Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2023 Feb 1.
Published in final edited form as: Anim Reprod Sci. 2022 Jan 19;237:106927. doi: 10.1016/j.anireprosci.2022.106927

Effects of lipopolysaccharide on follicular estrogen production and developmental competence in bovine oocytes

KK Forrest a, VV Flores a, SC Gurule a, S Soto-Navarro a, CB Shuster b, CA Gifford c, JA Hernandez Gifford a,*
PMCID: PMC8928215  NIHMSID: NIHMS1773853  PMID: 35074697

Abstract

Reproductive efficiency and female fertility is essential for productive and sustainable beef cattle operations. Gram-negative bacterial infections cause release of the endotoxin lipopolysaccharide (LPS) which initiates immune responses shown to alter ovarian steroidogenesis and impair oocyte development. The current study was designed to investigate the impact of varying levels of naturally occurring infection and follicular LPS on estradiol (E2) production and oocyte maturation. Bovine ovary pairs were harvested from a slaughterhouse, and oocytes were aspirated from small follicles and matured in vitro. Meiotic events were evaluated on nuclear maturation and spindle morphology to classify oocytes as normal or abnormal. Follicular fluid LPS concentrations were measured and subsequently separated into Low or High LPS groups. A marked difference was detected between the percent of abnormal oocytes matured from Low LPS follicles, compared to the percent of abnormal oocytes matured from High LPS follicles (P = 0.1). Follicular E2 concentrations tended to be greater for high LPS follicles (P = 0.1), however, relative abundance of mRNA transcripts for aromatase (P = 0.93) and beta-catenin (P = 0.63) were similar between groups. No changes were detected in Toll-like Receptor 4 (P = 0.15), Myeloid Differentiation Factor-2 (P = 0.61), or cluster of differentiation 14 (P = 0.46) mRNA transcript abundance in follicles with high LPS, compared to low. Therefore, even Low levels of follicular LPS indicating a subacute infection is capable of impacting the ovarian milieu and may represent an unappreciated factor leading to reduced female fertility and decreased cow retention.

Keywords: Disease, Estradiol, Fertility, Follicle, Meiosis, Oocyte

1. Introduction

Impaired female fertility is detrimental to cow-calf operations who heavily depend on reproductive success to drive productivity and profitability. The primary reason females are culled from the herd is related to reproductive failure. Understanding the mechanism contributing to impaired fertility may improve herd productivity by minimizing the number of female replacements. This allows producers to retain a greater quantity of productive cows, reducing development costs associated with replacement heifers and increase profitability of their operation (Roberts et al., 2015).

Fertilization and ultimately birth of an offspring, is reliant on appropriate development of the oocyte inside an ovarian follicle. The unique follicular microenvironment surrounding the oocyte is essential for its developmental competence. The oocytes ability to undergo coordinated nuclear and cytoplasmic maturation, determines its potential to become fertilized and sustain embryonic development through parturition (Sirard et al., 2006). Optimal female fertility is dependent upon circulating estradiol (E2) concentrations produced by ovarian granulosa cells that contribute to numerous physiological events including female sexual receptivity (Simpson et al., 2005), folliculogenesis (Drummond and Findlay, 1999), granulosa cell proliferation and ovulation (Emmen et al., 2005), meiotic progression (Liu et al., 2017), oocyte competency and embryonic development (Wu et al., 1992). Biosynthesis of E2 is a tightly regulated molecular process dependent on expression of key steroidogenic enzymes by follicle-stimulating hormone (FSH) and intra-ovarian signaling molecules including beta-catenin, a co-transcription factor essential for maximal FSH induction of aromatase mRNA transcript (Parakh et al., 2006) and subsequent E2 production (Hernandez Gifford et al., 2009).

Previous studies have shown cattle are susceptible to a wide range of pathologies associated with Gram-negative bacteria that may manifest in subclinical or clinical infection. Clinical infections often present apparent and visible symptoms, whereas subclinical infections appear asymptomatic and therefore, may go unnoticed. Bacterial release of LPS is responsible for inducing an immune response in the infected host, negatively impacting fertility by disrupting ovarian steroidogenesis (Magata et al., 2014) and oocyte competency (Bromfield and Sheldon, 2011; Rincon et al., 2019). Lipopolysaccharide resulting from infections caused by Gram-negative bacteria have been detected in follicular fluid from randomly collected slaughterhouse ovaries, and subsequently demonstrated reduced aromatase transcript abundance and E2 production associated with increased amounts of LPS (Magata et al., 2014). These changes instigated by detectable LPS in the follicular fluid may present negative consequences to fertility, supporting reproductive failure as a leading cause of cow culling.

Granulosa cells challenged in vivo and in vitro with known amounts of LPS demonstrated diminished estradiol production (Herath et al., 2007; Guan et al., 2021). However, whether or not LPS accumulation at concentrations found innately in follicular fluid of cattle disrupts follicular E2 synthesis and impacts oocyte competency remains to be determined. Therefore, the experiments presented herein were aimed at investigating the impact of naturally occurring LPS accumulation in ovarian follicles collected from animals with no obvious clinical signs of infection on follicular estrogen biosynthesis, including consequences on the developing oocyte. We hypothesize varying concentrations of LPS present in follicular fluid collected from slaughterhouse ovaries may impact the beta-catenin, aromatase signaling axis resulting in a reduction of follicular estradiol concentrations, subsequently impairing oocyte competency.

