Introduction
Millions of people across all ages, genders, nationalities, and socioeconomic backgrounds suffer from overweight and obesity [1]. Although the pathophysiology of obesity is extremely complex, the primary cause of the disorder is a positive energy imbalance, which is further influenced by a variety of environmental and genetic factors [2]. Excess energy can be stored as fat in the body, potentially disrupting numerous physiological processes in the endocrine glands, immune system, and blood vessels [3], as well as increasing the body mass index (BMI). Fat deposition is also linked to various non-communicable diseases, such as type 2 diabetes and cancer [4,5]. Obesity not only increases the risk of non-communicable diseases but can also impact fertility. Parental obesity is a significant risk factor for obesity in children. This risk is likely triggered by genetic variations and early exposure to environmental factors; however, these mechanisms are not yet fully understood. Research findings suggest that obesity and overweight may have a hereditary component [6]. Metabolic changes caused by parental obesity can affect epigenetic markers in oocytes and sperm, potentially influencing epigenetic programming and reprogramming processes during embryogenesis. However, there is still much to learn about the fundamental processes of development and adipogenesis in the early stages of embryogenesis [7].
It is well established that obesity is associated with infertility, independent of ovulatory disorders, and increases the risk of spontaneous abortions [8,9]. Additionally, obesity is strongly linked to perinatal complications such as preeclampsia, preterm birth, and gestational diabetes [10]. Nowadays, obese women who struggle to conceive naturally may turn to assisted reproductive techniques (ART); however, the outcomes of these pregnancies remain unclear. While some studies indicate that obesity has no direct negative effects on conception rates [11,12], the presence of a higher proportion of poor responders in the obese population may skew the overall results [13]. Further data suggests that obesity is associated with lower rates of implantation, pregnancy, and live births [14]. Obesity also adversely affects the endocrine, follicular, embryonic, and uterine systems. It impairs the maturation of oocytes at the follicular level by increasing lipid concentrations and inflammatory markers in the follicular fluid, altering cumulus cell gene expression, and inducing endoplasmic reticulum stress [15].
While the impact of female obesity on embryologic parameters has been extensively studied, the adverse effects of male obesity on human embryos have garnered less attention. Obesity negatively affects sperm concentration, motility, and DNA quality. Additionally, obesity is linked to a systemic and chronic inflammatory response. Fat cells continuously release inflammatory chemicals, contributing to a pro-inflammatory state and heightened oxidative stress, which further exacerbates sperm DNA damage [16]. Prolonged exposure to reactive oxidative species during spermatogenesis leads to changes in DNA methylation patterns and chromatin structure, potentially affecting paternal epigenetic contributions to embryogenesis later on. The effects of paternal obesity on the development of early human embryos have not been thoroughly investigated. Researchers have shown a keen interest in in vitro pre-implantation embryo development, as it provides a unique opportunity to observe the direct impacts of paternal factors, uninfluenced by the mother's uterine environment [17].
Although lower pregnancy and birth rates have been observed following ART in obese parents [18], studies examining the impact of BMI on embryos have yielded contradictory results. Some research suggests that parental BMI does not influence embryo development [19], while other studies report adverse effects on fertilization and embryo cleavage rates [20]. This inconsistency may stem from the inaccuracy of static morphology-based methods commonly employed in in vitro fertilization (IVF) laboratories [21,22]. Recently, the adoption of time-lapse technology has facilitated a more accurate analysis by incorporating morphokinetic parameters, which have been shown to enhance pregnancy outcomes [23]. The current study investigated the relationship between parental BMI, sperm and oocyte quality, and embryo culture up to day 5. It also examined the impact of parental BMI on embryo implantation potential and ART outcomes following embryo transfer.
