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Article

Uterine Myometrial Distension Augments the Production of Angiogenic and Proinflammatory Factors

by
Maurizio Mandalà
1,2,*,
Matthew E. Poynter
3,4,
Benjamin T. Suratt
3,4 and
George Osol
1
1
Department of Obstretrics, Gynecology and Reproductive Sciences, Larner College of Medicine, University of Vermont, Burlington, VT 05405, USA
2
Department of Biology, Ecology and Earth Sciences, University of Calabria, 87036 Rende, Italy
3
Pulmonary Disease and Critical Care Medicine, Department of Medicine, Larner College of Medicine, University of Vermont, Burlington, VT 05405, USA
4
Vermont Lung Center, University of Vermont, Burlington, VT 05405, USA
*
Author to whom correspondence should be addressed.
Targets 2025, 3(1), 3; https://doi.org/10.3390/targets3010003
Submission received: 19 October 2024 / Revised: 17 December 2024 / Accepted: 9 January 2025 / Published: 15 January 2025
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Abstract

:
We recently found that myometrial distension stimulates maternal uterine vascular remodeling, and hypothesized that this may be a previously unrecognized mechanism for inducing arterial growth during pregnancy. The aim of this study was to further characterize a recently developed surgical model in which medical-grade silicone was infused into one uterine horn of a non-gravid rat to induce acute myometrial stretch, followed by an additional, gradual distension due to the secretion of an exudate into the uterine lumen. Our objectives were to better understand the effects of stretch on the myometrium and to look for the expression of proangiogenic and proinflammatory factors that may stimulate uterine vascular remodeling. Morphometric analysis showed hypertrophy of the uterine corpus that was primarily due to axial growth since the myometrial cross-sectional area was unchanged due to a thinning of the uterine wall secondary to stretch. This finding was supported by significantly increased myometrial smooth muscle cell mitosis. There was also an increase in the concentration of myometrial elastin but not collagen. The analysis showed modest increases in neutrophils, activated and unactivated macrophages, and the proinflammatory cytokines RANTES, MIP-3α, GRO-KC, and TNFα. The most dramatic change was the extremely high level of VEGF in the exudate, which was increased >900× above circulating levels.

1. Introduction

Remodeling and growth of the uterine vasculature during pregnancy occurs in all mammalian species and is requisite for a normal pregnancy outcome. The net effect of the remodeling process is a decrease in uteroplacental vascular resistance, which facilitates significant (10- to 100-fold) increases in uteroplacental blood flow (UPBF) during gestation in humans and experimental animals [1,2,3,4,5,6,7,8,9]. Uterine volume increases progressively throughout gestation in all species secondary to fetoplacental growth [8,10].
The growth of the maternal pre-placental vasculature during gestation is three-dimensional, as vessels are found to remodel both axially and circumferentially [8,10]. Attenuation of the vascular growth and remodeling process is associated with gestational pathologies such as preeclampsia and intrauterine growth restriction (IUGR) [8]. We and others have found that nitric oxide (NO) plays a key role in circumferential remodeling since systemic inhibition of nitric oxide synthase (NOS) mitigates the process, and an attenuation of vascular growth has been observed in genetic mouse models lacking the endothelial isoform of this enzyme (eNOS) [8,10,11,12].
Conversely, the mechanisms underlying arterial lengthening are largely unknown. However, unlike the circumferential changes mentioned above, arterial lengthening was completely unaffected by NOS inhibition [11]. The remodeling process is highly localized, as it only occurs in uterine horns that contain implantation sites [13]. There is a strong correlation between arterial lengthening and the number of implanted fetuses present in the uterine horn [10,13]. Based on this observation, we hypothesized that myometrial distension may actively contribute to the process of vascular elongation through stretch- or deformation-induced production and secretion of cytokines [10,14].
