Skip to main content
Download PDF

Associations between testicular development and fetal size in the pig

Journal of Animal Science and Biotechnology volume 13, Article number: 24 (2022) Cite this article

Abstract

Background

Impaired reproductive performance is the largest contributing factor for the removal of boars from commercial systems. Intrauterine growth restricted piglets represent 25% of the total number of piglets born and have impaired reproductive performance. This study aimed to improve the understanding of temporal changes in testicular gene expression during testes development in fetuses of different size. The lightest and closest to mean litter weight (CTMLW) male Large White × Landrace littermates were collected at gestational days (GD) 45, 60 and 90 (n = 5–6 litters/GD).

Results

Testes weight and testes weight as a percentage of fetal weight were not associated with fetal size at GD60 or 90. Fetal plasma testosterone was not associated with fetal size at GD90. There was no association between fetal size and seminiferous tubule area and number, number of germ or Sertoli cells per tubule. The lightest fetuses tended to have wider seminiferous tubules compared to the CTMLW fetuses at GD90 (P = 0.077). The testicular expression of KI67 (P ≤ 0.01) and BAX:BCL2 ratio (P = 0.058) mRNAs decreased as gestation progressed. Greater SPP1 mRNA expression was observed at GD60 when compared with GD45 and 90 (P ≤ 0.05). Lower expression of DMRT1 and SPP1 (P < 0.01) mRNAs was observed in testes associated with the lightest fetuses compared to the CTMLW fetuses at GD90.

Conclusions

These findings provide novel insights into the expression profiles of genes associated with testicular development and function. Further, these data suggest that programming of reproductive potential in IUGR boars occurs late in gestation, providing a platform for further mechanistic investigation.

Background

Artificial insemination is extensively utilised to improve herd genetics and fertility, which has proved to be a successful tactic to increase the profitability of the pig industry [1, 2]. To maximise the profitability of a commercial pig system, it is essential that the boars utilised as semen donors have their fertility monitored. Impaired reproductive performance is the largest contributing factor for the removal of boars from commercial systems therefore, to maximise productivity, it is essential that boars with poor reproductive performance are identified as early as possible. Significant developmental changes must occur during development of the mammalian testes [3, 4], and perturbations in these processes prenatally can have significant consequences for fertility postnatally [5, 6]. Therefore, improving the understanding of the mechanisms regulating fetal testes development is critical to improve the understanding of the regulation of postnatal boar fertility, and is vital to improve the success of the pig industry.

It has been proposed that birth weight could be an easily applicable criterion for the selection of boars with the greatest reproductive potential. An association between birth weight and testicular weight and/or volume has been reported in both neonatal and adult boars [7,8,9]. Auler et al. [7] demonstrated that heavier male littermates have greater numbers of somatic and germ cells compared to their lightest male littermates on postnatal day 8, despite no alterations in circulatory testosterone or testicular 17α-hydroxylase expression. Further, despite a lack of differences in circulatory testosterone or testicular 17α-hydroxylase expression at 8 months of age, the lighter birthweight males had decreased numbers of pachytene spermatocytes and round spermatids, accompanied by decreased total numbers of elongated spermatids and daily sperm production.

To increase pig production, genetic selection for favourable traits such as litter size and ovulation rate has been utilised to increase the number of piglets weaned per sow per year [10]. Whilst this strategy has increased the number of live born piglets, there is substantial evidence that uterine capacity has not increased as anticipated, which hinders both litter size and piglet weight due to inadequate placental growth and efficiency [11, 12]. This has led to substantial within-litter variation in piglet weight and a high prevalence of low birthweight piglets; with approximately 25% of the total number of piglets born considered intrauterine growth restricted (IUGR) [13]. It has been suggested that IUGR in the pig may be programmed from an early stage of gestation, with significant within-litter variation in fetal size observed from as early as gestational day 30–35 [14,15,16,17,18]. IUGR in the pig severely impacts neonatal and adult development, with piglets exhibiting increased morbidity and mortality [12]. In addition, this phenotype has been suggested to be associated with impaired reproductive performance in boars, although this concept remains under-investigated. IUGR boars have been reported to have impaired testicular development, with decreased numbers of germ cells, testicular volume, semen volume, and fewer spermatozoa per ejaculate postnatally when compared to normally-grown piglets [8, 9]. Considering the high prevalence and severity of the phenotype, it is essential to improve the understanding of the mechanisms associated with impaired testis development in the lightest fetuses compared to their normal-sized littermates [19].

Whilst Pontelo et al. [20] recently described temporal changes in testicular size and histological structure during porcine fetal development, the expression profiles of genes associated with testicular development and function remain poorly understood. Significant changes in the structure of the testes occur during gestation to prepare for their essential role in male fertility later in life. In this study, the temporal expression profile of mRNAs with central roles in apoptosis, proliferation, and the testicular extracellular matrix was determined. Despite the appreciation that angiogenesis is essential for organ development and growth, and the essential role of angiogenesis in the postnatal testes, little is known regarding vascularisation of the fetal testes. Considering this, several of the candidate genes investigated in this study have central roles in the regulation of angiogenesis. Given the high prevalence of growth-restricted piglets, and the postnatal reproductive phenotype associated with IUGR, it is essential that we improve our understanding of the mechanisms governing fetal testicular development to improve the reproductive efficiency of the pig industry.