Methods
1. Calculation of BMI and categorization of patients
Patients were selected from those referred to the infertility treatment center. All participants provided signed consent forms. The women included in this study used their own eggs during their first intracytoplasmic sperm injection (ICSI) cycle, and the BMI of their husbands was calculated and recorded in the electronic medical record system. Height and weight measurements were taken using a standardized method, and BMI was calculated as weight in kilograms divided by height in meters squared (kg/m2). Based on the most recent World Health Organization (WHO) classification, both male and female patients were categorized into nine BMI groups (Figure 1):
Group 1: underweight male and female (G1; n=94)
Group 2: underweight male, normal-weight female (G2; n=111)
Group 3: underweight male, overweight female (G3; n=79)
Group 4: normal-weight male, underweight female (G4; n=71)
Group 5: normal-weight male and female (G5; n=255)
Group 6: normal-weight male, overweight female (G6; n=229)
Group 7: overweight male, underweight female (G7; n=89)
Group 8: overweight male, normal-weight female (G8; n=228)
Group 9: overweight male and female (G9; n=164).
(underweight: BMI <18.50 kg/m2; normal weight: 18.50≤ BMI <24.99 kg/m2; overweight: BMI ≥25 kg/m2)
1) Collection and semen preparation
Sperm samples were collected following 2 to 5 days of abstinence. After liquefaction (approximately 30 minutes at 37 °C), semen samples were prepared using the density gradient technique with 40% and 80% silane-coated silica in gamete buffer (Gradient Kit from Cook Medical). An 1 mL from the bottom layer (80% Sil-Select) was transferred to a centrifuge tube, and 1 mL from the upper layer (40% Sil-Select) was gently added on top. The liquefied sample was then placed over the upper layer, and the tube was centrifuged at 1,400 rpm for 14 minutes. Subsequently, approximately 3 mL of sperm medium was added to the pellet, followed by centrifugation at 1,200 rpm for 10 minutes. Finally, the sample was resuspended in 1 mL of sperm medium (Origio) and stored in an incubator at room temperature until needed.
2) Evaluation of semen parameters
The WHO guidelines (2019) were used to measure sperm concentration, motility, and morphology. Diff-Quik (Idevarzan-e-Farda Co.) was then utilized to stain the dried smears in order to assess sperm morphology. According to Kruger strict criteria, at least 200 spermatozoa were counted and classified as having either normal or abnormal morphology. Sperm defects are categorized as follows: head defects: large or small, conical, amorphous, vacuolated (more than two vacuoles or more than 20% of the head area occupied by unstained vacuolar areas), small or large acrosome areas; defects of the middle segment: bending, cytoplasmic remnants; tail defects: twisted, numerous [24]. Eosin-nigrosine staining was used to assess sperm viability. After evaluating 200 sperm, the results were expressed as a proportion of colored or pink sperm.
3) Evaluation of sperm DNA damage
To evaluate the amount of DNA damage, the sperm DNA fragmentation assay (SDFA) kit (Idevarzan-e-Farda Co.) was used according to the manufacturer's instructions. In brief, the following procedure was performed: 60 μL of semen sample was added to a tube with agarose, and a drop was placed on a glass slide. The coverslip was removed after 4 minutes at 4 °C. Then, acid denaturation for 7 minutes and lysis for 20 minutes were performed consecutively. Subsequently, the slide was dehydrated in 70%, 90%, and 100% ethanol, each for 2 minutes. After Wright staining, a total of 500 spermatozoa were counted per slide using a bright field microscope. Sperm chromatin dispersion (SCD) values were then calculated to evaluate sperm DNA integrity. An SCD value below 30% is considered normal.
4) Toluidine blue staining
Chromatin integrity was assessed using the Toluidine blue (TB) method [25]. TB, a cationic dye, binds to negatively charged DNA phosphate residues in loose chromatin and/or damaged DNA. In each sample, 200 sperm were randomly selected and examined under high magnification. Cells were classified into two groups: dark purple cells (TB+ cells: abnormal chromatin structure) and light blue cells (TB˗ cells: normal chromatin structure).
5) Measurement of reactive oxygen species
Reactive oxygen species (ROS) formation was measured using a chemiluminescence assay in semen. In this method, we used 10 μL of 5 mM luminol (5-amino-2,3-dihydro-1,4-phthalazinedione) as a probe, incubating the sample for 15 minutes in a Berthold luminometer (AutoLumat Plus 953). The results were expressed as relative light units per second per million sperm (RLU/sec/106 sperm).
