Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2022 Oct 5.
Published in final edited form as: Chemosphere. 2022 Mar 14;298:134296. doi: 10.1016/j.chemosphere.2022.134296

High-fat diet aggravates prenatal low-dose DEHP exposure induced spermatogenesis disorder: Characterization of testicular metabolic patterns in mouse offspring

Yun Fan a,b,c,1, Qiaoqiao Xu a,b,1, Hong Qian a,b,1, Chengzhe Tao a,b, Tingya Wan a,b, Zhi Li a,b, Wenkai Yan a,b, Rui Niu a,b, Yuna Huang a,b, Minjian Chen a,b, Qiujin Xu a,d, Elizabeth M Martin e, Xinru Wang a,b, Yufeng Qin b,c,**, Chuncheng Lu a,b,*
PMCID: PMC9533191  NIHMSID: NIHMS1819055  PMID: 35301995

Abstract

Di-(2-ethylhexyl) phthalate (DEHP) is a widely used plasticizer and has been identified as a male prenatal reproductive toxicant. A high fat diet (HFD) has also been suggested as another potential disruptor of male reproductive function. Despite this potential synergism between DEHP exposure and HFD, little is known about the concomitant effects of prenatal DEHP and a subsequent HFD exposure on male offspring reproductive injury. Here we established a mouse model of prenatal exposure to DEHP (0.2 mg/kg/day) to assess the testicular development and spermatogenesis in offspring subjected to obesogenic diet during the pubertal period. Gross phenotype, hormone profiles and the testicular metabolome were analyzed to determine the underlying mechanism. We found that prenatal exposure to low-dose DEHP resulted in decreased sperm density, decreased testosterone (T) levels, increased luteinizing hormone (LH) levels and testicular germ cell apoptosis. Furthermore, these injury phenotypes were aggravated by pubertal HFD treatment. Testicular riboflavin and biotin metabolites were enriched implying their roles in contributing HFD to exacerbate offspring spermatogenesis disorders due to prenatal low-dose DEHP exposure. Our findings suggest that pubertal HFD exacerbates reproductive dysfunction associated with prenatal exposure to low-dose DEHP in male adult offspring.

Keywords: DEHP, HFD, WCNA, Male adult offspring, Reproductive dysfunction

Graphical Abstract

graphic file with name nihms-1819055-f0008.jpg

1. Introduction

Phthalates (PAEs) are commonly used in the manufacture and processing of plastic products (Gao and Wen, 2016). Almost 300 million pounds of phthalates are produced on average per year in the United States (Hannon et al., 2016). Di-(2-ethylhexyl) phthalate (DEHP), one of the most extensively used phthalates worldwide, is found in many common items including flooring, carpets, cable protectors, cosmetics, and food packaging materials (Heudorf et al., 2007; Wu et al., 2017). Humans are readily exposed to DEHP through leaching into the environment during production, storage, transportation, use and processing (Fay et al., 1999; Fromme et al., 2013). In 2014, the U.S. Environmental Protection Agency (EPA) had included DEHP in its priority pollutant list. The level of daily human exposure to DEHP has been estimated between 3 and 30 μg/kg/day (Doull et al., 1999). Complicating the study of DEHP is its ubiquity in human samples and the environment (Zolfaghari et al., 2014).

DEHP is classified as endocrine disrupting chemicals in both male and female reproductive systems (Zhang et al., 2013; Richardson et al., 2018). Epidemiological data showed that maternal DEHP exposure was associated with impaired gonadal development and fertility in human males (Hauser et al., 2007; Pant et al., 2008). Furthermore, DEHP and its metabolites have been identified in amniotic fluid, breast milk, blood, umbilical cord blood, ovarian follicular fluid and reproductive tissues (Silva et al., 2004; Hogberg et al., 2008), suggesting that DEHP exposure occurs during the prenatal period (Adibi et al., 2009). In animal studies, DEHP was recognized as the most potent phthalates that affected the male reproductive tract development (Gray et al., 2000). Specifically, DEHP has been shown to pass through the placental barrier and reduce sperm number as well as adversely affect semen quality (Abd-Ellah et al., 2016). Additionally, exposure to high-dose of DEHP has been reported to negatively affect spermatogenesis (Andrade et al., 2006; Vo et al., 2009); however, the reproductive effects of prenatal DEHP exposure at an environmentally relevant dose are unclear. It is reported that exposure to environmental chemicals could cause metabolic disturbances on testicular functions (Yeung et al., 2011). Some EDCs can alter the function of the reproductive axis by disrupting the pathways involved in hormone synthesis and metabolism (Rato and Sousa, 2021). EDCs were considered to be metabolic modulators of Sertoli cells in testis (Rato et al., 2012). As mentioned in other studies, DEHP could interfere with the intricate signaling and metabolizing networks in testis (Walker et al., 2021). Therefore, it is necessary to assess the effects of prenatal DEHP exposure at a low dose on male reproductive health.

As a plasticizer for synthetic polymers including those used in food containers, obese people may have increased levels of exposure to DEHP. This is of particular importance as previous studies showed that HFD could impair male fertility via reducing sperm concentration and motile sperm count (Mu et al., 2017; Crean and Senior, 2019). Further, rodent studies showed that HFD resulted in not only weight gain, but also decreasing levels of testosterone (Vigueras-Villasenor et al., 2011; Zhao et al., 2018). It is reported that HFD leads to abnormal sperm development through testicular metabolic molecules (Crisóstomo et al., 2021). Besides, testicular metabolic abnormalities caused by HFD were often accompanied by non-reversible spermatogenic cell damage (Crisóstomo et al., 2019 ). Furthermore, studies proposed that the effects of obesity on spermatogenesis was highly dependented on testicular metabolism (D’Cruz et al., 2012; Rato and Sousa, 2021). Current literature suggests that DEHP may augment the adverse effects of obesity-induced damage to the male reproductive system via inducing excessive oxidative stress (Zhao et al., 2018), and that prenatal exposure to DEHP can result in a lasting effect on obesity phenotype in offspring (Lee et al., 2016; Fan et al., 2020). Because of this potentially synergistic relationship, we set out to investigate the relationship between exposure to DEHP and HFD on male reproductive injury.

To investigate the harmful male offspring reproductive effects of prenatal DEHP exposure, we utilized a low dose (0.2 mg/kg/day) prenatal exposure model. Next, male offspring were placed on a HFD during the pubertal period to determine whether there was increased reproductive function damage and spermatogenesis disorder. Testicular metabolomic profiling and weighted co-expression network analysis (WCNA) was applied to investigate potential correlations between phenotypes and the highly correlated modules clustered by the obtained key metabolites for further verification and interpretation. Our findings provide novel insights into the mechanism of spermatogenesis disorder and highlight the importance of evaluating early life exposure to adverse environmental factors and lifestyle on offspring reproductive health.

2. Materials and methods

2.1. Animals and dosing regimen

Six-week-old ICR mice were provided by SLAC Laboratory Animal Co, Ltd. (Shanghai, China). Animal handling and procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Nanjing Medical University. Mice were acclimatized before the study for 1 week and maintained in a temperature- and humidity-controlled (23 ± 1 °C, 53 ± 2%) room on a 12 h light-dark cycle. All mice used in this study were provided with free food and water.

DEHP (99% purity) was obtained from Sigma-Aldrich (St. Louis, USA). US Environmental Protection Agency (EPA) reference dose for human exposure reported that the reference dosage (RfD) of DEHP was 20 μg/kg/day (Lee et al., 2016). Actually, accumulative human evidence estimated the daily intake of DEHP was up to be about 2.7 mg/kg/day (Lee et al., 2016). Previous studies showed that DEHP negatively affected spermatogenesis in animals at high doses, but functioned on the order of 2 mg/kg/day (Andrade et al., 2006). Seven-week-old female mice were assigned to the control and treatment groups (N = 14, n = 7 per group) randomly. Female mice were dosed daily by oral gavage with 1% DMSO or DEHP (0.2 mg/kg/day) dissolved in 1% DMSO (vehicle) beginning 7 days prior to parental mating and gavaged continuously for 28 days. Glass water bottles were applied in administration of DEHP to avoid potential leakage of related compound from plastic bottles. Eight-week-old female mice were individually caged with aged-match males for fertility. Pregnant mice and male mice were treated with a standard laboratory diet (12% calories from fat, 20.6% calories from protein, and 67.4% calories from carbohydrates, 3.62 kcal/g; Cooperative Medical Biological Engineering Co, Ltd, China).

After weaning, two group male pups were divided into four offspring groups (N = 28, n = 7 per group) for the establishment of high-fat diet-induced obese (DIO) mouse model. DIO mice were fed with high fat diet (HFD, 60% calories from fat, 20% calories from protein, and 20% calories from carbohydrates, 5.24 kcal/g; Cooperative Medical Biological Engineering Co, Ltd, China). Lean mice were fed with a standard laboratory diet (12% calories from fat, 20.6% calories from protein, and 67.4% calories from carbohydrates, 3.62 kcal/g; Cooperative Medical Biological Engineering Co, Ltd, China). At 12-week-old, mice were sacrificed by CO2 euthanasia and all samples were stored in nitrogen.