2. Materials and methods

2.1. Sample collection

Ovary pairs (n = 19) of mixed-breed non-pregnant cows and heifers were collected from a slaughterhouse immediately following time of harvest. Ovaries were washed twice with 0.9% saline, once with 70% ethanol, and stored in 0.9% saline containing 1% penicillin-streptomycin (10,000 U/mL; Gibco) and transported back to the laboratory on ice (~6 h). All animals were considered healthy as the reproductive tracts had no gross evidence of infection. Cumulus-oocyte complexes (COC, n = 74), granulosa cells, and follicular fluid from small developing follicles (2 to 9 mm) were manually aspirated using a 5cc pyrogen-free syringe with an 18-G needle and pooled for each ovary pair. Follicular fluid supernatant was removed and stored at −20°C for future radioimmunoassay (RIA) and lipopolysaccharide (LPS) endotoxin assay analysis. Remaining granulosa cells were washed with phosphate-buffered saline (PBS), resuspended in TRIzol reagent (Invitrogen), and stored at −80°C until isolation of RNA as described below.

2.2. In vitro maturation of oocytes

A temperature of 38.5°C was maintained throughout COC collection and washes using a benchtop slide warmer (Premiere, XH-2002). Oocytes surrounded by a minimum of three compact cumulus cell layers were selected for maturation. Oocyte maturation media were prepared as described by Rivera and Hansen (2001) with slight modifications. Briefly, after the last wash, COC were placed in a pre-equilibrated multi-well culture plate (Sigma-Aldrich), previously prepared 3 to 6 h prior to maturation with Medium 199 (Sigma-Aldrich) oocyte maturation media. Additional supplements including gentamicin (5 mg/mL; Sigma-Aldrich), 10% FBS, folltropin (20 mg/mL; Vetoquinol; USA, Inc.), β-estradiol (1 mg/mL; Sigma-Aldrich) and sodium pyruvate (Sigma-Aldrich) were added to oocyte-maturation media. Oocytes were matured at 38.5°C, 5% CO2 for 21 h.

2.3. Denuding and fixation of oocytes

Following 21 h maturation, COC were washed in PBS for 10 min and incubated for 5 min in a denuding stock including hyaluronidase (1 mg/mL; Sigma-Aldrich) in pre-warmed HEPES with Tyrode’s Albumin Lactate Pyruvate (HEPES-TALP; Cassion Laboratories). Oocytes were vortexed vigorously for 4 min to ensure removal of tightly adhered cumulus cells. Denuded oocytes were fixed at room temperature in 3.7% paraformaldehyde (Fisher Scientific) in PBS for 30 min and permeabilized in phosphate-buffered saline with 0.1% Triton-X 100 (PBST, Sigma-Aldrich) overnight at 4°C. Oocytes were subsequently blocked overnight in 3% bovine serum albumin (BSA, Fisher Scientific) in PBS at 4°C.

2.4. Immunofluorescence

Oocytes were removed from blocking solution and incubated overnight with a primary antibody, rat anti-α tubulin (YL1/2) (1:1000, Santa Cruz Biotechnology) in 3% BSA at 4°C. Oocytes were washed three consecutive times in PBS for 10 min and stored overnight at 4°C in Alexa Flour-labeled secondary antibodies diluted in 3% BSA. Secondary antibodies included: 488 donkey anti-rat (1:1000, Life Technologies), 568 Phalloidin (Thermo Fisher Scientific) to visualize filamentous actin, and Hoescht (1:10,000, Invitrogen) to visualize DNA. Oocytes were mounted over 40μl of 90 % glycerol (Sigma-Aldrich) in PBS and stored at 4°C until imaging.

2.5. Image acquisition and analysis

Fixed oocytes were evaluated for significant meiotic events characterized by nuclear maturation and spindle arrangement using an Andor Dragonfly 550 spinning disc confocal microscope mounted on an Olympus IX83 inverted microscope, and a 60x N.A. 1.3 silicone objective. Z-stacks images of individual oocytes were acquired using an iXon 888 EMCCD camera driven by Fusion Software. Bitplane Imaris (version 8.1–9.1.2) software was used to generate three-dimensional renderings from the acquired image stacks. Montages were prepared in Adobe Photoshop (CS). Oocytes were categorized through evaluation of chromatin condensation and orientation, spindle bipolarity, and presence of a polar body. Only oocytes with a bipolar spindle, condensed chromatids aligned along the metaphase plate, and presence of a polar body were classified as normal. Oocytes also demonstrating proper nuclear maturation but which were removed from maturation before arresting in metaphase II were also considered normal. All remaining oocytes signifying irregular developmental characteristics were classified abnormal. Three different experienced observers evaluated oocytes without reference to previously collected data.

2.6. Quantitation of lipopolysaccharide in follicular fluid

Concentrations of LPS in follicular fluid collected from bovine ovaries were measured using the Pierce LAL Chromogenic Endotoxin Quantitation Assay Kit following the manufacturer’s instructions (88282; Thermo Fisher Scientific) including minor modifications. Samples were thawed at room temperature, vortexed vigorously using a bench top vortexer (Standard Vortex Mixer 120V; Fisher Scientific) for 3 min and diluted 1:10 in PBS (Gibco). To eliminate interference of the limulus amebocyte lysate (LAL) assay by endogenous factors, samples were heat-treated in a water bath (Isotemp Heating Block, Fisher Scientific) at 75°C for 30 min following as previously described (Herath et al., 2007).

Samples were incubated in duplicate with the LAL substrate over thermal beads set at 37°C ± 1°C, using 96-well endotoxin-free microplates (Endosafe, Charles River Laboratories Inc.). A standard curve was prepared using endotoxin-free water with concentrations of 0.1, 0.25, 0.5, and 1 endotoxin units (EU)/mL. Internal recovery of assay as determined using positively spiked samples (0.5 EU/mL) was > 80%, and the limit of detection was 0.1 EU/mL or approximately 0.01 ng/mL. Immediately following assay completion, the optical density of 405 nm was read for samples at 37°C ± 1°C and plotted against the standard curve to quantify existing follicular endotoxin concentration. The intra- and inter-assay coefficient of variation were 4.0 and 5.5%, respectively. A follicular fluid LPS concentration below 1.6 EU/mL were categorized as ‘Low LPS’ (1.32 ± 0.22 EU/mL, n = 15), while follicular fluid LPS concentrations greater than 1.6 EU/mL were categorized as ‘High LPS’ (1.98 ± 0.20 EU/mL, n = 4)