6) Measurement of seminal plasma malondialdehyde and total antioxidant capacity levels
Lipid peroxidation and total antioxidant capacity (TAC) levels in seminal plasma were evaluated by measuring malondialdehyde (MDA) and TAC utilizing MDA and TAC assay kits (ZB-MDA-96A and ZB-TAC-96A; ZellBio GmbH) according to the manufacturer's instructions.
2. Ovarian stimulation protocol and oocyte collection
Gonadotropin-releasing hormone antagonists (Ganirelix; Merck Sharp & Dohme) and human recombinant follicle stimulating hormone (Puregon, Merck Sharp & Dohme; or Gonal-F, Merck) were used to stimulate the ovaries. The initial dose and subsequent adjustment during the treatment were tailored individually for each patient based on their gonadotropin response. Ovulation was triggered with 10,000 IU of human chorionic gonadotropin (hCG), administered 36 hours prior to oocyte retrieval (Ovitrelle Merck), when follicles measuring at least 17 to 18 mm in diameter were observed.
1) Oocyte evaluation and intracytoplasmic sperm injection
After retrieval, cumulus-oocyte complexes were cultured in fertilization medium (Sequential Fert; Origio). Cumulus cells were then removed through brief exposure to hyaluronidase, followed by mechanical action using a stripper (denuding pipettes; Vitromed). Subsequently, oocyte diameter was measured and analyzed to assess the quality of mature oocytes. Prior to ICSI, two perpendicular measurements of the oocyte ooplasm of the patients were performed. ICSI was carried out using standard procedures. For this process, the oocytes were oriented with the first polar body at the 12 o'clock position, and the injection needle was inserted at the 3 o'clock position.
2) Embryo culture in a time-lapse system
Injected oocytes were transferred to time-lapse dishes (Vitrolife) in 30 μL microdrops of culture medium (SAGE 1-Srep, Origio or GT RONI), which had been covered with paraffin oil (Origio) Embryos were cultured in a time-lapse device (EmbryoScope Timelapse System; Vitrolife) to evaluate morphokinetic parameters. During the study, no changes were made to the embryo culture conditions. Image capture was set for every 10 minutes at seven different focal levels for each embryo. Depending on the day of transfer, embryos were morphologically evaluated on day 3 or day 5 and classified by quality grade (1=poor; 2=moderate; 3=good). On day 3, the primary parameters evaluated were the degree of fragmentation, number of blastomeres, and symmetry between blastomeres. On day 5, the main parameters considered were the degree of expansion, hatching status, intracellular mass, and trophoectoderm quality.
3) Evaluation of time-lapse images
Embryo imaging was performed using software (Unisense FertiliTech). Cleavage time is defined as the point at which cell division is complete, with two cells fully separated and each surrounded by its own cytoplasmic membrane. The time points evaluated in this study are defined as follows: time of pronuclei appearance (tPNa); time of pronuclei fading (tPNf); t2 is the point at which the embryo displays two separate and distinct cells; t3 occurs when the embryo consists of three blastomeres; t4 is the time at which a four-blastomere embryo is formed; t5 marks the formation of a five-blastomere embryo; and t8 is when the embryo has eight blastomeres. The t8 time point was the final parameter assessed, even though embryos are often cultured until day 5 for transfer. Additional parameters evaluated include the intervals between t3 and t2 (cc2) and between t5 and t3 (cc3), which indicate the duration of the second and third cell divisions, respectively. The parameters s2 and s3 measure the simultaneity of cell divisions from two to four blastomeres (s2) and from five to eight blastomeres (s3).
4) Embryo transfer
The strategy for embryo transfer (ET) was determined by considering the mother's age, the couple's history, and the quality of the embryos. The transfer was conducted on either the third or fifth day following ICSI, guided by abdominal ultrasound in accordance with the guidelines set by the American Society for Reproductive Medicine. For younger patients (≤35 years) who did not have significant male infertility factors and had at least three high-quality embryos, a single embryo was transferred on the fifth day, taking into account both morphological scores and morphokinetic parameters. In other scenarios, either one or two embryos were transferred on the third day, with the remaining embryos being cultured and subsequently frozen.