2.2. Fertility test

Seven-week-old female ICR mice were obtained from SLAC Laboratory Animal Co, Ltd. (Shanghai, China) and adapted to 1 week. Fertility test was carried out by mating within each aged-match male mouse with a female for 1 week until vaginal sperm plug were observed. We calculated and recorded the litter size (number of pups per litter), percent of fertility (number of males that produce litter/total number of males × 100%; number of females that produce litter/total number of females × 100%), and the sex of the pups (number of female or male little size each pregnant mouse) as previously presented (Barakat et al., 2017).

2.3. Epididymalsemen analysis

The left side cauda epididymis was carefully dissected away from the fat after the mice was sacrificed. The isolated cauda epididymis were immediately transferred and excised into 200 μL pre-warmed 37 °C phosphate buffered saline (PBS). The sperm suspension was collected following 10 min incubation at 37 °C to allow spermatozoa to swim out the minced epididymis. The sperm suspension was put in a pre-warmed chamber slide (2X-CEL Chamber, 20 μL, Hamilton Thorne). Semen quality including the sperm count and motility, the curvilinear velocity (VCL; lm/sec), average path velocity (VAP; lm/s), beat cross frequency (BCF; Hz), straightness (STR; VSL/VAP × 100), straight line velocity (VSL; lm/s), amplitude of lateral head displacement (ALH; lm), as well as linearity (LIN; VSL/VCL × 100) were then examined from two chambers covering at least 20 different fields by the CASA system, IVOS (Hamilton-Thorne Research, Inc.) immediately. The above protocol was conducted according to previous studies (Zhao et al., 2018).

2.4. Determination of serum hormone levels

To obtain serum samples, peripheral blood (about 2 mL) was centrifuged at 3000×g for 10 min. Then serum samples were collected and preserved at − 80 °C for further analyses. The concentrations of testosterone, luteinizing hormone (LH) and follicle-stimulating hormone (FSH) levels were detected by an enzyme-linked immunosorbent assay (ELISA) kit (BioHJ Biotech, Xiamen, China) under the guidance of manufacturer’s introduction. All serum samples were analyzed by multi-function microplate reader (InfiniteM200 PRO, Tecan Co, Ltd, Switzerland) and the results were expressed as ng/L or IU/L.

2.5. Histological analyses and immunofluorescence

The testis and epididymis tissues were collected at 12-week-old and routinely fixed in Bouin’s solution, dehydrated, and embedded in paraffin. Each tissue was sectioned at 5 μm and stained by hematoxylin and eosin (H&E). In each pathological section, three different parts of testis were randomly selected to count different spermatogenic cells in the intact lumen. Spermatogenic tubules can be divided into different developmental stages according to the type of spermatogenic cells. The count of different types of spermatogenic cells was calculated by the standardization of Sertoli cell numbers in the same lumen.

For immunofluorescence, paraffin embedded testis sections were conducted to antigen retrieval using boiling in citrate buffer after dewaxing and hydration, then washed with PBS for three times. After blocking in 5% bovine serum albumin (BSA) for 2 h, samples were stained overnight at 4 °C with SOX9 antibodies (dilution 1:250, ab185230, abcam). Next, secondary antibodies were labeling to the samples for 2 h at room temperature after washing by PBS four times. The samples were washed and incubated with DAPI (ab104139, abcam) for 5 min at room temperature.

The histological features of H&E staining were observed and captured under a light microscope (Carl Zeiss). The histological features of immunofluorescence were scanned by a laser scanning confocal microscope (LSM710, Zeiss).

2.6. TUNEL assay

For TUNEL apoptosis detection, TUNEL Apoptosis Detection Kit (Vazyme) was applied to detect apoptotic cells following with the manufacturer’s instructions. The histological features of apoptotic cells were scanned by a laser scanning confocal microscope (LSM710, Zeiss).

2.7. Testicular metabolomic profiling

According to our previous study (Hu et al., 2017), we randomly selected seven testis samples per group to perform metabolomics analysis. For the preparation testis samples, 50 mg of representative testis was mixed with 750 μL ultra-pure water in a 1.5 ml Eppendorf tube. The mixture was fragmented and then sonicated for 5 min at 70% power and 5/3 pulses. The supernatant (150 μL) was then mixed 450 μL methanol, and centrifugalized at 16 000×g for 15 min. The supernatant was dried and the residue was dissolved in 20 μL ultra-pure water for metabolomics analysis.

LC-HRMS analysis was carrid out on an UPLC Ultimate 3000 system (Dionex, Germering, Germany) and combined with a Q-Exactive mass spectrometer in both positive and negative modes simultaneously (Thermo Fisher Scientific, Bremen, Germany). The instrument used a 1.9 μm Hypersile Gold C18 column (100 mm × 2.1 mm) to conduct the chromatographic separation (Thermo Fisher Scientific). A multistep gradient was coupled with a mobile phase A of 0.1% formic acid in ultra-pure water and a mobile phase B of acetonitrile acidified with 0.1% formic acid. Over a run time of 15 min, the gradient operated at a flow rate of 0.4 ml/min. Each sample was injected 10 μL volume per sample and analyzed randomly to avoid bias from the injection order. After full-scan acquisition ranging from 70 to 1050 m/z, the instrument operated at the resolution of 700 000. The metabolite identification was defined according to the accurate mass and the retention time of commercial standards. In this study, the first principal component of variable importance (VIP) in the projection was obtained exceeding 1 were first selected as changed metabolites.

2.8. Weighted co-expression network analysis (WCNA)

Weighted co-expression network analysis (WCNA, also known as weighted gene co-expression network) was conducted for further analysis. WCNA was a robust method for identification of the hub genes in the gene network and analyzing the correlations of gene co-expression pattern with traits of samples. It could also be used for metabolome data for providing functional explanations of metabolome with related traits (Bhargava et al., 2017; Mahajan et al., 2021). Briefly, the common analytic procedure of WCNA included 1) excluding the outlier samples considering the results of clustering analysis; 2) Selecting the appropriate soft threshold power and transformed the matrix into the topological overlap matrix (TOM); 3) Clustering for dissimilarity TOM and identifying the modules; 4) Estimating the correlations of each module with selected traits and calculating the hub metabolites for each module; 5) Validating the significant correlations between traits and modules (Langfelder and Horvath, 2008; Long et al., 2021).

For identification of key metabolites involved in the reproductive-disorder period in our study, WCNA was established to identify trait-related modules and hub metabolites based on our metabolomic profile (Pei et al., 2017). Appropriate soft thresholding power (β) were determined to build scale-free topology. We excluded the outlier according to our clustering analysis based on metabolomic profile of each sample. The weighted adjacencies were then transformed into a TOM. Cluster procedure was performed to construct metabolite module for subsequent analysis. Next, we linked trait data of the mice to the metabolite modules. We calculated the Pearson correlation coefficient between trait data (including sperm parameters, related treatments, sex hormone concentrations) and metabolite modules. Modules which were significantly correlated to specific trait would be focused based on our prior results. Hub metabolites were defined as metabolites highly conncected with nodes of the selected modules and were determined via calculating the correlations between metabolites and selected modules (Langfelder and Horvath, 2008). To identify key pathways that may be related to reproductive damage caused by HFD and DEHP exposure, we then put the top 20 hub metabolites in related modules for KEGG pathway analysis. We additionally validated the correlations between specific trait and related modules by calculating correlations between metabolite significance (MS) and module memberships (MM) (Langfelder and Horvath, 2008; Guo et al., 2021).

2.9. Statistical analysis

Differences between groups were examined for statistical significance by the ANOVA analysis or Student’s t-test. Group comparisons of quantitative data were assessed with two-way ANOVA analysis followed by Sidak post-hoc analysis performed for pair wise comparisons as appropriate. All statistical tests were performed by GraphPad Prism (Version 6.01) software, and all analysis data are represented as mean ± SEM. For all tests, P < 0.05 was defined as significant.

3. Results

3.1. HFD aggravates prenatal low-dose DEHP exposure induced impairments of gonadal and subfertility in male adult offspring

To explore synergistic effects of prenatal exposure to low-dose DEHP and pubertal HFD on male offspring reproductive injury, female mice were treated with DEHP (0.2 mg/kg/day) for 4 weeks or a Vehicle Control 1% DMSO (Control). After weaning, the male offspring were treated with HFD or SD until 12 weeks of age (Fig. 1). As shown in Fig. 2A, significant differences in body weight were observed between the SD group and the HFD group as was expected. In line with our previous study (Fan et al., 2020), prenatal low-dose DEHP (0.2 mg/kg/day) exposure increased male offspring body weight in SD group significantly (P < 0.05). However, there was no significant change between Control + HFD subgroup and DEHP + HFD subgroup. As shown in Fig. 2B, prenatal exposure to low-dose DEHP resulted in a decrease in the testis organ coefficients in SD group, which was significantly exacerbated by HFD treatment. While there was an observed decrease in epididymis organ coefficients in the DEHP + SD group as compared to the Control + SD group, no significant statistical difference was detected, but the epididymis organ coefficients in the DEHP + HFD (Fig. 2C).

Fig. 1.

Fig. 1.

Open in a new tab

The experimental scheme of mouse model. 7-week-old female mice received daily oral administration DEHP (0.2 mg/kg/day) dissolved in 1% DMSO (vehicle) and started 7 days prior to parental mating continuously for 28 days. After weaning, the two groups of offspring male mice were given high-fat diet and standard diet for 9 weeks. Offspring at 12-weeks-old underwent further analysis.

Fig. 2.

Fig. 2.