This study aimed to improve the understanding of temporal changes in testicular gene expression during fetal testes development. Further, it was hypothesised that the developmental trajectory of testicular development deviates between the lightest and normal sized fetuses.

Methods

All procedures were performed with approval from The Roslin Institute (University of Edinburgh) Animal Welfare and Ethical Review Board and in accordance with the U.K. Animals (Scientific Procedures) Act, 1986.

Experimental animals and sample collection

Large White × Landrace gilts (age 11–14 months) were observed daily for signs of oestrus and were housed in groups of 6–8 animals per pen. Oestrous cyclicity and ovarian function were controlled in accordance with routine normal practice at The Roslin Institute Large Animal Unit (Roslin, Midlothian, Scotland). All gilts were inseminated twice daily for the duration of oestrus with semen from one of four Large White sires. The sires used were equally distributed between gilts at the different gestational days (GD) of interest to minimise any effect of sire. The first day of insemination was assigned as GD0. Gilts were humanely killed with sodium pentobarbitone 20% w/v (Henry Schein Animal Health, Dumfries, Dumfries and Galloway, U.K.) at a dose of 0.4 mL/kg by intravenous injection via a cannula inserted in the ear vein at GD45, 60 and 90 (n = 5–6 gilts/GD).

Following confirmation of death, mid-ventral incision revealed the reproductive tract which was lifted from the body cavity and placed in a dissecting tray. Both uterine horns were dissected, from the ovary towards the cervix. All fetuses were weighed to identify the lightest and closest to mean litter weight (CTMLW) fetuses, with sex determined morphologically and confirmed by PCR for the SRY region of the Y chromosome as described previously [21].

For all developmental stages, the testes were dissected for analyses. At GD45, the testes were not weighed, but at GD60 and 90 the testes were weighed. One testis from each fetus was snap frozen in liquid nitrogen and stored at – 80 °C until required, and the other testis was fixed in Bouin’s for histological analysis (GD60 and 90 only). In addition, cardiac puncture was performed using an EDTA coated syringe to collect blood samples from the lightest and CTMLW male fetuses at GD90 (n = 5 litters). Plasma was obtained from fetal blood samples by centrifugation and samples were stored at − 20 °C until required.

Plasma testosterone quantification

Testosterone concentrations were determined in fetal plasma samples from the lightest and CTMLW male fetuses at GD90 (n = 5 CTMLW fetuses; 5 lightest fetuses) using an ELISA kit that has been validated for use with porcine samples (Abscitech, College Park, MD, USA; EK0379), as per the manufacturer’s instructions. The detection range of the assay was 0.1–20 ng/mL, with a sensitivity of 0.05 ng/mL.

Histological analysis

Testes samples from GD60 (n = 5 CTMLW fetuses; 5 lightest fetuses) and 90 (n = 6 CTMLW fetuses; 6 lightest fetuses) were fixed with Bouin’s overnight at room temperature and changed daily for approximately 1 week in 70% ethanol (Genta Medical, York, U.K.). Testes were processed by passing through graded ethanol (70%, 95%, and 99%; Genta Medical) and xylene (Genta Medical). The samples were then embedded in paraffin wax (Fisher Scientific, Loughborough, Leicestershire, U.K.) and 5 μm sections were cut and placed on polysine microscope slides (Fisher Scientific).

Following dewaxing and heat-induced epitope retrieval in 0.01 mol/L sodium citrate (Vector Laboratories), endogenous peroxidase activity was blocked by incubating slides with 0.3% hydrogen peroxide (Sigma Aldrich) in methanol. Non-specific binding sites were blocked by incubation with normal goat serum (Vectastain Elite ABC kit; Vector Laboratories, Orton Southgate, Peterborough, U.K.). Sections were incubated with a primary antibody for GATA binding protein 4 (GATA4) (sc-9053; Santa Cruz, Heidelberg, Germany) at a 1:200 dilution, or with rabbit immunoglobulin G (RIgG) (Vector Laboratories) (equivalent total protein concentration) as a negative control (Additional file 1: Supplementary Fig. 1). The slides were incubated in a humidified chamber at 4 °C overnight, washed in phosphate buffered saline (PBS), and incubated for 30 min at room temperature with a biotinylated anti-rabbit IgG secondary antibody (Vectastain Elite ABC kit; Vector Laboratories) at a dilution of 1:200 in PBS containing 1.5% normal goat serum. Sections were incubated with Vectastain Elite ABC reagent (Vectastain Elite ABC kit; Vector Laboratories) for 30 min, before incubation with the Novared peroxidase substrate (Vector Laboratories) for 5 min. Sections were then counterstained with haematoxylin and dehydrated in a graded series of ethanol and xylene (70%, 95%, and 99% ethanol; 99% ethanol 1:1 with xylene, and absolute xylene; Genta Medical). The sections were imaged using the NanoZoomer slide scanner (Hamamatsu, Shizuoka, Japan).

Image analysis

All image analyses were performed using ImageJ. Six images were taken at × 20 magnification for each section. For each testis, 2 sections were analysed which were a minimum of 5 serial sections apart from one another. The number of tubules were quantified and expressed as number per 0.1 mm2. For each tubule, the mean diameter was calculated by taking the average of 3 measurements. The total area for each tubule, and the number of germ cells and Sertoli cells per tubule were quantified.

