected to exhibit low TE. GV-stage SLE oocytes
   had 3936 genes with low TE (TE < 0.5) and only 465 genes with high TE
   (TE > 2). In contrast, GV-stage WT oocytes had 1506 genes with low TE
   and 3726 genes with high TE ([91]Fig. 4C). A similar trend in gene
   overall transcriptional status and TE was observed between the SLE and
   WT oocytes at the MII stage, although the differences were less
   pronounced than those at the GV stage ([92]Fig. 4, D to F). These
   findings indicated that the differences in gene expression in SLE mouse
   oocytes were primarily concentrated at the GV stage and that gene
   translation in SLE mouse oocytes was more negatively affected than
   transcription.

Fig. 4. Translational defects in oocytes from SLE mice.

   [93]Fig. 4.
   [94]Open in a new tab

   (A) TE cumulative curve of GV oocytes. The red line denotes the SLE
   group, and the blue line represents the WT group. (B) Scatter plot
   showing the RNA TE alterations of SLE GV oocytes compared with the WT
   group. Red and blue dots denote up- and down-regulated genes,
   respectively. Up-regulated, FC > 1; down-regulated, FC < 0.67. (C) Bar
   plots showing the numbers of high-TE genes (TE > 2) and low-TE genes
   (TE < 0.5) in the SLE group and the WT group GV oocytes. (D) TE
   cumulative curve of MII oocytes. The red line denotes the SLE group,
   and the blue line represents the WT group. (E) Scatter plot showing the
   RNA TE alterations of SLE MII oocytes compared with the WT group. Red
   and blue dots denote up- and down-regulated genes, respectively.
   Up-regulated, FC > 1; down-regulated, FC < 0.67. (F) Bar plots showing
   the numbers of high-TE genes (TE > 2) and low-TE genes (TE < 0.5) in
   the SLE group and the WT group MII oocytes (n = 3/3).

   To further understand the effect of gene translation dysfunction in
   GV-stage SLE oocytes, we performed gene enrichment analysis on the
   genes with down-regulated TE in GV-stage SLE oocytes. Kyoto
   Encyclopedia of Genes and Genomes (KEGG) analysis showed that the most
   enriched pathways involved metabolism and environmental information
   processing (fig. S4A). Most of the genes were enriched in metabolic
   pathways, indicating that balanced energy metabolism is crucial for
   optimal oocyte development ([95]25). Autophagy, one of the primary
   modes of oocyte death, suggests that SLE oocytes may undergo autophagic
   cell death following developmental impairment ([96]26). Cell cycle and
   DNA replication pathways were also enriched, suggesting that abnormal
   expression of these genes may be a notable contributor to
   SLE-associated spindle morphological abnormalities and meiotic defects.
   Multiple protein synthesis and degradation pathways, processes vital
   for normal oocyte development and zygotic transition ([97]27, [98]28),
   were enriched as well. Mitophagy, the selective autophagic removal of
   excess or damaged mitochondria, is essential for mitochondrial
   homeostasis and cell survival in oocytes ([99]29), which may be related
   to the reduced mitochondrial activity observed in SLE oocytes. We also
   identified enrichment in cellular senescence pathways, and previous
   studies have confirmed a genetic relationship between SLE and premature
   ovarian failure ([100]30), suggesting that SLE may lead to oocyte aging
   and premature depletion (fig. S4B). In addition, through gene set
   enrichment analysis, we found significant down-regulation of
   extracellular signal–regulated protein kinases, mitogen-activated
   protein kinase, and transforming growth factor–β pathways. Previous
   studies have demonstrated that dysregulation of these pathways can lead
   to meiotic arrest, reduced ovulation rates, and decreased fertility in
   mice (fig. S4C) ([101]31–[102]33). These pathways play crucial
   regulatory roles during oocyte development. Therefore, SLE can lead to
   disturbances in oocyte gene translation, which, in turn, impairs oocyte
   development and fertility.

Dysfunction of oocyte translation in SLE mice is related to the
NAT10-mediated ac^4C modification

   According to previous reports, m^6A and ac^4C are the main epigenetic
   modifications that affect gene TE in oocytes ([103]34, [104]35). We
   divided the genes of GV-stage oocytes into four groups based on the
   presence or absence of m^6A and ac^4C modifications ([105]36, [106]37).
   We found that genes with ac^4C and m^6A modifications showed a greater
   decrease in TE than genes without these modifications. Notably, the TE
   of genes with ac^4C modification decreased most significantly, with
   GV-stage SLE oocytes having the lowest TE for ac^4C-modified genes
   ([107]Fig. 5A). To identify the specific modification abnormalities in
   SLE mouse oocytes, we compared the m^6A and ac^4C modification levels
   in GV-stage oocytes between the SLE and WT mice using
   immunofluorescence staining. The results revealed a significant
   reduction in ac^4C modification levels in GV-stage oocytes from the SLE
   mice compared with those from the WT controls ([108]Fig. 5, B and C).
   Further omics data analysis confirmed that while ac^4C modification
   enhanced gene TE in the WT mice, this efficiency was markedly decreased
   in the SLE mice ([109]Fig. 5D), indicating that aberrant ac^4C
   modification in SLE mouse oocytes was a primary factor contributing to
   the reduced translational efficiency. In addition, there were no
   significant differences in m^6A modification levels between the SLE and
   WT mouse oocytes ([110]Fig. 5, E and F), suggesting that m^6A
   modification was not the main cause of the translational abnormalities
   in SLE mouse oocytes.

Fig. 5. Effect of NAT10-mediated ac^4C modification on gene translation in
SLE mouse GV oocytes.

   [111]Fig. 5.
   [112]Open in a new tab

   (A) Comparison of GV oocyte gene TE log[2]FC (SLE versus WT) in four
   groups with or without m6A and ac^4C modifications. The top and bottom
   borders of the bars indicate the maximum and minimum values, and the
   middle horizontal line identifies the median. (B and C)
   Immunofluorescence images showing ac^4C (red) and nuclei (DAPI; blue)
   of the WT and SLE GV oocytes and the statistical histogram (n = 11/18).
   Scale bars, 10 μm. (D) Comparison of GV oocyte gene TE in groups with
   or without m^6A and ac^4C modifications from SLE and WT mice. The
   dashed red line indicates the quartile, and the solid red line
   identifies the median. (E and F) Immunofluorescence images showing m^6A
   (red) and nuclei (DAPI; blue) of the WT and SLE GV oocytes and the
   statistical histogram (n = 16/16). Scale bars, 10 μm. n.s., not
   significant. (G to I) Transcriptional expression, translational
   expression, and TE levels of Nat10 in mouse GV oocytes (n = 3/3). Data
   are shown as the mean ± SEM. (J) Quantitative reverse transcription
   polymerase chain reaction (RT-PCR) results showing the relative
   expression levels of mouse Nat10 mRNA in SLE and WT GV oocytes. Using
   enhanced green fluorescent protein (EGFP) expression levels as a
   reference for each sample (n = 3/3). Data were presented as mean ± SEM.
   (K and L) Immunofluorescence images showing NAT10 (red) and nuclei
   (DAPI; blue) of the WT and SLE GV oocytes and the statistical histogram
   (n = 15/10). Scale bars, 10 μm. Data are shown as the mean ± SD.