Open in a new tab

HFD aggravates prenatal DEHP exposure induced impairments of gonadal and subfertility in male adult offspring. (A) The body weights of male offspring were measured for 12 weeks. (B)-(C) The testis organ coefficient and epididymis organ coefficient of male offspring were measured at 12 week. (D) The percentage of vaginal plugs of female mice. (E) The fertility rate of female mice. (F) The number of litter size of mice were measured at 12 week. Data shown are mean ± SEM. The data were analyzed by unpaired two-tailed Student’s ttest; *: P < 0.05, **: P < 0.01, compared with the control group. N = 7 mice/group. *: Control + SD subgroup vs. DEHP + SD subgroup, &: Control + SD subgroup vs. Control + HFD subgroup, #: DEHP + SD subgroup vs. DEHP + HFD subgroup.

To further evaluate the fertility of male offspring, 8-week-old male mice from four treatment subgroups (Control + SD, Control + HFD, DEHP + SD, DEHP + HFD) were mating with aged-match female mice for one week. As shown in Fig. 2D, no obvious change of the percentage of vaginal plugs was found among the four subgroups. However, fertility rate analysis showed that DEHP + SD subgroup had a reduction in fertility of about 20% when compared with the Control + SD subgroup. In support of the hypothesis that HFD treatment could further harm male fertility following prenatal DEHP exposure, the DEHP + HFD subgroup displayed a further 20% reduction in fertility rate when compared to DEHP + SD subgroup (Fig. 2E). The litter size for both the prenatal DEHP + SD subgroup and DEHP + HFD decreased compared with the Control + SD and Control + HFD, but there were not statistically different (Fig. 2F). Taken together, our results suggest that pubertal HFD treatment aggravates testicular injury caused by prenatal exposure to low-dose DEHP which led to a reduction in fertility rate.

3.2. HFD aggravates prenatal low-dose DEHP exposure induced spermatogenesis disorder in male adult offspring

In order to explore whether pubertal HFD treatment exacerbates the impact of prenatal exposure to low-dose DEHP on the sperm quality of the male offspring mice, we used computer assisted sperm analyzer (CASA) to assess the sperm quality. The results suggested that both prenatal exposure to low-dose DEHP and HFD treatment could significantly reduce sperm concentration in mice. Compared to Control + HFD subgroup, sperm concentration of DEHP + HFD subgroup and DEHP +SD subgroup were significantly reduced (Fig. 3A). Besides, sperm progressive and sperm motility were also significantly reduced in the DEHP + HFD subgroup beyond the effect of either HFD or DEHP exposure alone (Fig. 3B and C). These data provide further support for a synergistic influence of prenatal exposure to low-dose DEHP and a pubertal HFD treatment on sperm content and quality.

Fig. 3.

Fig. 3.

Open in a new tab

HFD aggravates prenatal low-dose DEHP exposure induced spermatogenesis disorder in male adult offspring. (A)-(C) Results of sperm concentration, sperm motility and sperm progressive. (D) Representative photomicrographs depicting H&E staining (20 ×) of testis. (E) The ratio of Pachytene/Sertoli cell at VII-VIII stage. (F) The ratio of Leptotene/Sertoli cell at IX-X stage. (G) The ratio of Diplotene/Sertoli cell at XI stage. (H) The ratio of Zygotene/Sertoli cell at XI-XII stage. (I) Representative photomicrographs depicting H&E staining (20 ×) of epididymis. Data shown are mean ± SEM. The data were analyzed by unpaired two-tailed Student’s t-test; *: P < 0.05, **: P < 0.01, ***: P < 0.001.

To explore the reasons for the decline in sperm quality, the testicular histology was evaluated. No significant differences were observed in lumen integrity and spermatogenic cells at the same stages based on the H&E staining (Fig. 3D). According to the sperm development process, we further counted the spermatogenic cells (pachytene (P), leptotene (L), diplotene (D) and zygotene (Z) cells) at different stages in the testicular section. Compared with Control + SD subgroup, the ratio of P/Sertoli cell at stage VII-VIII showed a decreasing trend in the DEHP + SD subgroup, and with a further decrease being observed in the DEHP + HFD (Fig. 3E). The ratio of L/Sertoli cell at stage IX-X, the ratio of L/Sertoli cell at stage XI and the ratio of Z/Sertoli cell at stage XI-XII showed a similar trend in the DEHP + HFD subgroup with the lowest ratios among the four subgroups (Fig. 3F–H). These results suggested that the number of spermatogenic cells was decreased at different stages. H&E staining of epididymis showed that the mature sperm decreased in DEHP + SD and DEHP + HFD groups when compared with the Control + SD group. However, no significant changes were observed in the Control + HFD group. These results suggested that HFD exacerbated the phenotype induced by DEHP exposure (Fig. 3I). Similar findings were also observed on spermatogenic cells. Taken together, these data suggest that HFD treatment aggravates prenatal low-dose DEHP exposure inducing spermatogenesis disorder.

3.3. HFD aggravates prenatal low-dose DEHP exposure induced apoptosis of germ cells and sex hormone disorders in male adult offspring

To further explore the potential mechanism of spermatogenesis dysfunction as a consequence of prenatal exposure to low-dose DEHP in male offspring treated with HFD, we evaluated the testicular apoptosis and serum hormone levels. Increased testicular cell apoptosis has been previously reported to directly cause a decrease in spermatogenic cells and testicular atrophy (Awoniyi et al., 1989). In our study, prenatal exposure to low-dose DEHP increased the number of TUNEL positive cells (i.e. apoptotic cells) per testis tube significantly when the mice treated with SD (Fig. 4A). Furthermore, HFD in conjunction with DEHP exposure leads to TUNEL positive cells increasing significantly relative to DEHP + SD subgroup (Fig. 4A).

Fig. 4.

Fig. 4.

Open in a new tab

HFD aggravates prenatal low-dose DEHP exposure induced apoptosis of germ cells and sex hormone disorders in male adult offspring. (A) Representative immunofluorescence confocal microscopic images and positive cells per testis tube of TUNEL stain. (Red represents TUNEL staining; Blue represents nuclear DNA staining by DAPI. (B)-(D) The quantification of plasma testosterone levels, follicle stimulating hormone levels, and luteinizing hormone levels by ELISA analysis. (E) The ratio of luteinizing hormone/testosterone. Data shown are mean ± SEM. The data were analyzed by unpaired two-tailed Student’s t-test; *: P < 0.05, **: P < 0.01, ***: P < 0.001, compared with the control group.

Given that previous research has shown that germ cell death is associated with androgen withdrawal (El Chami et al., 2005), we investigated the serum hormone levels in each of the four subgroups. ELISA assay analysis demonstrated that serum testosterone (T) levels were decreased in DEHP + SD subgroup compared to Control + SD subgroup while a more significant decline was observed in DEHP + HFD subgroup (Fig. 4B). Follicle stimulating hormone (FSH) levels decreased in both DEHP treatment but no statistical difference was observed (Fig. 4C). Serum luteinizing hormone (LH) levels were significantly elevated in both HFD groups when compared with SD groups, with the greatest increase being observed in the DEHP + HFD subgroup (Fig. 4D). This is of particular interest as LH stimulates the production of testosterone which is the key to sperm production (McLachlan et al., 2002). To directly assess this relationship, the ratio of LH/T was assessed and was found to increase in the DEHP + HFD subgroup (Fig. 4E).

Given that Sertoli cells regulate the production of sperm and provide necessary nutrients for the germ cells (Liu et al., 2020), we then tested to see whether levels of Sertoli cells differed between the four subgroups. Sertoli cells were identified by immunofluorescence staining of SOX9. We did not find significant differences in SOX9 positive cells between SD and HFD control groups (Figure S1). However, a significant decrease in Sertoli cells was observed in the DEHP + HFD subgroup (Figure S1). Taken together, our results suggest that HFD treatment aggravates male reproductive dysfunction induced by prenatal low-dose DEHP exposure through disturbing sex hormone levels and promoting testicular apoptosis.

3.4. Contribution effect of testicular metabolome altered by prenatal low-dose DEHP exposure was greater than diet in male adult offspring

The metabolome is considered to be closer to the phenotype (ter Kuile and Westerhoff, 2001) and dynamic, quantitative complement of small-molecules present in testicular metabolome are better to unveil the phenotype in our system. To understand how diet and prenatal low-dose DEHP exposure change the testicular metabolome, we applied ultra-high performance liquid chromatography (UPLC) combined with high resolution mass spectrometry (HRMS) based metabolomics of testis from four subgroups male offspring (Control + SD, Control + HFD, DEHP + SD, DEHP + HFD). As shown in Fig. 5A–C, partial least squares discriminant analysis (PLS-DA) demonstrated that clear separation in the testicular metabolome between each two subgroups. The metabolome data of Control + SD subgroup and DEHP + SD subgroup, Control + HFD subgroup and DEHP + HFD subgroup and DEHP + SD subgroup and DEHP + HFD subgroup were analyzed respectively. There were forty significant testicular metabolites were changed in Control + SD subgroup and DEHP + SD subgroup, nineteen significant metabolites in Control + HFD subgroup and DEHP + SD subgroup, and fifty-seven significant metabolites in DEHP + SD subgroup and DEHP + HFD subgroup (P < 0.05, VIP > 1) (Table S1-3). The top ten differential metabolic molecules were shown in Fig. 5D–F, including guanine, Xanthosine, 8-Hydroxy-deoxyguanosine, riboflavin, all-trans-retinoic acid (Table S1-3). These results suggest that prenatal DEHP exposure and HFD treatment may result in an increase of oxidative stress in the testis (Safarinejad, 2011; Huang et al., 2017).