2.7. Semi-quantitative endpoint PCR

Total RNA was isolated from pooled granulosa cells of small developing follicles using TRIzol reagent according to the manufacturer’s protocol and quantitated using a NanoDrop spectrophotometer (ND-2000 Spectrophotometer; Thermo Fisher Scientific). Following DNase treatment (Invitrogen), total RNA (1 μg) was reverse transcribed into cDNA using oligo (dT) primers and Superscript II Reverse Transcriptase (Invitrogen). Prior to analysis of mRNA transcript abundance, cDNA was diluted 1:10 with nuclease-free water. Gene-specific primer sequences are listed in Table 1. Primer sets were optimized for appropriate temperatures and a linear signal range was determined for each gene. The optimal number of cycles determined for each primer set was identified so that the amplification product was clearly visible on the agarose gel and could be quantified. The optimal number of cycles for each gene was identified from 22 to 40 cycles to ensure the amplification was in the exponential range and a plateau had not been reached. Polymerase chain reaction was performed using 2μl of 1:10 cDNA and resolved by gel electrophoresis on either a 1% or 2.5% agarose gel containing ethidium bromide (Fisher Bioreagents). Images were collected using a ChemiDoc XRST imager (Bio-Rad Laboratories). Fluorescent intensity of bands was analyzed using Image Studio software (Bio-Rad Laboratories) and determined in relation to the PPIA control. Total RNA was isolated from bovine peripheral blood leukocytes (PBL) collected from whole blood, reverse transcribed into cDNA similar to the method explained above, to generate an external control to ensure optimal PCR conditions for genes encompassing the LPS-signaling receptor complex.

Table 1.

Primer sequences used in semi-quantitative endpoint PCR.

Sequences of primer (5’- 3’)
Gene Accession No. Forward Reverse
PPIA 1 BC105173.1 GAGCTTCCCTTGCCGTTTAG TGTCCACAGTCAGCAATGGT
CTNNB1 2 NM_001076141.1 CAAGATGATGGTGTGCCAAG CTGCACAAACAATGGAATGG
CYP19A1 3 NM_174305.1 TGCATGGCAAGCTCTCCTTCTCAAACCA TGACCAGGTCCACAACGGGCTGG
TLR4 4 NM_174198.6 CTTGCGTACAGGTTGTTCCTAA CTGGGAAGCTGGAGAAGTTATG
MD2 5 DQ319076.1 GGGAAGCCGTGGAATACTCTAT CCCCTGAAGGAGAATTGTATTG
CD14 6 NM_174008.1 TGCCGTCGAGGTGGAGATCA ATTGGAGGGCCGGGAACTTG
Open in a new tab
1

PPIA = cyclophilin A

2

CTNNB1 = beta-catenin

3

CYP19A1 = aromatase

4

TLR4 = toll-like receptor 4

5

MD2 = myeloid differentiation factor 2

6

CD14 = cluster of differentiation 14

2.8. Radioimmunoassay

Follicular fluid samples were analyzed for E2 concentrations by solid-phase RIA using a commercial kit manufactured by MP Biomedicals (Santa Ana, CA) using methods described by (Castañon et al., 2012). The assay was run using polypropylene tubes coated with an antibody against E2, and a radiolabeled E2 (125I-E2) was used as the tracer. Two stock standard solutions were prepared by suspending E2 (Sigma-Aldrich) at 100 pg/mL and 1 ng/mL in assay buffer (0.01 M PBS plus 0.1% gelatin, pH 7.0), and were pipetted into the tubes coated with antibody to generate a standard curve of 0, 2.5, 5, 10, 20, 40, 80, 160, and 320 pg of E2 per tube. Follicular fluid (25μl) were diluted 1:100 in PBS and assayed in duplicate. Each tube received 1 mL of tracer, after which tubes were vortexed and incubated at room temperature for approximately 24 h. Tubes were decanted and counted for 1 min in a Packard Cobra II auto-gamma counter (Packard Instrument Co. Inc., Downers Grove, IL, USA). The specific binding was 78%. Detection limit (95% of maximum binding) of the assay was 2 pg/mL. Follicular fluid E2 had an intra-assay CV of 2.4%.

2.9. Statistical analyses

All statistical analysis was performed using SAS (Version 9.3; SAS Institute, Inc., Cary, NC). The main effects of LPS concentration (Low and High) on follicular fluid E2 concentrations and gene abundance were analyzed using the generalized linear model (GLM) procedure in SAS for a complete randomized design. Comparison between oocyte morphology (normal and abnormal) categorized in High or Low LPS was determined as a percent of oocytes for each animal and analyzed using the GLM procedure of SAS. The correlation between follicular fluid LPS concentration and follicular E2 concentrations, granulosa cell gene abundance of CTNNB1, CYP19A1, CD14, MD2 and TLR4 were estimated using the PROC REG procedure of SAS. P-values ≤ 0.05 were considered statistically significant, P-values > 0.05 and <0.1 reflect trends.

3. Results

3.1. Lipopolysaccharide and estradiol concentration in follicular fluid

Lipopolysaccharide was detected in all pools of follicular fluid aspirated from bovine ovaries collected from a slaughterhouse. The concentration of LPS measured in the follicular fluid of developing follicles ranged from 0.92 to 2.31 EU/mL (1.5 ± 0.4 EU/mL) and were separated into Low and High levels of LPS based on a clear separation of the LPS concentrations at 1.6 EU/mL. Follicular E2 concentrations tended to be greater in in follicles with High LPS compared to Low LPS follicles in the present study (P = 0.1; Fig. 1A). This was further demonstrated when evaluating the association between LPS concentration and E2 (Fig. 1B).

Fig. 1.

Fig. 1.