5) Outcome measures
A positive β-hCG test confirmed implantation biochemically 2 weeks after ET. Ultrasound examinations at 6 and 12 weeks showed a gestational sac and the embryo's heartbeat, thereby confirming the pregnancy. Pregnancy is established between the 6th and 12th weeks if an embryo's heartbeat is detected. Additionally, spontaneous abortion is characterized as the loss of a clinical pregnancy before the 20th week of gestation.
3. Statistical analysis
Data analysis was performed using SPSS software ver. 23 (IBM Co.). The baseline characteristics of the various BMI groups were compared using the Kruskal-Wallis test. To evaluate the effect of BMI status on morphokinetic parameters, a general linear regression model was employed, in which the normal-weight group served as the reference, and other BMI groups were treated as variables. A p-value of less than 0.05 was considered to indicate statistical significance. Results were expressed as the mean±standard deviation.
Results
1. Basic characteristics of men
Of the 1,320 male participants in this study, 284 were underweight, 585 were normal weight, and 481 were overweight. The participants were categorized into three groups based on their BMI: (1) underweight (BMI <18.50 kg/m2), (2) normal weight (18.50≤ BMI ≥24.99 kg/m2), and (3) overweight (BMI ≥25 kg/m2). There were no statistically significant differences in baseline characteristics among the groups, with the exception of orchidopexy (Table 1).
2. Evaluation of semen parameters
Density gradient centrifugation is a technique used to prepare sperm by separating motile sperm with good morphology from dead and abnormal sperm, immature germ cells, and non-sperm cells [26]. This method was utilized in the current study to explore the impact of paternal BMI on common sperm characteristics. As shown in Table 2, sperm concentration was significantly lower in the overweight group than in the normal- and underweight groups. Notably, the percentage of sperm with normal morphology also differed statistically in the overweight group compared to the normal- and underweight groups. However, there were no statistically significant differences in motility, sperm lifespan, liquefaction time, volume, pH, or white blood cells/high-power field among the groups (Table 2).
3. Sperm chromatin integrity assessment
Although the exact mechanism by which a high paternal BMI influences the epigenetic state of sperm remains unclear, previous studies have linked paternal obesity to changes in sperm chromatin [27]. In order to assess sperm chromatin integrity and breakage, the sperm nucleus DNA integrity kit and TB staining were employed, respectively. The results showed that overweight males had a significantly higher level of DNA fragmentation (30.86%±3.54%) compared to underweight (12.40%±1.05%) and normal-weight men (16.71%±1.4%) (p<0.001) (Table 3). Additionally, the percentage of TB+ sperm, indicative of altered chromatin integrity, was higher in the semen of overweight men (29.45±0.73) compared to that of normal-weight (17.51±0.49) and underweight individuals (15.65±0.42) (p<0.001).
4. Measurement of oxidative stress parameters
Obesity and systemic oxidative stress have been found to have a strong association [28]. This study aimed to examine ROS levels in motile sperm across different paternal BMI categories. To achieve this, ROS, MDA, and TAC levels were measured using a chemiluminescence assay and a kit. The average ROS levels were 1.98 in overweight males, 1.63 in normal-weight males, and 1.60 in underweight males. ROS levels were significantly higher in overweight men compared to those in normal-weight and underweight men. Additionally, ROS levels in normal-weight men did not significantly differ from those in underweight men. The results also showed that overweight men exhibited higher MDA levels and lower TAC levels in their seminal plasma compared to the other groups (Table 3).