   Given that NAT10 is the sole enzyme responsible for ac^4C modification,
   we further compared Nat10 expression at the transcriptional,
   translational, and protein levels in GV-stage oocytes between the SLE
   and WT mice. While there were no significant differences in Nat10
   transcription levels between the two groups ([113]Fig. 5G), SLE mouse
   oocytes exhibited significantly decreased Nat10 translation levels and
   translational efficiency ([114]Fig. 5, H and I). In agreement with this
   finding, quantitative real-time polymerase chain reaction (PCR)
   validated the similar expression trend of Nat10 mRNA levels ([115]Fig.
   5J). Immunofluorescence staining further confirmed the reduced protein
   levels ([116]Fig. 5, K and L). These findings demonstrated that
   down-regulated Nat10 translation in SLE mouse GV-stage oocytes led to
   decreased protein levels, consequently resulting in reduced ac^4C
   modification.

   We further evaluated the transcriptional and translational expression
   levels of m^6A-related modification enzyme genes in oocytes from the
   SLE and WT mice. Notably, most m^6A writers, erasers, and
   readers—including Wtap, Mettl14, Mettl3, Fto, Alkbh5, Ythdc1, Ythdc2,
   Ythdf2, and Ythdf3—showed no significant differences in their
   transcriptional and translational levels between the two groups. In the
   SLE mouse oocytes, only the reader protein Ythdf1 exhibited
   significantly decreased translation levels (fig. S5, A to J). Our
   previously published research has demonstrated that depleting Ythdf1
   expression in oocytes through trim-away technology does not affect
   oocyte maturation ([117]15), indicating that the reduced Ythdf1
   translation levels in SLE mouse oocytes are not responsible for their
   developmental defects. Collectively, these findings suggest that m^6A
   modification plays a less notable role than ac^4C in SLE mouse oocyte
   development. Subsequently, our research focused on investigating the
   regulatory mechanisms of ac^4C modification and its impact on oocyte
   development.

Inhibition of ac^4C reduces oocyte TE, leading to developmental disorders

   To verify whether ac^4C inhibition affects oocyte development, we
   pretreated oocytes with remodelin to inhibit NAT10 activity in vitro
   while maintaining their GV stage using 3-isobutyl-1-methylxanthine
   (IBMX). After 24 hours of treatment, a subset of oocytes was randomly
   collected for immunofluorescence staining and T&T-seq, while the
   remaining oocytes proceeded to in vitro maturation (IVM). The
   experimental workflow is illustrated ([118]Fig. 6A). First, to ensure
   sufficient inhibition of the ac^4C modification level in oocytes, we
   used IBMX to inhibit oocyte development, and remodelin was added to the
   culture medium for 24 hours in vitro. Immunofluorescence confirmed that
   the ac^4C modification level of oocytes significantly decreased after
   the treatment ([119]Fig. 6, B and C). The mitochondrial membrane
   potential of oocytes down-regulated by ac^4C modification was also
   reduced, similar to that observed in SLE ([120]Fig. 6, D and E). We
   then transferred the oocytes to an IBMX-free maturation medium to
   initiate the developmental process. Immunofluorescence showed that the
   normal spindle rate of remodelin-treated oocytes was decreased
   ([121]Fig. 6, F and G) and that the first polar body ejection rate was
   also reduced ([122]Fig. 6H), indicating maturation disorders in both
   the nucleus and the cytoplasm of oocytes. These findings suggest that
   ac^4C inhibition significantly impairs oocyte development, further
   supporting the role of ac^4C modifications in oocyte quality and
   developmental competence.

Fig. 6. Phenotypic and omics landscape of remodelin-treated mouse oocytes in
vitro.

   [123]Fig. 6.
   [124]Open in a new tab

   (A) Flow chart depicting remodelin treatment. (B and C)
   Immunofluorescence images showing ac^4C (green) and nuclei (DAPI; blue)
   of the remodelin-treated and the control GV oocytes and the statistical
   histogram (n = 17/20). Scale bars, 10 μm. (D and E) Immunofluorescence
   images showing mitochondrial membrane potential (TMRM; red) of the
   remodelin-treated and the control GV oocytes and the statistical
   histogram (n = 23/23). Scale bars, 10 μm. (F and G) Immunofluorescence
   images showing chromosome morphology (DAPI; blue) and microtubules
   (α-tubulin; red) of the remodelin-treated and the control MII oocytes
   and the statistical histogram (n = 90/59). Scale bars, 10 μm. (H) The
   in vitro first polar body (PB1) emission rates of mouse oocytes from
   the remodelin-treated and the control groups (n = 47 to 64). Each dot
   represents a single experiment replicate. (I and J) Volcano plots
   showing DEGs of the remodelin-treated and the control oocytes
   identified by T&T-seq. P < 0.05, Wald test, log[2]FC > 1.5. (K) TE
   cumulative curve of GV oocytes. The red line denotes the remodelin
   group, and the blue line represents the control group. (L) Scatter plot
   showing the RNA TE alterations of remodelin-treated oocytes compared
   with the control group. Red and blue dots denote up- and down-regulated
   genes, respectively. Up-regulated, FC > 1; down-regulated, FC < 0.67.
   (M) Bar plots showing the numbers of high-TE genes (TE > 2) and low-TE
   genes (TE < 0.5) in the remodelin-treated and the control group GV
   oocytes. Data are shown as the mean ± SD.

