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. 2019 Nov 29;97(12):4965–4973. doi: 10.1093/jas/skz343

Differentiating between the effects of heat stress and lipopolysaccharide on the porcine ovarian heat shock protein response1

Jacob T Seibert 1, Malavika K Adur 1, Ronald B Schultz 1, Porsha Q Thomas 1, Zoe E Kiefer 1, Aileen F Keating 1, Lance H Baumgard 1, Jason W Ross 1,✉
PMCID: PMC6915215  PMID: 31782954

Abstract

Heat stress (HS) negatively affects both human and farm-animal health and undermines efficiency in a variety of economically important agricultural variables, including reproduction. HS impairs the intestinal barrier, allowing for translocation of the resident microflora and endotoxins, such as lipopolysaccharide (LPS), from the gastrointestinal lumen into systemic circulation. While much is known about the cellular function of heat shock proteins (HSPs) in most tissues, the in vivo ovarian HSP response to stressful stimuli remains ill-defined. The purpose of this study was to compare the effects of HS or LPS on ovarian HSP expression in pigs. We hypothesized that ovarian HSPs are responsive to both HS and LPS. Altrenogest (15 mg/d) was administered per os for estrus synchronization (14 d) prior to treatment and three animal paradigms were used: (i) gilts were exposed to cyclical HS (31 ± 1.4 °C) or thermoneutral (TN; 20 ± 0.5 °C) conditions immediately following altrenogest withdrawal for 5 d during follicular development; (ii) gilts were subjected to repeated (4×/d) saline (CON) or LPS (0.1 μg/kg BW) i.v. infusion immediately following altrenogest withdrawal for 5 d; and (iii) gilts were subjected to TN (20 ± 1 °C) or cyclical HS (31 to 35 °C) conditions 2 d post estrus (dpe) until 12 dpe during the luteal phase. While no differences were detected for transcript abundances of the assessed ovarian HSP, the protein abundance of specific HSP was influenced by stressors during the follicular and luteal phases. HS during the follicular phase tended (P < 0.1) to increase ovarian protein abundance of HSP90AA1 and HSPA1A, and increased (P ≤ 0.05) HSF1, HSPD1, and HSPB1 compared with TN controls, while HS decreased HSP90AB1 (P = 0.01). Exposure to LPS increased (P < 0.05) HSP90AA1 and HSPA1A and tended (P < 0.1) to increase HSF1 and HSPB1 compared with CON gilts, while HSP90AB1 and HSPD1 were not affected by LPS. HS during the luteal phase increased (P < 0.05) abundance of HSPB1 in corpora lutea (CL), decreased (P < 0.05) CL HSP90AB1, but did not impact HSF1, HSPD1, HSP90AA1, or HSPA1A abundance. Thus, these data support that HS and LPS similarly regulate expression of specific ovarian HSP, which suggest that HS effects on the ovary are in part mediated by LPS.

Keywords: corpus luteum, heat stress, ovary, pigs

Introduction

Heat stress (HS) is a global environmental problem, claiming thousands of human lives in recent years (Kosatsky, 2005; Russo et al., 2015). In addition to its impact on human health, HS threatens global food security for the growing population (World Bank, 2007; United Nations, 2017), undermining efficiency at every stage of livestock production (Baumgard and Rhoads, 2013; Ross et al., 2017). Many species, including pigs, limit their feed intake during HS to minimize the thermic effect of feeding, resulting in reduced growth (Bianca, 1976; Blaxter, 1989; Collin et al., 2001; Baumgard and Rhoads, 2013). However, despite the reduced nutrient intake, heat-stressed animals actually retain more adipose tissue when compared with thermoneutral (TN) counterparts on the same plane of nutrient intake (Close et al., 1971; Collin et al., 2001; Baumgard and Rhoads, 2013; Pearce et al., 2013a). While contradicting what is energetically predicted, hyperthermia also results in increased circulating insulin (Pearce et al., 2013a), stemming from increased pancreatic beta cell secretion (Sanz Fernandez et al., 2015a). Additionally, whole-body insulin sensitivity is increased (Sanz Fernandez et al., 2015b), which appears to be explained primarily by increased glucose utilization by the immune system (Kvidera et al., 2017). Immunoactivation is in response to compromised intestinal integrity and concomitant increase in circulating lipopolysaccharide (LPS) during HS (Pearce et al., 2013b). Collectively, the aforementioned immune activation and endocrine changes contribute to the overall metabolic dysfunction during HS (Baumgard and Rhoads, 2013).