Total RNA extraction and cDNA synthesis

RNA was extracted from the whole snap-frozen testis sample from each fetus as described previously [22,23,24]. The RNA was quantified, and the quality assessed spectrophotometrically using a Nanodrop ND-1000 (Labtech International Ltd., Heathfield, East Sussex, U.K.) and electrophoretically using a Tapestation 2200 (Agilent Technologies, Edinburgh, U.K.) (RNA Integrity Number Equivalent (RINe); 9.048 ± 0.092). Extracted RNA was stored at – 80 °C until required.

Complementary DNA (cDNA) was prepared from 0.3 μg of RNA with SuperScript III reverse transcriptase (Life Technologies, Paisley, Renfrewshire, U.K.) following the manufacturer’s instructions. Each reaction contained 250 ng random primers (Promega, Chilworth, Hampshire, U.K.) and 40 units RNaseIn (Promega). Negative controls without reverse transcriptase were included to check for genomic contamination and all cDNA was stored at – 20 °C until required.

Relative expression of candidate genes in testicular samples

The expression of candidate mRNAs was quantified  by qPCR in testes samples associated with the lightest and CTMLW male fetuses at GD45, 60 and 90. Quantitative PCR was performed on a Stratagene MX3000 instrument using SensiFAST® SYBR Lo-ROX (Bioline, London, U.K.) and cDNA from fetal testes at GD45, 60 and 90. All qPCRs were carried out at an annealing temperature of 60 °C and dissociation curves consisting of single peaks were generated. Candidate mRNAs were quantified: BCL-2-associated X protein (BAX), b-cell lymphoma 2 (BCL2), platelet endothelial cell adhesion molecule (CD31), doublesex and mab-3 related transcription factor 1 (DMRT1), GATA binding protein 4 (GATA4), hypoxia inducible factor 1 alpha subunit (HIF1A), KI67, tumour suppressor protein 53 (P53), secreted phosphoprotein 1 (SPP1) and vascular endothelial growth factor A (VEGFA). The reference genes, TATA box binding protein 1 (TBP1) and tyrosine 3-monooxygenase/tryptophan 5-monooxygenase activation protein, zeta polypeptide (YWHAZ) had stable expression in testes samples as determined by geNORM V3.5 (Ghent University Hospital, Centre for Medical Genetics, Ghent, Belgium). The expression of these genes was not affected by gestational day or fetal size (P > 0.1). The primer sequences for all genes investigated are detailed in Table 1 [25,26,27,28,29]. Serial dilutions of pooled cDNA ranging from 1:5 to 1:640 in nuclease-free water were used as standards. Sample cDNA was diluted 1:25 and 2 μL of sample, standard or control were added per well. Each plate contained duplicate wells of a no template control, standards, sample cDNA and reverse transcriptase blanks. SensiFAST® SYBR Lo-ROX supermix (5 μL), 10 μmol/L forward and reverse primer stock (0.4 μL each) and water (2.2 μL). Data were analysed using qbase+ software V3.0 (Biogazelle, Ghent, Belgium). A target and run specific strategy was employed and the results, normalised to the two reference genes, were scaled to the minimum sample. The mean slope, intercept, PCR efficiency and R2 values are detailed in Additional file 2: Supplementary Table 1.

Table 1 Primer sequence details for qPCRs
Full size table

Statistical analysis

All statistical analyses were performed using GenStat 13.1 (VSN International Ltd., Hemel Hempstead, Hertfordshire, U.K.). Mean values were calculated for each individual sample for each parameter investigated and the normality of the distribution of the data was assessed by an Anderson-Darling test. If a P value of ≤ 0.05 was obtained, then the data were not considered to have a normal distribution. Log10 and square root transformations were carried out to achieve normality of the distribution of the data where required. Outliers identified by a ROUT outlier test were excluded.

Where data had a normal distribution, one and two-way ANOVA for GD and fetal size was performed, with a block for gilt to account for the common maternal environment within litter. A post-hoc Tukey test was then performed. Where data did not have a normal distribution, Kruskal-Wallis and Mann Whitney tests were performed. In all cases results were considered significant when P ≤ 0.05, and trending towards significant when 0.05 < P < 0.1.

Results

Temporal changes in fetal and testes weight, the number of seminiferous tubules, seminiferous tubule diameter, and number of germ cells per seminiferous tubule

As anticipated, fetal and testes weights increased with advancing gestational day (Table 2; P < 0.001). The mean number of tubules per 0.1 mm2 was decreased at GD90 compared to GD60 (Table 2; P < 0.001), which was unsurprising given that tubule diameter tended to be larger at GD90 compared to GD60 (Table 2; P = 0.09). Further, there were more germ cells per tubule at GD90 compared to GD60 (Table 2, P < 0.05). Seminiferous tubule area and the number of Sertoli cells per tubule was not significantly affected by GD (Table 2).

Table 2 Fetal testes weights and histological analyses
Full size table

Fetal size was associated with altered tubule diameter at GD90

The lightest fetuses were significantly lighter than the CTMLW fetuses at GD90 (Fig. 1A; P < 0.001). Interestingly, despite the significant decrease in fetal weight in the lightest fetuses compared to the CTMLW fetuses, neither testes weight (Fig. 1B) or testes weight as a percentage of fetal weight (Fig. 1C) were significantly associated with fetal size at GD60 or GD90. The number of tubules per 0.1 mm2 (Fig. 1D), tubule area (Fig. 1F), and number of germ cells (Fig. 1G) and Sertoli cells (Fig. 1H) per tubule were comparable between the lightest and the CTMLW fetuses at GD60 and 90. There was a gestational day × fetal size interaction for tubule diameter, with the lightest fetuses tending to have a wider tubule diameter compared to the CTMLW fetuses at GD90 (Fig. 1E; P = 0.077). At GD90, fetal plasma testosterone concentration was not associated with fetal size (CTMLW 0.335 ± 0.102 ng/mL. Lightest 0.299 ± 0.096 ng/mL. P > 0.1).