   To explore whether ac^4C affects mouse oocyte development by regulating
   RNA TE, we performed T&T-seq on GV-stage oocytes from the
   remodelin-treated and control groups. Similar to SLE oocytes, the
   remodelin-treated oocytes exhibited more pronounced genetic changes in
   the translation process, with 645 genes down-regulated and 278 genes
   up-regulated in the translatome, compared with 382 genes down-regulated
   and 183 genes up-regulated in the transcriptome ([125]Fig. 6, I and J).
   The cumulative frequency curve of RNA TE showed that ac^4C inhibition
   decreased oocytes’ TE ([126]Fig. 6K). Notably, the TE of most RNAs in
   the remodelin group was down-regulated (2855 genes), more than twice
   the number of RNAs with up-regulated TE (1382 genes) ([127]Fig. 6L).
   The genes with down-regulated TE in the remodelin group were also
   enriched in various metabolism and environmental information processing
   pathways (fig. S6A). Autophagy, cell cycle, protein synthesis, and
   oocyte maturation pathways were significantly enriched (fig. S6B). In
   addition, the number of genes with high TE decreased (1516 versus
   2210), while the number of genes with low TE increased (646 versus 220)
   ([128]Fig. 6M). All these results indicate that remodelin suppression
   of ac^4C modification levels could produce SLE-like oocyte
   developmental disorder phenotypes and gene translation disorders.

Zar1 is one of the targets of ac^4C modification in SLE mice

   To investigate the dysregulated genes involved in ac^4C modification in
   oocytes, we performed acRIP-seq on mouse GV-stage oocytes. The
   distribution patterns of ac^4C signal peaks from three independent
   biological replicates showed high consistency across the genome, with
   notable signal enrichment near transcription start sites (TSS) and
   transcription end sites (TES) ([129]Fig. 7A). Among the three
   replicates, 2757 transcripts were commonly captured ([130]Fig. 7B).
   Differences between the acRIP replicates may stem from biological
   differences between samples due to the limited level of ac^4C
   modification in SLE mouse oocytes used in sequencing. Further analysis
   of the detected peaks revealed enrichment of the ac^4C consensus
   sequence “CXX” ([131]Fig. 7C), indicating that our acRIP-seq experiment
   accurately captured the ac^4C-modified genes. Upon examining the
   transcriptional and translational changes of the ac^4C-modified genes
   in GV- and MII-stage SLE oocytes across the genome, we generated nine
   quadrant plots depicting gene transcription and translation levels. The
   most significant differences were observed during the GV stage, which
   were characterized by low translation levels but high transcription
   levels for most genes (368 genes in total; located in the top left
   quadrant) ([132]Fig. 7, D and E). This suggests that the ac^4C-modified
   genes in the SLE mice primarily exhibited translational impediments
   during the GV stage. Subsequent KEGG pathway enrichment analysis
   revealed significant enrichment of these ac^4C genes in pathways
   related to apoptosis, tight junction, gap junction, neurotrophin
   signaling, ribosome, FoxO signaling, regulation of actin cytoskeleton,
   sphingolipid signaling, and protein synthesis ([133]Fig. 7, F and G).
   These pathways influence crucial biological processes such as protein
   synthesis and meiotic progression in oocytes, thereby profoundly
   influencing normal oocyte development. Therefore, ac^4C modification
   plays a critical regulatory role in SLE oocyte development, and its
   down-regulation leads to impaired oocyte maturation.

Fig. 7. acRIP-seq landscape of SLE GV oocyte.

   [134]Fig. 7.
   [135]Open in a new tab

   (A) Distribution and heatmap of ac^4C modifications along substrate
   mRNA from three biological replicates of SLE GV oocytes. (B) Venn
   diagram portraying the overlap of target genes among three independent
   acRIP-seq biological replicates. (C) Motif identified by HOMER within
   acRIP-seq peaks in oocytes. (D and E) Scatter plot showing the
   acRIP-seq target genes in transcriptome and translatome of GV and MII
   oocytes (SLE versus WT; log[2]FC > 1). (F) KEGG analysis of ac^4C genes
   identified in acRIP-seq. (G) Representative KEGG pathways of ac^4C
   genes identified in acRIP-seq.

   To decipher how NAT10 interprets ac^4C modifications and modulates mRNA
   translation in SLE oocytes, we overlapped the candidate genes
   identified from acRIP-seq with the genes with down-regulated TE in SLE
   GV-stage oocytes and remodelin-treated oocytes to pinpoint downstream
   targets. Among the potential 325 downstream targets, our selection
   criteria were based on two main aspects, namely, (i) high expression
   levels in oocytes and (ii) previously documented roles in oocyte
   maturation and early embryonic development according to the existing
   literature. Therefore, Zygote arrest 1 (Zar1) caught our attention
   ([136]Fig. 8A) because it has been proven to be closely related not
   only to maternal transcriptome and translational activation, assembly
   of the mitochondria-associated ribonucleoprotein domain and
   mitochondrial clustering in oocyte maturation, but also to embryo
   development ([137]38–[138]40). We further investigated the ac^4C
   modification sites of the ZAR1 gene in both mice and humans. We found
   that the human ZAR1 gene also has potential ac^4C modification sites,
   although these sites are not conserved between the two species
   ([139]Fig. 8B). Moreover, in acRIP-seq of oocytes from the WT and SLE
   mice, Zar1 was also significantly reduced in the SLE group ([140]Fig.
   8C). Quantitative real-time PCR validated that Zar1 showed similar
   transcriptional expression levels but decreased translation in SLE
   GV-stage oocytes compared with WT GV-stage oocytes ([141]Fig. 8D).
   Immunofluorescence also confirmed that the expression of ZAR1 was
   decreased in the SLE GV-stage oocytes and remodelin-treated oocytes
   ([142]Fig. 8, E to H). These results demonstrate that Nat10 binds to
   the Zar1 mRNA and promotes the translation of Zar1 in an
   ac^4C-dependent manner, which may be one of the mechanisms involved in
   the regulation of oocyte development in SLE mice.

Fig. 8. Zar1 is one of the target genes of ac^4C modification in SLE oocytes.