Using a novel surgical approach in which medical-grade silicone was injected into the uterine lumen of a non-gravid rat to induce distension, our initial study established that significant remodeling of the uterine vasculature occurred secondary to myometrial stretch [10]. Specifically, there was significant axial remodeling of both the main uterine artery (MUA) and smaller, segmental (radial) vessels [10]. Additionally, there was a modest circumferential remodeling of the radial vessels, but not of the MUA [10]. It was also found that the initial distension induced by silicone infusion was followed by an accumulation of an exudate into the uterine lumen. Its formation produced further progressive swelling of the affected uterine horn, resulting in an additional eight- to ten-fold increase in uterine volume over a period of 4 weeks [10].
Studies investigating the effects of obstructive uropathy on rat and lamb bladders reported changes in the smooth muscle composition, extracellular matrix (e.g., elastin and collagen), and volume; secondary to a combination of hypertrophy and hyperplasia [15,16,17,18,19]. The aim of the present study was to apply the observations made in other systems to the surgical model by determining the composition of the uterine luminal exudate, with the purpose of examining its cellular and acellular components. Our working hypothesis was that understanding the composition of the exudate may help identify molecular factors responsible for the previously documented induction of vascular growth. We were also interested in better understanding how distension affects the myometrium itself by determining cellular mitotic indices, changes in overall structure, and matrix collagen and elastin content.

2. Materials and Methods

Animals: Virgin cycling Sprague Dawley rats were obtained at 11 weeks of age from Charles River (St. Constant, QC, Canada). Animals (n = 9) were housed at the University of Vermont Small Animal Facility, which is fully accredited by the Association for Assessment and Accreditation of Laboratory Animal Care. Feed and water were provided ad libitum. All experimental protocols were approved by the University’s Institutional Animal Care and Use Committee.
Surgical Protocol: Prior to surgery, vaginal smears were conducted on all rats to select an animal that was in the estrus or proestrus phase of the rat estrous cycle, when uterine volume is greatest. A surgical plane of anesthesia was induced with 4% isoflurane prior to the start of all surgical procedures and maintained with continued administration of isoflurane at 2–2.5%. All surgical procedures were conducted under sterile operating conditions. The abdomen was shaved and cleaned with Betadine Surgical Scrub (7.5% povidone–iodine) and alcohol prep pads (70% isopropyl alcohol) in an alternating fashion, to minimize the chance of infection.
The cervical and ovarian ends of the uterine corpus were each accessed through separate incisions. First, an oblique incision was made on the right side, just below the ribcage, and a portion of the uterine horn was exteriorized through it. A single suture (3-0 silk, Ethicon) was tied around the uterine corpus just below the ovary, with extra care taken to ensure that the uterine vasculature was not disrupted. This was facilitated by the manifold-like arrangement of the mesometrial vessels, which generally penetrate the uterus at an angle perpendicular to its corpus. A sterile normal saline solution (Vedco) was applied as needed to keep the abdominal contents moist. The abdominal cavity was closed with interrupted vertical mattress sutures (4-0 chromic gut, Ethicon), and a dosage of 0.25% Bupivicane (1:1 dilution of 0.5% Bupivicane with sterile normal saline solution) was applied around the incision before closing the skin with an interrupted stitch (4-0 Ethilon, Ethicon).
A second, inch-long incision was made above the vaginal opening. The lower (cervical) end of the uterine horn was exteriorized, and a single suture (3-0 silk, Ethicon) was loosely tied around the body of the uterus (also without disrupting the vasculature) approximately 1 cm from the cervix. A 1 cc syringe fitted with a 21-gauge needle prefilled with sterile medical-grade silicone (MDM Silicone Fluid 1000, Santa Paula, CA, USA) was inserted proximal to the suture, and the suture was then tightened around the needle. Silicone (0.15–0.20 mL) was infused into the lumen until the uterus was visually distended. After the needle was removed, the suture was double-tied to prevent leakage. A sham procedure was carried out on the contralateral horn by inserting and then withdrawing the needle and thread to mimic the trauma of puncturing the mesometrium and physically touching the myometrium, although the suture was not tied and shortly withdrawn. The abdominal cavity was closed in the same fashion as the oblique incision. All animals were given injections of the analgesic buprenorphine (0.05 mg/kg, Hospira, Inc., Lake Forest, IL, USA) immediately after surgery, 3–4 h after surgery, and the following morning. Povidone–iodine swab sticks (Medline) were used on all closed incisions, and animals were monitored for several days post-operatively for any signs of distress.