Fig. 5.

Fig. 5.

Open in a new tab

Contribution effect of testicular metabolome altered by prenatal low-dose DEHP exposure was greater than diet in male adult offspring. (A)-(C) PCA score plots of the testis from Control + SD subgroup and DEHP + SD subgroup, Control + HFD subgroup and DEHP + HFD subgroup, and DEHP + HFD subgroup and DEHP + SD subgroup. (D)-(F) The top10 VIP value of testis from Control + SD subgroup and DEHP + SD subgroup, Control + HFD subgroup and DEHP + HFD subgroup, and DEHP + HFD subgroup and DEHP + SD subgroup. (G)-(I) Summary of pathway analysis between Control + SD subgroup and DEHP + SD subgroup, Control + HFD subgroup and DEHP + HFD subgroup, and DEHP + HFD subgroup and DEHP + SD subgroup. N = 6–7/group.

Next, pathway enrichment analysis of the differential metabolites was performed (Fig. 5G–I). In Control + SD subgroup and DEHP + SD subgroup, the metabolites were enriched in riboflavin metabolism, biotin metabolism, retinol metabolism and tryptophan metabolism (Pathway Impact >0.1) (Fig. 5G). Similarly, differentially metabolites were enriched in riboflavin metabolism, retinol metabolism, tryptophan metabolism and pyrimidine metabolism in Control + HFD subgroup and DEHP + HFD subgroup (Pathway Impact >0.1) (Fig. 5H). Highly similar patterns of testicular metabolism subject to prenatal DEHP exposure were found regardless of SD or HFD treatment. In addition, the analysis results of DEHP + SD subgroup and DEHP + HFD subgroup showed that differentially metabolites were enriched in the retinol metabolism, biotin metabolism, nicotinate and nicotinamide metabolism, serine and threonine metabolism, glycine metabolism and glyoxylate and dicarboxylate metabolism (Pathway Impact > 0.1) (Fig. 5I). Altogether, these findings suggest that prenatal exposure to DEHP potentially induce the phenotype of spermatogenesis disorder by affecting the testicular metabolism of male adult offspring, and HFD treatment mainly aggravates the degree of its disorder.

3.5. Riboflavin and biotin metabolites contribute to HFD exacerbated spermatogenesis disorder phenotypes due to prenatal low-dose DEHP exposure

WCNA is a well-established method to establish scale-free gene co-expression networks enabling a better understanding of the correlations between the functionally similar modules. This method was applied to analyze testicular metabolites co-expression networks and complex phenotypic traits, like semen quality, subgroup, hormone profile. Firstly, we numbered samples of Control + SD group as K1–7, DEHP + SD group as L1–7, Control + HFD group as E1–6 and DEHP + HFD group as F1–6. Samples of all groups were clustered based on metabolomic profile to exclude the outliers. The results of clustering analysis were shown in Fig. 6A. K3 and F3 were identified as outliers and were not included in following analysis (Fig. 6A). The network was constructed using the soft-thresholding power as 12 based on a scale-free R2 (R2 = 0.8). The topology model fit index (R2) and mean connectivity for various soft-thresholding powers were presented in Fig. 6B and C. The metabolites were clustered into different modules according to the correlations between metabolites and related phenotypes or the treatments. Heatmap displayed the correlations between metabolites and traits. This analysis derived 6 modules, with metabolites that failed to be clustered in gray color (Fig. 6D). We listed the hub metabolites of each functional module in Supplementary materials (Table S4–S5). Pearson correlations analysis was used to calculate the Pearson correlation coefficient between trait data (including sperm parameters, related treatments, sex hormone concentrations) and metabolite modules. From the heatmap of the module-trait correlation, no significant correlation was found between brown module and DEHP, HFD or DEHP + HFD traits, respectively. The green module was observed positively correlated to DEHP trait while yellow module was negatively correlated to (green: r = 0.063, P = 0.00042, yellow: r = − 0.542, P = 0.0062). The correlations between the two modules and HFD trait were also found (green: r = − 0.634, P = 0.00088, yellow: r = 0.684, P = 0.00023). No significance of the correlations between green module or yellow module and DEHP + HFD trait was obtained in the heatmap (green: r = − 0.634, P = 0.00088, yellow: r = 0.684, P = 0.0002). The green and yellow modules were not correlated to sperm quality (sperm concentration, sperm progression and sperm motility), respectively. In the 6 cluster modules, the heatmap of the module-trait correlation showed that the blue module was significantly correlated to DEHP trait (r = − 0.76, P = 1.6e-0.5) and turquoise module was significantly correlated to HFD trait (r = − 0.698, P = 0.00015) (Fig. 6E). Meanwhile, the two modules were also significantly correlated to DEHP + HFD trait (blue: r = − 0.428, P = 0.037, turquoise: r = − 0.488, P = 0.015). Notably, significant correlations between sperm concentrations and blue module were observed (r = 0.622, P = 0.0012). We also observed significant relations between turquoise module and traits including testosterone (r = 0.579, P = 0.003), LH (r = − 0.493, P = 0.014) and sperm concentrations (r = 0.484, P = 0.017), while these correlations were consistent with the previous results. Hence, the blue and turquoise modules were considered as two modules of interest for subsequent analyses. In the turquoise module, the hub metabolites were enriched in phenylalanine, tyrosine and tryptophan biosynthesis, glycine metabolism and biotin metabolism pathway (Fig. 6F). In the blue module, the hub metabolites were identified to be related to steroid hormone biosynthesis and riboflavin metabolism (Fig. 6G). In addition, we also validated the significant correlations between modules and traits. The association plots of metabolite significance (MS) and module memberships (MM) were performed to validate the relationships between the specific traits and turquoise (Figure S2) and blue modules (Figure S3). All the correlations were consistent with the module-trait relationships. These data suggest that cluster modules of the testicular metabolites are significantly associated with the accompanying complex phenotypes altered by prenatal DEHP exposure and diet.

Fig. 6.

Fig. 6.

Open in a new tab

WCNA procedure in metabolites and related traits. (A) Clustering of selected samples detected one outlier sample. (B) Analyses of network topology model fit index for various soft-thresholding powers, scale-free topology was set as 0.8 roughly. Higher soft-thresholding powers representsed that the network was more in line with the distribution of the scale-free networks. (C) Distributions of mean connection level of the network corresponding to the soft-thresholding powers. (D) Metabolite dendrograms and modules with different colors derived from WCNA based on metabolomic profiles. Heatmap displayed the correlations between metabolites and traits. (E) Correlations between modules with different colors and related traits. (F) Pathway enrichment analysis of top 20 hub metabolites in turquoise module. (G) Pathway analysis of top 20 hub metabolites in blue module.

Next, we further identified the potential key metabolites involved in the relationships between the specific traits and the cluster modules (blue and turquoise). As shown in Fig. 7A, Venn diagrams were used to show the overlapped differentially metabolites in each subgroup (Control + SD vs DEHP + SD, Control + HFD vs DEHP + HFD) and the hub metabolites in blue or turquoise cluster modules. We found that there was a total of 11 overlapped metabolites that included xanthosine, riboflavin, indoleacrylic acid, thiamine, 3-Indoleacetonitrile, 3-Methylindole, L-Tryptophan, thymine, 2-Hydroxycaproic acid, biotin, cytidine and then enrolled in the subsequent analysis (blue: 9; turquoise: 2). The heatmap showed the expression pattern of the 11 key metabolites in each subgroup (Fig. 7B). Interestingly, we observed that the expression pattern of 11 testicular metabolites was overall decreased in DEHP + SD group. Notably, HFD aggravated its decreasing pattern in DEHP + HFD compared with DEHP +SD subgroups. KEGG enrichment analysis of these key metabolites found that riboflavin metabolism related to DEHP trait and biotin metabolism related to HFD trait had the largest pathway impact (Fig. 7C). We also found that a decreasing trend of relative concentrations in the two key metabolites (riboflavin and biotin) among four subgroups, and the lowest concentrations were observed in DEHP + HFD subgroup. Generally, riboflavin was significantly lower in DEHP + SD subgroup compared with Control + SD subgroup (P < 0.05). Similar findings were observed in DEHP + HFD subgroup compared with Control + HFD subgroup. (P <0.01), Meanwhile, the concentration of biotin was significantly decreased in DEHP + SD subgroup compared with Control + SD subgroup. HFD treatment accelerated the decrease of biotin concentration. Significant difference of biotin was found in DEHP + HFD subgroup compared with DEHP + SD subgroup (P < 0.05) (Fig. 7E and F). Collectively, these results indicate that the important roles of riboflavin and biotin involved in spermatogenesis disorder induced by prenatal exposure to low-dose DEHP and diet. Meanwhile, our results also provide the clue that the alterations of riboflavin and biotin potentially contribute HFD to exacerbate testicular development and spermatogenesis disorders due to prenatal exposure to low-dose DEHP in male offspring.

Fig. 7.

Fig. 7.