   [143]Fig. 8.
   [144]Open in a new tab

   (A) Venn diagram portraying the overlap of acRIP-seq target genes,
   down-regulated TE genes in SLE GV oocytes, and down-regulated TE genes
   in remodelin-treated oocytes. (B) The diagram displayed ac^4C-modified
   potential sites of ZAR1 in Homo sapiens and Mus musculus. Sites were
   predicted by online tools [[145]www.rnanut.net/paces/ ([146]61)]. CDS,
   coding sequences; UTR, untranslated regions. (C) Fold enrichment levels
   of Zar1 in acRIP-seq from WT and SLE mouse GV oocytes (n = 3/3). (D)
   Quantitative RT-PCR results showing the relative expression levels of
   mouse Zar1 mRNA in SLE and WT GV oocytes. Using EGFP expression levels
   as a reference for each sample (n = 3/3). Data were presented as
   mean ± SEM. (E and F) Immunofluorescence images showing ZAR1 (red) and
   nuclei (DAPI; blue) of the WT and SLE GV oocytes and the statistical
   histogram (n = 27/21). Scale bars, 10 μm. Data are shown as the
   mean ± SD. (G and H) Immunofluorescence images showing ZAR1 (red) and
   nuclei (DAPI; blue) of the remodelin-treated and the control GV oocytes
   and the statistical histogram (n = 21/23). Scale bars, 10 μm. Data are
   shown as the mean ± SD.

Nicotinamide improves oocyte quantity and quality in SLE mice by enhancing
ac^4C modification

   Sirtuin 1 (Sirt1) can inhibit the activity of NAT10 by mediating its
   deacetylation, while nicotinamide (NIA) can block the inhibitory effect
   of Sirt1 on NAT10 and increase the occupancy of NAT10 on rRNA
   ([147]41). Therefore, we hypothesized that NIA could enhance NAT10
   activity and ac^4C modification levels in oocytes of SLE mice. Previous
   studies have predominantly used doses ranging from 50 to 500 mg/kg,
   without toxic damage in mice ([148]42, [149]43). On the basis of this,
   we performed preliminary experiments with different NIA concentration
   gradients (50, 100, 200, and 500 mg kg^–1 day^–1 of body weight). In
   consideration of animal welfare and ethical guidelines, our preliminary
   study included six mice per group, with three mice allocated for
   GV-stage oocyte collection and three for MII-stage oocyte collection,
   to identify the optimal concentration for improving SLE oocyte
   quantity. Our results indicated that 100 mg/kg was the minimum
   effective concentration that enhanced SLE oocyte numbers (fig. S7, A
   and B). Subsequently, a concentration of 100 mg/kg was chosen for
   formal experiments. The experimental workflow is illustrated in
   [150]Fig. 9A. Notably, supplementation with NIA notably increased ac^4C
   modification levels and Zar1 expression in oocytes from the SLE mice
   ([151]Fig. 9, B to D). These findings highlight the potential role of
   NIA in improving oocyte quality and female reproductive life span in
   animals with SLE.

Fig. 9. Nicotinamide can improve the quantity and quality of oocytes in SLE
mice by increasing ac^4C modification.

   [152]Fig. 9.
   [153]Open in a new tab

   (A) A timeline scheme for nicotinamide (NIA) treatment. ig,
   intragastric administration; ip, intraperitoneal; h, hours. (B)
   Immunofluorescence images showing ac^4C (green), ZAR1 (red), and nuclei
   (DAPI; blue) of the SLE oocytes with and without NIA treatment and the
   WT GV oocytes. Scale bars, 10 μm. (C and D) Statistical histogram of
   fluorescence intensity for ac^4C and ZAR1 (n = 17/13/16). (E)
   Comparison of the number of GV oocytes collected from the SLE mice with
   and without NIA treatment and the WT mice after superovulation
   (n = 10/10/10). COH, controlled ovarian hyperstimulation. (F and G)
   Immunofluorescence images showing mitochondrial membrane potential
   (TMRM; red) of the SLE GV oocytes with and without NIA treatment and
   the WT GV oocytes and the statistical histogram (n = 11/13/14). Scale
   bars, 10 μm. (H) Comparison of the number of MII oocytes collected from
   the SLE mice with and without NIA treatment and the WT mice after
   superovulation and trigger (n = 10/10/10). (I and J) Immunofluorescence
   images showing mitochondrial membrane potential (TMRM; red) of the SLE
   MII oocytes with and without NIA treatment and the WT MII oocytes and
   the statistical histogram (n = 17/11/14). Scale bars, 10 μm. (K and L)
   Immunofluorescence images showing chromosome morphology (DAPI; blue)
   and microtubules (α-tubulin; red) of the SLE MII oocytes with and
   without NIA treatment and the WT MII oocytes and the statistical
   histogram (n = 48/50/43). Scale bars, 10 μm. (M) The in vitro PB1
   emission rates of mouse oocytes from the SLE mice with and without NIA
   treatment and the WT mice (n = 23 to 40). Each dot represents a single
   experiment replicate. Data are shown as the mean ± SD.

   Next, we performed superovulation stimulation and found that GV-stage
   oocytes from the SLE mice treated with NIA showed a significant
   increase in number ([154]Fig. 9E) and a notable improvement in the
   mitochondrial membrane potential ([155]Fig. 9, F and G). Consistently,
   after superovulation and hCG trigger, more MII-stage oocytes were
   obtained from the NIA treatment SLE mice ([156]Fig. 9H). The
   mitochondrial membrane potential of the SLE MII-stage oocytes after NIA
   treatment also improved ([157]Fig. 9, I and J). Immunofluorescent
   staining and confocal imaging revealed that most oocytes from the NIA
   treatment SLE mice had barrel-like spindle apparatuses with
   well-aligned chromosomes, resembling those in the WT mice. In contrast,
   oocytes from the untreated SLE mice exhibited various morphologically
   aberrant spindles with misaligned chromosomes ([158]Fig. 9, K and L).
   To determine whether NIA could directly improve the maturation rate of
   GV-stage oocytes, we removed granulosa cells from the GV-stage oocytes
   of the SLE mice and cultured them in a medium with or without NIA. We
   conducted preliminary IVM experiments using SLE GV-stage oocytes with
   nicotinamide concentrations of 0.1, 1, and 5 mM ([159]44). Our
   observations revealed that 1 mM showed a promising trend in improving
   SLE oocyte maturation rates (fig. S7C). We found that the maturation
   rate of SLE oocytes in vitro improved after NIA treatment ([160]Fig.
   9M).

