Abstract

Background

   Sterigmatocystin (STE) is a mycotoxin widely found in contaminated food
   and foodstuffs, and excessive long-term exposure to STE is associated
   with several health issues, including infertility. However, there is
   little information available regarding the effects of STE toxin on the
   female reproductive system, particularly concerning oocyte maturation.

Methods

   In the present study, we investigated the toxic effects of STE on mouse
   oocyte maturation. We also used Western blot, immunofluorescence, and
   image quantification analyses to assess the impact of STE exposure on
   the oocyte maturation progression, mitochondrial distribution,
   oxidative stress, DNA damages, oocyte ferroptosis and asymmetric
   division defects.

Results

   Our results revealed that STE exposure disrupted mouse oocyte
   maturation progression. When we examined the cellular changes following
   100 µM STE treatment, we found that STE adversely affected polar body
   extrusion and induced asymmetric division defects in oocytes.
   RNA-sequencing data showed that STE exposure affects the expression of
   several pathway-correlated genes during oocyte meiosis in mice,
   suggesting its toxicity to oocytes. Based on the RNA-seq data, we
   showed that STE exposure induced oxidative stress and caused DNA damage
   in oocytes. Besides, ferroptosis and α-tubulin acetylation were also
   found in STE-exposed oocytes. Moreover, we determined that STE exposure
   resulted in reduced RAF1 protein expression in mouse oocytes, and
   inhibition of RAF1 activity also causes defects in asymmetric division
   of mouse oocytes.

Conclusions

   Collectively, our research provides novel insights into the molecular
   mechanisms whereby STE contributes to abnormal meiosis.

Supplementary Information

   The online version contains supplementary material available at
   10.1186/s13048-024-01499-w.

   Keywords: Sterigmatocystin, RAF1, Oxidative stress, Ferroptosis, Oocyte

Introduction

   The prerequisite to obtaining a healthy embryo is the production of a
   high-quality oocyte [[36]1], and oocyte maturation with appropriate
   meiotic resumption is important for the acquisition of high-quality
   oocytes [[37]2]. In mammalian oocytes, meiotic maturation is
   characterized by an asymmetric division, which produces a large egg and
   a small polar body [[38]3]. In order to maintain maternal stores,
   oocytes expel a minimal amount of cytoplasm with half chromosomes,
   which is essential to the early embryonic development [[39]4]. In
   addition, mammalian oocyte maturation is characterized by multi-stage,
   accurately orchestrated and systematic process [[40]5]. As a hallmark
   of meiotic resumption, germinal vesicle breakdown (GVBD) represents the
   initiation of oocyte maturation [[41]6]. After GVBD, the first meiotic
   spindle is assembled at the center of the oocyte and subsequently
   migrates to one side where an actin-rich cortical domain is formed that
   overlies the metaphase I (MI) spindle [[42]7]. The spindle subsequently
   migrates to the cortex, and first polar body (Pb1) extrusion takes
   place, with the oocyte reaching and remaining at the metaphase II (MII)
   stage until fertilization [[43]8].

   There is an extensive and serious problem of mycotoxins contaminating
   food in many countries, particularly developing countries [[44]9].
   Sterigmatocystin (STE) is a common food contaminant produced by fungi
   and exerts potently negative economic effects on the biotechnological,
   agricultural, and food industries [[45]10]. The mycotoxin STE is
   produced principally by Aspergillus fungi [[46]11] and is hepatotoxic
   and nephrotoxic in animals; given this, STE has been classified as a
   possible human carcinogen (group 2B) by the International Agency for
   Research on Cancer [[47]12, [48]13]. There are indications that dairy
   cattle suffer from diarrhea and death as a result of substantial levels
   of STE in their feed [[49]14]. Epidemiological evidence also highlights
   the existence of associations between STE exposure and the risk of
   cancer development [[50]12]. In vivo studies have revealed that STE
   induces tumors in several species of animals such as mice [[51]15],
   rats [[52]16], monkeys [[53]17], and fish [[54]18]. Exposure to STE has
   been additionally shown to induce oxidative stress-related impairment
   in rat liver and kidneys [[55]19]. STE has been reported to markedly
   reduce cell proliferation in different mammalian cells [[56]20] and can
   generate genotoxicity as it is able to form DNA-adducts and induce DNA
   damage [[57]21]. Elevated incidence rates for liver and stomach cancers
   have been reported in areas with high levels of STE contamination
   [[58]22]; teratogenic effects, embryonic morality, and embryonic
   developmental disorders resulting from STE exposure have been reported
   in chickens [[59]23]. Nonetheless, the effects and underlying
   mechanisms of STE exposure on oocytes remain unclear.

   In light of this, the aim of the present study was to investigate the
   negative effects of STE on the maturation progress of mouse oocytes. We
   assessed several critical indicators during oocyte nuclear and
   cytoplasmic maturation such as first polar body extrusion, defects in
   asymmetric division, cytoskeletal assembly, and oxidative stress.

Results

STE toxin exposure reduces the polar body extrusion rate of mouse oocytes

   Oocytes were collected from adolescent mice that had been injected with
   PMSG 48 h previously. The collected oocytes were then cultured in M16
   medium for 3, 8, and 14 h, by which time they had reached the germinal
   vesicle breakdown (GVBD), metaphase I (MI), and metaphase II (MII)
   stages, respectively. To study the STE toxicity on mice oocyte
   maturation progress, five groups of oocytes were treated with a
   concentration gradient of STE solution: 0 µM, 25 µM, 50 µM, 100 µM and
   200 µM. We selected oocytes cultured in vitro for 14 h to observe the
   first polar body (Pb1) extrude, as shown in Fig. [60]1A, we noted that
   almost all oocytes matured and extruded their first polar body in the
   control group, however, exposure to STE inhibited oocyte maturation.
   Compared with the control group (82.7 ± 3.68% [number of
   replicates = 3, number of oocytes = 145, and number of mice = 6,
   p < 0.01]), the PB1 extrusion rates were markedly declined after STE
   treatment in a dose-dependent manner. And the proportion of PB1 was
   53.0 ± 6.16% [number of replicates = 3, number of oocytes = 132, and
   number of mice = 6], 44.2 ± 5.14% [number of replicates = 3, number of
   oocytes = 157, and number of mice = 6], 37.5 ± 4.08% [number of
   replicates = 3, number of oocytes = 128, and number of mice = 6],
   35.0 ± 4.08% [number of replicates = 3, number of oocytes = 116, and
   number of mice = 6] in 25 µM, 50 µM, 100 µM and 200 µM STE groups
   (Fig. [61]1A and B), respectively. We also observed a large proportion
   of oocytes that extruded first polar body (Pb1) that was larger than
   1/3 of the oocyte’s diameter in the 100 µM treated oocytes compared
   with other group oocytes at the MII stage (Fig. [62]1C). The large
   polar body and symmetric division of oocytes constituted the chief
   phenotype related to failure of spindle migration during meiosis I
   [[63]24]. These results suggested that STE caused the failure of oocyte
   maturation and induced the asymmetric division defects. For subsequent
   experiments, we chose 100 µM STE was chosen as the treatment
   concentration.