Hyperthermia, similar to other hyperinsulinemic conditions (e.g., polycystic ovarian syndrome and obesity), can compromise many aspects of female reproduction and reduce fertility (Ross et al., 2017; Bidne et al., 2018a). Moreover, just as with HS (Tompkins et al., 1967; Omtvedt et al., 1971; Madan and Johnson, 1973), LPS exposure can also alter gonadotropin secretion (Shakil et al., 1994) and impair embryonic survival (Deb et al., 2004). In the ovary specifically, HS and LPS both influence the transcript abundance of steroidogenic enzymes (Herath et al., 2007; Nteeba et al., 2015; Bidne et al., 2018a) and disrupt in vitro gamete development (Tseng et al., 2006; Bromfield and Sheldon, 2011). Interestingly, however, while in vivo LPS exposure has pro-apoptotic effects in the ovary (Perez et al., 1996; Besnard et al., 2001), in vivo HS induces ovarian autophagy and promotes anti-apoptotic signaling (Hale et al., 2017). Thus, LPS and HS have similarities and differences with respect to molecular mechanisms underlying how each contributes to cellular function and female infertility.

Originally discovered in Drosophila for their involvement in the heat shock response (HSR; Tissières et al., 1974), heat shock proteins (HSP) are stress responsive proteins that maintain intracellular homeostasis (Fulda et al., 2010). Heat shock factor 1 (HSF1) is a critical transcription factor in the HSR that acts via binding to regulatory regions containing heat shock elements, ultimately promoting the transcription of the affiliated HSP gene (Fulda et al., 2010; Tetievsky and Horowitz, 2010). In response to a stress stimulus, HSF1 self-oligomerizes and translocates into the nucleus enabling DNA binding (Sarge et al., 1993; Westwood and Wu, 1993). Heat shock protein family A (HSP70) member 1A (HSPA1A) and HSP family B (small) member 1 (HSPB1) are stress-induced proteins involved in protein refolding, the latter specifically participating in cytoskeleton maintenance (Bakthisaran et al., 2015; Seo et al., 2016). In the mitochondria, HSP family D (HSP60) member 1 (HSPD1) is important for protein import and stress-responsive protein refolding (Martin et al., 1992; Levy-Rimler et al., 2001). Heat shock protein 90 (HSP90) chaperones have a variety of functions with two major isoforms, HSP90 α family class A member 1 (HSP90AA1) and HSP90 α family class B member 1 (HSP90AB1), and reportedly have differing responses to thermal stimuli (Schopf et al., 2017). Interestingly, specific HSPs have been implicated in cytokine production (Kol et al., 2000; Triantafilou et al., 2001) and are involved in normal reproductive processes, including estrogen receptor α regulation (Dhamad et al., 2016), luteolysis (Khanna et al., 1995a, 1995b; Kim et al., 1996), and CYP19A1 control (Driancourt et al., 1999).

Although HSPs are involved in cellular pathways responsive to both HS and LPS, their involvement in ovarian stress response pathways remain ill-defined. Furthermore, it is uncertain as to whether the direct and/or indirect effects (e.g., HS-induced “leaky gut”) of HS influence the in vivo ovarian HSP response and how that corresponds with ovarian dysfunction. We hypothesized that the porcine ovarian HSP response is similarly regulated during both in vivo HS and LPS challenges. Thus, our objective herein was to investigate and compare the effects of HS or LPS during specific phases of the estrous cycle on the ovarian HSP response.

Materials and Methods

Animals and Experimental Design

All procedures involving animals were approved by the Iowa State University Institutional Animal Care and Use Committee. This study consisted of three independent live-phase experiments (Figure 1), all of which have been described previously (Bidne, 2017; Hale et al., 2017; Bidne et al., 2018b;). In all experiments, altrenogest (15 mg/d; Matrix; Merck Animal Health; Summit, NJ) was administered per os for estrus synchronization 14 d prior to either HS or LPS administration. In the first experiment (Hale et al., 2017), postpubertal gilts (126 ± 22 kg) were exposed to TN (20.3 ± 0.5 °C; n = 6) or cyclical HS (25 to 31°C ± 1 °C; n = 6) conditions for 5 d immediately following altrenogest withdrawal (the follicular phase). Similarly, in Exp. 2 (Bidne et al., 2018b), gilts (163 ± 3 kg) were subjected to 5 d of repeated (4×/d) saline (CON; n = 3) or LPS (0.1 μg/kg BW; from Escherichia coli O55:B5, L2880; n = 6) via i.v. administration immediately following altrenogest withdrawal. In Exp. 3 (Bidne, 2017), gilts (167 ± 10 kg) were subjected to TN (20 ± 1 °C; n = 7) or cyclical HS (31 to 35 °C; n = 7) conditions 2 d post estrus (dpe) until 12 dpe during the luteal phase. At the end of each experiment, all animals were euthanized using captive bolt followed by exsanguination. Ovaries were removed immediately following exsanguination and corpora lutea (CL) were excised from the ovaries, flash frozen in liquid nitrogen, and stored at −80 °C until utilized for nucleic acid and protein extraction.