   To evaluate whether NIA could enhance early embryonic development in
   SLE mice, we performed IVF using MII-stage oocytes collected from the
   WT, SLE, and SLE + NIA mice fertilized with sperm from normal male
   mice. Each stage of embryonic development from the SLE + NIA mice
   showed an increasing trend (fig. S8A). Next, we conducted natural
   mating experiments to assess the reproductive outcomes. Successful
   mating was confirmed through vaginal plug observation, after which
   females were individually housed to monitor delivery outcomes. Our
   findings revealed that the SLE mice demonstrated compromised fertility
   compared with the WT controls, aligning with our human population data
   ([161]Fig. 1H). Notably, NIA supplementation showed encouraging trends
   in improving live birth rates and offspring body weight in the SLE
   mice, although litter size differences did not reach statistical
   significance (fig. S8, B to D). Histological examination of major
   organs (heart, lung, liver, spleen, and kidney) showed no significant
   differences among the offspring from all three groups (fig. S8, E to
   I), indicating that SLE offspring, when successfully delivered, develop
   normal organ structures. This observation also confirmed that our NIA
   supplementation protocol demonstrated no apparent toxicity to the
   offspring. Collectively, on the basis of the comprehensive analyses of
   oocytes, embryos, and reproductive outcomes, our findings demonstrate
   that NIA supplementation improves both the quantity and quality of
   oocytes in SLE mice.

   Aside from fertility rescue, the impact of NIA on SLE disease itself
   also deserves attention. Our goal is to safely improve fertility in
   SLE. Therefore, we observed the SLE indicators in SLE mice after NIA
   treatment. Notably, NIA treatment also improved SLE inflammation
   levels. NIA treatment significantly improved glomerular morphology and
   matrix deposition in SLE mice (fig. S9, A and B). Inflammatory
   infiltrates in the lungs were also reduced (fig. S9C). In addition,
   megakaryocyte hyperplasia (fig. S9D) and splenomegaly (fig. S9, E and
   F) were resolved. Inflammatory infiltration of the liver (fig. S9G) and
   hepatomegaly (fig. S9H) were also reduced. Notably, serum
   anti–double-stranded DNA (dsDNA) antibody levels were significantly
   reduced in the NIA treatment group (fig. S9I). In conclusion, our data
   show that supplementation with NIA improves the quality and quantity of
   oocytes in SLE mice and aids in the recovery from SLE disease.

DISCUSSION

   Given the adverse fertility outcomes in women with SLE, there is a
   pressing need to elucidate the underlying mechanisms that reduce oocyte
   quality and competence ([162]45). However, the small number of oocytes
   available for study, coupled with the limitations of various omics
   techniques, has left the gene transcription and translation landscapes
   in SLE oocytes largely unexplored. In this study, we used
   ultrasensitive translatomics and transcriptomics to document mRNA
   translation patterns in SLE mouse oocytes. Our findings revealed
   notable down-regulation of gene translation during the GV stage, which
   was modified by ac^4C. Furthermore, through microcellular acRIP-seq of
   SLE mouse GV-stage oocytes, we identified ZAR1 as one of the ac^4C
   target genes involved in regulating oocyte quality in SLE. We found
   that NIA was able to safely and effectively enhance both the quantity
   and quality of oocytes in the SLE mice by promoting ac^4C modification.
   This suggests that NIA supplementation could serve as a potential
   therapeutic approach for improving oocyte quality and achieving better
   fertility outcomes in women with SLE who are preparing for pregnancy.

   Traditionally, gene transcription has been viewed as the primary
   mechanism governing GV-stage oocyte development, with translation
   regarded as a secondary process that becomes more prominent after GV
   breakdown ([163]46). However, our study challenges this conventional
   perspective, demonstrating that translation is critical during the GV
   stage. We observed a substantial reduction in gene TE in GV-stage
   oocytes of SLE mice compared with their normal counterparts. This
   impairment adversely affects metabolic processes, protein synthesis,
   and meiosis, ultimately diminishing oocyte quality. Although the
   overall transcriptional differences between SLE and WT mouse oocytes
   were less pronounced than translational changes, there were still some
   DEGs between the two transcriptome groups. Transcriptomics analysis
   revealed 532 down-regulated and 338 up-regulated genes in SLE GV-stage
   oocytes compared with WT controls. At the MII stage, 456 genes showed
   decreased expression, whereas 598 genes were up-regulated. These DEGs
   may contribute to the developmental defects in SLE oocytes, warranting
   further investigation.

   The gene expression differences between SLE and WT oocytes were less
   pronounced at the MII stage than at the GV stage, which can be
   attributed to two main factors. First, the inherent biological
   heterogeneity of SLE pathology, where disease manifestations vary among
   individuals and may result in some SLE oocytes displaying
   characteristics more similar to WT oocytes and the mixed samples might
   cause mild differences in affected genes. Second, the decreased oocyte
   yield at the MII stage compared with the GV stage ([164]Fig. 2, A and
   D) suggests a natural selection process during oocyte maturation,
   whereby severely affected oocytes may have failed to progress through
   development and were eliminated. Consequently, the MII-stage oocytes
   available for sequencing likely represented a subset that had
   successfully overcome developmental barriers, potentially explaining
   the reduced transcriptional divergence observed at this stage.

   Our findings indicated that GV-stage oocytes in SLE mice exhibited a
   marked decrease in NAT10 translation, leading to reduced ac^4C
   modification. This reduction contributes to widespread gene translation
   disorders. Notably, similar reductions in NAT10 expression and ac^4C
   modification have been reported in peripheral blood CD4^+ T cells of
   patients with SLE ([165]47), suggesting that these mechanisms may also
   be relevant in humans. Moreover, a comparable decrease in NAT10
   expression has been observed in ankylosing spondylitis ([166]48), while
   rheumatoid arthritis shows NAT10 hyperactivation ([167]49). This
   variability underscores the complex, disease-specific roles of NAT10 in
   autoimmune conditions, highlighting the need for further research to
   elucidate its multifaceted roles and therapeutic potential.

   Our study revealed that NIA could enhance ac^4C modification levels in
   oocytes. The mechanistic link between nicotinamide, NAT10 activation,
   and ac^4C modifications has been documented in previous studies. SIRT1
   is a classical deacetylase protein, and NIA can directly inhibit SIRT1
   activity and its deacetylation function in vitro ([168]50). Oral
   administration of NIA in mice can inhibit systemic SIRT1 function and
   increase overall protein acetylation levels ([169]42). Moreover,
   acetylation plays a regulatory role in NAT10 activity, and
   SIRT1-mediated deacetylation of NAT10 protein can lead to its activity
   suppression ([170]41). According to previous reports, NAT10 is the sole
   writer protein for ac^4C modification ([171]37). Therefore,
   nicotinamide may enhance NAT10 protein activity by inhibiting
   SIRT1-mediated deacetylation of NAT10, thereby increasing ac^4C
   modification levels. In addition, other studies have shown that
   nicotinamide, a precursor of nicotinamide adenine dinucleotide
   (oxidized form), has direct metabolic improvement effects ([172]51).
   During ac^4C modification, acetyl–coenzyme A provides acetyl groups,
   and adenosine 5′-triphosphate/guanosine 5′-triphosphate hydrolysis
   provides energy ([173]52, [174]53). Thus, NIA may also improve SLE
   oocyte quality by increasing the availability of substrates required
   for ac^4C modification.