Fig. 1.

   [64]Fig. 1
   [65]Open in a new tab

   Effects of STE on maturational competence of mouse oocytes. (A)
   Representative images of oocytes from control and STE-treated groups.
   Red arrowheads denote oocytes with apparent symmetrical division and
   blue arrowheads indicate oocytes that fail to extrude polar bodies
   (scale bar, 80 μm). (B) The rate of polar body extrusion was
   significantly lower after STE exposure than in the control group. And
   the proportion of PB1 was (82.7 ± 3.68% [number of replicates = 3,
   number of oocytes = 145, and number of mice = 6], 53.0 ± 6.16% [number
   of replicates = 3, number of oocytes = 132, and number of mice = 6],
   44.2 ± 5.14% [number of replicates = 3, number of oocytes = 157, and
   number of mice = 6], 37.5 ± 4.08% [number of replicates = 3, number of
   oocytes = 128, and number of mice = 6], 35.0 ± 4.08% [number of
   replicates = 3, number of oocytes = 116, and number of mice = 6],
   p < 0.01) in control, 25 µM, 50 µM, 100 µM and 200 µM STE groups,
   respectively. (C) The percentage of oocytes with large polar bodies
   after STE-treated. And the proportion of large polar body was
   (9.6 ± 1.72% [number of replicates = 3, number of oocytes = 145, and
   number of mice = 6], 18.3 ± 2.35% [number of replicates = 3, number of
   oocytes = 132, and number of mice = 6], 20.9 ± 1.31% [number of
   replicates = 3, number of oocytes = 157, and number of mice = 6],
   25.6 ± 2.33% [number of replicates = 3, number of oocytes = 128, and
   number of mice = 6], 18.1 ± 1.32% [number of replicates = 3, number of
   oocytes = 116, and number of mice = 6], p < 0.01) in control, 25 µM, 50
   µM, 100 µM and 200 µM STE groups, respectively. The Figure depicts the
   mean percentage ± SD of the results obtained from three independent
   experiments. The bars labeled with completely different letters
   indicate significant difference at p < 0.01

RNA sequencing for the STE-exposed oocytes

   We attempted to explore the potential mechanism of STE toxicity on mice
   oocyte maturation through RNA sequencing. Considering that oocytes
   cultured for 8 h in vitro represents an intermediate stage between
   meiosis recovery (GVBD) and meiosis maturation (MII), the changes of
   oocytes at this stage will have an impact on the first polar body
   extrusion, so we chose to observe oocytes cultured for 8 h in vitro.
   The control and STE group oocytes were cultured in medium without or
   containing 100 µM STE, and oocytes were collected after 8 h of culture.
   Each group has 3 replicates, 40 oocytes (derived from 2 mice) per
   replicate. As the results shown in Fig. [66]2A and B, transcriptome
   analysis showed a total of 357 genes expression were changed due to 100
   µM STE treatment compared with the control group. Among these genes
   with altered expression, 229 (60.15%) genes were upregulated,
   nevertheless, 128 (35.85%) genes were downregulated. In addition, we
   conducted GO term analysis on these differentially expressed genes. And
   the results showed that STE exposure mainly affected the metabolic
   process, intracellular component, and RNA binding function
   (Fig. [67]2C). Moreover, as shown in Fig. [68]2D and Supplementary
   Material 1, the results for KEGG pathway analysis indicated that STE
   exposure significantly affected the fatty acid biosynthesis (Acsl4,
   Fasn), fatty acid metabolism (Acsl4, Scd3, Hacd4, Fasn), ferroptosis
   (Acsl4, Ncoa4), cell cycle (Ywhah, Ccnb1, Ccnh and Cdc25a), and
   regulation of actin cytoskeleton (Vav3, Mos, Raf1 and Pik3cd).
   Collectively, the results for RNA-seq provided the possible mechanisms
   for the STE toxicity on oocyte meiosis.

Fig. 2.

   [69]Fig. 2
   [70]Open in a new tab

   Transcriptome analysis for the effects of STE exposure on oocyte
   maturation. (A) The heat map for the altered expressed genes after STE
   treatment. (B) The numbers of altered expressed genes and volcano plot
   analysis after STE exposure. (C) The differential gene GO enrichment
   analysis for the differentially expressed genes after STE treatment.
   (D) The KEGG pathway enrichment analysis for the differentially
   expressed genes after STE exposure

STE causes mitochondrial distribution and membrane-potential defects in
oocytes

   We first assessed the effects of STE on mitochondria-related functions
   in oocytes. MitoTracker was deployed to evaluate the distribution of
   mitochondria in oocytes that at the 8 h of the IVM (in vitro
   maturation). Based on fluorescence staining, mitochondria accumulated
   at the spindle periphery in control oocytes, while the mitochondrial
   distribution was principally scattered or aggregated in clumps in the
   cytoplasm of the oocytes treated with STE (Fig. [71]3A). It was also
   apparent that the proportion of mitochondria surrounding the spindle in
   oocytes at the MI stage was lower than that in the control group
   (control group, 25.3 ± 2.87% [number of replicates = 3, number of
   oocytes = 51, and number of mice = 3]; STE group, 55.4 ± 9.09% [number
   of replicates = 3, number of oocytes = 65, and number of mice = 3];
   p < 0.01) (Fig. [72]3B). From the aforementioned results, we discerned
   that STE affected the mitochondrial distribution in MII oocytes. To
   determine whether STE also exerted toxic effects on the mitochondrial
   function of oocytes, we selected JC-1 as the membrane-potential stain
   to detect alterations in oocyte mitochondrial membrane potential. As
   shown in Fig. [73]3C, the fluorescence intensity of the red channel
   with mitochondria JC-1 in the STE group decreased relative to the
   control group, while the intensity of the green channel increased. We
   calculated the ratio of the red-to-green channel fluorescence intensity
   in the control and STE groups, and the data confirmed these findings
   (control group, 1.08 ± 0.10 [number of replicates = 3, number of
   oocytes = 43, and number of mice = 4]; STE group, 0.54 ± 0.14 [ number
   of replicates = 3, number of oocytes = 51, and number of mice = 4];
   p < 0.01) (Fig. [74]3D). These results revealed that STE was toxic to
   the mitochondrial functions of oocytes.