Figure 1.

Figure 1.

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Experimental scheme for the study. Gilts were subjected to TN (n = 6) or HS (n = 6) conditions during the follicular phase (5 d; top), saline (CON; n = 3), or LPS (n = 6) infusion via indwelling jugular catheter during the follicular phase (5 d; middle), or TN (n = 7) or HS (n = 7) during the luteal phase (12 d; bottom). Each live-phase experiment was conducted independently (Bidne, 2017; Hale et al., 2017; Bidne et al., 2018b). In all experiments, altrenogest was administered per os to all gilts to facilitate estrus synchronization prior to thermal or IV treatments during the follicular phase; gilts underwent estrus detection (behavioral estrus) prior to thermal treatment during the luteal phase.

Quantitative One-Step RT-PCR

Frozen ovary (Exps. 1 and 2) or CL tissue representing several CL randomly selected from the same ovary (Exp. 3) were homogenized in QIAzol Lysis Reagent using the TissueLyser II and total RNA isolated via the miRNeasy Mini kit (Qiagen, Hilden, Germany) according to the manufacturer’s protocol for each animal. Quantitative RT-PCR analysis of transcripts of interest from both tissues was conducted using the QuantiTect SYBR Green RT-PCR Kit (Qiagen) and measured on an Eppendorf Mastercycler (Eppendorf, Hamburg, Germany). All primer sequences utilized for quantitative analysis for each target gene are presented in Table 1. RNA input (10 ng) was DNase-treated according to the manufacturer’s instructions (AM1906, Ambion, Austin, TX) and assayed for each sample in triplicate. Thermal cycling conditions for SYBR Green detection were 50 °C for 30 min (reverse transcription), 95 °C for 15 min, followed by 40 repetitive cycles of denaturation at 94 °C for 15 s, annealing at 57 to 60 °C (depending on the primer set) for 30 s, and extension at 72 °C for 30 s followed by fluorescent data acquisition. Melting curve analysis was subsequently conducted following the completion of the PCR protocol. Additionally, to determine possible genomic DNA contamination, a pooled sample was assayed in the absence of reverse transcriptase. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA was included as an endogenous normalization control to correct for loading discrepancies (Kuijk et al., 2007; Li et al., 2011; Knapczyk-Stwora et al., 2019). Relative quantification of target gene expression was evaluated with the comparative cycle threshold (Ct) method. The ΔCt value was determined by subtracting the target Ct of each sample from its respective GAPDH Ct value. Calculation of ΔΔCt involved using the single greatest sample ΔCt (the sample with the lowest expression) as an arbitrary constant to subtract from all other sample ΔCt values (Ashworth et al., 2006). Relative differences for each sample were calculated assuming an amplification efficiency of 2 during the geometric region of amplification and applying the equation 2ΔΔCt. Statistical analysis for each assay was performed on the ΔCt values. HSPD1 was not statistically analyzed due to the presence of multiple amplicons in the qRT-PCR reaction.

Table 1.