5. Basic characteristics of women
A total of 1,320 women undergoing their first IVF cycle at our fertility center were analyzed based on their BMIs. The participants included 254 underweight women (n=1,265 embryos), 594 women of normal weight (n=3,810 embryos), and 472 overweight women (n=2,772 embryos). The embryos from these patients were assessed for histological and morphokinetic parameters. Table 4 presents the baseline characteristics of the women across different BMI categories. There were no statistically significant differences in female age, percentage of mature oocytes, fertilization rate, or embryo quality score. However, educational levels varied significantly; the overweight group had lower educational attainment compared to the underweight and normal-weight groups (32.5% vs. 63.9% and 55.6%, respectively). The causes of infertility were consistent across all BMI categories (Table 4). In contrast, the number of oocytes retrieved from overweight women was significantly lower compared to normal-weight women (7.89±4.2 vs. 12.65±4.7, respectively; p<0.05). We also measured the diameter of oocytes among the different BMI groups, as it is considered an indicator of embryo developmental competence. Our findings revealed that overweight women had smaller oocytes (118±0.4, p<0.01), which are less likely to develop successfully after fertilization (Table 4).
6. Morphokinetic parameters
Morphokinetic parameters and pre-implantation embryo quality were evaluated, as these parameters can be strong predictors of implantation and ongoing pregnancy rates [29]. Table 5 illustrates the statistically significant influence of parental BMI on various morphokinetic parameters. This impact persisted even after adjusting for paternal and maternal age and the cause of infertility. Specifically, embryos from overweight women reached the tPNf stage more quickly than those from women of normal weight. Additionally, significant differences were observed in two morphokinetic measurements, t5 and t8, across the BMI groups (Table 5).
The results indicate that cleavage time 5 occurred later in the embryos of overweight women (G3 and G6) compared to the control group (G5), with times of 50.97 and 51.08 hours versus 49.25 hours, respectively (p<0.05). A similar trend was observed for cleavage time 8, where the times were 57.91 and 57.98 hours compared to 55.56 hours in the control group (p<0.01).
Obesity affects the epigenome and molecular levels of motile sperm. Considering the critical role of the sperm epigenome in early embryonic development, our study found that the tPNa was shorter in groups G7 and G8 compared to the control group (6.2 and 6.32 hours vs. 8.6 hours, p<0.05). Additionally, the tPNf occurred later in the control group (23.3 and 23.42 hours vs. 25.4. p<0.01). In overweight men (G7 and G8), the parameters t2 and t8 were delayed relative to the control group (G5) (t2: 27.88 and 28.14 hours vs. 26.09 hours; t8: 58.25 and 58.31 hours vs. 55.56 hours, respectively). Overweight males also experienced longer first and second embryonic cell cycles (cc1, cc2) compared to the control group (p<0.0001). However, there was no statistically significant difference between the groups. Additionally, there was a non-significant delay (p>0.05) in the third embryonic cell cycle (cc3), s2, and s3, between the groups of overweight men and the control group. Moreover, analyses indicated that embryos in the G9 group (overweight men and women) reached tPNa and tPNf earlier than those in the control group, and the morphokinetic parameters t2, t5, and t8 were reached later in G9 than in G5 (Table 5).
7. Clinical treatment outcomes before and after embryo transfer
We evaluated the individual and combined effects of maternal and paternal BMI on clinical treatment outcomes before and after ET. The results of the multilevel regression analysis, based on the patients' BMI, are presented in Table 6. There was a significant disparity in the fertilization rate between the overweight men's groups (G7, G8, and G9) and the control group; however, no significant differences were observed between the remaining groups and the control group (p>0.05). Embryo morphology was evaluated using quality scores, with no notable differences across BMI groups. The clinical parameter significantly influenced by maternal BMI was the abortion rate, which was higher in overweight women (groups G3, G6) and overweight couples (group G9) compared to the normal-weight group. Additionally, the clinical pregnancy rate was lower in overweight men and women, with a notable decrease observed in overweight couples (G9) compared to the control group (46.9% vs. 59.5%, respectively) (Table 6).