Fig. 3.

   [75]Fig. 3
   [76]Open in a new tab

   STE exposure disrupts mitochondrial redistribution and membrane
   potential in oocytes. Control and STE-exposed oocytes were labeled with
   MitoTracker Red to visualize mitochondrial localization and
   counterstained with Hoechst 33,342 to show meiotic stages. We evaluated
   mitochondrial distribution patterns using confocal microscopy. (A)
   Representative images of mitochondrial distribution patterns in MI
   oocytes: polarized distribution, homogeneous distribution, and
   clustered distribution. White arrowheads represent mitochondria
   distributed in clusters. (B) Quantification of control and STE-exposed
   oocytes at the MI stage with each mitochondrial distribution pattern
   (control group, 25.3 ± 2.87% [number of replicates = 3, number of
   oocytes = 51, and number of mice = 3]; STE group, 55.4 ± 9.09% [number
   of replicates = 3, number of oocytes = 65, and number of mice = 3];
   p < 0.01). (C) Mitochondrial membrane potential in control and
   STE-exposed oocytes as measured using JC-1 fluorescence; the green
   fluorescence shows inactive mitochondria, and the red fluorescence
   shows active mitochondria. (D) Histogram depicting the JC-1 red/green
   fluorescence ratio in the control and STE-exposed groups (control
   group, 1.08 ± 0.10 [number of replicates = 3, number of oocytes = 43,
   and number of mice = 4]; STE group, 0.54 ± 0.14 [number of
   replicates = 3, number of oocytes = 51, and number of mice = 4]). Error
   bars indicate mean ± SD (** p < 0.01; scale bar, 25 μm)

Exposure to STE toxin induces an increase in oxidative stress and DNA damage
in mouse oocytes

   Next, we investigated whether STE exposure influenced ROS levels in
   mouse oocytes. To address this line of inquiry, oocytes cultured for
   8 h in vitro (MI) were collected from the control and STE-treatment
   groups and stained with CM-H2DCFDA fluorescent dye for the assessment
   of ROS generation. Compared with the control group, STE treatment
   significantly augmented the levels of ROS in mouse oocytes
   (Fig. [77]4A) as determined by mean fluorescence intensity (control
   group, 19.5 ± 2.95 [number of replicates = 3, number of oocytes = 32,
   and number of mice = 3]; STE group, 58.3 ± 2.98 [number of
   replicates = 3, number of oocytes = 38 and number of mice = 3];
   p < 0.01) (Fig. [78]4B). These findings implied that STE exposure
   disrupted redox homeostasis during oocyte maturation.

Fig. 4.

   [79]Fig. 4
   [80]Open in a new tab

   STE exposure induces an increase in oxidative stress and DNA damage in
   mouse oocytes. (A) Representative images of CMH2DCFAD fluorescence in
   control and STE-exposed oocytes (scale bar, 80 μm). (B) Quantitative
   analysis of fluorescence intensity in control and STE-exposed oocytes
   (control group, 19.5 ± 2.95 [number of replicates = 3, number of
   oocytes = 32, and number of mice = 3]; STE group, 58.3 ± 2.98 [number
   of replicates = 3, number of oocytes = 38 and number of mice = 3];
   p < 0.01). (C) Representative images of DNA damage in control and
   STE-exposed oocytes (scale bar, 20 μm). Arrows represent aggregated
   γH2AX signals. (D) Fluorescence intensity of γH2A.X signals was
   quantified in control and STE-exposed oocytes (control group,
   10.1 ± 1.60 [number of replicates = 3, number of oocytes = 35, and
   number of mice = 3]; STE group, 29.1 ± 1.93 [number of replicates = 3,
   number of oocytes = 41, and number of mice = 3]). Data are expressed as
   the mean ± SD from three independent experiments (**significantly
   different at p < 0.01)

   We also assessed the level of DNA damage by γ-H2AX staining of
   STE-exposed GV oocytes since excessive ROS may attack macromolecules in
   cells (including DNA). Additionally, our fluorescence imaging and
   intensity measurement data revealed that STE exposure generated a
   greater occurrence of DNA damage in oocytes (arrows) (control group,
   10.1 ± 1.60 [number of replicates = 3, number of oocytes = 35, and
   number of mice = 3]; STE group, 29.1 ± 1.93 [number of replicates = 3,
   number of oocytes = 41, and number of mice = 3]; p < 0.01) (Fig. [81]4C
   and D). Thus, these results implied that STE exposure induced the
   accumulation of excessive ROS to generate DNA damage, consequently
   producing poorer quality oocytes.

STE exposure disrupts assembly of the spindle/chromosome in mice oocytes

   To explore the reason for oocyte maturation defects in the STE-exposed
   oocytes, we examined spindle morphology in vitro culture for 8 h after
   STE treatment. Immunofluorescence data revealed that a large number of
   control oocytes have a regular spindle with a conventional shape and
   well-aligned chromosomes at the MI stage (Fig. [82]5A). In
   contradistinction, a higher incidence of spindle assembly abnormalities
   (arrows) and unconventional chromosomal alignments was observed in the
   STE-exposed oocytes (Fig. [83]5A), and we noted that the proportion of
   STE-treated oocytes with spindle/chromosome defects was remarkably
   higher than in control oocytes (control group, 10.5 ± 2.20% [number of
   replicates = 3, number of oocytes = 53, and number of mice = 3]; STE
   group, 36.3 ± 7.43% [number of replicates = 3, number of oocytes = 57,
   and number of mice = 3]; p < 0.01) (Fig. [84]5B), indicating that STE
   exposure induces a defective spindle/chromosome structure in mouse
   oocytes. We finally examined whether STE exposure affects α-tubulin
   acetylation levels in mouse oocytes, we found that STE treatment
   increases oocyte ac-tubulin levels (Fig. [85]5C and D).