Primer information for quantitative amplification of each target gene

Gene Description Primers, 5′ to 3′
GAPDH Glyceraldehyde-3-phosphate dehydrogenase F: TGGTGAAGGTCGGAGTGAAC R: GAAGGGGTCATTGATGGCGA
HSP90AA1 Heat shock protein 90 α family class A member 1 F: CTGGTCAAGAAATGCTTGGAG R: TGGTCCTGGGTCTCACCTGT
HSP90AB1 Heat shock protein 90 α family class B member 1 F: AACACTGCGGTCAGGGTATC R: ACATTCCCTCTCCACACAGG
HSF1 Heat shock transcription factor 1 F: AGCTCAGTGACGTCGGAGAT R: AGCATGGATTCCAAACTGCT
HSPA1A Heat shock protein family A (Hsp70) member 1A F: TCAAGGGCAAGATCAGCGAG R: TCAAACTCGTCCTTCTCGGC
HSPB1 Heat shock protein family B (small) member 1 F: TCGAAAATACACGCTGCCCC R: TTCCGGGCTTTTCCGACTTT
HSPB8 Heat shock protein family B (small) member 8 F: GTCTGGCAAACACGAGGAGA R: TGGGGAAAGCGAGGCAAATA
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Western Blot Analysis

Whole-ovarian (Exps. 1 and 2) or CL (Exp. 3) tissues were homogenized in tissue lysis buffer (1% Triton X-100, 50 mM HEPES, 150 mM NaCl, 10% glycerol, 50 mM NaF, 2 mM EDTA, 1% SDS), incubated on ice for 30 min, and centrifuged at 10,621 × g at 4 °C for 15 min. The resulting supernatant was centrifuged again at 10,621 × g at 4 °C for 15 min. The protein concentration of the clarified supernatant was determined using Pierce BCA Protein Assay Kit (Thermo Scientific, Rockford, IL) and quantified using a microplate photometer (Hycult Biotech, Uden, the Netherlands). Samples were denatured in loading buffer (50 mM Tris–HCl, pH 6.8, 2% SDS, 10% glycerol, 1% BME, 12.5 mM EDTA, 0.02% bromophenol blue) at 95 °C for 5 min, placed immediately on ice for 1 min, then stored at −80 °C until used. Protein samples were loaded into a 4% to 20% gradient Tris glycine gel (Lonza PAGEr Gold Precast Gels) with 50 µg per lane. The Bio-Rad Mini PROTEAN Tetra System was used to run the gel at 60 V for 30 min followed by 120 V for 90 min at room temperature (RT). After size separation, the proteins were transferred to a nitrocellulose membrane for 1.5 h at 100 V at 4 °C. Equal protein loading and transfer efficiency was confirmed by Ponceau S staining of the nitrocellulose membranes, followed by washing (with shaking) in Tris-buffered saline (TBS) with 0.1% Tween-20 (TBST) then washed with shaking in TBST three times for 10 min at RT. Membranes were blocked for 1 h with shaking at RT in 5% milk in TBST. Membranes were incubated overnight with primary antibodies specific for human HSP90AA1 (Cell Signaling, Danvers, MA, 8165; 1:1000), human HSP90AB1 (Cell Signaling, 7411; 1:1000), human HSF1 (Cell Signaling, 12972; 1:1000), chicken HSPA1A (Novus Biologicals, NB110-96427; 1:5000), human HSPD1 (Cell Signaling, 12165; 1:1000), and human HSPB1 (Novus Biologicals, NBP2-25149; 1:10000) at 4°C in 5% milk TBST. All primary antibodies used in the study have demonstrated reactivity or are predicted to detect homologous proteins in the pig. Negative controls were also evaluated using a pooled sample, representing a portion of all protein samples utilized in the western blot analysis, in which the membrane was incubated with rabbit IgG (Cell Signaling, 2729; 1:1,000), mouse IgG (Cell Signaling, 5415; 1:1,000), or no primary antibody. Membranes were washed with shaking in TBST three times for 10 min at RT and incubated with the appropriate secondary antibody for 1 h at RT and washed three times for 10 min each in TBST at RT. Horseradish peroxidase substrate (Millipore, Billerica, MA) was added to the membrane for 40 s in the dark. Membrane images were captured using a ChemiImager 5500 (Alpha Innotech, San Leandro, CA) with Alpha Ease FC software (version 3.03 Alpha Innotech) and densitometry was used to quantify protein band intensities corresponding to the primary antibody. Densitometric analysis was also conducted with Ponceau S staining for each blot using Image Studio Lite (Li-Core), confirming equal protein loading and used for normalization of specific proteins of interest. Two technical replications were also performed for each protein of interest. HSPB8 protein was not evaluated due to the lack of chemiluminescent signal compared with the negative controls from antibodies from various suppliers.

Statistical Analysis

Western blot and qRT-PCR data were analyzed using PROC TTEST in SAS University Edition software, version 9.4 (SAS Institute Inc., Cary, NC). Data are reported as means and statistical significance (P ≤ 0.05) and tendency thresholds (0.05 < P ≤ 0.10) were utilized for interpretation.