Experimental Protocol: On the day of analysis, animals were anesthetized with Sleepaway (26% sodium pentobarbital, 7.8% isopropyl alcohol (Fort Dodge Animal Health, Fort Dodge, IA, USA)). Once the rat reached a surgical plane of anesthesia (no visible reflexes), the abdominal and thoracic cavities were opened. Blood obtained via cardiac puncture was collected into a BD Vacutainer PPT Plasma Preparation Tube (Fisher). Animals were then decapitated with a small animal guillotine. The PPT tube was then centrifuged at 1300× g for 10 min (Sorvall, Germany) and plasma was removed, placed into a 2 mL centrifuge tube, snap-frozen in liquid nitrogen, and placed into a −80 °C freezer for future analysis.
The entire uterus was removed and pinned in a Sylgard-lined Petri dish filled with room-temperature physiologic saline solution (HEPES-PSS) that contained 0.1 mM Papaverine (Sigma-Aldrich, St. Louis, MO, USA) and 10 µM Diltazem (Sigma-Aldrich, St. Louis, MO, USA) to fully relax the vasculature. Each uterus was visually examined; there was no evidence of bacterial infection in any of the specimens. Measurements were made of uterine height, width, and length, and uterine volume was calculated based on the formula for the volume of a cylinder (π r2h, where h = length). The main uterine artery (MUA) and vein (MUV) length and segmental artery length (length of the vessels from the main uterine artery to the uterine wall, SA) [10] were measured, as were the main uterine artery inner and outer diameters, and the main uterine vein outer diameter using a flexible ruler (for vascular length) and an eyepiece micrometer calibrated to a dissecting scope (Zeiss, Oberkochen, Germany) for inner and outer diameters.
After all morphological measurements were obtained, the exudate in the ligated horn was withdrawn using a 19 gauge needle (BD Biosciences, San Jose, CA, USA) and a 5 mL syringe (BD Biosciences, San Jose, CA, USA). The exudate was centrifuged in a microcentrifuge (Sorvall, Langenselbold, Germany) at 2400× g for 10 min to remove any silicone particles. The exudate was then vortexed to thoroughly resuspend all cellular matter (American Scientific Products, Livonia, MI, USA).
The blood sample in the K2 EDTA tube and the exudate were then run on an Advia 120 Automated Hematology System (Siemens, Munich, Germany) to determine the cellularity of the samples. Cytospins of the exudate sample were taken using a Cytospin 4 at 80 rpm for 8 min (ThermoFisher, Waltham, MA, USA), and were then stained with Protocol Hema 3 (Fisher, Manchester, UK). Cytospin slides were photographed with AMG Evos XL Cells and hand-counted under an inverted light microscope (Zeiss, Oberkochen, Germany) to ensure the accuracy of the Advia.
Transverse sections of the non-stretched and stretched uterine horns were taken and fixed in 10% neutral buffered formalin and were submitted to the Surgical Pathology Laboratory at the University of Vermont Medical Center (Burlington, VT, USA) for tissue processing, sectioning, and staining. The histological techniques involve standardized tissue fixation/dehydration/embedding/staining procedures and are therefore not described in detail.
Tissue sections were stained with hematoxylin and eosin (H&E), Ki67 (MIB-1), Masson’s Trichrome, and Elastin Van Giesen. Additionally, portions of the non-stretched and stretched uterine horns and the exudate were snap-frozen in liquid nitrogen and placed in a −80 °C freezer for long-term storage and future analysis.