Fig. 5.

   [86]Fig. 5
   [87]Open in a new tab

   STE exposure disrupts assembly of the spindle/chromosome apparatus in
   mouse oocytes. (A) Oocytes were immunostained with anti-α-tubulin-FITC
   antibody to visualize spindle morphology (green) and counterstained
   with propidium iodide (PI) to show chromosomal alignment (red). (a)
   Control oocytes exhibit the characteristic barrel-shaped spindle and
   well-aligned chromosomes. (b-d) Three examples illustrate the
   disorganized spindles (arrows) and misaligned chromosomes (arrowheads),
   which were frequently observed in STE-exposed oocytes (scale bar,
   20 μm). (B) The proportion of aberrant spindles and chromosome
   alignment were quantified in control and STE-exposed oocytes (control
   group, 10.5 ± 2.20% [number of replicates = 3, number of oocytes = 53,
   and number of mice = 3; STE group, 36.3 ± 7.43% [number of
   replicates = 3, number of oocytes = 57, and number of mice = 3]). (C)
   Expression of acetylated tubulin levels in control and STE-exposed
   oocytes. Proteins from 200 oocytes were loaded for each sample. (D)
   Relative-intensity results for AC-Tubulin protein expression in the
   control and STE-treated oocytes. Data are presented as mean percentage
   (mean ± SD) from at least three independent experiments
   (**significantly different at p < 0.01)

Exposure to STE toxin affects meiotic progression and perturbs the actin caps
during mouse oocyte maturation

   Next, we used phalloidin-TRITC to stain oocytes that cultured in vitro
   for 8 h (MI) after STE treatment and determined whether the
   STE-exposure affected actin polymerization. According to the
   Fig. [88]6A and B, actin caps were visible in approximately 81% and 40%
   of control and STE-treatment oocytes, respectively. (control group,
   81.0 ± 1.96% [number of replicates = 3, number of oocytes = 32, and
   number of mice = 3]; STE group, 40.8 ± 2.16% [number of replicates = 3,
   number of oocytes = 37, and number of mice = 3]; p < 0.01). In
   addition, nuclear staining and quantitative analysis revealed that the
   proportion of oocytes arrest in GVBD stage increased significantly
   after 8 h of STE treatment. (control group, 18.9 ± 1.90% [number of
   replicates = 3, number of oocytes = 32, and number of mice = 3]; STE
   group, 40.1 ± 1.62% [number of replicates = 3, number of oocytes = 37,
   and number of mice = 3]; p < 0.01) (Fig. [89]6C). In summary, we
   demonstrated that STE treatment affects the development process of
   oocytes and perturbs the formation of actin cap.

Fig. 6.

   [90]Fig. 6
   [91]Open in a new tab

   Exposure to STE toxin perturbs the actin cytoskeleton during mouse
   oocyte meiosis. (A) Representative images of actin filaments on the
   plasma membrane in control and STE-exposed oocytes (scale bar, 20 μm).
   (B) Quantification of control and STE-exposure oocytes with formation a
   normal actin cap (control group, 81.0 ± 1.96% [n = 32]; STE group,
   40.8 ± 2.16% [n = 37]; p < 0.01). (C) Percentage of GVBD-arrested
   oocytes after STE exposure (control group, 18.9 ± 1.90% [number of
   replicates = 3, number of oocytes = 32, and number of mice = 3]; STE
   group, 40.1 ± 1.62 [number of replicates = 3, number of oocytes = 37,
   and number of mice = 3]; p < 0.01). Data are presented as mean
   percentage (mean ± SD) from at least three independent experiments
   (**significantly different at p < 0.01)

STE exposure induces ferroptosis and reduces pERK1/2 expression in mouse
oocytes

   Our RNA-seq data revealed that STE exposure induces oocyte ferroptosis
   process. And we observed that Fe^2+ level was increased in oocytes
   cultured in vitro for 8 h (MI) after STE treatment (control group,
   32.3 ± 3.86 [number of replicates = 3, number of oocytes = 33, and
   number of mice = 3]; STE group, 19.0 ± 1.63 [number of replicates = 3,
   number of oocytes = 36, and number of mice = 3]; p < 0.01)
   (Fig. [92]7A, B). While ferroptosis-related protein expression levels
   of GPX4 and ACSL4 were significantly upregulated (Fig. [93]7C-F). Next,
   we examined the effects of STE supplementation on GSH levels in
   oocytes. Figure [94]7G shows weak GSH fluorescence signals were
   generated in STE exposure oocytes (control group, 31.7 ± 2.8 [number of
   replicates = 3, number of oocytes = 35, and number of mice = 3]; STE
   group, 16.0 ± 2.94 [number of replicates = 3, number of oocytes = 33,
   and number of mice = 3]; p < 0.01). Thus, these data suggested that STE
   exposure induced ferroptosis, consequently resulting in poorer quality
   oocytes.

Fig. 7.

   [95]Fig. 7
   [96]Open in a new tab

   STE exposure induces ferroptosis and reduces pERK1/2 expression in
   mouse oocytes. (A) Representative images of Fe^2+ fluorescence in
   control and STE-exposed oocytes (scale bar, 20 μm). (B) Quantitative
   analysis of Fe^2+ fluorescence intensity in control and STE-exposed
   oocytes (control group, control group, 32.3 ± 3.86 [number of
   replicates = 3, number of oocytes = 33, and number of mice = 3]; STE
   group, 19.0 ± 1.63 [number of replicates = 3, number of oocytes = 36,
   and number of mice = 3]). (C) Expression of GPX4 levels in control and
   STE-exposed oocytes. Proteins from 200 oocytes were loaded for each
   sample. (D) Relative-intensity results for GPX4 protein expression in
   the control and STE-treated oocytes. (E) Expression of ACSL4 levels in
   control and STE-exposed oocytes. Proteins from 200 oocytes were loaded
   for each sample. (F) Relative-intensity results for ACSL4 protein
   expression in the control and STE-exposed oocytes. (G) Representative
   images of GSH fluorescence in control and STE-exposed oocytes (scale
   bar, 80 μm). (H) Quantitative analysis of GSH fluorescence intensity in
   control and STE-exposed oocytes (control group, 31.7 ± 2.87 [number of
   replicates = 3, number of oocytes = 35, and number of mice = 3]; STE
   group, 16.0 ± 2.94 [number of replicates = 3, number of oocytes = 33,
   and number of mice = 3]). Experiments were repeated three times, and
   the results are presented as mean ± SD (*significantly different at
   p < 0.05 and **significantly different at p < 0.01)