   The increase in ac^4C modification in SLE oocytes correlated with an
   improvement in systemic inflammatory symptoms associated with SLE. This
   finding suggests that NIA may alleviate SLE-related inflammation,
   potentially through improved ac^4C modification in immune cells,
   although this requires further confirmation. Nicotinamide has also
   shown a direct metabolic improvement effect in other studies ([175]51),
   so it may also improve SLE oocyte quality by improving metabolism,
   which warrants further investigation. Our findings align with those of
   “Imaruoka et al. ([176]54),” who reported that nicotinamide ameliorated
   glomerular damage and pregnancy outcomes in MRL/MpJ-Faslpr/lpr
   spontaneous lupus mice. Their study showed that NIA treatment led to
   prolonged gestation, fewer stillbirths, and increased fetal weight in
   SLE mice. These outcomes may also be attributable to improved oocyte
   quality, which enhanced embryo quality and overall pregnancy outcomes.
   Together, the promising application of NIA in our study and previous
   research underscores its potential to positively influence reproductive
   health in patients with SLE.

   ZAR1 is a conserved gene essential for the transition from oocyte to
   embryo, expressed in vertebrate ovaries ([177]55). Whole-exome
   sequencing studies, including one by Chen and colleagues ([178]56)
   involving 1030 patients with POI, identified ZAR1 as highly associated
   with POI. Misexpression of ZAR1 in aging oocytes correlates with poor
   cleavage rates and embryo quality ([179]57). These findings suggest
   that aberrant ZAR1 expression contributes to oocyte and ovarian aging,
   mirroring the situation seen in patients with SLE. Previous studies
   reported that Zar1 knockout female mice exhibit complete embryonic
   arrest at the one-cell stage postfertilization ([180]40). In contrast,
   SLE mice demonstrate a milder phenotype, with a small proportion of
   embryos capable of further development. This less severe phenotype
   likely stems from the residual Zar1 expression in SLE mouse oocytes, as
   opposed to the complete absence of Zar1 in knockout models. Currently,
   nuclear transfer is the only reported method for rescuing oocytes with
   ZAR1 deficiency ([181]58), but this technique faces substantial
   clinical and ethical challenges. Our study demonstrated that
   ac^4C-mediated epigenetic modifications regulate the translation level
   of ZAR1. Notably, oral administration of NIA, a form of vitamin B3,
   effectively increases ac^4C levels in oocytes and consequently boosts
   ZAR1 expression. In addition, given that NIA is water-soluble and
   easily excreted in urine when excessive, its safety profile is
   favorable ([182]59). Its effectiveness, simplicity, and safety present
   a promising approach for managing POI in clinical settings. Future
   studies should focus on elucidating the detailed mechanisms by which
   nicotinamide influences ZAR1 and determining its efficacy in patients
   with POI.

MATERIALS AND METHODS

Ethics statement

   All procedures performed in studies involving human participants were
   approved by the Medical Ethical Committee of the Sixth Affiliated
   Hospital, Sun Yat-sen University (2024ZSLYEC-230), and all the related
   data were extracted from the electronic medical record.

Retrospective study design and population

   It was a single-center retrospective cohort study. All women undergoing
   IVF/ICSI cycles from 2014 to 2023 in the Reproductive Medicine Center
   of the Sixth Affiliated Hospital, Sun Yat-sen University in China were
   reviewed.

   Fifty women with a history of rheumatologically confirmed SLE were
   included in the SLE group. All included patients with SLE were
   clinically diagnosed with SLE in the department of rheumatology before
   they were treated with assisted reproductive technique (ART). The
   classification criteria for patients with SLE were based on the 2012
   Systemic Lupus International Collaborating Clinics criteria ([183]17),
   in which patients met at least four classification criteria, including
   at least one clinical criterion and one immunological criterion, to be
   diagnosed with SLE. Included patients with SLE had been evaluated by
   rheumatologists and reproductive clinicians before performing IVF
   techniques to ensure they were in remission (clinical remission or
   complete remission) for at least 6 months. The criteria for exclusion
   were as follows: (i) diagnosed with other rheumatological diseases,
   (ii) women with ovarian tumors, (iii) preimplantation genetic testing
   cycles, (iv) oocyte donation cycles, (v) women with chromosomal
   abnormalities, (vi) congenital or acquired absence of ovaries, (vii)
   women with a history of ovarian surgery, and (viii) fertility
   preservation. All had been evaluated by clinicians to ensure that they
   were in the rest phase of the disease before IVF/ICSI cycles.
   Considering the complexity of etiology in patients with SLE undergoing
   ART, to eliminate the potential confounders in the patients with SLE
   and comparisons, a PSM at a 1:4 ratio was performed to balance the
   distribution of sample and clinical characteristics including female
   age (years), female BMI (kilograms per square meter), infertility
   duration (years), infertility type (primary and secondary), ovarian
   endometriosis, and polycystic ovarian syndrome. Details about the
   patient’s enrollment and comparison are shown in fig. S1.

Mice

   The female wild-type C57BL/6 mice were purchased from the GemPharmatech
   Co. Ltd. (Guangdong, China) and underwent a 1-week adaptation period
   before experimentation. All animal care and experimental procedures
   were performed following the guidelines of the Institutional Animal
   Care and Use Committee (IACUC) of Tsinghua University, Beijing, China.
   The mouse facility maintained a 12-hour/12-hour dark/light cycle, 20°
   to 26°C temperature range, and 40 to 70% humidity level, with ad
   libitum access to water and food. All procedures were approved by the
   Animal Care and Use Committee of the Sixth Affiliated Hospital, Sun
   Yat-sen University (Guangzhou, China) (ethical approval numbers:
   IACUC-2021041401, IACUC-2023042302, and IACUC-2023083001). The skin on
   the right ears of the mice (6-week-old females) was treated with 5%
   imiquimod cream epicutaneously, three times a week for 6 weeks as
   previously described, to induce a lupus phenotype ([184]20–[185]22).
   Control mice were age-matched mice without treatment.