Composition of HEPES-PSS: The HEPES physiologic salt solution was prepared using deionized water and contained the following (in mmol/L): sodium chloride, 141.8; potassium chloride, 4.7; magnesium sulfate, 1.7; calcium chloride, 2.8; potassium phosphate, 1.2; HEPES, 10.0; ETDA, 0.5; and dextrose, 5.0. The solution was titrated with sodium hydroxide to a physiologic pH of 7.4 at 37 °C. All chemicals were obtained from Fisher Scientific (Agawam, MA, USA) and Sigma-Aldrich (St. Louis, MO, USA).
Microscopy: The H&E-stained slides of uterine sections were taken to the Microscopy Imaging Center at the University of Vermont Medical Center hospital (Burlington, VT, USA). These slides were imaged using an Olympus BX50 Light Microscope (Center Valley, PA, USA) at 2× magnification, and pictures were taken using a QImaging Retiga 2000R digital camera (Surry, BC, Canada).
Using the saved images, the unstressed cross-sectional area of the uterine lumen was calculated using MetaMorph Meta Imaging Series 7.7 (Molecular Devices Inc., Sunnyvale, CA, USA) for both stretched and non-stretched horns. Using MetaMorph software, luminal and total circumference were measured. The luminal diameter (LD) and the outer diameter (OD) were then calculated by entering previously measured circumferences into the equation D = C/π. The inner and outer circumferences delineate the inner and outer edges of the myometrium, and total myometrial areas were then automatically calculated by the MetaMorph software. Based on this delineation, myometrial wall thickness was determined using the equation WT = (OD − LD)/2 in each of the corresponding horns.
Smooth muscle cell mitotic indices were calculated as follows: Four randomly selected non-contiguous fields from each myometrial section were viewed under 40× magnification. The total number of nuclei per field was determined using MetaMorph software gated to correspond to visually visible nuclei. These values (33 total image views from 7 matched pairs) were averaged to provide the total number of nuclei per image area. The mitotic index was derived by dividing the number of MIB-1-stained nuclei (which were brown rather than blue) by the total number of nuclei, and multiplied by 100 so as to be expressed as a percentage.
Random images per uterine section were taken under 40× magnification for each stretched and non-stretched Masson’s Trichrome, and Elastin Van Giesen-stained slides (n = 26 image views from n = 5 matched pairs). Analysis was performed using MetaMorph software to calculate the percentage of elastin and collagen fibers in the extracellular matrix per image area.
Molecular Analyses: Three different molecular techniques (antibody array, ELISA, Bio-Plex) were used to determine the cytokine and growth factor protein content in the stored samples (plasma/serum, exudate, and tissue). The tissue samples were homogenized prior to beginning the assays. The samples were weighed individually to determine the proper amount of diluted 10× lysis buffer (50 mM Tris, 0.1% SDS, 1% Triton X-100) to add to the samples (8.333 µL buffer/mg of tissue). The 10× lysis buffer was prepared by diluting to 1X with deionized water and adding a Roche Complete Mini Protease Inhibitor Cocktail Tablet (Indianapolis, IN, USA) for every 10 mL of lysis buffer. The tissues were then homogenized using a Polytron Tissue Homogenizer (Kinematica, Eschbach, Germany), and centrifuged at 800× g for 20 min at 4 °C in order to collect the supernatant for the assay.
BCA: A bicinchoninic acid protein assay (BCA) was completed on the tissue samples to quantify the total amount of protein in the samples that were used for the protein array and the ELISAs (Pierce). A standard curve was created using albumin protein standards (1000, 500, 250, 125, 62.5, 31.25, 0), and 25 µL of each standard was pipetted into wells in a microplate. Working Reagent (WR) was prepared by mixing 10 mL of Reagent A with 200 µL of Reagent B. Once prepared, 200 µL of WR was added to each well (standards and samples), and the microplate was covered at placed on an orbital shaker (The Belly Dancer Stovall, IBI Scientific, Peosta, IA, USA) for 30 s. The plate was then incubated in the plate reader at 37 °C for 30 min and read at 562 nm (BioTek Synergy HT, Winooski, VT, USA).