STE exposure inhibits RAF1 activity and induced oocyte asymmetric division
defects

   As a serine/threonine protein kinase, RAF1 activity is required for
   progesterone-induced oocyte maturation [[97]25]. According to the
   results from RNA-sequencing, we found that STE exposure significantly
   reduced Raf1 mRNA levels (Supplementary Material 1). When RAF1 protein
   expression was then assessed by western blotting in oocytes exposed to
   STE for 8 h in vitro, we found it to be significantly reduced
   (Fig. [98]8A and B). In addition, we noted that STE treatment
   significantly reduced the phosphorylation of ERK1/2 (Fig. [99]8C-E).
   Considering the important role of RAF1 during oocyte development, mouse
   oocytes were treated with a RAF1 specific inhibitor (10 µM), WB results
   revealed that RAF1 protein expression significantly reduced after RAF1
   inhibitor treatment for 8 h in vitro (Fig. [100]8F and G). After 14 h
   in vitro culture, we observed the first polar body extrusion rate of
   oocytes in the control and RAF1-inhibited groups, and we noted that
   when the activity of RAF1 was inhibited, the asymmetric defects (large
   polar body) of oocytes significantly increased (Fig. [101]8H and I).
   Collectively, our results demonstrated that the inhibition of RAF1
   activity by STE is one of the factors responsible for the defect in
   oocyte asymmetric division.

Fig. 8.

   [102]Fig. 8
   [103]Open in a new tab

   STE exposure inhibits RAF1 activity and induced oocyte asymmetric
   division defects. (A) Expression of RAF1 levels in control and
   STE-exposed oocytes. Proteins from 100 oocytes were loaded for each
   sample. (B) Relative-intensity results for RAF1 protein expression in
   the control and STE-treated oocytes. (C) Expression of ERK1/2 levels in
   control and STE-exposed oocytes. Proteins from 100 oocytes were loaded
   for each sample. (D) Expression of pERK1/2 levels in control and
   STE-exposed oocytes. Proteins from 100 oocytes were loaded for each
   sample. (E) Relative-intensity results for pERK1/2 protein expression
   in the control and STE-exposed oocytes. (F) Expression of RAF1 levels
   in control and RAF1-Inhibitor oocytes. Proteins from 100 oocytes were
   loaded for each sample. (G) Relative-intensity results for RAF1 protein
   expression in the control and RAF1-Inhibitor oocytes. (H)
   Representative images of oocytes from control and RAF1-Inhibitor
   groups. Red arrowheads denote oocytes with apparent symmetrical
   division and blue arrowheads indicate oocytes that fail to extrude
   polar bodies (scale bar, 80 μm). (I) The percentage of oocytes with
   large polar bodies after STE-treated. And the proportion of large polar
   body was (10.3 ± 1.86% [number of replicates = 3, number of
   oocytes = 105, and number of mice = 6], 19.0 ± 2.67% [number of
   replicates = 3, number of oocytes = 121, and number of mice = 6],
   p < 0.01) in control and RAF1 Inhibition groups. (J) Proposed model for
   the role of STE in oocyte maturation. STE exposure caused abnormal
   expression of GPX4 and ACSL4, which induced ferroptosis in oocytes. In
   addition, STE exposure inhibited RAF1 activity, which is one of the
   reasons for the defects in STE-induced asymmetric division in mice
   oocytes. Experiments were repeated three times, and the results are
   presented as mean ± SD (*significantly different at p < 0.05 and
   **significantly different at p < 0.01)

Discussion

   Toxic chemicals can disrupt oocyte growth and embryonic development
   [[104]26]. As a mycotoxin, STE is widely present in human environments,
   including foods, feeds, and indoor dust [[105]27]. The STE is generally
   recognized as a possible carcinogen, mutagen, and teratogen [[106]28].
   Extensive evidence demonstrated that exposure to STE induced adverse
   effects on organisms and in particular its reproductive toxicity has
   been attracted public concern [[107]29]. In the present study, we
   focused on the effects of STE exposure on mouse oocyte maturation and
   our findings revealed that STE exposure reduced the emission of the
   first polar body as well as disrupted the asymmetric division in mouse
   oocytes.

   Our results showed that STE exposure significantly decreased the
   developmental capacity of mouse oocytes, showing by a reduction in the
   rate of the first polar body extrusion and asymmetric division defects
   in a dose dependent manner. A key indicator of oocyte maturation is the
   extrusion of the first polar body (Pb1) [[108]30]. This may also
   suggest that STE exposure could cause the decline of oocyte quality.
   Oocyte maturation is a complex process. Mouse oocytes reach maturity
   within 14 h during in vitro maturation, but it takes much longer in
   vivo [[109]31]. In our study, we examined the impact of exposure dose
   and duration on the maturation of oocytes, finding that a higher
   concentration of STE (100 µM, 32.42 ng/µL) inhibited the maturation of
   most oocytes while enabling a minority to reach maturity. Furthermore,
   intrabdominal injection of STE has been shown to reduce the growth rate
   of chicks [[110]32]. The levels of STE were detected in human breast
   milk (1.2 ng/L) [[111]33] and plasma (1.0–30.0 ng/µL) [[112]34]. Our
   results confirmed that exposure to 100 µM (32.42 ng/µL) caused meiosis
   failure in mouse oocytes, this concentration is slightly higher than
   that found in human plasma. In addition, our results also demonstrated
   that STE could induce oocyte meiosis abnormalities even at 25 µM (8.1
   ng/µL). Although the concentration of STE in mammalian and human
   follicular fluid has not been reported, our results show that STE can
   interfere with the meiosis process of mouse oocytes even at low levels
   of exposure.