Open in a new tab

Estradiol (E2) concentrations measured in pooled follicular fluid of developing follicles aspirated from bovine ovaries. A. Concentrations of E2 were quantified by radioimmunoassay. Follicular LPS concentrations of less than 1.6 EU/mL were categorized as ‘Low LPS’ (n = 15) and follicular LPS concentrations greater than 1.6 EU/mL were categorized as ‘High LPS’ (n = 4). High LPS follicles tended to have greater follicular E2 concentrations compared to follicles with Low LPS (P = 0.1). B. Correlation between follicular LPS concentrations and follicular E2 concentrations of all follicles (n = 19).

3.2. Granulosa cells express lipopolysaccharide receptor signaling complex

No differences were quantified for the amount of mRNA transcribed for aromatase (P = 0.93) and beta-catenin (P = 0.63) between follicles with Low and High LPS (Fig. 2). Similarly, granulosa cells isolated from follicles with High LPS demonstrated no differences in TLR4 (P = 0.15), MD2 (P = 0.61) or CD14 (P = 0.46) mRNA transcript abundance, compared to granulosa cells isolated from Low LPS follicles (Fig. 3). Additionally, no demonstrated association between concentrations of LPS and TLR4, MD2, or CD14 was detected.

Fig. 2.

Fig. 2.

Open in a new tab

Granulosa cell driven estradiol synthesis is dependent on key steroidogenic enzymes and intra-ovarian signaling molecules including aromatase and beta-catenin. A. Representative gel demonstrating aromatase and beta-catenin mRNA expression in bovine granulosa cells of follicles of Low LPS (<1.6 EU/mL; n = 15) and High LPS (>1.6 EU/mL; n = 4). Cyclophilin A (PPIA) was used as a loading control. B. Semi-quantitative PCR analysis demonstrates no change in mRNA expression of aromatase (CYP19A1; P = 0.93) and beta-catenin (CTNNB1; P = 0.63) between follicles of Low and High follicular LPS concentrations. C. Correlation between follicular LPS concentrations and CYP19A1 (P = 0.74) and CTNNB1 (P = 0.76) mRNA expression in bovine granulosa cells of all follicles (n = 19).

Fig. 3.

Fig. 3.

Open in a new tab

Bovine granulosa cells express the receptor complex capable of responding to LPS. A. Representative gel demonstrating toll-like receptor 4 (TLR4), cluster-differentiation 14 (CD14) and myeloid differentiation factor-2 (MD2) mRNA expression in granulosa cells of follicles of Low LPS (<1.6 EU/mL; n = 15) and High (>1.6 EU/mL; n = 4). Cyclophilin A (PPIA) was used as a loading control. B. Semi-quantitative PCR analysis demonstrated no change in TLR4 (P = 0.15), MD2 (P = 0.61) and CD14 mRNA expression between follicles with Low and High LPS. C. Correlation between follicular LPS concentrations and TLR4 (P = 0.83), MD2 (P = 0.61), and CD14 (P = 0.46) mRNA expression in bovine granulosa cells of all follicles (n = 19).

3.3. Lipopolysaccharide and in vitro maturation of bovine oocytes

To examine potential effects of LPS on oocyte maturation, oocytes were matured in vitro, and morphological characteristics were assessed for 74 oocytes by confocal microscopy (Fig. 4). Fully mature oocytes could be observed with a bipolar barrel-shaped spindle, bioriented chromosomes aligned along the metaphase plate, and a first polar body, and were classified as normal (Fig. 5A), however, by 21 hours in maturation media, only 24% of scored oocytes had reached metaphase arrest in meiosis II. Given the observed delay in maturation, oocytes that displayed normal meiosis I spindle formation, orientation and chromosome alignment but had not matured completely to meiosis II were also considered normal (Fig. 5B). Oocytes classified as abnormal demonstrated significantly perturbed meiotic structures contributing to meiotic failure including failed GVBD (Fig. 6A), aberrant spindle formation, mis-aligned chromosomes, parthenogenic activation (Fig. 6B), or failed polar body extrusion (Fig. 6C). High LPS follicles demonstrated a tendency for increased percentage of abnormal oocytes matured compared to the Low LPS follicles (P = 0.1, Fig. 4), suggesting a greater percent of competent oocytes were matured from follicles with Low follicular LPS concentrations.

Fig. 4.

Fig. 4.

Open in a new tab

Morphology of bovine oocytes was assessed in 74 oocytes visualized by confocal microscopy to determine meiotic competency. Oocytes metaphase or anaphase of MI were judged to be normal if the spindle was fully formed and correctly oriented relative to the cortex, and chromosomes are correctly aligned along the metaphase plate. Bovine oocytes were categorized within Low LPS (<1.6 EU/mL; n = 63) and High LPS (>1.6 EU/mL; n = 11). For each category High or Low, abnormal oocyte percentages were calculated for each animal. High LPS follicles had an increase in abnormal oocyte morphology (P = 0.1) compared to Low LPS animals. Data are presented as LSM ± S.E.M.

Fig. 5.

Fig. 5.

Open in a new tab

Representative oocytes in Meiosis I and II. Oocytes were collected and cultured in maturation media 21 h before being denuded, fixed, and probed for tubulin (cyan), F-actin (red) and DNA (white). Confocal microscopy was used to acquire 3D renderings. A. Representative of a mature oocyte in metaphase of meiosis II. The chromatin is aligned along the metaphase plate, oriented by a meiotic bipolar shape spindle and the presences of a polar body. B. Representative of metaphase of meiosis I. The chromatin is aligned along the metaphase plate, oriented by a meiotic bipolar shaped spindle. Scale bar = 20μm.

Fig. 6.

Fig. 6.