Fig. 1
figure 1

Histological analyses of fetal testes at GD60 and GD90. Fetal weights at gestational day (GD) 45, 60, and 90 (A), paired fetal testes weight at GD60 and 90 (B), and paired testes weight as a percentage of body weight at GD60 and 90 (C) were compared between fetuses of different size. The number of seminiferous tubules (D), seminiferous tubule diameter (E), seminiferous tubule area (F), number of germ cells per tubule (G), and number of sertoli cells per tubule (H) were compared between fetuses of different sizes at GD60 and 90. Mean values presented. Error bars represent S.E.M. CTMLW = closest to mean litter weight. n = 4–6 per fetal size per gestational day.

Full size image

Temporal changes in testicular expression of KI67 and SPP1 mRNAs

There were temporal changes in expression of several of the candidate mRNAs investigated (Table 3). The expression of KI67 mRNA decreased with advancing gestation (P < 0.01), with a statistically significant decrease at GD90 when compared with GD45. Similarly, there was a trend towards a significantly lower BAX: BCL2 ratio with advancing gestation (P = 0.058). Testicular SPP1 mRNA expression peaked at GD60 (P < 0.05) when compared with GD45 and 90. There were no significant temporal changes in the testicular expression of BAX, BCL2, CD31, DMRT1, GATA4, HIF1A, P53, or VEGFA mRNAs (Table 3).

Table 3 Candidate testicular mRNA expression at days 45, 60 and 90 of gestation
Full size table

Testes from the lightest fetuses had lower expression of DMRT1 and SPP1 mRNAs compared to the CTMLW fetuses at GD90

The mRNA expression of the candidate genes was quantified and compared between testes from the lightest and the CTMLW fetuses within GD (Fig. 2). At GD45, there was a trend towards increased expression of DMRT1 mRNA (P = 0.09; Fig. 2E) in testes from the lightest fetuses compared to the CTMLW fetuses. Testes from the lightest fetuses had lower expression of DMRT1 (P < 0.01; Fig. 2E), and SPP1 (P < 0.01; Fig. 2J) mRNAs compared to the CTMLW fetuses at GD90. There were no statistically significant associations between fetal size and expression of BAX, BCL2, BAX: BCL2 Ratio, CD31, GATA4, HIF1A, KI67, P53, or VEGFA mRNAs (Fig. 2).

Fig. 2
figure 2

Candidate testicular mRNA expression in fetuses of different sizes at days 45, 60 and 90 of pregnancy. mRNA expression of BAX (A), BCL2 (B), BAX: BCL2 Ratio (C), CD31 (D), DMRT1 (E), GATA4 (F), HIF1A (G), KI67 (H), P53 (I), SPP1 (J) and VEGFA (K) in the lightest and closest to mean litter weight (CTMLW) porcine fetal testes at gestational days 45, 60 and 90. Error bars represent S.E.M. **P < 0.01 n = 5–6 per fetal size per gestational day

Full size image

Discussion

Improved understanding of the mechanisms and timing of developmental changes in the fetal testes is essential to improve knowledge of how male pigs reach their reproductive potential postnatally. Whilst Pontelo et al. [20] recently described temporal changes in testicular size and structure during porcine fetal development, the expression profiles of genes associated with testicular development and function remains poorly understood. In this study, the temporal expression profile of selected mRNAs with central roles in apoptosis (BAX and BCL2), proliferation (KI67), angiogenesis (CD31, HIF1A and VEGFA), and the testicular extracellular matrix (SPP1) and function (DMRT1 and GATA4) was determined.

Striking changes in the structure of the testes occur during gestation to prepare the testes for its important and essential role in male fertility in postnatal life. SPP1, also known as osteopontin, is an extra-cellular matrix (ECM) protein which functions by binding to integrin receptors expressed on the cell surface to promote cellular adhesion and communication [30]. Given the important role of this extracellular matrix protein in other tissues [31,32,33], the observed gestational stage-related changes in SPP1 mRNA expression are perhaps unsurprising. SPP1 protein is localised to the germ cells within seminiferous tubules, supporting cells in the seminiferous tubules and the interstitial Leydig cells of boar testes [34]. Given the striking change in SPP1 mRNA expression observed during late gestation, further studies should be performed to characterise the role of SPP1, integrins, and other ligands of the integrin receptors such as fibronectin, in the fetal testes, to gain better understanding of the mechanisms involved in testicular development.

In this study, neither testes weight or testes weight as a percentage of fetal weight were associated with fetal size at GD60 or GD90, suggesting that gonadal growth may be spared in these growth restricted fetuses, consistent with our previous findings in female fetuses at the same stages of gestation [26]. Similar findings have been reported in boars at 8 days and 8 months postnatally, suggesting that whilst overall testicular weight may be reduced in IUGR relative to normal weight boars, testicular weight is proportional to body size throughout development [7]. Whilst it is known that reproductive function is impaired postnatally in boars that are lighter at birth, and this study has demonstrated that there are alterations in testicular mRNA expression during fetal life, the idea that relative testicular growth is proportional to fetal size interesting and warrants further investigation.