   Studies have demonstrated that STE could induce oxidative stress,
   mitochondrial dysfunction, apoptosis, and cell cycle arrest [[113]12].
   The status of mitochondria in the experimental mouse oocytes became our
   first object of assessment. The distribution pattern of mitochondria
   has been correlated with the quality and developmental potential of
   mammalian oocytes and embryos in scientific literature [[114]35,
   [115]36]. In this study, we focused on the mitochondrial distribution
   patterns of oocytes in the control and STE treated oocytes cultured in
   vitro for 8 h. The mitochondrial distribution pattern of most oocytes
   in the control group showed polarized distribution, however, oocytes in
   the STE-treated group showed a high proportion of clustered
   distribution. In addition, we noted that STE-exposed oocytes exhibited
   a markedly reduced mitochondrial membrane potential in mouse oocytes,
   diminished mitochondrial membrane potential indicates mitochondrial
   dysfunction [[116]37]. It was previously noted that STE administration
   generates free radicals, induces lipid peroxidation and changes in
   cellular antioxidant status [[117]38]. We observed that ROS levels were
   dramatically augmented with STE exposure, suggesting mitochondrial
   dysfunction. Increased production of reactive oxygen species (ROS) by
   mitochondria has deleterious effects on various cellular components
   such as DNA, RNA, proteins, and lipids, leading to disruptions in key
   biological processes including cellular metabolism, apoptosis, and
   senescence [[118]39]. The MAPK signaling pathway, which includes p38
   MAPK, extracellular signal kinase (ERKs), and c-Jun-terminal kinases
   (c-JNKs) [[119]40]. The ERK signaling pathway is fundamentally known as
   a regulator of cell proliferation, and is downregulated by oxidative
   stress [[120]41]. ERK1/2-depleted oocytes had poorly-assembled
   metaphase II (MII) spindles [[121]42]. Our results also showed that STE
   treatment reduced phosphorylation of ERK1/2 in oocytes. Inhibition of
   the ERK activation increased mitochondrial ROS levels and the cell
   death rate of T-ALL cells [[122]43], our RNA-seq results showed that
   STE exposure decreased Dusp1 gene expression in mouse oocytes
   (Supplementary material 1). It has been reported that inhibition of
   DUSP1 can induce apoptosis in human MGC803 and MKN45 gastric cancer
   (GC) cell lines by inactivating MAPK [[123]44]. However,
   Dusp1-Knockdown could activate the ERKs in HEK293T cell [[124]45],
   which may be related to the differences in the use of cell lines.
   Inhibition of DUSP1 in cumulus cells caused abnormal cell cycle
   progression [[125]46] and DUSP1 might be a potential biomarker to
   diagnose oocyte quality [[126]47] .

   Moreover, ROS constituted an important factor causing DNA lesions in
   cells [[127]48]. The accumulation of γH2AX signal in STE-exposed
   oocytes indicates the STE can induce oocyte DNA damages. When DNA
   damage occurs in cells, it triggers cell cycle arrest and DNA repair
   responses [[128]49], which to some extent explains why STE causes
   oocyte development arrest. Mitochondrial function and redox state are
   important for the proper assembly of meiotic structures during oocyte
   meiosis, and increased oxidative stress is tied to spindle
   disorganization and chromosomal misalignment in oocytes [[129]50].
   Female reproductive disorders have been linked to ferroptosis
   [[130]51]. During ferroptosis, granulosa cells, oocytes and embryos can
   be damaged [[131]52]. Through its ability to scavenge lipid peroxides,
   GPX4 is one of the inhibitors of ferroptosis [[132]53], and Gpx4
   knockout causes cell death [[133]54]. In our research, RNA sequencing
   results showed that Acsl4 mRNA expression was downregulated in
   STE-treated oocytes, however, ACSL4 protein was upregulated. Previous
   research has indicated that while protein is more closely linked to the
   target of natural selection, gene expression is commonly quantified
   through mRNA levels. However, the assumption that mRNA levels
   accurately reflect protein levels has been challenged by various
   studies, suggesting that mRNA levels do not consistently predict
   protein levels [[134]55]. In this study, we cultured oocytes in vitro
   for 8 h after STE treatment for RNA-Seq and Western blot. When oocytes
   were treated with STE, the intracellular ferroptosis process was
   activated, leading to an increase in ACSL4 protein expression. Since
   cells need a stable environment, they become stressed when protein
   levels rise. To maintain homeostasis, cells will inevitably reduce gene
   transcription [[135]56].

   In addition to its role in polyunsaturated fatty acid (PUFA)
   biosynthesis, ACSL4 promotes the accumulation of lipid peroxidation
   products [[136]57]. Investigators have shown that STE caused lipid
   peroxidation [[137]58] and that STE treatment attenuated cell
   proliferation and augmented apoptosis in SH-SY5Y cells [[138]59]. It is
   believed that microtubules (MTs) play a significant role in organelle
   transport and positioning, regulation cell shape, and motility
   [[139]60]. Acetylation of α-tubulin on lysine 40 is one of the major
   microtubular posttranslational modifications [[140]61], and α-tubulin
   acetylation serves as a marker for the presence of stable MTs
   [[141]62]. The function(s) of acetylated MTs during meiosis remain
   unknown, although acetylated MTs have been detected in mouse oocytes.
   Reduced tubulin acetylation has been found to decrease microtubule
   stability, but high tubulin acetylation did not maintain microtubule
   stability [[142]63]. During mouse oocyte meiosis, altered tubulin
   acetylation causes spindle assembly defects and chromosome misalignment
   [[143]64]. The elevated acetylation of tubulin by HDAC3 damages the
   stability of microtubules during oocyte maturation [[144]65],
   nevertheless, reduced tubulin acetylation by RASSF1A impairs the
   stability of microtubules as well [[145]66]. There is still much to
   learn about how tubulin acetylation affects microtubule stability.