Open in a new tab

Representative images demonstrating abnormal morphology of bovine oocytes. Oocytes were collected and cultured in maturation media 21 h before being denuded, fixed, and probed for tubulin (cyan), F-actin (red) and DNA (white). Confocal microscopy was used to acquire 3D renderings. A. Representative of an oocyte that failed to undergo germinal vesicle breakdown (GVBD) and arrested in the germinal vesicle (GV) or most immature stage of maturation. B. Representative of an oocyte that extruded all chromatin resulting in two polar bodies. C. Representative of an oocyte considered a parthenote that failed polar body extrusion in meiosis I, resulting in formation of two anaphase meiotic II spindles. Scale bar = 20μm.

4. Discussion

Bacterial LPS released into circulation can be detected at tissues distant from the site of infection as demonstrated by detection of LPS in plasma (Mateus et al., 2003) and follicular fluid (Herath et al., 2007) of cows experiencing metritis, and plasma and milk of cows with mastitis (Hakogi et al., 1989). All follicles collected from slaughterhouse ovaries in the present study had measurable amounts of LPS. This is consistent with a previous report that also collected a similar amount of ovaries and aspirated follicles collected from a slaughterhouse (Magata et al., 2014). Lipopolysaccharide concentrations measured in the follicular fluid in the present study were varied only slightly from those reported by Magata et al. (2014). The slight differences in the endotoxin assay kits utilized in each study may provide an explanation to variation in the LPS ranges detected (Bidne et al., 2018). The discrepancy in measurable LPS may also be attributed to the stage of follicular development, as the follicles measured in the current study were developing follicles 2 to 9 mm, whereas, the follicles evaluated by Magata and colleagues were greater than 8 mm (Magata et al., 2014). Nevertheless, both studies demonstrate the ability to detect varying concentrations of LPS in follicular fluid collected randomly from females which are sent to a slaughterhouse. However, the follicular fluid LPS concentrations collected from the slaughterhouse ovaries in these studies are markedly less than the reported LPS concentrations for dairy cattle classified as having subclinical endometritis (Herath et al., 2007; Piersanti et al., 2019). One explanation which may account for this dramatic difference is that although the animals were considered subclinical, they did present with mild to moderate inflammation of the uterus (Herath et al., 2007) or presence of neutrophils in endometrial cytology (Piersanti et al., 2019). Therefore, since LPS concentrations were evaluated at the time when subclinical disease was evident it is likely that the detectable LPS is more abundant. Conversely, it is not possible to know the time point at which a bacterial insult resulting in LPS accumulation may have occurred in the current study and in the study by Magata et al. (2014). Another difference between the studies likely contributing to the discrepancy is the stage of production of the animals. Ovaries collected from the slaughterhouse in the current study would likely be distinct from postpartum ovaries evaluated studies by Herath et al. (2007) and Piersanti et al. (2019).

Pathogenic Gram-negative bacteria release of the endotoxin LPS are responsible for diseases affecting cattle. Most notably, Gram-negative Escherichia coli is the principal pathogen causing metritis and mastitis in dairy cattle (Hertl et al., 2010; Sheldon et al., 2010). Likewise, bovine respiratory disease in beef cattle is attributed at least in part to Gram-negative bacterial pathogens (Duff and Galyean, 2007). Infections may present as clinical cases with apparent symptoms or as subclinical cases, often appearing asymptomatic and causing them to go unnoticed. Given the ability to detect LPS in the follicular fluid of ovaries in the present study, it is tempting to speculate that these females experienced some type of Gram-negative bacterial infection and mounted an immune response that may have attributed to the reason they were culled. However, since evaluations of clinical diseases were not assessed there is no way to be certain of the origin of LPS in the follicular fluid.

The lower rate of meiotic competence observed may indicate that a direct effect of LPS on the oocyte is underlying the failure of oocyte maturation. Soto et al. (2003) demonstrated that oocytes matured in 1 ng/mL LPS or higher reduced the percentage of oocytes that developed to the blastocyst stage at day 8 after fertilization. However, when LPS was added after fertilization, there was no noted reduction in development to the blastocyst stage. Although a relatively high concentration of LPS was required to disrupt oocyte maturation in the study by Soto et al. (2003), it is possible that the follicle has an increased sensitivity to the molecule.

A previous study investigating the induction of an acute systemic inflammatory response by intravenous injection of LPS on bovine granulosa cell function, demonstrated no significant differences in the amount of mRNA transcribed for aromatase or subsequent E2 production between LPS-treated cows, when compared to the saline-treated controls (De Campos et al., 2017). The investigators speculated the inability to detect differences may have been associated with the short interval between the LPS injection and sample collection, and suggested increasing the time point of collection to mimic a longer chronic inflammatory response, similar to various infections in cattle that often appear subclinical and asymptomatic. Similarly, no differences were quantified for mRNA transcript abundance of aromatase and beta-catenin between follicles with Low and High LPS in the present study. However, there was a tendency for E2 to be greater in High LPS follicles compared to Low LPS follicles. Likewise, in a study evaluating the impact of low doses of LPS on ovarian E2 signaling, heifers receiving subcutaneous injects of 2 μg/kg BW LPS were found to have increased E2 production in large follicles of LPS-treated heifers when compared to the controls (Ferranti et al., 2021). These data suggest that LPS given at a dose intended to replicate an undetectable disease state may be modulating beta-catenin induction of aromatase transcript abundance allowing for E2 synthesis to increase. Although a tendency for changes were detected in follicular fluid E2 concentrations the present study, extended length of timing of a prior infection could allow for partial ovarian estradiol signaling to return and thus prevent greater detectable changes in E2 concentrations. This idea is supported by findings of Lavon et al. (2011) demonstrating that acute mastitis induced by intramammary injection of LPS resulted in immediate, short term disruption of follicular steroidogenesis but did not result in carryover effects of LPS on follicular steroidogenesis when evaluated 2 weeks after endotoxin exposure. Therefore, despite the absence of these differences, follicular concentrations with High detectable LPS suggests those animals may have experienced a more recent or higher degree of infection. Cows who are not performing reproductively are generally culled from the herd, and follicular LPS accumulation might implicate a reason these females failed reproductively.