IUGR boars postnatally have impaired testicular development, with decreased numbers of germ cells, testicular volume, semen volume, and fewer spermatozoa per ejaculate postnatally when compared to normally-grown piglets [8, 9]. Neonatal testes from the lightest boars have fewer total Sertoli, Leydig, and germ cells compared to the heaviest boars [7, 8]. Yet, at 10 months of age there was no association between seminiferous tubule diameter and birth weight [8]. At 8–10 months of age, there were fewer pachytene spermatocytes, round spermatids, elongated spermatids, and lower semen volume and number of spermatozoa per ejaculate in the lightest boars compared to normal or heavy birthweight boars [7, 9]. Whilst Smit et al. [8] reported lower total numbers of germ, Sertoli and Leydig cells, they did note that the number of each of these cell types per gram of testis weight was comparable. The developmental trajectory of testicular development in the lightest fetuses compared to their normally grown littermates remains poorly understood. In the present study, no association between fetal size and the number of tubules per 0.1 mm2, tubule area, germ cell number per tubule, or Sertoli cell number per tubule were observed at GD60 or 90. Pontelo et al. [20] reported a significant correlation between fetal weight and Sertoli cell number in testes from days 50, 80 and 106 of gestation. However, these correlations appear to have been performed on samples combined from multiple gestational days (GD50, 80 and 106), so they are perhaps not directly comparable with the data in the present study, which presumably more accurately reflect the relationships between fetal size and testicular development within gestational day. It has been postulated that testicular cell proliferation is greatest during the final trimester of pregnancy in the pig [20]. Given this, it could be hypothesised that the reproductive phenotype of IUGR boars postnatally originates due to alterations in proliferation and/or apoptosis during the final stages of fetal development, perhaps between day 90 and parturition.

Despite the appreciation that testosterone is vital for both the regulation of fetal development and postnatal reproduction, the relationship between plasma testosterone and boar size both prenatally and postnatally remains under investigated. In the current study, no association between fetal size and plasma testosterone concentration was observed at GD90. This is consistent with data from neonatal boars where plasma testosterone and the mRNA expression of 17α-hydroxylase were not associated with birth weight at 8 days and 8 months postnatally [7]. Similarly, Lin et al. [9] suggested that there was no association between circulating testosterone concentrations and birth weight in boars at postnatal days 82 and 242.

Given the striking differences that have been reported in the literature regarding variation in porcine fetal size from an early stage of gestation [14,15,16,17,18], and the significant development of the testes during early and mid-gestation, we hypothesized that significant differences in the testicular expression of candidate mRNAs involved in extracellular-matrix remodelling, proliferation and apoptosis would be observed across gestational stages and between littermates of different weight. Given the postnatal phenotype of IUGR boars, it could be speculated that enhanced apoptosis and impaired cellular proliferation would be observed in the testes of the lightest compared to the CTMLW fetuses. In this study, no association between fetal size and the expression of mRNAs involved in apoptosis and proliferation were observed at any stage investigated. However, it has been suggested that testicular cellular proliferation is greatest during the final trimester of pregnancy in the pig [20]. Given this, it could be hypothesised that the reproductive phenotype of IUGR boars observed postnatally originates due to alterations in cellular apoptosis and proliferation either during the initial stages of testes development or during the final stages of fetal development.

As described earlier, the ECM protein SPP1 is important for regulation of cellular adhesion and communication, and is known to be expressed in new-born boar testes [34]. In the present study, the lightest fetuses had lower testicular expression of SPP1 mRNA than the CTMLW fetuses at GD90. Considering the localisation of this protein in the testes to the germ cells within seminiferous tubules, supporting cells in the seminiferous tubules and the interstitial Leydig cells of new-born boar testes [34], and its vital role in the regulation of ECM structure, further assessment of the role of integrins and their ligands may provide interesting insights into the mechanisms regulating the impaired reproductive phenotype observed in IUGR boars postnatally.

Sex determination occurs early in embryonic development in mammalian species, whereby the bipotential gonad differentiates into either a testis or an ovary depending upon the sex chromosomes present in the embryo [35]. In the male embryo, the Y-linked gene SRY is critical for the development of the testes, with SRY expression in pre-Sertoli cells inducing upregulation of SOX9, which then induces the expression of many testes determining genes, including DMRT1. DMRT1 is a highly conserved gene which is expressed in greater abundance in the testis compared to the ovary in the embryo [36, 37]. At GD90, the expression of DMRT1 mRNA in testes was lower in the lightest compared to the CTMLW fetuses. Whilst it is well-established that DMRT1 has an important role in early testes development, it is also essential for the maintenance of Sertoli cells, both in fetal and postnatal life, and thus ultimately it is important for the maintenance of testicular function [38]. The finding of decreased DMRT1 mRNA expression at GD90 in the testes of the lightest fetuses compared to the CTMLW fetuses warrants further investigation. Reduced expression of DMRT1 by the testes from the lightest fetuses may be indicative of less maintenance of the Sertoli cell population, which may in turn lead to decreased Sertoli cell number in postnatal life. As Sertoli cells play an essential role in the regulation of spermatogenesis, and given the reported decrease in spermatocytes, round spermatids, elongated spermatids, and lower semen volume and number of spermatozoa per ejaculate in IUGR boars [7, 9], reduced DMRT1 signalling could in this way lead to the impaired reproductive phenotype observed postnatally in these animals. In IUGR rats, Taqman arrays have revealed alterations in genes associated with Sertoli cell and Leydig cell function during early postnatal life [39]. Further, RNASeq experiments comparing testicular RNA expression between Meishan and Duroc boars at postnatal days 20, 75, and 270 identified many genes associated with attainment of puberty and spermatogenesis [40]. Further investigation of some of the candidate genes validated in these studies could provide interesting insights into the mechanisms governing altered postnatal reproductive performance in IUGR boars.