Open in a new tab

Integration analysis of WCNA and testicular metabolome. (A) Venn diagram of integration of results. (B) Heatmap of key metabolites in each sample. (C) Pathway analysis of key metabolites in blue module. (D) Pathway analysis of key metabolites in turquoise module. (E) Concentrations of Riboflavin among groups. (F) Concentrations of Biotin among groups. Students’t-test was performed to estimate the significant differences of metabolites between selected two groups. *: P < 0.05, **: P < 0.01, ns: no significance.

4. Discussion

In our previous study, prenatal low-dose DEHP (0.2 mg/kg/day) exposure causes an increase in the weights of male offspring mice significantly (Fan et al., 2020). This study demonstrates that prenatal low-dose DEHP exposure affects male spermatogenesis, alters gonad histology, reduces fertility rate, disturbs sex hormones, promotes testicular apoptosis and alters testicular metabolism in prenatally exposed males. In addition, pubertal HFD treatment exacerbates the observed reproductive function damage and spermatogenesis disorder following prenatal DEHP exposure.

Our study adds to the growing awareness of the important roles of environment in health and disease susceptibility regardless of early and late in life. According to the hypothesis for developmental origins of health and disease (DOHaD) (Barker, 2007), early life is a critical period of body development, and it is highly susceptible to external stimuli. Exposure to endocrine disrupting chemicals (EDCs) during early developmental windows can lead to long-term physiological changes in the body and increase the risk of diseases in adults (Guillette et al., 1995). Studies on the effects of prenatal and postnatal EDCs exposure showed that EDCs exposure at the sensitive window disrupted the development of their offspring reproductive function (Johnston et al., 2004; Hutchison et al., 2008). In this study, we found that prenatal exposure to low-dose DEHP caused the change of weight, sperm concentration, and testosterone at adulthood in offspring. Studies have shown that prenatal DEHP exposure can lead to male sperm damage, increased testicular apoptosis, and sex hormone disorders (Jones et al., 2014; Balci et al., 2020). However, another study showed that the prenatal exposure to DEHP had no adverse health problems in male offspring until the age of 23 months (Barakat et al., 2017). The difference in the temporal window of DEHP administration is a reason for the observed differences in those studies. We conducted DEHP management from one week before pregnancy to the entire pregnancy.

The reprogramming of the environment to early life and the emergence of patterns that stimulate late-life responses to the environment are central themes of DOHaD and have been observed in organs other than the liver (Greathouse et al., 2008; Wang et al., 2016). In later life, interactions between metabolome and environment can uncover the effect of this reprogramming with the host exhibiting aberrant responses to environmental challenges (ex. a Western-style diet). To further explore the effect of pubertal HFD on reproductive function in offspring following with prenatal DEHP exposure, male offspring were subjected to HFD treatment from postnatal day 21 to postnatal day 84. Notably, HFD treatment was found to aggravate these susceptibilities of reproductive function damage and spermatogenesis disorder following prenatal DEHP exposure in offspring. Previous study revealed that DEHP exposure created a vulnerability to obesity-induced damage to the male reproductive system (Zhao et al., 2018), which is consistent with our results. Our study highlights that both prenatal and postnatal stages are the sensitive windows for external stimuli on long-term reproductive health.

DEHP is a peroxisome proliferator and is considered a non-classical endocrine disruptor (Ward et al., 1998). Contrary to the classic endocrine disruptor that interferes with the endocrine process at the receptor level, the current hypothesis is that DEHP can damage reproductive function by antiandrogenic effects (Akingbemi et al., 2004). Considering the detrimental effect of obesity on endocrine potentials and the homeostatic reactivity, male obesity and DEHP exposure have a combined effect on plasma testosterone levels (Zhao et al., 2018; Moradi-Ozarlou et al., 2021). Testosterone (T) is generated by Leydig cells stimulated by LH, which involved in the initiation and maintenance of spermatogenesis in the Sertoli cells (Wang et al., 2017). Similar to prior studies, we observed that exposure to DEHP caused a decrease in T levels (Jones et al., 1993) and an concomitant increase of LH levels (Araki et al., 2017). Furthermore, we demonstrated that this dysfunction was exacerbated by a pubertal HFD exposure. The dynamic of LH/T is also important in initiating and maintaining sperm production (Ramaswamy and Weinbauer, 2014). Elevated biosynthesis rates of Leydig cells triggered by LH subsequently stimulate the reproduction T and eventually damage the spermatogenesis in mouse testis. Abnormal reduction in T levels can induce germ cell apoptosis (Sofikitis et al., 2008), spermatogenic cells reduction (Awoniyi et al., 1989), and even cause testicular atrophy (Huber et al., 1989). These findings confirmed our present results, suggesting that prenatal low-dose DEHP exposure and HFD treatment jointly induced spermatogenesis disorders via disturbing the synthesis or release of T and promoting testicular apoptosis in male adult offspring.

Either DEHP exposure or obesity can influence reactive oxygen species (ROS) production, sperm DNA damage, markers of inflammation, and alterations in reproductive hormones (Mu et al., 2017; Sun et al., 2018). In the process of spermatogenesis, unrepaired DNA accumulation will lead to arrest and apoptosis of spermatogenesis when ROS-triggered DNA damage in spermatogenic cells not repaired in time and effectively (Habas et al., 2017). Previous study believed that the phthalate metabolite mono-(2-ethylhexyl) phthalate (MEHP) selectively induced the oxidative stress response of germ cells and the release of mitochondrial cytochrome C, causing the expression of Fas receptor expression, and early start the apoptosis of germ cells (Erkekoglu et al., 2010). In the present study, the average numbers of apoptotic cells in the testis of the DEHP + SD subgroup increased compared with that of the Control + SD subgroup. In DEHP + HFD subgroup, the average number of apoptotic cells in the testis increased significantly, suggesting that HFD markedly increases apoptosis of testicular cells induced by prenatal low-dose DEHP exposure via elevating the level of oxidative stress in the testicles.

Testicular metabolome and WCNA indicated that prenatal exposure to low-dose DEHP altered the riboflavin metabolism and biotin metabolism, and HFD treatment further aggravated the decreasing trends of two key metabolites (riboflavin and biotin). Riboflavin is an essential vitamin B2 with powerful antioxidant and anti-inflammatory effects (Sanches et al., 2014). Riboflavin deficiency can result in biological oxidation, leading to a decline in antioxidant capacity (Tang et al., 2019). Lack of riboflavin could also lead to insufficient adenosine triphosphate (ATP) production. The insufficient ATP disturbs the spermatozoa energy generation and motility which affects the male fertility (Mukai and Travis, 2012; Kuang et al., 2021). Also, feeding rat with a riboflavin-deficient diet demonstrated a markable decrease in sperm motility (Pinto and Cooper, 2014). The disruption of riboflavin transport resulted in male infertility through altering flavoenzyme-mediated bioenergetic metabolism (Kuang et al., 2021). Biotin is a coenzyme for decarboxylase required for gluconeogenesis and fatty acid oxidation. Biotin deficiency was reported to result in increasing cellular reactive oxygen species (ROS) levels (Depeint et al., 2006; Madsen et al., 2015). Higher ROS level has been known to be a trigger to cellular death during spermatogenesis (Landenberger et al., 2004; Nandi and Chowdhuri, 2021). The abnormal concentrations of biotin could also affect spermatozoa motility and morphology (Pastén-Hidalgo et al., 2020). DEHP, a peroxisome proliferator, can induce a large amount of ROS in vivo and in vitro (Erkekoglu et al., 2011). Also, previous study showed that HFD could increase the emission of ROS and reduce the activity of antioxidant enzymes (Ruegsegger et al., 2019). Excessive ROS production and insufficient antioxidant defense mechanism can lead to the occurrence of oxidative stress (Sies, 1997), which in turn affects the interaction of the hypothalamic-pituitary-gonad axis (HPG) and/or other hormone axes, disrupts hormone secretion, and causes testicular damage (Darbandi et al., 2018). These findings supported our study results, suggesting that prenatal low-dose DEHP exposure and HFD treatment potentially triggered oxidative stress by altering the riboflavin and biotin metabolites, thus contributing to exacerbate spermatogenesis disorder in male adult offspring.

5. Conclusion

Collectively, the findings of the present study indicated that prenatal low-dose DEHP exposure triggered a series of adverse effects on the function of reproduction and spermatogenesis and pubertal HFD aggravated the susceptibility of these features in male offspring. Also, we found that these harmful changes were related to the alterations of riboflavin and biotin metabolites through the testicular metabolome. Our study provides new insights into long-term reproductive toxicity of DEHP and highlights the importance for the early control and management of EDCs.

Supplementary Material

Supplemental Figures and Tables

HIGHLIGHTS.

  • Pubertal HFD exacerbates the male offspring reproductive dysfunction resulting from prenatal low-dose DEHP exposure.

  • Pubertal HFD exacerbates sex hormone alterations and testicular apoptosis in offspring exposed to prenatal low-dose DEHP.

  • Testicular metabolites are significantly associated with the phenotypes altered by prenatal DEHP exposure and pubertal HFD.

  • Riboflavin and biotin contribute to HFD exacerbated spermatogenesis disorders due to prenatal low-dose DEHP exposure.

Acknowledgements

This work was supported by the National Natural Science Foundation of China 81973080 (to X.Wang), 81671461 (to C.Lu), 82173548 (to Y. Qin). Priority Academic Program for the Development of Jiangsu Higher Education Institutions (Public Health and Preventive Medicine).