Toll-like receptors are a family of cellular receptors that function in detecting and initiating the innate immune defense against pathogens including bacteria and viruses, that are primarily located on immune cells (Takeuchi and Akira, 2010). These specific receptors initiate an inflammatory response to pathogen-associated molecular patterns (PAMPs) (Takeuchi and Akira, 2010). Gram-negative derived LPS is a PAMP that specifically binds TLR4, initiating a complex with the co-receptors CD14 and MD2 (Takeuchi and Akira, 2010). The cell-mediated response initiated by LPS binding to its receptor complex initiates recruitment of a cytoplasmic protein MYD88, resulting in activation of MYD88 dependent and independent pathways (Kawai et al., 2001; Laird et al., 2009). Activation of these signaling pathways initiates numerous intracellular responses, including activation of the PI3K/AKT pathway, responsible for production of nuclear factor kappa B (NF-κB) (Lee et al., 2003). Recent research has implicated cross-communication between the NF-κB signaling pathway with the Wingless-Type Mammary Tumor Virus Integration-Site (WNT)/beta-catenin signaling axis during inflammation in a variety of cell types (Ma and Hottiger, 2016). Numerous studies have investigated the cross-regulation of these signaling pathways by the impact of NF-κB on regulation of target genes that affect the function or stability of beta-catenin (Ma and Hottiger, 2016). This may be a possible mechanism by which LPS signaling is modulating ovarian E2 signaling in follicular granulosa cells.

Ovarian follicles lack the presence of immune cells, however, bovine granulosa cells have been shown to modulate the inflammatory response induced by LPS signaling in vitro and in vivo (Bromfield and Sheldon, 2011; De Campos et al., 2017). This is demonstrated throughout different stages of follicular growth, as granulosa cells express the receptor complex, TLR4, CD14, and MD2 cumulatively required for LPS recognition and its subsequent inflammatory response (Herath et al., 2007). In the present study, the entire LPS recognition receptor complex was detected in bovine granulosa cells isolated from follicles 2 to 9 mm. This contradicts a previous experiment using follicles collected from slaughterhouse ovaries 2 to 8 mm, that reported bovine granulosa cells did not express CD14 (Magata and Shimizu, 2017). Granulosa cells demonstrate the capability of mounting an immune response indicated by an increased concentration of pro-inflammatory cytokine production after exposure to LPS (Price et al., 2013). Pro-inflammatory cytokines have also been shown to significantly reduce E2 production in bovine granulosa cells of small follicles in vitro (Spicer and Alpizar, 1994). This inflammatory response to bacterial infection is potentially the mechanism by which LPS accumulation in the follicular environment may perturb follicular steroidogenesis and consequently impact the developing oocyte.

Production of a competent oocyte requires a series of tightly regulated maturational events in preparation for subsequent fertilization and embryonic development that can be disrupted in the event of infection by the presence of LPS accumulation in the follicular environment. Protocols in the literature involving maturation of bovine oocytes includes a common range of 21 to 24 h. A series of preliminary experiments were conducted in the present study allowing a decision for 21 h of maturation time. After reviewing the data in the present study, it was suggested oocytes may have been pulled too early from maturation, represented by several oocytes observed in metaphase of meiosis I. As an artifact of culture time, oocytes demonstrating normal meiotic structures and proper nuclear maturation were classified as normal. Future analysis will require further preliminary studies to conclude the appropriate maturation time. However, all remaining oocytes signifying irregular developmental characteristics or failed GVBD were classified as abnormal.

A study conducted by Shabankareh et al. (2015) correlated developmental competence of bovine oocytes collected from slaughterhouse ovaries following in vitro maturation, to the size of the follicle it was aspirated from. Of the total oocytes collected from small follicles (3 to 5 mm, n = 435), approximately 72% oocytes successfully matured in vitro (Shabankareh et al., 2015). Additionally, oocytes collected from medium follicles (6 to 9 mm; n = 494) demonstrated an in vitro maturation success rate of 81% (Shabankareh et al., 2015). Throughout the literature associated with maturing bovine oocytes, it is commonly reported that an estimated average 80% of oocytes successfully mature to metaphase of meiosis II (Bromfield and Sheldon, 2011; Zhao et al., 2017). This indicates approximately 20% of oocytes matured in vitro do not undergo successful maturation. A previous experiment collected bovine ovaries from a slaughterhouse, aspirated small developing follicles (2 to 8 mm), and treated the oocyte maturation media with 1 or 5 μg/mL of LPS (Rincon et al., 2019). Of the oocytes matured, 63% arrested in metaphase of meiosis II with no additional treatment in the culture media, indicating normal maturation (Rincon et al., 2019). The percent of oocytes that successfully matured in the media supplemented with the low-doses of LPS was significantly reduced to 39% (Rincon et al., 2019).

This was similar to the present study, where 73.9% of the oocytes in the Low LPS follicles successfully matured and were considered normal, whereas, 26.1% were classified as abnormal. Additionally, of the total oocytes matured from follicles with High LPS, 58.2% were classified as abnormal. Oocytes classified as abnormal demonstrated significantly perturbed meiotic structures contributing to meiotic failure including aberrant spindle formation, mis-aligned chromosomes, parthenogenic activation, retained polar body extrusion, or failed GVBD. Similar results were observed in previous experiments when LPS was supplemented during in vitro maturation of bovine oocytes, demonstrated by the increased rate of meiotic failure by delayed maturation progression, reduced polar body extrusion rates and significantly disrupted meiotic structures (Bromfield and Sheldon, 2011; Zhao et al., 2017).