Conclusions

This study provides novel insights into the temporal histological and transcriptomic changes that occur during normal fetal testicular development. Based on morphological and gene expression analyses, our results suggest that the developmental trajectory of IUGR testes does not deviate from that of normally sized fetuses until late in gestation. This study suggests that the expression of SPP1 and DMRT1 mRNAs is lower in testes from the lightest fetuses at GD90, providing a platform for further mechanistic investigations. Given the key role of SPP1 in the regulation of the extracellular matrix, and of DMRT1 in sex determination and the maintenance of Sertoli cells, further studies are warranted to elucidate their involvement in the impaired reproductive potential of IUGR boars. Improving the understanding of the mechanisms regulating fetal testes development is critical to improve the understanding of the regulation of postnatal boar fertility and is vital to improve the success of the pig industry.

Availability of data and materials

The authors confirm that the data supporting the findings of this study are available within the article and its supplementary materials.

Abbreviations

CTMLW:

Closest to mean litter weight

ECM:

Extracellular matrix

ELISA:

Enzyme-linked immunoassay

GD:

Gestational day

IUGR:

Intrauterine growth restricted

PBS:

Phosphate buffered saline

PCR:

Polymerase chain reaction

qPCR:

Quantitative polymerase chain reaction

RINe:

RNA integrity number equivalent

SEM:

Standard error of the mean

References

  1. Knox RV. Artificial insemination in pigs today. Theriogenology. 2016;85(1):83–93. https://doi.org/10.1016/j.theriogenology.2015.07.009.

    Article  CAS  PubMed  Google Scholar 

  2. Knox RV. Impact of swine reproductive technologies on pig and global food production. Adv Exp Med Biol. 2014;752:131–60. https://doi.org/10.1007/978-1-4614-8887-3_7.

    Article  PubMed  Google Scholar 

  3. Mäkelä JA, Koskenniemi JJ, Virtanen HE, Toppari J. Testis Development. Endocr Rev. 2019;40(4):857–905. https://doi.org/10.1210/er.2018-00140.

    Article  PubMed  Google Scholar 

  4. Svingen T, Koopman P. Building the mammalian testis: origins, differentiation, and assembly of the component cell populations. Genes Dev. 2013;27(22):2409–26. https://doi.org/10.1101/gad.228080.113.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. Cargnelutti F, Di Nisio A, Pallotti F, Sabovic I, Spaziani M, Tarsitano MG, et al. Effects of endocrine disruptors on fetal testis development, male puberty, and transition age. Endocrine. 2021;72(2):358–74. https://doi.org/10.1007/s12020-020-02436-9.

    Article  CAS  PubMed  Google Scholar 

  6. Skakkebæk NE, Rajpert-De Meyts E, Main KM. Testicular dysgenesis syndrome: an increasingly common developmental disorder with environmental aspects. Hum Reprod. 2001;16(5):972–8. https://doi.org/10.1093/humrep/16.5.972.

    Article  PubMed  Google Scholar 

  7. Auler PA, Moreira GHFA, Hogg CO, Ashworth CJ, Bortolozzo FP, Chiarini-Garcia H, et al. Testicular parameters and spermatogenesis in different birthweight boars. Reprod Fertil Dev. 2017;29(9):1720–8. https://doi.org/10.1071/RD16164.

    Article  CAS  PubMed  Google Scholar 

  8. Smit MN, Spencer JD, Almeida FRCL, Patterson JL, Chiarini-Garcia H, Dyck MK, et al. Consequences of a low litter birth weight phenotype for postnatal lean growth performance and neonatal testicular morphology in the pig. Animal. 2013;7(10):1681–9. https://doi.org/10.1017/S1751731113001249.

    Article  CAS  PubMed  Google Scholar 

  9. Lin Y, Cheng X, Sutovsky P, Wu D, Che LQ, Fang ZF, et al. Effect of intra-uterine growth restriction on long-term fertility in boars. Reprod Fertil Dev. 2017;29(2):374–82. https://doi.org/10.1071/RD15130.

    Article  PubMed  Google Scholar 

  10. Bee G. Birth weight of litters as a source of variation in postnatal growth, and carcass and meat quality. Advances in Pork Production. 2007;18:191–6.

    Google Scholar 

  11. Town SC, Putman CT, Turchinsky NJ, Dixon WT, Foxcroft GR. Number of conceptuses in utero affects porcine fetal muscle development. Reproduction. 2004;128(4):443–54. https://doi.org/10.1530/rep.1.00069.

    Article  CAS  PubMed  Google Scholar 

  12. Wu G, Bazer FW, Wallace JM, Spencer TE. Board-invited review: intrauterine growth retardation: implications for the animal sciences. J Anim Sci. 2006;84(9):2316–37. https://doi.org/10.2527/jas.2006-156.