Results

HS or LPS Have Marginal Effects on Ovarian HSP Transcript Abundance

Quantitative RT-PCR was conducted to analyze changes in mRNA transcript abundance of several HSP genes (Figure 2). There was a tendency for increased ovarian HSPB8 (1.67-fold; P = 0.09; Figure 2B) in response to LPS exposure during the follicular phase and numerical reductions for ovarian HSPB1 (1.50-fold; P = 0.17; Figure 2A) and CL HSPA1A (3.50-fold; P = 0.13; Figure 2C) in response to HS during the follicular or luteal phase, respectively. The transcript abundance of all other assessed HSP genes were not influenced by the stressor in each experiment (P ≥ 0.27; Figure 2A to C).

Figure 2.

Figure 2.

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Effects of HS or LPS on ovarian transcript abundance of HSPs genes. Heat stress during the follicular phase (n = 6) had a marginal impact on the whole ovary transcript abundance of HSP-related genes (HSF1, HSP90AA1, HSP90AB1, HSPA1A, and HSPB8; P ≥ 0.35; A) relative to TN (n = 6) conditions; HSPB1 was numerically reduced (P = 0.17; A). Similarly, LPS during the follicular phase (n = 6) had little influence on whole ovary HSP transcript abundance (P ≥ 0.51; B) relative to saline-infused controls (CON; n = 3), but HSPB8 tended to be increased due to LPS (P = 0.09; B). Heat stress during the luteal phase (n = 7) had a similar influence on CL transcript abundance of most HSP-related genes (P ≥ 0.26; C) and only numerically reduced HSPA1A transcript abundance (P = 0.13; C) relative to TN conditions (n = 7). Relative differences of normalized transcript abundance are represented as fold change between the treatments in each experiment. The # symbol indicates a statistical tendency (0.05 < P ≤ 0.10).

The Protein Abundance of Ovarian HSPs Is Selectively Influenced by HS and LPS

Despite little influence on transcript abundance of HSP-related genes, HS during the follicular phase tended to increase ovarian HSP90AA1 (70%, P = 0.07; Figure 3B) and HSPA1A (101%, P = 0.10; Figure 3E) protein abundance. A similar pattern was observed in response to LPS during the follicular phase as HSP90AA1 (61%, P = 0.01; Figure 4B) and HSPA1A (86%, P = 0.01; Figure 4E) protein abundances were increased. However, the protein abundances of CL HSP90AA1 (P = 0.42; Figure 5B) and HSPA1A (P = 0.51; Figure 5E) were not influenced by HS. Thus, ovarian HSP90AA1 and HSPA1A are responsive to both HS and LPS during the follicular phase, but not influenced by HS in the CL during the luteal phase.

Figure 3.

Figure 3.

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HS during the follicular phase alters HSP abundance in the ovary. Gilts either underwent 5 d of cyclical HS (n = 6) or TN (n = 6) conditions during the follicular phase after estrus synchronization. Western blotting of ovarian protein lysates for each gilt with antibodies directed toward HSPs (A) demonstrated a tendency for increased protein abundance due to HS for HSP90AA1 (B). HS decreased the protein abundance of HSP90AB1 (C), but increased and tended to increase HSF1 (D) and HSPA1A (E) protein abundance, respectively. The protein abundance of HSPD1 (F) and HSPB1 (G) was increased due to HS. Protein band intensities corresponding to the primary antibody were normalized to Ponceau S staining for each respective lane following densitometric analysis.

Figure 4.

Figure 4.

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LPS during the follicular phase alters HSP abundance in the ovary. Gilts were infused with either saline (CON; n = 3) or LPS (n = 6) via indwelling jugular catheter for 5 d during the follicular phase after estrus synchronization. Western blotting of ovarian protein lysates for each gilt with antibodies directed toward HSPs (A) demonstrated increased protein abundance due to LPS for HSP90AA1 (B), but HSP90AB1 (C) was unaltered. Lipopolysaccharide tended to increase HSF1 (D) and increased HSPA1A (E). The protein abundance of HSPD1 (F) and HSPB1 (G) was not influenced and tended to be increased by LPS, respectively. Protein band intensities corresponding to the primary antibody were normalized to Ponceau S staining for each respective lane following densitometric analysis.

Figure 5.

Figure 5.