The ovary pairs used in this study were collected from a slaughterhouse with no additional information as to why these animals were culled from the herd. The origin of the oocytes recovered from ovaries obtained from slaughterhouse animals is unknown, suggesting possible variability in oocyte quality. Regardless, the results of the current study demonstrated an ability to detect varying concentrations of LPS in the follicular fluid, suggesting a previous infection and subsequent inflammatory response prior to being removed from the herd. This inflammatory response may be a mechanism by which increased LPS accumulation in the follicular environment may be disrupting follicular E2 biosynthesis and consequently impacting the developing oocyte.

5. Conclusions

The follicular microenvironment surrounding the oocyte is essential for its developmental competence. The granulosa cells ability to express the signaling receptor complex that recognizes LPS, may contribute to the follicle’s susceptibility of LPS accumulation and subsequent inflammatory response. Understanding the cellular mechanism contributing to perturbed follicular steroidogenesis and impaired oocyte competency, may provide insight into the reason fertility is reduced in response to infection. Maximal fertility is dependent on circulating E2 to regulate many aspects of female reproduction, including follicular maturation and production of a competent oocyte. Compromised oocyte maturation prevents successful fertilization and subsequent embryonic development. This may be a contributing factor to open females that are sent to the slaughterhouse, resulting in a loss of productivity and profitability for the producer. This research demonstrates the capacity to measure LPS accumulation in the follicular environment and the potential impacts on oocyte maturation. Additional investigation into this mechanism may help with understanding how long the detrimental impacts of an infection will negatively impact the cow, especially in situations involving subclinical infections that appear asymptomatic and likely go unnoticed.

Highlights.

  • Low levels of detectable endotoxin within developing follicles can impact ovarian function

  • Bovine granulosa cells express signaling receptor complex that recognizes lipopolysaccharide

  • Higher concentrations of follicular LPS has a greater impact on oocyte competency

  • Follicular hormonal milieu was similar between follicles with Low and High lipopolysaccharide

Acknowledgements

Funding: This research was supported by the NM Agric. Exp. Sta. Las Cruces, NM (NMGifford-A19) from the USDA National Institute of Food and Agriculture; an Institutional Development Award (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant number P20GM103451; and National Science Foundation under grant number MCB1917983 to C.B.S. The authors would like to thank Caviness Beef Packers (Hereford, TX) for the generous donation of cattle ovaries.

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Declarations of interest: none.

Competing interest statement

The authors declare that there is no conflict of interest.