Discussion
Classical morphological parameters commonly used to assess embryo quality may not be sufficiently accurate to detect the potential effects of maternal or paternal BMI on oocytes and embryo development ability [17]. To address this limitation, previous studies have highlighted the effectiveness of time-lapse technology in enhancing the accuracy of embryo selection for ET. These studies have identified a strong correlation between varying implantation success rates and the temporal patterns of embryo cleavage [30,31]. In this study, we evaluated the impact of combined maternal and paternal BMI on embryo development using time-lapse technology. Previous research has independently assessed the effects of male and female BMI on static morphology or morphokinetic parameters. However, only a few studies have suggested a synergistic effect of obesity in couples on ART outcomes [32,33]. Here, we report findings indicating delayed development in cleavage-stage embryos, reduced clinical pregnancy rates, and increased miscarriage rates among overweight couples, as observed through time-lapse technology.
Our research findings demonstrated that paternal overweight is associated with changes in the molecular composition of the motile sperm as well as the morphokinetics of the pre-implantation embryo. We evaluated 481 overweight men, 585 normal-weight men, and 284 underweight men to assess the impact of obesity on semen parameters, characteristics of motile sperm, and embryo morphokinetics. Our findings strongly support previous reports that excessive BMI affects sperm parameters and can subsequently reduce male reproductive potential [34,35]. It is important to note that sperm motility and morphology do not necessarily correspond to their molecular composition [36]. Therefore, the primary objective of this study was to evaluate how paternal obesity affects the molecular composition of motile sperm. Following a density gradient analysis, we observed no significant variation in sperm motility among the overweight, normal weight, and underweight groups. However, the overweight group exhibited a significantly lower percentage of sperm with normal morphology compared to the other groups. Poor fertility outcomes, often characterized by low fertilization rates, poor embryo quality, repeated ART failures, and miscarriage, are frequently associated with issues in chromatin compaction and DNA integrity [37]. Our investigation highlighted the impact of male obesity on the decline in sperm quality and the outcomes following ART. Moreover, compared to normal-weight males, overweight men had a significantly higher percentage of motile spermatozoa with altered chromatin, suggesting a greater susceptibility to obesity-related changes. Studies have indicated that the increased production of ROS due to elevated paternal BMI impairs the integrity of sperm chromatin. There is a significant negative correlation between paternal BMI and the levels of ROS, MDA, TAC, DNA fragmentation index, and TB. Elevated paternal BMI compromises sperm chromatin integrity and density, leading to a higher percentage of immature sperm due to increased ROS production. Our findings corroborate previous data showing that BMI is linked to increased sperm DNA damage and ROS production [28]. Thus, it can be concluded that paternal BMI negatively affects the molecular constitution of motile sperm.
Similarly, we found that an increase in paternal BMI is associated with reduced fertilization and clinical pregnancy rates following ART cycles. Consequently, issues such as chromatin compaction and compromised DNA integrity are linked to adverse reproductive outcomes, including low fertilization rates, poor embryo quality, frequent ART failures, and miscarriages [38]. It is believed that post-fertilization, the sperm genome undergoes extensive epigenetic modifications, which involve the removal of the sperm nuclear envelope and the decondensation of chromatin through the reduction of protamine disulfide bonds. From this, we can infer that even minor alterations in the sperm profile can adversely affect embryo development [37]. Hence, in this present study, we have demonstrated the detrimental effects of high paternal BMI on early embryo development.
The outcomes of IVF in obese women and men have been documented in several studies. While some research indicates no adverse effects [21], other studies have identified alterations in several parameters, including fertilization rates [28], cleavage rates [20], and embryo quality [39]. In our observations, overweight men exhibited earlier pronuclei appearance and disappearance compared to the control group. Additionally, we found a significant association between paternal overweight and the duration of the first and third embryonic cell cycles, indicating delayed cell division [40,41]. This suggests that high paternal BMI negatively affects the developmental capabilities of human embryos. Our findings regarding embryonic morphokinetics are likely due to epigenetic changes, such as alterations in DNA methylation and/or chromatin structure, which are thought to influence cleavage divisions up to the 4-cell stage [17,42]. Not all obese men share a specific pathophysiological mechanism, and future research should aim to explore the roles of these mechanisms.