Rat Cytokine Antibody Array: A Rat Cytokine Antibody Array was obtained from RayBiotech, Inc. (Norcross, GA, USA) to determine the expression profiles of cytokine proteins in the plasma, homogenized tissues, and exudate of the stored samples. Array membranes were placed into the provided eight-well tray. Blocking buffer (2 mL) was added to each well and the plate was incubated for 30 min at room temperature on an orbital shaker. Samples were pooled to obtain the 1 mL volume necessary to run the assay. Samples were diluted (10-fold for tissue lysates and plasma), and the assay was run according to kit instructions. Each membrane was imaged on a BioRad ChemiDoc XRS System (Philadelphia, PA, USA) to determine the cytokines present in the plasma, tissue lysate, and exudate through the intensity of chemiluminescent signals.
Mouse PlGF-2 ELISA and Rat PDGF-AA ELISA: Mouse PlGF-2 ELISA and Rat PDGF-AA ELISA kits were obtained from RayBio (Norcross, GA, USA) to quantify the protein levels in the plasma, tissues, and exudate. The mouse PlGF-2 kit has cross-reactivity with rat PlGF-2. The tissues were homogenized, and protein content determined using the BCA was run on all tissue samples as described above. All samples were diluted and the standard was prepared and plated according to kit instructions. The samples were then plated in duplicates (diluted and undiluted) and the plate was incubated overnight in a cold room at 4 °C on an orbital shaker. The following day the ELISA was run according to kit instructions. The plate was read at 450 nm (BioTek Synergy HT, Winooski, VT, USA). Calculations were made of the mean absorbance for each set of duplicate standards and samples and the average zero standard optical density was subtracted. The standard curve was plotted in SigmaPlot 11 (Systat Software, San Jose, CA, USA).
Bio-Plex Pro Rat Cytokine 23-Plex Assay: A Bio-Plex Pro Rat Cytokine 23-Plex Assay was obtained from BioRad (Hercules, CA, USA) to quantify protein levels of 23 different cytokines (IL-1α, IL-1β, IL-2, IL-4, IL-5, IL-6, IL-7, IL-10, IL-12p70, IL-13, IL-17, IL-18, EPO, G-CSF, GM-CSF, GRO KC, IFNγ, M-CSF, MIP-3α, RANTES, TNFα, and VEGF) in stored plasma, exudate, and tissue samples. The assay was run on a BioRad Bio-Plex 200 machine (Hercules, CA, USA) that was calibrated prior to running the assay. Tissue samples were homogenized and prepared as described above. The standard dilution series was prepared according to kit instructions. Plasma samples were diluted 1:4, and the exudate and tissue homogenates were run undiluted. The plate was run according to kit instructions. The plate outline and dilutions were entered into the Bio-Plex Manager software and the plate was subsequently read on the Bio-Plex 200. Outliers were removed by the Bio-Plex Manager Software by selecting the assay optimization option to generate an accurate standard curve. In the initial run, VEGF was found to be above the upper limit of quantification in the exudate and the samples were run again at a 1:2 dilution using the same procedure described above.
Statistical Analyses: For each uterine morphological and vascular measurement, the stretched horn was compared to the non-stretched horn, which served as an intra-animal control. Data were analyzed using SigmaPlot statistical software (Systat Software, Inc., San Jose, CA, USA). Data are expressed as mean ± standard error of means, with paired t-test used for all comparisons between groups. T-tests for the ELISAs and BioPlex were made between the tissue (NS vs. ST) group and the plasma vs. exudate. Groups are graphed together for visualization purposes. Data for the ELISAs are represented in pg/mL. Data for the BioPlex are represented in pg/mL for plasma and exudate and in pg/mg for the myometrial tissue (ST and NS). p values of <0.05 were considered significant.