Histological analysis

   Hematoxylin and eosin (H&E) staining was conducted according to
   previously established standard procedures ([186]60). Briefly, freshly
   collected organ samples were fixed in 4% paraformaldehyde at room
   temperature for 24 hours. Paraffin-embedded samples were sectioned into
   slides with a thickness of 4 μm. The slides underwent deparaffinization
   with xylene and rehydration, followed by H&E staining. The slides were
   then dehydrated and mounted with neutral resins.

Oocyte collection

   For the collection of GV-stage oocytes, mice were injected
   intraperitoneally with 10 U of pregnant mare serum gonadotropin (PMSG)
   (Ningbo Second Hormone Factory, Hangzhou, China). After 48 hours,
   oocytes at the GV stage were harvested from ovaries by puncturing
   antral follicles with a sterile needle in M2 medium (Sigma-Aldrich, MO,
   United States, catalog no. M7167). Oocytes and granulosa cells were
   isolated with 0.1% hyaluronidase (Sigma-Aldrich, catalog no.
   37326-33-3), and oocytes were moved to maturation medium (Nanjing Aibei
   Biotechnology, M2115) covered with mineral oil (Nanjing Aibei
   Biotechnology, M2460) at 37°C in 5% CO[2] for further culture. In TMRM
   staining experiments, IBMX (MedChemExpress, Shanghai, China, catalog
   no. HY-12318) was added to the maturation medium to inhibit the
   occurrence of germinal vesical breakdown to maintain the GV state of
   oocytes.

   For the collection of MII-stage oocytes, mice were injected with 10 U
   of PMSG and, after 48 hours, with 10 U of hCG (Ningbo Second Hormone
   Factory). Cumulus-enclosed oocytes were isolated from the oviduct 14
   hours after hCG stimulation. MII-stage oocytes with a first polar body
   were separated from the cumulus cells with hyaluronidase and repeatedly
   pipetting in M2 medium.

In vitro maturation

   The GV oocytes stripped of granulosa cells were transferred to
   maturation medium covered with mineral oil at 37°C in 5% CO[2] for 13
   hours. The ejection of the first polar body was observed under a
   stereoscope.

In vitro fertilization

   Cauda epididymides from C57BL/6 adult male mice were lanced in 200 μl
   of equilibrated Human Tubal Fluid medium (Nanjing Aibei Biotechnology,
   M1135) to release sperm, followed by capacitation for 1 hour in an
   incubator at 37°C with 5% CO[2]. Then, capacitated sperm were added to
   cumulus-enclosed oocytes in 100 μl of HTF for 6 hours at 37°C with 5%
   CO[2]. To facilitate the observation of embryonic form, excess sperm
   were washed away in KSOM (Nanjing Aibei Biotechnology, M1435), and
   embryos were transferred to the KSOM droplets covered with mineral oil
   at 37°C in 5% CO[2] for culture. Embryo development was observed by
   microscope: 18 hours (2 cells), 42 hours (4 cells), 54 hours (8 cells),
   66 hours (morula), and 68 hours (blastocyst).

Immunofluorescence staining

   Denuded oocytes were fixed in 4% paraformaldehyde (Servicebio, G1101)
   at room temperature for 30 min. The fixing solution was removed, and
   the oocytes were incubated in phosphate-buffered saline (PBS) with 0.5%
   Triton X-100 at room temperature for 30 min to permeabilize.
   Subsequently, the oocytes underwent three washes with 3% bovine serum
   albumin (BSA) in PBS, followed by a 1-hour incubation with 3% BSA at
   room temperature. The primary antibody (diluted with 3% BSA; 1:200) was
   then applied to the oocytes, and they were placed in a wet box
   overnight at 4°C. After being washed three times with 0.3% BSA, the
   oocytes were incubated with a secondary antibody (diluted with 0.3%
   BSA; 1:500) at room temperature for 1 hour before being placed in a
   light-free wet box. For 4′,6-diamidino-2-phenylindole (DAPI)
   counterstaining of the nucleus, the oocytes underwent three washes with
   0.3% BSA and were then incubated with DAPI solution at room temperature
   for 15 min, out of light. Following another three washes with 0.3% BSA,
   the oocytes were placed in an aliquot of 20 to 30 μl of 0.3% BSA at
   4°C, out of light. The oocytes were observed, and pictures were
   captured under an inverted confocal microscope. Antibodies used in this
   study are listed as follows: α-tubulin (Abcam, ab7291), NAT10
   (Proteintech, 13365-1-AP), ac^4C (Abcam, ab252215), ZAR1 (HuaBio,
   ER1918-78), CY3 goat anti-mouse immunoglobulin G (IgG; EarthOx,
   E031610-01), and CY3 goat anti-rabbit IgG (EarthOx, E031620-01).

   In ac^4C and ZAR1 costaining experiments, a fluorescent-labeled
   antibody technique was used to prevent fluorescent secondary antibodies
   from binding to both primary antibodies at the same time due to the
   same species origin of two primary antibodies. The antibody label
   experiments were performed almost according to the manufacturer’s
   instructions of LinKine AbFluor 488/555 Labeling Kit (LinKine, KTL0520;
   Linkine, KTL0530). Subsequent fluorescence staining experiments can be
   directly photographed without the addition of a secondary antibody.

Enzyme-linked immunosorbent assay

   Concentrations of anti-dsDNA antibody in mouse serum were detected by
   enzyme-linked immunosorbent assay kits (Alpha Diagnostic International,
   5120) according to the manufacturer’s instructions.