   RAF1 is a serine/threonine protein kinase that plays an essential role
   in cell proliferation [[146]67]. The Raf1^−/− mice die by embryonic day
   e12.5 [[147]68]. The interaction between RAF1 and cytoskeletal actin is
   critical for its redistribution and proliferation in the airways
   [[148]67]. In our research, we noted that RAF1 inhibition affected RAF1
   expression, this may be because the inhibition of RAF1 interferes with
   meiotic process of oocytes, it means that the oocytes obtained at 8 h
   of RAF1 inhibitor treatment were at different developmental stages. In
   mouse granulosa cells, RAF1 stimulates ERK phosphorylation to synthesis
   and secrete estradiol [[149]69]. RAF1 kinase activity is required for
   Xenopus oocyte maturation [[150]25]. Our data revealed that STE
   exposure inhibits the kinase activity of RAF1, and oocytes with
   inhibitors of RAF1 kinase can lead to asymmetric division defects in
   oocytes. This can at least partially explain the development
   abnormalities of oocytes caused by STE exposure.

   Collectively, our results provided evidence that STE exposure induced
   mitochondrial dysfunction, DNA damage, and ferroptosis in mouse
   oocytes. In addition, STE exposure inhibited RAF1 activity, which is
   one of the reasons asymmetric division defects (Fig. [151]8J).

Materials and methods

   All chemicals and culture media were purchased from Sigma (St. Louis,
   MO, USA) unless stated otherwise.

Mice

   The mice used in this study were purchased from Beijing Sibefu
   Biotechnology Co., Ltd, and 6–8-week-old ICR female mice were used in
   all experiments. All experimental animal protocols were performed
   following relevant ethical guidelines and regulations and approved by
   the First Affiliated Hospital of Anhui Medical University.

Antibodies and chemicals

   Mouse monoclonal anti-α-tubulin-FITC antibody (Cat#: F2168), propidium
   iodide (Cat#: P4170), phalloidin-TRITC (Cat#: P1951), and RAF1 specific
   inhibitor (Cat#: 220904-83-6) were purchased from Sigma (St. Louis, MO,
   USA); mouse polyclonal anti-GPX4 (Cat#: 67763-1-Ig); rabbit polyclonal
   anti-ACSL4 (Cat#: 22401-1-AP), mouse monoclonal anti-RAF1 (Cat#:
   66592-1-Ig), rabbit polyclonal anti-pERK1/2 (CAT#:28733-1-AP), and
   mouse monoclonal anti-GAPDH antibody (Cat#: 60004–1 g) were bought from
   Proteintech (Wuhan, China); mouse monoclonal anti-acetyl-tubulin
   (Lys-40) (Cat#: sc-23950) antibody were obtained from Santa Cruz
   Biotechnology, rabbit polyclonal anti-ERK1/2 (Cat#: abs130092) was
   obtained from Absin (Shanghai, China). We obtained HRP-conjugated
   rabbit (Cat#: 7074 S) and mouse (Cat#: 7076 S) secondary antibodies
   from Cell Signaling Technology (Danvers, MA, USA).

Oocyte collection and culture

   To obtain fully grown GV oocytes, 6–8-week-old ICR mice were
   superovulated with 10 IU of pregnant mare serum gonadotropin (PMSG;
   SANSHENG BIOLOGICAL TECHNOLOGY, Ningbo, China) by intraperitoneal
   injection; 48 h later, GV oocytes were harvested from the mouse ovaries
   and placed in M2 (Sigma, St. Louis, MO, USA; Cat#: M7167) medium.
   Oocytes were subsequently cultured in mini-drops of M16 (Sigma, St.
   Louis, MO, USA; Cat#: MR-016) medium covered with mineral oil (Sigma,
   St. Louis, MO, USA; Cat#: M8410) at 37 °C in an atmosphere of 5% CO[2]
   in compressed air and high humidity.

STE treatment

   STE (CAT#: S102406, Aladdin, Shanghai, China) was dissolved in dimethyl
   sulfoxide (DMSO) after centrifugation and was diluted in M16 medium to
   produce final concentrations of 25, 50, 100 and 200 µM. The control
   oocytes were grown in M16 medium with the same concentration of DMSO
   (0.05%). The final DMSO concentration was 0.05% in each experiment
   group. After culture 3 h and 14 h, the germinal vesicle breakdown
   (GVBD) and metaphase II (MII) oocytes were used for the subsequent
   experiments.

RNA – sequencing (RNA-seq) and analysis

   In both control and STE exposure groups, total RNA was extracted
   oocytes after 8 h of culture. We collected 40 oocytes for each group
   for disposal by the Beijing Allawe Gene Technology Co. Ltd. The total
   RNA was first extracted using the TRIzol method (Invirtrogen, CA, USA)
   and treated with RNase-free DNase I (Takara, Kusatsu, Japan). RNA
   degradation and contamination were monitored on 1% agarose gels. RNA
   was quantified using Agilent 2100 Bioanalyzer (Agilent Technologies,
   CA, USA), the quality and integrity were assessed by NanoDrop
   spectrophometer (Thermo Scientific, DE, USA). RNA samples with
   concentrations higher than 200 ng/µL were considered eligible. A total
   of 1.5 µg of RNA per sample was utilized for RNA sample preparations.
   Sequencing libraries were constructed using the NEBNext^® Ultra™ RNA
   Library Prep Kit for Illumina^® (NEB, USA) in accordance with the
   manufacturer’s instructions. To isolate cDNA fragments ranging from 200
   to 250 bp in length, the library fragments were purified using the
   AMPure XP system (Beckman Coulter, Beverly, USA). The library
   preparations were sequenced on an Illumina Novaseq 6000 platform by the
   Beijing Allwegene Technology Company Limited (Beijing, China),
   resulting in the generation of paired-end 150 bp reads.

   Gene expression levels were quantified using fragments per kilobase of
   transcript per million fragments mapped (FPKM). Differential expression
   analysis between two groups was conducted utilizing the DESeq R package
   (version 1.10.1). To account for multiple testing, the resulting P
   values were adjusted using the Benjamini and Hochberg method to control
   the false discovery rate. Genes with an adjusted P-value < 0.05 as
   determined by DESeq were classified as differentially expressed. The
   GOseq R packages were utilized to conduct Gene Ontology (GO) enrichment
   analysis on the differentially expressed genes (DEGs), employing the
   Wallenius non-central hyper-geometric distribution to account for gene
   length bias.

   KEGG serves as a database resource for comprehending the overarching
   functions and utilities of biological systems, ranging from cellular to
   ecosystem levels, by leveraging molecular-level data, particularly from
   large-scale molecular datasets produced through genome sequencing and
   other high-throughput experimental methodologies
   ([152]http://www.genome.jp/kegg/). We used KOBAS software to test the
   statistical enrichment of differential expression genes in KEGG
   pathways.