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Heat stress (HS) during the luteal phase alters HSP abundance in the corpus luteum (CL). Gilts either underwent 12 d of cyclical HS (n = 7) or TN (n = 7) conditions during the luteal phase after estrus synchronization and subsequent estrus detection. Western blotting of CL protein lysates for each gilt with antibodies directed toward HSPs (A) demonstrated that HS did not influence HSP90A1A (B), but reduced HSP90AB1 (C) protein abundance. HS numerically increased HSF1 (D), but did not alter HSPA1A (E). The protein abundance of HSPD1 (F) and HSPB1 (G) was numerically increased and significantly increased due to HS, respectively. Protein band intensities corresponding to the primary antibody were normalized to Ponceau S staining for each respective lane following densitometric analysis.

Interestingly, ovarian protein abundance of HSP90AB1 was decreased in response to HS (45%, P = 0.01; Figure 3C) and CL (53%, P = 0.02; Figure 5C) during both the follicular and luteal phases, respectively. Inversely, HS increased ovarian HSPD1 during the follicular phase (95%, P < 0.01; Figure 3F) and numerically increased CL HSPD1 (69%, P = 0.17; Figure 5F). Unlike HS, LPS exposure during the follicular phase did not alter the protein abundance of ovarian HSP90AB1 (P = 0.86; Figure 4C) or HSPD1 (P = 0.89; Figure 4F). Collectively, HSP90AB1 and HSPD1 are altered in response to HS, but not influenced by LPS exposure.

The protein abundance of ovarian HSF1 was increased in response to HS (53%, P = 0.05; Figure 3D) and tended to be increased by LPS (200%, P = 0.08; Figure 4D) during the follicular phase. In the CL, HS numerically increased HSF1 protein abundance (47%, P = 0.17; Figure 5D). Ovarian HSPB1 protein abundance was increased due to HS (101%; P = 0.01; Figure 3G) and tended to increase in response to LPS (57%, P = 0.09; Figure 4G) during the follicular phase. HS also increased CL HSPB1 protein abundance (175%; P = 0.03; Figure 5G). Taken together, ovarian HSF1 and HSPB1 are increased in response to both HS and LPS exposure.

Discussion

HS results in female infertility through alterations to cyclicity, pregnancy, and ovarian function (De Rensis et al., 2017; Ross et al., 2017). Many of these HS-induced reproductive consequences are seemingly akin to those caused by LPS exposure (Ross et al., 2017; Bidne et al., 2018a). The aforementioned parallels are likely not coincidental as HS compromises intestinal barrier function, resulting in endotoxemia and immune activation (Hall et al., 1999; Baumgard and Rhoads, 2013). Thus, it remains unclear whether direct and/or indirect effects of HS mediate alterations to ovarian intracellular signaling pathways and little is known regarding the ovarian molecular stress response machinery during HS. Interestingly, while marginal effects were observed for transcript levels, both HS and LPS had a major impact on the protein abundances of assessed HSPs. Thus, intraovarian HSP-related pathways during HS may partially be the result of HS-induced “leaky gut”.

Differential effects of HS and LPS on specific HSP with regard to stress type occurred during the follicular phase. While both HS and LPS increased ovarian HSP90AA1, HSPA1A, HSPB1, and HSF1 protein abundances, HSP90AB1 and HSPD1 were only responsive to HS and not influenced by LPS administration. Non-ovarian tissues have increased HSP90AA1 and HSP90AB1 transcript abundance in response to in vivo LPS exposure (Kaucsár et al., 2014), which may indicate overall differences in tissue responses to LPS. Regarding thermal stimuli, HSP90AA1 is inducible (Zhang et al., 1999), and HSP90AB1 is unresponsive (Meng et al., 1993), which may be due to the constitutive expression of the latter (Csermely et al., 1998; Sreedhar et al., 2004; Schopf et al., 2017). Although HS increased HSP90AA1 in this study, it also unexpectedly decreased protein abundance of HSP90AB1. Reasons for this differential regulation are unclear but could be due to differences between in vitro and in vivo experiments, cell types, and/or species-specific responses to HS.

We have previously demonstrated that the protein abundance of ovarian TLR4, a major receptor involved in LPS stimulation of innate immune pathways (Hoshino et al., 1999) is increased in response to both HS and LPS during the follicular phase (Bidne et al., 2018b; Dickson et al., 2018). Interestingly, HSP90AA1 is implicated in the LPS response and subsequent cytokine production (Triantafilou et al., 2001). Whether HSP90AA1 responds to LPS independently or in concert with ovarian TLR4, as well as the possible involvement of other HSPs (Kol et al., 2000; Triantafilou et al., 2001), in ovarian immune-related pathways during HS is of further interest.