    Article  CAS  PubMed  Google Scholar 

  13. Wu G, Bazer FW, Burghardt RC, Johnson GA, Kim SW, Li XL, et al. Impacts of amino acid nutrition on pregnancy outcome in pigs: mechanisms and implications for swine production. J Anim Sci. 2010;88(suppl_13):E195–204. https://doi.org/10.2527/jas.2009-2446.

    Article  CAS  PubMed  Google Scholar 

  14. Finch AM, Anitpatis C, Pickard AR, Ashworth CJ. Patterns of fetal growth within large white X landrace and Chinese Meishan gilt litters at three stages of gestation. Reprod Fertil Dev. 2002;14(7):419–25. https://doi.org/10.1071/RD01131.

    Article  CAS  PubMed  Google Scholar 

  15. Foxcroft GR, Dixon WT, Novak S, Putman CT, Town SC, Vinsky MDA. The biological basis for prenatal programming of postnatal performance in pigs. J Anim Sci. 2006;84(suppl_13):E105–12. https://doi.org/10.2527/2006.8413_supplE105x.

    Article  PubMed  Google Scholar 

  16. Pettigrew J, Cornelius S, Moser R, Heeg T, Hanke H, Miller K, et al. Effects of oral doses of corn oil and other factors on preweaning survival and growth of piglets. J Anim Sci. 1986;62(3):601–12. https://doi.org/10.2527/jas1986.623601x.

    Article  CAS  PubMed  Google Scholar 

  17. Wise T, Roberts AJ, Christenson RK. Relationships of light and heavy fetuses to uterine position, lacental weight, gestational age, and fetal cholesterol concentrations. J Anim Sci. 1997;75(8):2197–207. https://doi.org/10.2527/1997.7582197x.

    Article  CAS  PubMed  Google Scholar 

  18. Foxcroft GR, Town SC. Prenatal programming of postnatal performance – the unseen cause of variance. Adv Pork Prod. 2004;15:4–10.

    Google Scholar 

  19. Ashworth CJ, Finch AM, Page KR, Nwagwu MO, McArdle HJ. Causes and consequences of fetal growth retardation in pigs. Reprod Suppl. 2001;58:233–46.

    CAS  PubMed  Google Scholar 

  20. Pontelo TP, Miranda JR, Felix MAR, Pereira BA, da Silva WE, Avelar GF, et al. Histological characteristics of the gonads of pig fetuses and their relationship with fetal anatomical measurements. Res Vet Sci. 2018;117:28–36. https://doi.org/10.1016/j.rvsc.2017.11.005.

    Article  PubMed  Google Scholar 

  21. Stenhouse C, Tennant P, Duncan WC, Ashworth CJ. Doppler ultrasound can be used to monitor umbilical arterial blood flow in lightly sedated pigs at multiple gestational ages. Reprod Fertil Dev. 2018;30(11):1402–11. https://doi.org/10.1071/RD17298.

    Article  PubMed  Google Scholar 

  22. Stenhouse C, Hogg CO, Ashworth CJ. Association of foetal size and sex with porcine feto-maternal interface integrin expression. Reproduction. 2018;157:317–28.

    Article  Google Scholar 

  23. Stenhouse C, Hogg CO, Ashworth CJ. Associations between fetal size, sex and both proliferation and apoptosis at the porcine feto-maternal interface. Placenta. 2018;70:15–24. https://doi.org/10.1016/j.placenta.2018.08.006.

    Article  PubMed  PubMed Central  Google Scholar 

  24. Stenhouse C, Hogg CO, Ashworth CJ. Associations between fetal size, sex and placental angiogenesis in the pig. Biol Reprod. 2018;100(1):239–52. https://doi.org/10.1093/biolre/ioy184.

    Article  PubMed Central  Google Scholar 

  25. Zhao YY, Wang W, Li XX, Chen X, Yu P, Wang JJ, et al. Effect of BMPRIB gene silencing by siRNA on apoptosis and steroidogenesis of porcine granulosa cells. Genet Mol Res. 2014;13(4):9964–75. https://doi.org/10.4238/2014.November.28.1.

    Article  CAS  PubMed  Google Scholar 

  26. Stenhouse C, Cortes-Araya Y, Hogg CO, Donadeu FX, Ashworth CJ. Associations between foetal size and ovarian development in the pig. Anim Reprod Sci. 2020;221:106589.

    Article  CAS  Google Scholar 

  27. Oliver GA, Novak SA, Patterson JLA, Pasternak JAA, Paradis FA. Restricted feed intake in lactating primiparous sows . II . Effects on subsequent litter sex ratio and embryonic gene expression. Reprod Fertil Dev. 2011;23(7):899–911. https://doi.org/10.1071/RD11013.

    Article  CAS  PubMed  Google Scholar 

  28. Hernández SC, Hogg CO, Billon Y, Sanchez MP, Bidanel JP, Haley CS, et al. Secreted phosphoprotein 1 expression in endometrium and placental tissues of hyperprolific large white and meishan gilts. Biol Reprod. 2013;88(5):1–7. https://doi.org/10.1095/biolreprod.112.104679.

    Article  Google Scholar 

  29. Nygard AB, Jørgensen CB, Cirera S, Fredholm M. Selection of reference genes for gene expression studies in pig tissues using SYBR green qPCR. BioMed Cent Mol Biol. 2007;8(1):67. https://doi.org/10.1186/1471-2199-8-67.