Ultrasensitive T&T-seq

   The ultrasensitive T&T-seq procedures were performed according to a
   previously published protocol ([187]11, [188]15). Each mouse oocyte
   sample contained four oocytes, which were lysed in 10 μl of sample
   buffer consisting of 9 μl of lysis buffer and 1 μl of RNase inhibitor
   (Vazyme, n712) on ice for 10 min. The lysates were then divided into
   two parts, with 1.5 μl used for transcriptome analysis and 8.5 μl for
   translatome analysis. For translatome analysis, the RiboLace beads
   (Immagina, RL001) were functionalized following the manufacturer’s
   instructions and then divided into 10 μl of aliquots in separate 1.5 ml
   of Eppendorf tubes. The solution of functional RiboLace beads was
   removed and resuspended with 8 μl of lysates, 1 μl of RNase inhibitor,
   and 8 μl of binding buffer [RNase-free water supplemented with
   cycloheximide at a concentration of 100 μg/μl, dithiothreitol at a
   concentration of 1 mm, MgCl[2] at a concentration of 5 mm, NaCl at a
   concentration of 150 mm, and tris-HCl (pH adjusted to 8.0)], followed
   by incubation on a rotator at 3 rpm at 4°C for 1 hour. The solution was
   discarded after incubation, and the beads were washed twice with
   W-buffer (Immagina, RL001) using a magnet. The beads were resuspended
   with 12 μl of RLT (QIAGEN,74 004) supplemented with 10%
   β-mercaptoethanol and 1% glycol. Incubated at room temperature for 5
   min, the solution was collected in a new PCR tube, and the beads were
   discarded. Fifteen microliters of 2 M LiCl and 54 μl of VAHTS RNA Clean
   Beads (Vazyme, N412) were added to the abovementioned solution. The
   ribosome binding full-length RNA was isolated and purified according to
   the manufacturer’s instructions. Both total RNA (the 2 μl of lysates
   described above) and ribosome binding full-length RNA were reverse
   transcribed to cDNA and amplified for 18 PCR cycles, according to the
   protocol from Single Cell Full Length mRNA-Amplification Kit (Vazyme,
   n712). VAHTS DNA Clean Beads (Vazyme, N411) were used to purify the
   cDNA amplification products. Furthermore, the concentration of cDNA was
   assessed by Qubit (Invitrogen, USA), and the peak was detected by
   Bioanalyzer 2100 (Agilent, CA, USA), which was ≈2000 base pairs (bp).
   The indexed libraries were constructed using TruePrep DNA Library Prep
   Kit V2 for Illumina (Vazyme, TD502). VAHTS DNA Clean Beads (Vazyme,
   N411) were used to fractionate and purify the amplified product to 250
   to 450 bp. The quantities of libraries were identified by Qubit
   (Invitrogen, USA) and Bioanalyzer 2100 (Agilent, CA, USA). Paired-end
   sequencing was conducted with the Illumina NovaSeq 6000 platform
   (Azenta Life Sciences, Hangzhou, China), with the sequencing mode of
   PE150.

Quantitative reverse transcription PCR

   The pcDNA3.1-EGFP plasmid was used to overexpress enhanced green
   fluorescent protein (EGFP) DNA, followed by purification using StarPrep
   DNA Gel Extraction Kit (GenStar, D205-04). Linearized DNA was
   transcribed into RNA in vitro using HyperScribe T7 High Yield RNA
   Synthesis Kit (APExBIO, K1047) and subsequently purified. The EGFP RNA
   was incorporated into the T&T-seq system after oocyte lysis for reverse
   transcription and library construction. Quantitative reverse
   transcription PCR (RT-PCR) was then performed using the ABI 7500
   instrument. The primers are listed in table S2.

acRIP sequencing

   For ultrasensitive acRIP in oocytes, the acRIP procedure was optimized
   as follows: Protein A coupled with HaloTag was purified using HaloTag
   beads (Promega, G7281). Anti-ac^4Cantibody (2.5 μl; Abcam, ab252215)
   was incubated with 50 μl of protein A–HaloTag beads in the purification
   buffer with rotation for 2 hours at room temperature. After incubation,
   the beads were washed with a purification buffer. A total of 200
   oocytes were lysed in 25 μl of purification buffer with 0.5% NP-40 and
   0.3% RNase inhibitor on ice for 20 min. The lysates were mixed and
   divided into two parts, of which 1.5 μl of lysates were used for input
   and 23.5 μl for acRIP. For acRIP, the purified protein–ac^4C antibody
   compounds were incubated with 23.5 μl of lysates at 1400 rpm for 2
   hours at room temperature. Then, the beads were washed with a
   purification buffer, and the solution of HaloTag beads was removed. The
   beads were resuspended with 12 μl of RLT (QIAGEN,74 004) supplemented
   with 10% β-mercaptoethanol and 1% glycoblue. Incubated at room
   temperature for 5 min, the solution was collected in a new PCR tube,
   and the beads were discarded. A total of 15 μl of 2 M LiCl and 54 μl of
   VAHTS RNA Clean Beads (Vazyme, N412) were added to the solution
   described above. Input RNA and acRIP RNA were reverse transcribed to
   cDNA, and their concentrations were detected. Then, indexed libraries
   were constructed, and paired-end sequencing was conducted.

T&T-seq data analysis

   Raw reads were processed to remove adaptor contaminants and low-quality
   bases using fastp. The clean reads were aligned to the mouse genome
   (mm10) using HISAT2, and uniquely mapped reads were counted with
   featureCounts. We quantified gene expression levels with fragments per
   kilobase of exon model per million mapped fragments. DEGs were assessed
   with the DESeq2 package with a cutoff: adjusted P < 0.05 and fold
   change (FC) ≥ 1.5. TE was calculated using the FPKM + 1 of the
   transformant divided by FPKM + 1 of the transcriptome. Statistical
   analyses were performed using R software (version 4.3.1).

acRIP-seq data analysis

   First, to obtain high-quality clean FASTQ reads, fastp software was
   first used to remove the adaptor and low-quality reads. Clean reads
   were then aligned to the reference genome (mm10) using bowtie2
   software. After alignment, the Sequence Alignment/Map format files were
   sorted and converted to Binary Alignment/Map format (BAM) files. Last,
   the BAM-format reads for each paired IP + input sample were processed
   by MACS3 software to identify significant acetylation peaks in each IP
   sample. HOMER software was used for the discovery of the acetylation
   motif. R package ChIPseeker was used to characterize the interval
   annotation analysis of ac^4C peaks. Visualization of the ac^4C peak of
   ZAR1 was performed using the Integrated Genome Viewer.

Statistical analysis

   All experiments were performed at least in biological triplicates
   unless otherwise indicated. Animal experiments included six animals per
   group, and animals that died accidentally, such as due to manipulation
   during the experiment, were excluded. Data were analyzed and presented
   using Statistical Package for Social Sciences software (version 26.0,
   International Business Machines Corporation, United States) and R
   (version 4.3.1). Continuous data do not conform to the normal
   distribution of the Kruskal-Wallis nonparametric method, in accordance
   with normal distribution by Student’s t test. Categorical data were
   presented as the number of cases and frequency (percentage), and a
   chi-square test was used to assess the group differences. Values of
   P < 0.05 were deemed statistically significant. Statistically
   significant values of P < 0.05, P < 0.01, and P < 0.001 were indicated
   by one, two, and three asterisks, respectively.

Acknowledgments