Immunofluorescence microscopy

   Oocytes were first fixed in 4% paraformaldehyde for 30 min at 4℃ and
   then permeabilized with 0.5% Triton X-100 for 20 min. Oocyte protein
   binding sites were then blocked in 1% BSA-supplemented
   phosphate-buffered saline (PBS) for 1 h and stained with different
   first antibodies (1:250 for γH2AX) at 4 °C overnight. After extensive
   washing, the samples were then stained with rabbit secondary antibodies
   (CST, Cat#: 7074 S, 1:300) at 37 °C for 1 h. For spindle staining,
   oocytes were immunolabeled with anti- α- tubulin- FITC antibody (Sigma,
   Cat#: F2168, 1:300) to visualize the spindle. Oocytes were stained with
   PI (10 mg/mL in PBS) for 10 min at room temperature and ultimately
   examined with a laser-scanning confocal microscope (Zeiss LSM 800 META;
   Jena, Germany). To assess mitochondrial staining in oocytes, the cells
   were incubated in M2 medium supplemented with 200 nM MitoTraker Red
   (Invitrogen, Cat#: M7152) for 30 min at 37℃. Subsequently, the oocytes
   underwent three washes with M2 medium before being examined using a
   confocal laser scanning microscope. Evaluation of mitochondrial
   membrane potential (MMP) was conducted by culturing oocytes in M2
   medium containing 2 µM JC-1 for 30 min at 37℃. The mitochondrial
   membrane potential was determined by quantifying the ratio of red
   fluorescence, indicative of activated mitochondria (J-aggregates), to
   green fluorescence, indicative of less activated mitochondria
   (J-monomers). For F-actin staining, the samples were fixed with 4%
   paraformaldehyde for 5 min and then blocked with 1% BSA for 1 h.
   Following a 1 h incubation period with FITC-conjugated phalloidin
   (5 µg/mL in PBS), the DNA was subsequently counterstained with PI for
   10 min. Subsequently, the oocytes were washed three times and
   co-stained with propidium iodide (10 mg/mL in PBS) for 10 min and then
   mounted on glass slides.

Assays of intracellular ROS and GSH levels

   Denuded oocytes were added to medium that contained 5 uM
   2’,7’-dichlorofluorescin diacetate (2’,7’-DCFHDA) or 10 µM Cell Tracker
   Blue to measure ROS or GSH at 37 °C for 30 min. After three washes with
   M2, the oocytes were placed on glass slides for further detection under
   a laser scanning confocal microscope (LSM 800, Zeiss, Germany). The
   fluorescence of each oocyte was analyzed by Zeiss software (ZEISS,
   Germany).

Ferrous ion staining

   Consistent with the oocyte developmental stage of the previous RNA
   sequencing, intracellular Fe^2+ levels were examined at the 8 h of the
   IVM period. The oocytes in each group were thoroughly washed in
   prewarmed M2 medium and assessed using the 1 µM fluorescent probe Ferro
   Orange (Dojindo, Cat#: F374) for 30 min. After three washes, the
   oocytes were placed on confocal dishes for further detection under a
   laser scanning confocal microscope (LSM 800, Zeiss, Germany).

Analysis of fluorescence intensity

   Oocytes in the control and experimental groups were mounted on the same
   glass slide, and the same previous scan settings were used for sample
   scanning with a Zeiss LSM 800 META confocal system. The same
   immunostaining procedure was performed on control and STE-exposed
   oocytes to measure fluorescence intensity and the same parameters were
   used as with confocal microscopes. Image J software was utilized to
   delineate a region of interest (ROI) and calculate the average
   fluorescence intensity per unit area within the ROI. The resulting
   average values were then compared between the control and STE groups to
   assess differences in fluorescence intensity.

Western blot analysis

   A total of 200 mouse oocytes in each group were collected and lysed in
   Laemmli sample buffer (95% Laemmli sample buffer and 5%
   β-mercaptoethanoll). The samples were first boiled at 100 °C for
   10 min, and then subjected to SDS-polyacrylamide gel electrophoresis
   (PAGE) using 10% gels in Tris/Glycine/SDS Running Buffer (Epizyme,
   China, Cat: # PS105S). After electrophoretic separation, proteins were
   transferred onto polyvinylidene fluoride membranes (Millipore,
   Billerica, MA, USA); membranes were blocked with Tris-buffered saline
   (TBS) that contained 0.1% (w/w) Tween 20 (TBST) and 5% nonfat dry milk
   at 37 °C for at least 1 h. After a brief wash in TBST, the membranes
   were incubated with a mouse monoclonal anti-GPX4 antibody (1:1000),
   rabbit monoclonal anti-ACSL4 antibody (1:1000), mouse monoclonal
   anti-RAF1 antibody (1:2000), mouse monoclonal anti-acetyl-tubulin
   (Lys-40) (1:200), rabbit polyclonal anti-pERK1/2 (1:2000), rabbit
   polyclonal anti-ERK1/2 (1:2000) and a mouse monoclonal anti-GAPDH
   antibody (1:5000) at 4 °C overnight. After washing three times in TBST
   (10 min each), membranes were incubated for 1 h with secondary
   anti-rabbit or anti-mouse HRP-conjugated antibodies (1:5000) in 5%
   nonfat dry milk in TBST. The membranes were then exposed to an enhanced
   chemiluminescence reagent (Epizyme, Shanghai, China). For quantitation
   of Western blot band intensity, densitometric analysis was performed
   using Image J software and normalized to either the levels of GAPDH.

Statistical analysis

   We expressed all percentages or values from at least three independent
   replicates as means ± SD. The normal distribution of the data was
   performed using Kolmogorov-Smirnov in SPSS 20.0 software. Unpaired,
   two-tailed Student’s t tests were applied to assess differences between
   two groups, and comparisons among more than two groups were analyzed
   with a one-way ANOVA and Tukey’s multiple-comparison test using Prism
   GraphPad 8. A p value of < 0.05 was considered to be statistically
   significant.

Electronic supplementary material

   Below is the link to the electronic supplementary material.
   [153]Supplementary Material 1^ (63KB, xls)
   [154]Supplementary Material 2^ (522.3KB, docx)

Acknowledgements