3. Results

Uterine morphological changes: Unless otherwise noted, nine matched intra-animal pairs were used for all morphological comparisons between the stretched (ST) and non-stretched (NS) horns. As seen in Figure 1, the surgical procedure caused a significant (25×) increase after 28–30 days in the calculated uterine volume of the stretched (ST) vs. non-stretched (NS) horn (0.13 ± 0.052 mL NS vs. 3.32 ± 0.96 mL ST, p < 0.001).
Uterine vascular changes: Remodeling of the uterine vasculature in the stretched horn was also observed when compared to the non-stretched horn (Figure 2 and Table 1). As noted in an earlier study [10], there were no significant changes in the average unstressed inner diameters of the main uterine arteries of the stretched vs. non-stretched horns. Circumferential remodeling was observed in the venous circulation, however, as evidenced by the significant increase in the main uterine vein diameter. There were also significant differences between the two horns in the axial remodeling of the main uterine artery and vein length, and in the length of the segmental arteries (SA; Table 1).
Histological analysis: Uterine myometrial morphological changes were evaluated using H&E staining of eight matched pairs (n = 8). Metamorph analysis showed the average smooth muscle area to be similar between non-stretched and stretched horns (227,586 ± 20,735 µm2 NS vs. 252,628 ± 8036 µm2 ST). Further analysis confirmed that the similarity of areas was due to the combination of a 3-fold increase in average luminal diameter (304 ± 28.5 µm NS vs. 962 ± 75.3 µm ST, p < 0.001) combined with a ~2-fold decrease in average wall thickness (335 ± 22.0 µm NS vs. 191 ± 15.1 µm ST, p < 0.001).
The mitotic index was calculated using MIB-1-stained slides for seven matched pairs (n = 7). There was a small, but significant (1.3-fold) increase in the mean mitotic index (% dividing cells) between the NS and ST uterine horns (4.0 ± 0.2%, NS vs. 5.5 ± 0.3%, ST, p < 0.0004).
Changes in uterine elastin and collagen composition were calculated and compared for five matched pairs (n = 5). There was a significant (1.3-fold) increase in the elastin concentration of stretched uterine horn sections when compared with the non-stretched control sections (51.8 ± 3.6% NS vs. 69.8 ± 1.8% ST, p < 0.0001). There was no significant difference in collagen content between non-stretched and stretched uterine sections (59.6 ± 4.6% NS vs. 59.3 ± 3.9% ST).

4. Cellular and Molecular Analyses

Cellularity: A total of eight animals were used for the Advia analysis of the luminal exudate and whole blood. There was a ~4.6-fold difference in the number of white blood cells present in whole blood when compared to the exudate. Examination of the exudate, using a cytospin, revealed the presence of both unactivated and activated macrophages, along with neutrophils (Figure 3) [20].
Rat Cytokine Bio-Plex: Of the 23 cytokines and growth factors that were tested using the Bio-Plex, only 5 were elevated above the lower limit of quantification: GRO-KC, MIP3α, RANTES, TNFα, and VEGF (Table 2).
The most significant increase was that of VEGF in the exudate, which was >900-fold above the plasma levels (Table 2). There was also a significant (~5.3-fold) increase in VEGF in the stretched vs. non-stretched myometrium (Table 2).
Exudate GRO-KC concentrations were increased ~10-fold relative to plasma but were not detectable in the ST and NS myometrium (Table 2). The concentrations of MIP-3α in the exudate were ~1.7-fold greater than in the plasma and were not detectable in the ST and NS myometrium (Table 2). On the other hand, RANTES was undetectable in the exudate, but measurable in the plasma. Additionally, a ~3.5-fold increase in RANTES was observed in the ST myometrium when compared to the NS myometrium (Table 2). There was a ~1.2-fold increase in the level of TNFα in the ST vs. NS myometrium, but it was undetectable in the plasma and exudate (Table 2).
Rat Cytokine Antibody Array: The data obtained from this non-quantitative screening assay were used to determine which cytokines and growth factors to test for in the ST and NS myometrium, plasma, and exudate samples with subsequent ELISAs and the Bio-Plex assays.