    Article  CAS  Google Scholar 

  30. Humphries JD. Integrin ligands at a glance. J Cell Sci. 2006;119(19):3901–3. https://doi.org/10.1242/jcs.03098.

    Article  CAS  PubMed  Google Scholar 

  31. Fisher LW, Torchia DA, Fohr B, Young MF, Fedarko NS. Flexible structures of SIBLING proteins, bone sialoprotein, and osteopontin. Biochem Biophys Res Commun. 2001;280(2):460–5. https://doi.org/10.1006/bbrc.2000.4146.

    Article  CAS  PubMed  Google Scholar 

  32. Johnson GA, Burghardt RC, Bazer FW, Spencer TE. Osteopontin: roles in implantation and placentation. Biol Reprod. 2003;69(5):1458–71. https://doi.org/10.1095/biolreprod.103.020651.

    Article  CAS  PubMed  Google Scholar 

  33. Johnson GA, Burghardt RC, Bazer FW. Osteopontin: a leading candidate adhesion molecule for implantation in pigs and sheep. J Anim Sci Biotechnol. 2014;5:56. https://doi.org/10.1186/2049-1891-5-56.

  34. Kim S, Shin T. Immunohistochemical study of osteopontin in boar testis. J Vet Sci. 2007;8(2):107–10. https://doi.org/10.4142/jvs.2007.8.2.107.

    Article  PubMed  PubMed Central  Google Scholar 

  35. Wilhelm D, Palmer S, Koopman P. Sex determination and gonadal development in mammals. Physiol Rev. 2007;87(1):1–28. https://doi.org/10.1152/physrev.00009.2006.

    Article  CAS  PubMed  Google Scholar 

  36. Huang S, Ye L, Chen H. Sex determination and maintenance: the role of DMRT1 and FOXL2. Asian J Androl. 2017;19(6):619–24. https://doi.org/10.4103/1008-682X.194420.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Ferguson-Smith M. The evolution of sex chromosomes and sex determination in vertebrates and the key role of DMRT1. Sex Dev. 2006;1(1):2–11. https://doi.org/10.1159/000096234.

    Article  CAS  Google Scholar 

  38. Matson CK, Murphy MW, Sarver AL, Griswold MD, Bardwell VJ, Zarkower D. DMRT1 prevents female reprogramming in the postnatal mammalian testis. Nature. 2011;476(7358):101–4. https://doi.org/10.1038/nature10239.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Pampanini V, Germani D, Puglianiello A, Stukenborg JB, Reda A, Savchuk I, et al. Impact of uteroplacental insufficiency on postnatal rat male gonad. J Endocrinol. 2017;232(2):247–57.

    Article  CAS  Google Scholar 

  40. Ding H, Luo Y, Liu M, Huang J, Xu D. Histological and transcriptome analyses of testes from Duroc and Meishan boars. Sci Rep. 2016;6:20758.

    Article  CAS  Google Scholar 

Download references

Acknowledgements

We thank the staff of The Roslin Institute Large Animal Unit for their skilled assistance.

Funding

The Roslin Institute receives Institute Strategic Grant funding from the BBSRC (BB/J004316/1). CS was in receipt of a studentship from the University of Edinburgh. YC was funded by the National Agency for Research and Development (ANID) / Scholarship Program / DOCTORADO BECAS CHILE/2016–72170349.

Author information

Authors and Affiliations

  1. Functional Genetics and Development Division, The Roslin Institute and Royal (Dick) School of Veterinary Studies, University of Edinburgh, Midlothian, UK

    Claire Stenhouse, Yennifer Cortes-Araya, F. Xavier Donadeu & Cheryl J. Ashworth

  2. Department of Animal Science, Texas A&M University, 440 Kleberg Center, College Station, TX, 77843-2471, USA

    Claire Stenhouse

Authors
  1. Claire Stenhouse
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Yennifer Cortes-Araya
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. F. Xavier Donadeu
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Cheryl J. Ashworth
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

CS, CJA, and FXD devised the experiment. CS and CJA collected the samples. CS and YCA performed the experiments. CS, YCA and CJA analysed the data. CS and CJA prepared the final manuscript.

Corresponding author

Correspondence to Claire Stenhouse.

Ethics declarations

Ethics approval and consent to participate

All procedures were performed with approval from The Roslin Institute (University of Edinburgh) Animal Welfare and Ethical Review Board and in accordance with the U.K. Animals (Scientific Procedures) Act, 1986.

Consent for publication

Not applicable.

Competing interests

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Supplementary Information

Additional file 1.

Supplementary Fig. 1: Representative Images of GATA4 Immunohistochemistry in Fetal Testes. Immunohistochemistry revealed that GATA4 protein is expressed by the somatic cells in both the lightest (B and D) and closest to mean litter weight (CTMLW) (A and C) at both gestational day (GD) 60 (A and B) and 90 (C and D). Rabbit IgG controls at an equivalent protein concentration were utilised as a negative control (E). Scale bars represent 100 μm.

Additional file 2.

Supplementary Table 1: Quantitative polymerase chain reaction calibration curve data.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article's Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article's Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Stenhouse, C., Cortes-Araya, Y., Donadeu, F.X. et al. Associations between testicular development and fetal size in the pig. J Animal Sci Biotechnol 13, 24 (2022). https://doi.org/10.1186/s40104-022-00678-3

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s40104-022-00678-3

Keywords

Journal of Animal Science and Biotechnology

ISSN: 2049-1891