HS decreased HSP90AB1 and increased HSPB1 protein abundance in the ovary and CL during the follicular phase and luteal phase, respectively. Heat-induced numerical increases for HSF1 (P = 0.17) and HSPD1 (P = 0.17) were also observed in the CL. Interestingly, and perhaps insightful to their biological roles, HSP90AA1 and HSPA1A were only altered by HS in the ovary during the follicular phase and not in the CL. Cell-specific responses to HS regarding ovarian HSP machinery have been demonstrated in pigs (Pennarossa et al., 2012), thus the possibility that HS-induced alterations to HSP90AA1 and HSPA1A are cell-type specific during the luteal phase is plausible. Interestingly, however, the abundance of certain HSP transcripts and proteins in porcine cumulus cells are not influenced by in vitro or in vivo HS (Pennarossa et al., 2012), suggesting specific HSPs could become active after granulosa and thecal cell differentiation based on our CL results. Considering HSPs are functionally involved in luteinization and CL regression in rodents (Khanna et al., 1995a, 1995b; Kim et al., 1996), further investigation into how the assessed HSPs in this study may alter ovarian function during HS are warranted.

Altered ovarian HSPs abundance and/or post-translational modifications could have critical consequences during HS. The thermal responsiveness of the cytoskeleton has been determined in non-ovarian cell types, thus increases in HSPB1 and HSF1 may indicate an attempt to stabilize cellular structures during HS (Welch and Suhan, 1985; Gavrilova et al., 2013; Baird et al., 2014; Bakthisaran et al., 2015). The HS-induced HSPD1 increase may indicate possible mitochondrial-specific responses as HSPD1 is a critical component for protein import and stress responses within mitochondria (Martin et al., 1992; Levy-Rimler et al., 2001) and HS negatively affects mitochondrial function (e.g., altered morphology, oxidative stress, and cytochrome c release) (Welch and Suhan, 1985; Davidson and Schiestl, 2001; Qian et al., 2004). If localized to the oocyte specifically, this could alter bioenergetics for the developing gamete, considering the high lipid content and the emphasized use of mitochondrial β oxidation in the porcine oocyte (Prates et al., 2014). Whether or not the functions of the aforementioned HSP are maintained in the ovary needs to be determined. Interestingly, HSP90AA1 and HSP90AB1 are highly abundant in the murine oocyte (Zhang et al., 2009) and selective chemical inhibition of the latter alters HSP protein abundance, including HSF1, in cancer cells (Khandelwal et al., 2018). Additionally, HSP90AB1 has been implicated in ovarian autoimmunity during premature ovarian failure (Pires and Khole, 2009). Further studies could establish if insulin’s regulatory ability of HSP expression (Li et al., 2006) is maintained or altered in the ovary during HS and determine if specific roles of ovarian HSPs and their association with HS-induced infertility.

Conclusion

In summary, this study identified consistent and differential ovarian HSP response stemming from either HS or LPS, suggesting that the intraovarian HSP response during HS appears to at least in part originate from HS-induced compromised intestinal barrier integrity. It is important to note that tissue collection occurred at the end of the respective treatment (5 to 10 d), and understanding the temporal pattern of the HSP response may reveal insight into the mechanism by which HS contributes to seasonal infertility. Additionally, how the observed alterations in HSP abundance corresponds with specific ovarian cell types and their impact on ovarian function and fertility during HS warrants further investigation.

Acknowledgment

The authors would like to thank Emily L. Fett for her design with Figure 1 in this manuscript.

Footnotes

1

Results described here within were supported by the National Pork Board, the Iowa Pork Producers Association, and Agriculture and Food Research Initiative competitive grant no. 2011-67003-30007 from the USDA National Institute of Food and Agriculture.

Funding

This project was supported by the Iowa Pork Producers Association, the National Pork Board, and the United States Department of Agriculture. Any opinion, findings, conclusions, or recommendations expressed in this publication are those of the authors and do not necessarily reflect the view of the Iowa Pork Producers Association, the National Pork Board, or the United States Department of Agriculture. No conflicts of interest, financial or otherwise are declared by the author (s).

Conflict of interest statement. None declared.

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