According to reports, a high maternal BMI may have significant effects on the embryo. However, the impact of maternal BMI on embryo quality remains unclear [31]. The current study found that both the number and diameter of oocytes in overweight women were significantly lower than those in the normal-weight group. The data also show a slower progression of cell division in the embryos of overweight women compared to those of normal-weight women. Significant differences were noted in embryos with four blastomeres (t4), five blastomeres (t5), and eight blastomeres (t8). Conversely, one study that acknowledged sample size as a potential limitation found that maternal BMI had no effect on cleavage-stage morphokinetics [43]. In the present study, in order to evaluate the combined effect of both maternal and paternal BMI, an extensive analysis of ICSI cycles was conducted using time-lapse technology and a relatively large sample size.
Early cleavage embryo development is delayed in obese mothers due to inherited genomic instability, which may be attributed to impaired regulation of embryogenesis. Indeed, embryos from overweight women tend to have fewer cells, and their glucose and amino acid metabolism is likely compromised [44]. The adverse effect of BMI on embryonic development is primarily linked to a decline in oocyte quality. Obese women have smaller oocytes, and oocyte diameter has been proven to be a reliable indicator of embryo development [45]. Obese women undergoing IVF exhibit higher levels of insulin, inflammatory markers, triglycerides, and non-esterified fatty acids (NEFA) in their follicular fluid [46]. The accumulation of triglycerides and NEFA indicates lipotoxicity, a major cause of organelle damage in oocytes. The influence of male BMI on ART outcomes has received less attention [47]. The first study on this topic revealed an association between high paternal BMI and a significant decrease in clinical pregnancy and live birth rates [48]. Additionally, a prospective study involving 114 couples who underwent 172 cycles of ICSI found that male obesity was linked to a reduced likelihood of achieving a live birth [49]. Male obesity can adversely affect hormonal balance, diminish semen quality, increase leptin and E2 levels, and disrupt spermatogenesis [50].
Except for the group of overweight couples (G9), there was no discernible variation in the pregnancy rate across the various BMI categories in the current study. However, embryos from obese groups exhibited slower growth and numerically lower ongoing pregnancy rates, supporting our hypothesis that maternal and paternal obesity may adversely affect embryo development. The impact of obesity on pregnancy rates remains a contentious issue in scientific literature. A recent multi-center retrospective analysis of 51,198 IVF cycles reported an overall negative effect of women’s BMI on ongoing pregnancy rates [14]. Another retrospective study evaluated the pregnancy outcomes of 9,587 oocytes donated by normal-weight donors and transferred to recipients of varying BMI. This study found decreased rates of implantation, pregnancy, and live births associated with higher BMI, indicating that maternal obesity might also affect uterine receptivity [51]. In contrast, other studies failed to show significant differences in live birth rates between normal-weight and obese patients, although these studies typically involved much smaller patient cohorts [52,53].
Our results indicated that male and female BMI had a negative synergistic effect on the outcomes of ART [32,54]. Additionally, obese women had a higher abortion rate compared to women of normal weight. Higher serum levels of leptin in obese women are associated with decreased endometrial receptivity and an increased risk of miscarriage [55,56]. The data from the present study also suggest that underweight status may adversely affect ART outcomes. However, the research on this phenomenon is limited and presents conflicting results [57,58]. Higher miscarriage rates in underweight women could be attributed to lower leptin levels, which have been previously implicated in impairing implantation and increasing the likelihood of recurrent miscarriages [43,59].
In conclusion, the current findings hold potential value in clinical practice. Given that obesity is linked to slower embryo development, particularly in terms of the time taken to reach five and eight blastomeres, these parameters could be utilized to enhance embryo selection and improve the prognosis for ET in obese patients. The time-lapse system, in contrast to static morphology, allows for a more accurate evaluation of embryonic development in relation to overweight and obesity. Overweight conditions can lead to metabolic issues that affect gamete quality and increase the risk of metabolic diseases in the offspring. Potential causes might include alterations in oocyte quality, DNA damage, and reduced cytoplasmic quality. Further research is necessary to elucidate the pathophysiological mechanisms by which maternal and paternal BMI influence pre-implantation development.
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