Footnotes

Declaration of competing interest

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

Supplemental Data

Supplemental Data includes three figures and five tables.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.chemosphere.2022.134296.

References

  1. Abd-Ellah MF, Aly HA, Mokhlis HA, Abdel-Aziz AH, 2016. Quercetin attenuates di-(2-ethylhexyl) phthalate-induced testicular toxicity in adult rats. Hum. Exp. Toxicol 35, 232–243. 10.1177/0960327115580602. [DOI] [PubMed] [Google Scholar]
  2. Adibi JJ, Hauser R, Williams PL, Whyatt RM, Calafat AM, Nelson H, Herrick R, Swan SH, 2009. Maternal urinary metabolites of Di-(2-Ethylhexyl) phthalate in relation to the timing of labor in a US multicenter pregnancy cohort study. Am. J. Epidemiol 169, 1015–1024. 10.1093/aje/kwp001. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Akingbemi BT, Ge R, Klinefelter GR, Zirkin BR, Hardy MP, 2004. Phthalate-induced Leydig cell hyperplasia is associated with multiple endocrine disturbances. Proc. Natl. Acad. Sci. U. S. A 101, 775–780. 10.1073/pnas.0305977101 [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Andrade AJ, Grande SW, Talsness CE, Gericke C, Grote K, Golombiewski A, Sterner-Kock A, Chahoud I, 2006. A dose response study following in utero and lactational exposure to di-(2-ethylhexyl) phthalate (DEHP): reproductive effects on adult male offspring rats. Toxicology 228, 85–97. 10.1016/j.tox.2006.08.020. [DOI] [PubMed] [Google Scholar]
  5. Araki A, Mitsui T, Goudarzi H, Nakajima T, Miyashita C, Itoh S, Sasaki S, Cho K, Moriya K, Shinohara N, Nonomura K, Kishi R, 2017. Prenatal di(2-ethylhexyl) phthalate exposure and disruption of adrenal androgens and glucocorticoids levels in cord blood: the Hokkaido Study. Sci. Total Environ 581–582, 297–304. 10.1016/j.scitotenv.2016.12.124. [DOI] [PubMed]
  6. Awoniyi CA, Santulli R, Sprando RL, Ewing LL, Zirkin BR, 1989. Restoration of advanced spermatogenic cells in the experimentally regressed rat testis: quantitative relationship to testosterone concentration within the testis. Endocrinology 124, 1217–1223. 10.1210/endo-124-3-1217. [DOI] [PubMed] [Google Scholar]
  7. Balci A, Ozkemahli G, Erkekoglu P, Zeybek ND, Yersal N, Kocer-Gumusel B, 2020. Histopathologic, apoptotic and autophagic, effects of prenatal bisphenol A and/or di(2-ethylhexyl) phthalate exposure on prepubertal rat testis. Environ. Sci. Pollut. Res. Int 27, 20104–20116. 10.1007/s11356-020-08274-6. [DOI] [PubMed] [Google Scholar]
  8. Barakat R, Lin PP, Rattan S, Brehm E, Canisso IF, Abosalum ME, Flaws JA, Hess R, Ko C, 2017. Prenatal exposure to DEHP induces premature reproductive senescence in male mice. Toxicol. Sci 156, 96–108. 10.1093/toxsci/kfw248. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Barker DJ, 2007. The origins of the developmental origins theory. J. Intern. Med 261, 412–417. 10.1111/j.1365-2796.2007.01809.x. [DOI] [PubMed] [Google Scholar]
  10. Bhargava P, Fitzgerald KC, Calabresi PA, Mowry EM, 2017. Metabolic alterations in multiple sclerosis and the impact of vitamin D supplementation. JCI Insight 2. 10.1172/jci.insight.95302. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Crean AJ, Senior AM, 2019. High-fat diets reduce male reproductive success in animal models: a systematic review and meta-analysis. Obes. Rev 20, 921–933. 10.1111/obr.12827. [DOI] [PubMed] [Google Scholar]
  12. Crisóstomo L, Jarak I, Rato LP, Raposo JF, Batterham RL, Oliveira PF, Alves MG, 2021. Inheritable testicular metabolic memory of high-fat diet causes transgenerational sperm defects in mice. Sci. Rep 11, 9444. 10.1038/s41598-021-88981-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Crisóstomo L, Rato L, Jarak I, Silva BM, Raposo JF, Batterham RL, Oliveira PF, Alves MG, 2019. A switch from high-fat to normal diet does not restore sperm quality but prevents metabolic syndrome. Reproduction 158, 377–387. 10.1530/REP-19-0259. [DOI] [PubMed] [Google Scholar]
  14. D’Cruz SC, Jubendradass R, Jayakanthan M, Rani SJA, Mathur PP, 2012. Bisphenol A impairs insulin signaling and glucose homeostasis and decreases steroidogenesis in rat testis: an in vivo and in silico study. Food Chem. Toxicol. : an international journal published for the British Industrial Biological Research Association 50, 1124–1133. 10.1016/j.fct.2011.11.041. [DOI] [PubMed] [Google Scholar]
  15. Darbandi M, Darbandi S, Agarwal A, Sengupta P, Durairajanayagam D, Henkel R, Sadeghi MR, 2018. Reactive oxygen species and male reproductive hormones. Reprod. Biol. Endocrinol 16, 87. 10.1186/s12958-018-0406-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Depeint F, Bruce WR, Shangari N, Mehta R, O’Brien PJ, 2006. Mitochondrial function and toxicity: role of the B vitamin family on mitochondrial energy metabolism. Chem. Biol. Interact 163. [DOI] [PubMed] [Google Scholar]
  17. Doull J, Cattley R, Elcombe C, Lake BG, Swenberg J, Wilkinson C, Williams G, van Gemert M, 1999. A cancer risk assessment of di(2-ethylhexyl)phthalate: application of the new U.S. EPA Risk Assessment Guidelines. Regul. Toxicol. Pharmacol 29, 327–357. 10.1006/rtph.1999.1296. [DOI] [PubMed] [Google Scholar]
  18. El Chami N, Ikhlef F, Kaszas K, Yakoub S, Tabone E, Siddeek B, Cunha S, Beaudoin C, Morel L, Benahmed M, Regnier DC, 2005. Androgen-dependent apoptosis in male germ cells is regulated through the proto-oncoprotein Cbl. J. Cell Biol 171, 651–661. 10.1083/jcb.200507076. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Erkekoglu P, Rachidi W, Yuzugullu OG, Giray B, Favier A, Ozturk M, Hincal F, 2010. Evaluation of cytotoxicity and oxidative DNA damaging effects of di(2-ethylhexyl)-phthalate (DEHP) and mono(2-ethylhexyl)-phthalate (MEHP) on MA-10 Leydig cells and protection by selenium. Toxicol. Appl. Pharmacol 248, 52–62. 10.1016/j.taap.2010.07.016. [DOI] [PubMed] [Google Scholar]
  20. Erkekoglu P, Rachidi W, Yuzugullu OG, Giray B, Ozturk M, Favier A, Hincal F, 2011. Induction of ROS, p53, p21 in DEHP- and MEHP-exposed LNCaP cells-protection by selenium compounds. Food Chem. Toxicol 49, 1565–1571. 10.1016/j.fct.2011.04.001. [DOI] [PubMed] [Google Scholar]
  21. Fan Y, Qin Y, Chen M, Li X, Wang R, Huang Z, Xu Q, Yu M, Zhang Y, Han X, Du G, Xia Y, Wang X, Lu C, 2020. Prenatal low-dose DEHP exposure induces metabolic adaptation and obesity: role of hepatic thiamine metabolism. J. Hazard Mater 385, 121534. 10.1016/j.jhazmat.2019.121534. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Fay M, Donohue JM, De Rosa C, 1999. ATSDR evaluation of health effects of chemicals. VI. Di(2-ethylhexyl)phthalate. Agency for Toxic Substances and Disease Registry. Toxicol. Ind. Health 15, 651–746. 10.1177/074823379901500801. [DOI] [PubMed] [Google Scholar]
  23. Fromme H, Lahrz T, Kraft M, Fembacher L, Dietrich S, Sievering S, Burghardt R, Schuster R, Bolte G, Volkel W, 2013. Phthalates in German daycare centers: occurrence in air and dust and the excretion of their metabolites by children (LUPE 3). Environ. Int 61, 64–72. 10.1016/j.envint.2013.09.006. [DOI] [PubMed] [Google Scholar]
  24. Gao DW, Wen ZD, 2016. Phthalate esters in the environment: a critical review of their occurrence, biodegradation, and removal during wastewater treatment processes. Sci. Total Environ 541, 986–1001. 10.1016/j.scitotenv.2015.09.148. [DOI] [PubMed] [Google Scholar]
  25. Gray LE Jr., Ostby J, Furr J, Price M, Veeramachaneni DN, Parks L, 2000. Perinatal exposure to the phthalates DEHP, BBP, and DINP, but not DEP, DMP, or DOTP, alters sexual differentiation of the male rat. Toxicol. Sci 58, 350–365. 10.1093/toxsci/58.2.350. [DOI] [PubMed] [Google Scholar]