References

  1. Bidne K, Dickson M, Ross J, Baumgard L, and Keating A. 2018. Disruption of female reproductive function by endotoxins. Reproduction 155(4):R169–R181. [DOI] [PubMed] [Google Scholar]
  2. Bromfield JJ, and Sheldon IM. 2011. Lipopolysaccharide initiates inflammation in bovine granulosa cells via the TLR4 pathway and perturbs oocyte meiotic progression in vitro. Endocrinology 152(12):5029–5040. doi: 10.1210/en.2011-1124. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Castañon BI, Stapp A, Gifford C, Spicer L, Hallford D, and Gifford JH. 2012. Follicle-stimulating hormone regulation of estradiol production: possible involvement of WNT2 and β-catenin in bovine granulosa cells. Journal of animal science 90(11):3789–3797. [DOI] [PubMed] [Google Scholar]
  4. De Campos FT, Rincon JAA, Acosta DAV, Silveira PAS, Pradieé J, Corrêa MN, Gasperin BG, Pfeifer LFM, Barros CC, and Pegoraro LMC. 2017. The acute effect of intravenous lipopolysaccharide injection on serum and intrafollicular HDL components and gene expression in granulosa cells of the bovine dominant follicle. Theriogenology 89:244–249. doi: 10.1016/j.theriogenology.2016.11.013 [DOI] [PubMed] [Google Scholar]
  5. Duff GC, and Galyean ML. 2007. Board-invited review: recent advances in management of highly stressed, newly received feedlot cattle. Journal of Animal Science 85(3): 823–840. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Ferranti E, Aloqaily B, Gifford C, Forrest K, Löest C, Wenzel J, and Gifford JH. 2021. Effects of lipopolysaccharide on beta-catenin, aromatase, and estrogen production in bovine granulosa cells in vivo and in vitro. Domestic Animal Endocrinology: 106652. [DOI] [PubMed] [Google Scholar]
  7. Guan H-Y, Xia H-X, Chen X-Y, Wang L, Tang Z-J, and Zhang W. 2021. Toll-Like Receptor 4 Inhibits Estradiol Secretion via NF-κB Signaling in Human Granulosa Cells. Frontiers in Endocrinology 12(225)(Original Research) doi: 10.3389/fendo.2021.629554 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Hakogi E, Tamura H, Tanaka S, Kohata A, Shimada Y, and Tabuchi K. 1989. Endotoxin levels in milk and plasma of mastitis-affected cows measured with a chromogenic limulus test. Vet Microbiol. 20(3):267–274. doi: 10.1016/0378-1135(89)90050-3. [DOI] [PubMed] [Google Scholar]
  9. Herath S, Williams EJ, Lilly ST, Gilbert RO, Dobson H, Bryant CE, and Sheldon IM. 2007. Ovarian follicular cells have innate immune capabilities that modulate their endocrine function. Reproduction 134(5):683–693. doi: 10.1530/REP-07-0229. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Hertl J, Gröhn Y, Leach J, Bar D, Bennett G, Gonzalez R, Rauch B, Welcome F, Tauer L, and Schukken Y. 2010. Effects of clinical mastitis caused by gram-positive and gram-negative bacteria and other organisms on the probability of conception in New York State Holstein dairy cows. Journal of dairy science 93(4): 1551–1560. [DOI] [PubMed] [Google Scholar]
  11. Kawai T, Takeuchi O, Fujita T, Inoue J.-i., Mühlradt PF, Sato S, Hoshino K, and Akira S. 2001. Lipopolysaccharide stimulates the MyD88-independent pathway and results in activation of IFN-regulatory factor 3 and the expression of a subset of lipopolysaccharide-inducible genes. J Immunol. 167(10):5887–5894. doi: 10.4049/jimmunol.167.10.5887. [DOI] [PubMed] [Google Scholar]
  12. Laird MH, Rhee SH, Perkins DJ, Medvedev AE, Piao W, Fenton MJ, and Vogel SN. 2009. TLR4/MyD88/PI3K interactions regulate TLR4 signaling. J Leukoc Biol. 85(6):966–977. doi: 10.1189/jlb.1208763. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Lavon Y, Leitner G, Moallem U, Klipper E, Voet H, Jacoby S, Glick G, Meidan R, and Wolfenson D. 2011. Immediate and carryover effects of Gram-negative and Gram-positive toxin-induced mastitis on follicular function in dairy cows. Theriogenology 76(5):942–953. [DOI] [PubMed] [Google Scholar]
  14. Lee JY, Ye J, Gao Z, Youn HS, Lee WH, Zhao L, Sizemore N, and Hwang DH. 2003. Reciprocal modulation of Toll-like receptor-4 signaling pathways involving MyD88 and phosphatidylinositol 3-kinase/AKT by saturated and polyunsaturated fatty acids. J Biol Chem. 278(39):37041–37051. doi: 10.1074/jbc.M305213200. [DOI] [PubMed] [Google Scholar]
  15. Ma B, and Hottiger MO. 2016. Crosstalk between Wnt/β-catenin and NF-κB signaling pathway during inflammation. Front Immunol. 7:378. doi: 10.3389/fimmu.2016.00378 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Magata F, Horiuchi M, Echizenya R, Miura R, Chiba S, Matsui M, Miyamoto A, Kobayashi Y, and Shimizu T. 2014. Lipopolysaccharide in ovarian follicular fluid influences the steroid production in large follicles of dairy cows. Animal Reprod Sci. 144(1-2):6–13. doi: 10.1016/j.anireprosci.2013.11.005. [DOI] [PubMed] [Google Scholar]
  17. Magata F, and Shimizu T. 2017. Effect of lipopolysaccharide on developmental competence of oocytes. Reprod Toxicol. 71:1–7. doi: 10.1016/j.reprotox.2017.04.001. [DOI] [PubMed] [Google Scholar]
  18. Mateus L, da Costa LL, Diniz P, and Ziecik A. 2003. Relationship between endotoxin and prostaglandin (PGE2 and PGFM) concentrations and ovarian function in dairy cows with puerperal endometritis. Animal Reprod Sci. 76(3-4): 143–154. doi: 10.1016/s0378-4320(02)00248-8. [DOI] [PubMed] [Google Scholar]
  19. Piersanti RL, Horlock AD, Block J, Santos JEP, Sheldon IM, and Bromfield JJ. 2019. Persistent effects on bovine granulosa cell transcriptome after resolution of uterine disease. Reproduction 158(1):35–46. doi: 10.1530/rep-19-0037 [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Price JC, Bromfield JJ, and Sheldon IM. 2013. Pathogen-associated molecular patterns initiate inflammation and perturb the endocrine function of bovine granulosa cells from ovarian dominant follicles via TLR2 and TLR4 pathways. Endocrinology 154(9):3377–3386. doi: 10.1210/en.2013-1102. [DOI] [PubMed] [Google Scholar]
  21. Rincon J, Gindri P, Mion B, Ferronato G, Barbosa A, Maffi A, Pradieé J, Mondadori RG, Corrêa MN, and Pegoraro L. 2019. Early embryonic development of bovine oocytes challenged with LPS in vitro or in vivo. Reproduction 1(aop)doi: 10.1530/REP-19-0316. [DOI] [PubMed] [Google Scholar]
  22. Rivera R, and Hansen P. 2001. Development of cultured bovine embryos after exposure to high temperatures in the physiological range. REPRODUCTION-CAMBRIDGE- 121(1): 107–115. [PubMed] [Google Scholar]
  23. Shabankareh HK, Shahsavari MH, Hajarian H, and Moghaddam G. 2015. In vitro developmental competence of bovine oocytes: Effect of corpus luteum and follicle size. Iran J Reprod Med. 13(10):615. [PMC free article] [PubMed] [Google Scholar]
  24. Sheldon IM, Rycroft AN, Dogan B, Craven M, Bromfield JJ, Chandler A, Roberts MH, Price SB, Gilbert RO, and Simpson KW. 2010. Specific strains of Escherichia coli are pathogenic for the endometrium of cattle and cause pelvic inflammatory disease in cattle and mice. PloS one 5(2):e9192. [DOI] [PMC free article] [PubMed] [Google Scholar]
  25. Soto P, Natzke R, and Hansen P. 2003. Identification of possible mediators of embryonic mortality caused by mastitis: actions of lipopolysaccharide, prostaglandin F2α, and the nitric oxide generator, sodium nitroprusside dihydrate, on oocyte maturation and embryonic development in cattle. American Journal of Reproductive Immunology 50(3):263–272. [DOI] [PubMed] [Google Scholar]
  26. Spicer L, and Alpizar E. 1994. Effects of cytokines on FSH-induced estradiol production by bovine granulosa cells in vitro: dependence on size of follicle. Domest Anim Endocrinol. 11(1):25–34. doi: 10.1016/0739-7240(94)90034-5. [DOI] [PubMed] [Google Scholar]
  27. Takeuchi O, and Akira S. 2010. Pattern recognition receptors and inflammation. Cell 140(6):805–820. doi: doi: 10.1016/j.cell.2010.01.022. [DOI] [PubMed] [Google Scholar]
  28. Zhao S-J, Pang Y-W, Zhao X-M, Du W-H, Hao H-S, and Zhu H-B. 2017. Effects of lipopolysaccharide on maturation of bovine oocyte in vitro and its possible mechanisms. Oncotarget 8(3):4656. doi: 10.18632/oncotarget.13965. [DOI] [PMC free article] [PubMed] [Google Scholar]

RESOURCES