PlGF-2 and PDGF-AA ELISAs: The PlGF-2 and PDGF-AA ELISA assays were run using samples of homogenized ST and NS myometrium, plasma, and exudate (n = 4 for all). PlGF-2 levels were highest in myometrium with no significant difference between the ST and NS groups (91.3 ± 19.5 pg/mL and 88.8 ± 22.4 pg, respectively; PlGF-2 was measurable in the plasma (45.6 ± 26.36 pg/mL), but could not be detected in the exudate.
PDGF-AA was also present in the myometrium (114 ± 74.7 pg and 133 ± 50.2 pg/mL, respectively), but there was no significant difference between the ST vs. NS groups. A marked elevation of PDGF-AA protein was observed in the exudate (1823 ± 505 pg, p = 0.011), but it was not detectable in the plasma.

5. Discussion

This novel surgical model enabled us to observe morphological changes in the myometrium caused directly by distension, without the confounding presence of fetuses and placentae. We were able to reaffirm the influence of mechanical stretch on uterine vascular remodeling in the non-gravid state through the observation of significant changes in axial and circumferential remodeling, including increases in the length of the main uterine artery and vein, as well as the increases in segmental artery length and diameter of the main uterine vein [10]. Excessive mechanical stretch causes alterations in cellular structure and function and composition of the extracellular matrix (ECM) associated with diseases such as obstructive uropathy [14,15,16,17,18,19,21,22,23,24]. During normal pregnancy, the uterus must stretch and grow considerably to accommodate fetoplacental growth causing changes in the ECM [21,22,23]. Through placental perfusion, the uterus must also adapt its circulation to meet the metabolic demands of the fetus and perfuse the increased myometrial mass in a process of expansive remodeling that involves significant increases in arterial and venous diameter and length [8]. This led us to characterize uterine myometrial morphological adaptations in response to distension using this model, which is unique in its combination of an acute stretch stimulus with a slower, more progressive distension, as is observed during pregnancy, which results from the formation of an exudate.
During pregnancy, the mechanical signaling within the myometrium promotes rapid and extensive hypertrophy of the uterine smooth muscle cells (SMCs) associated with increased expression of ECM proteins [22,25]. Based upon this, we originally hypothesized that mechanical stretch associated with surgery might also induce hypertrophy of the uterus. Hypertrophy of the myometrium takes place in the second half of gestation as smooth muscle cell size increases, leading to an increase in the thickness of uterine muscle layers, as well as significant changes in the mass and composition of the extracellular matrix [21,23,24,26,27].
In rats, a significant increase in myometrial smooth muscle cell mitotic activity has been observed after paraffin-induced distension of a uterine horn, particularly within the circular layer [17]. Although hypertrophy of individual smooth muscle cells was not observed in our model, the uterine corpus increased in both length and volume following stretch, such that the increased mitotic index may reflect axial growth. Notably, we also found previously that myometrial stretch primarily induced arterial elongation rather than widening [10]. Based on this, we predict that the remodeling mechanisms that operate in both the uterus and in the vasculature favor axial rather than circumferential growth since we were unable to find evidence for increased myometrial cross-sectional area in response to distension.