  26. Greathouse KL, Cook JD, Lin K, Davis BJ, Berry TD, Bredfeldt TG, Walker CL, 2008. Identification of uterine leiomyoma genes developmentally reprogrammed by neonatal exposure to diethylstilbestrol. Reprod. Sci 15, 765–778. 10.1177/1933719108322440. [DOI] [PubMed] [Google Scholar]
  27. Guillette LJ Jr., Crain DA, Rooney AA, Pickford DB, 1995. Organization versus activation: the role of endocrine-disrupting contaminants (EDCs) during embryonic development in wildlife. Environ. Health Perspect 103 (7), 157–164. 10.1289/ehp.95103s7157. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Guo H, Zhang H, Sheng N, Wang J, Chen J, Dai J, 2021. Perfluorooctanoic acid (PFOA) exposure induces splenic atrophy via overactivation of macrophages in male mice. J. Hazard Mater 407, 124862. 10.1016/j.jhazmat.2020.124862. [DOI] [PubMed] [Google Scholar]
  29. Habas K, Brinkworth MH, Anderson D, 2017. Diethylstilbestrol induces oxidative DNA damage, resulting in apoptosis of spermatogonial stem cells in vitro. Toxicology 382, 117–121. 10.1016/j.tox.2017.03.013. [DOI] [PubMed] [Google Scholar]
  30. Hannon PR, Niermann S, Flaws JA, 2016. Acute exposure to di(2-ethylhexyl) phthalate in adulthood causes adverse reproductive outcomes later in life and accelerates reproductive aging in female mice. Toxicol. Sci 150, 97–108. 10.1093/toxsci/kfv317. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Hauser R, Meeker JD, Singh NP, Silva MJ, Ryan L, Duty S, Calafat AM, 2007. DNA damage in human sperm is related to urinary levels of phthalate monoester and oxidative metabolites. Hum. Reprod 22, 688–695. 10.1093/humrep/del428. [DOI] [PubMed] [Google Scholar]
  32. Heudorf U, Mersch-Sundermann V, Angerer J, 2007. Phthalates: toxicology and exposure. Int. J. Hyg Environ. Health 210, 623–634. 10.1016/j.ijheh.2007.07.011. [DOI] [PubMed] [Google Scholar]
  33. Hogberg J, Hanberg A, Berglund M, Skerfving S, Remberger M, Calafat AM, Filipsson AF, Jansson B, Johansson N, Appelgren M, Hakansson H, 2008. Phthalate diesters and their metabolites in human breast milk, blood or serum, and urine as biomarkers of exposure in vulnerable populations. Environ. Health Perspect 116, 334–339. 10.1289/ehp.10788. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Hu W, Dong T, Wang L, Guan Q, Song L, Chen D, Zhou Z, Chen M, Xia Y, Wang X, 2017. Obesity aggravates toxic effect of BPA on spermatogenesis. Environ. Int 105, 56–65. 10.1016/j.envint.2017.04.014. [DOI] [PubMed] [Google Scholar]
  35. Huang YF, Wang PW, Huang LW, Lai CH, Yang W, Wu KY, Lu CA, Chen HC, Chen ML, 2017. Prenatal nonylphenol and bisphenol A exposures and inflammation are determinants of oxidative/nitrative stress: a Taiwanese cohort study. Environ. Sci. Technol 51, 6422–6429. 10.1021/acs.est.7b00801. [DOI] [PubMed] [Google Scholar]
  36. Huber MA, Hall EH, Rathbun WA, 1989. The role of the dentist in diagnosing infection in the AIDS patient. Mil. Med 154, 315–318. [PubMed] [Google Scholar]
  37. Hutchison GR, Scott HM, Walker M, McKinnell C, Ferrara D, Mahood IK, Sharpe RM, 2008. Sertoli cell development and function in an animal model of testicular dysgenesis syndrome. Biol. Reprod 78, 352–360. 10.1095/biolreprod.107.064006. [DOI] [PubMed] [Google Scholar]
  38. Johnston H, Baker PJ, Abel M, Charlton HM, Jackson G, Fleming L, Kumar TR, O’Shaughnessy PJ, 2004. Regulation of Sertoli cell number and activity by follicle-stimulating hormone and androgen during postnatal development in the mouse. Endocrinology 145, 318–329. 10.1210/en.2003-1055. [DOI] [PubMed] [Google Scholar]
  39. Jones HB, Garside DA, Liu R, Roberts JC, 1993. The influence of phthalate esters on Leydig cell structure and function in vitro and in vivo. Exp. Mol. Pathol 58, 179–193. 10.1006/exmp.1993.1016. [DOI] [PubMed] [Google Scholar]
  40. Jones S, Boisvert A, Duong TB, Francois S, Thrane P, Culty M, 2014. Disruption of rat testis development following combined in utero exposure to the phytoestrogen genistein and antiandrogenic plasticizer di-(2-ethylhexyl) phthalate. Biol. Reprod 91, 64. 10.1095/biolreprod.114.120907. [DOI] [PubMed] [Google Scholar]
  41. Kuang W, Zhang J, Lan Z, Deepak RNVK, Liu C, Ma Z, Cheng L, Zhao X, Meng X, Wang W, Wang X, Xu L, Jiao Y, Luo Q, Meng Z, Kee K, Liu X, Deng H, Li W, Fan H, Chen L, 2021. SLC22A14 is a mitochondrial riboflavin transporter required for sperm oxidative phosphorylation and male fertility. Cell Rep. 35, 109025. 10.1016/j.celrep.2021.109025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Landenberger A, Kabil H, Harshman LG, Zempleni J, 2004. Biotin deficiency decreases life span and fertility but increases stress resistance in Drosophila melanogaster. J. Nutr. Biochem 15, 591–600. [DOI] [PubMed] [Google Scholar]
  43. Langfelder P, Horvath S, 2008. WGCNA: an R package for weighted correlation network analysis. BMC Bioinf. 9, 559. 10.1186/1471-2105-9-559. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Lee KI, Chiang CW, Lin HC, Zhao JF, Li CT, Shyue SK, Lee TS, 2016. Maternal exposure to di-(2-ethylhexyl) phthalate exposure deregulates blood pressure, adiposity, cholesterol metabolism and social interaction in mouse offspring. Arch. Toxicol 90, 1211–1224. 10.1007/s00204-015-1539-0. [DOI] [PubMed] [Google Scholar]
  45. Liu Y, Fan J, Yan Y, Dang X, Zhao R, Xu Y, Ding Z, 2020. JMY expression by Sertoli cells contributes to mediating spermatogenesis in mice. FEBS J. 287, 5478–5497. 10.1111/febs.15328. [DOI] [PubMed] [Google Scholar]
  46. Long J, Huang S, Bai Y, Mao J, Wang A, Lin Y, Yang X, Wang D, Lin J, Bian J, Yang X, Sang X, Wang X, Zhao H, 2021. Transcriptional landscape of cholangiocarcinoma revealed by weighted gene coexpression network analysis. Briefings Bioinf. 22. 10.1093/bib/bbaa224. [DOI] [PubMed] [Google Scholar]
  47. Madsen CT, Sylvestersen KB, Young C, Larsen SC, Poulsen JW, Andersen MA, Palmqvist EA, Hey-Mogensen M, Jensen PB, Treebak JT, Lisby M, Nielsen ML, 2015. Biotin starvation causes mitochondrial protein hyperacetylation and partial rescue by the SIRT3-like deacetylase Hst4p. Nat. Commun 6, 7726. 10.1038/ncomms8726. [DOI] [PMC free article] [PubMed] [Google Scholar]
  48. Mahajan UM, Alnatsha A, Li Q, Oehrle B, Weiss F-U, Sendler M, Distler M, Uhl W, Fahlbusch T, Goni E, Beyer G, Chromik A, Bahra M, Klein F, Pilarsky C, Grützmann R, Lerch MM, Lauber K, Christiansen N, Kamlage B, Regel I, Mayerle J, 2021. Plasma metabolome profiling identifies metabolic subtypes of pancreatic ductal adenocarcinoma. Cells 10. 10.3390/cells10071821. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. McLachlan RI, O’Donnell L, Meachem SJ, Stanton PG, de Kretser DM, Pratis K, Robertson DM, 2002. Identification of specific sites of hormonal regulation in spermatogenesis in rats, monkeys, and man. Recent Prog. Horm. Res 57, 149–179. 10.1210/rp.57.1.149. [DOI] [PubMed] [Google Scholar]
  50. Moradi-Ozarlou M, Moshari S, Rezaei Agdam H, Nomanzadeh A, Shahmohamadlou S, Razi M, 2021. High-fat diet-induced obesity amplifies HSP70–2a and HSP90 expression in testicular tissue; correlation with proliferating cell nuclear antigen (PCNA). Life Sci. 279, 119633. 10.1016/j.lfs.2021.119633. [DOI] [PubMed] [Google Scholar]
  51. Mu Y, Yan WJ, Yin TL, Zhang Y, Li J, Yang J, 2017. Diet-induced obesity impairs spermatogenesis: a potential role for autophagy. Sci. Rep 7, 43475. 10.1038/srep43475. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Mukai C, Travis AJ, 2012. What sperm can teach us about energy production. Reproduction in domestic animals = Zuchthygiene 47 (4), 164–169. 10.1111/j.1439-0531.2012.02071.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Nandi A, Chowdhuri DK, 2021. Cadmium mediated redox modulation in germline stem cells homeostasis affects reproductive health of Drosophila males. J. Hazard Mater 402, 123737. 10.1016/j.jhazmat.2020.123737. [DOI] [PubMed] [Google Scholar]
  54. Pant N, Shukla M, Kumar Patel D, Shukla Y, Mathur N, Kumar Gupta Y, Saxena DK, 2008. Correlation of phthalate exposures with semen quality. Toxicol. Appl. Pharmacol 231, 112–116. 10.1016/j.taap.2008.04.001. [DOI] [PubMed] [Google Scholar]
  55. Pastén-Hidalgo K, Riverón-Negrete L, Sicilia-Argumedo G, Canul-Medina G, Salazar-Anzures T, Tapia-Rodríguez M, Hernández-González EO, Roa-Espitia AL, Cedillo-Peláez C, Fernandez-Mejia C, 2020. Dietary biotin supplementation impairs testis morphology and sperm quality. J. Med. Food 23, 535–544. 10.1089/jmf.2019.0137. [DOI] [PubMed] [Google Scholar]
  56. Pei G, Chen L, Zhang W, 2017. WGCNA application to proteomic and metabolomic data analysis. Methods Enzymol. 585, 135–158. 10.1016/bs.mie.2016.09.016. [DOI] [PubMed] [Google Scholar]
  57. Pinto JT, Cooper AJL, 2014. From cholesterogenesis to steroidogenesis: role of riboflavin and flavoenzymes in the biosynthesis of vitamin D. Advances in nutrition (Bethesda, Md 5, 144–163. 10.3945/an.113.005181. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Ramaswamy S, Weinbauer GF, 2014. Endocrine control of spermatogenesis: role of FSH and LH/testosterone. Spermatogenesis 4, e996025. 10.1080/21565562.2014.996025. [DOI] [PMC free article] [PubMed] [Google Scholar]
  59. Rato L, Alves MG, Socorro S, Duarte AI, Cavaco JE, Oliveira PF, 2012. Metabolic regulation is important for spermatogenesis. Nat. Rev. Urol 9, 330–338. 10.1038/nrurol.2012.77. [DOI] [PubMed] [Google Scholar]
  60. Rato L, Sousa ACA, 2021. The impact of endocrine-disrupting chemicals in male fertility: focus on the action of obesogens. J Xenobiot 11, 163–196. 10.3390/jox11040012. [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Richardson KA, Hannon PR, Johnson-Walker YJ, Myint MS, Flaws JA, Nowak RA, 2018. Di (2-ethylhexyl) phthalate (DEHP) alters proliferation and uterine gland numbers in the uteri of adult exposed mice. Reprod. Toxicol 77, 70–79. 10.1016/j.reprotox.2018.01.006. [DOI] [PubMed] [Google Scholar]
  62. Ruegsegger GN, Vanderboom PM, Dasari S, Klaus KA, Kabiraj P, McCarthy CB, Lucchinetti CF, Nair KS, 2019. Exercise and metformin counteract altered mitochondrial function in the insulin-resistant brain. JCI Insight 4. 10.1172/jci.insight.130681. [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Safarinejad MR, 2011. Effect of pentoxifylline on semen parameters, reproductive hormones, and seminal plasma antioxidant capacity in men with idiopathic infertility: a randomized double-blind placebo-controlled study. Int. Urol. Nephrol 43, 315–328. 10.1007/s11255-010-9826-4. [DOI] [PubMed] [Google Scholar]
  64. Sanches SC, Ramalho LN, Mendes-Braz M, Terra VA, Cecchini R, Augusto MJ, Ramalho FS, 2014. Riboflavin (vitamin B-2) reduces hepatocellular injury following liver ischaemia and reperfusion in mice. Food Chem. Toxicol 67, 65–71. 10.1016/j.fct.2014.02.013. [DOI] [PubMed] [Google Scholar]
  65. Sies H, 1997. Oxidative stress: oxidants and antioxidants. Exp. Physiol 82, 291–295. 10.1113/expphysiol.1997.sp004024. [DOI] [PubMed] [Google Scholar]
  66. Silva MJ, Reidy JA, Herbert AR, Preau JL Jr., Needham LL, Calafat AM, 2004. Detection of phthalate metabolites in human amniotic fluid. Bull. Environ. Contam. Toxicol 72, 1226–1231. 10.1007/s00128-004-0374-4. [DOI] [PubMed] [Google Scholar]
  67. Sofikitis N, Giotitsas N, Tsounapi P, Baltogiannis D, Giannakis D, Pardalidis N, 2008. Hormonal regulation of spermatogenesis and spermiogenesis. J. Steroid Biochem. Mol. Biol 109, 323–330. 10.1016/j.jsbmb.2008.03.004. [DOI] [PubMed] [Google Scholar]
  68. Sun Y, Shen J, Zeng L, Yang D, Shao S, Wang J, Wei J, Xiong J, Chen J, 2018. Role of autophagy in di-2-ethylhexyl phthalate (DEHP)-induced apoptosis in mouse Leydig cells. Environ. Pollut 243, 563–572. 10.1016/j.envpol.2018.08.089. [DOI] [PubMed] [Google Scholar]
  69. Tang J, Hu J, Xue M, Guo Z, Xie M, Zhang B, Zhou Z, Huang W, Hou S, 2019. Maternal diet deficient in riboflavin induces embryonic death associated with alterations in the hepatic proteome of duck embryos. Nutr. Metab 16, 19. 10.1186/s12986-019-0345-8. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. ter Kuile BH, Westerhoff HV, 2001. Transcriptome meets metabolome: hierarchical and metabolic regulation of the glycolytic pathway. FEBS Lett. 500, 169–171. 10.1016/s0014-5793(01)02613-8. [DOI] [PubMed] [Google Scholar]
  71. Vigueras-Villasenor RM, Rojas-Castaneda JC, Chavez-Saldana M, Gutierrez-Perez O, Garcia-Cruz ME, Cuevas-Alpuche O, Reyes-Romero MM, Zambrano E, 2011. Alterations in the spermatic function generated by obesity in rats. Acta Histochem. 113, 214–220. 10.1016/j.acthis.2009.10.004. [DOI] [PubMed] [Google Scholar]
  72. Vo TT, Jung EM, Dang VH, Jung K, Baek J, Choi KC, Jeung EB, 2009. Differential effects of flutamide and di-(2-ethylhexyl) phthalate on male reproductive organs in a rat model. J. Reprod. Dev 55, 400–411. 10.1262/jrd.20220. [DOI] [PubMed] [Google Scholar]
  73. Walker C, Garza S, Papadopoulos V, Culty M, 2021. Impact of endocrine-disrupting chemicals on steroidogenesis and consequences on testicular function. Mol. Cell. Endocrinol 527, 111215. 10.1016/j.mce.2021.111215. [DOI] [PubMed] [Google Scholar]
  74. Wang Q, Trevino LS, Wong RL, Medvedovic M, Chen J, Ho SM, Shen J, Foulds CE, Coarfa C, O’Malley BW, Shilatifard A, Walker CL, 2016. Reprogramming of the epigenome by MLL1 links early-life environmental exposures to prostate cancer risk. Mol. Endocrinol 30, 856–871. 10.1210/me.2015-1310. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Wang Y, Chen F, Ye L, Zirkin B, Chen H, 2017. Steroidogenesis in Leydig cells: effects of aging and environmental factors. Reproduction 154, R111–R122. 10.1530/REP-17-0064. [DOI] [PMC free article] [PubMed] [Google Scholar]
  76. Ward JM, Peters JM, Perella CM, Gonzalez FJ, 1998. Receptor and nonreceptor-mediated organ-specific toxicity of di(2-ethylhexyl)phthalate (DEHP) in peroxisome proliferator-activated receptor alpha-null mice. Toxicol. Pathol 26, 240–246. 10.1177/019262339802600208. [DOI] [PubMed] [Google Scholar]
  77. Wu W, Zhou F, Wang Y, Ning Y, Yang JY, Zhou YK, 2017. Phthalate levels and related factors in children aged 6–12 years. Environ. Pollut 220, 990–996. 10.1016/j.envpol.2016.11.049. [DOI] [PubMed] [Google Scholar]
  78. Yeung BH, Wan HT, Law AY, Wong CK, 2011. Endocrine disrupting chemicals: multiple effects on testicular signaling and spermatogenesis. Spermatogenesis 1, 231–239. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Zhang XF, Zhang T, Wang L, Zhang HY, Chen YD, Qin XS, Feng YM, Feng YN, Shen W, Li L, 2013. Effects of diethylhexyl phthalate (DEHP) given neonatally on spermatogenesis of mice. Mol. Biol. Rep 40, 6509–6517. 10.1007/s11033-013-2769-y. [DOI] [PubMed] [Google Scholar]
  80. Zhao J, Ren S, Liu C, Huo L, Liu Z, Zhai L, 2018. Di-(2-Ethylhexyl) phthalate increases obesity-induced damage to the male reproductive system in mice. Oxid. Med. Cell. Longev 1861984. 10.1155/2018/1861984,2018. [DOI] [PMC free article] [PubMed]
  81. Zolfaghari M, Drogui P, Seyhi B, Brar SK, Buelna G, Dube R, 2014. Occurrence, fate and effects of Di (2-ethylhexyl) Phthalate in wastewater treatment plants: a review. Environ. Pollut 194, 281–293. 10.1016/j.envpol.2014.07.014. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental Figures and Tables
NIHMS1819055-supplement-Supplemental_Figures_and_Tables.zip (817.2KB, zip)

RESOURCES