Using unilaterally pregnant rats, Shynlova and colleagues found that the myometrium of the non-gravid horns had a lower level of ECM gene expression than the gravid horns, which led them to conclude that there is a synergistic action between sex steroids, particularly progesterone (P4) and 17β-estradiol, and mechanical stretch during pregnancy [22]. It has been known for over 50 years that de novo collagen formation occurs in the rat uterus during pregnancy and that this process increases with the number of fetuses present in the horn [28]. In order to study collagen formation, Cullen and Harkness injected paraffin wax into the uterine lumen to determine the influence of mechanical stretch alone and with adjuvant hormonal treatment to mimic the average levels observed during pregnancy and reported a 3-fold increase in collagen content [17]. Conversely, we did not observe any appreciable change in collagen content between the distended and control horns, although elastin concentrations did increase significantly. Elastin content in gravid rats increases 3- to 8-fold above the baseline level by day 19 of pregnancy, as it is needed for accommodating the distension of the uterus [29,30]. In a study by Percival and Starcher, elastin fibers were observed in the pregnant rat uterus in a variety of random, partially extended configurations [31], similar to those observed in the stretched myometrium of our surgically altered rats. In addition to removing the confounding factors of pregnancy, our model enables us to simulate pregnancy through progressive distension of a single uterine horn without using a highly inflammatory stimulant, such as paraffin wax. Future studies will be directed toward determining if sex steroids play a role in the remodeling process through the use of ovariectomized rats.
Mechanical stretch itself has been found to affect cytokine and chemokine production by the cells of different tissues (fibroblasts, smooth muscle cells, and endothelial cells), and it is thought that the deformation of these cell types leads to the release of cytokines and proinflammatory mediators resulting in excessive immune system activation [14]. Distension itself, in the gravid and non-gravid state, has been found to induce changes in proinflammatory factor levels. In vascular cells, transcriptional activator and/or repressor molecules are sensitive to tensile stress and control stress-induced changes in proinflammatory gene expression [14]. Pregnancy is a mild inflammatory state, and many proinflammatory factors, as well as monocytes and neutrophils, are present at elevated levels during pregnancy [32,33]. It is thought that disturbances in the profile of proinflammatory factors during pregnancy may lead to certain pathologies, including severe preeclampsia [32,33].
Although we have not yet determined the exact cellular source(s) of the uterine exudate, we were able to characterize its cellular and molecular content using a combination of techniques. The level of VEGF—a factor known to induce hyperpermeability—was significantly elevated in the exudate, leading us to believe that the fluid may originate from the vasculature. Additionally, the exudate may also be a product of the uterine glands or the epithelium. Under the influence of VEGF, newly formed blood vessels have been reported to be leaky [34], although it is uncertain whether uterine distension stimulates intramyometrial angiogenesis. Increases in vascular permeability that led to the movement of the protein-rich exudate and blood cells into the uterine lumen have been noted in earlier surgical models that utilized paraffin to distend uterine horns in rats [17]. In addition to the increased transport of proteins and water, several different mechanisms have been invoked to explain why exudates form in response to inflammation. The most common cause is endothelial cell contraction, which leads to intercellular gaps in postcapillary venules, which can be attributed to the presence of TNFα [34]. In our study, TNFα concentrations were significantly elevated in the stretched vs. non-stretched myometrium, indicating a potential mechanism behind exudate formation.
We chose to use medical-grade silicone in this technique because it is used in a variety of surgeries. However, despite its relatively inert properties, some inflammation has been reported following its use—specifically, the release of proinflammatory cytokines and an inflammatory infiltrate that consists of macrophages, foreign body giant cells, and a variable number of lymphocytes and plasma cells [35,36]. The inflammatory process itself is associated with increases in blood flow, vascular permeability, and transendothelial leukocyte migration [37]. Increases in diameter and length in the mesometrial vasculature of the stretched horn are indicative of an increase in blood flow to the distended horn. The presence of cytokines and growth factors, such as VEGF, also suggests an increase in vascular permeability, and the presence of leukocytes in the uterine lumen indicates a transendothelial migration. The lymphatic system also participates in the inflammatory response by helping to drain edema fluid, leukocytes, and cell debris from the extravascular space [34]. It can also become secondarily inflamed, which can aid in the buildup of exudate because of reduced drainage [34]. Thus, we were not surprised to find evidence of inflammation in the stretched uterus. Moreover, although our primary focus was on the myometrium, it should be recognized that endometrial decidualization may also be activated by the presence of silicone and/or physical distention, and this may account for the exudate secretion and accumulation.
The observation of monocytes and leukocytes in the exudate is indicative of an inflammatory response. It is well known that most cytokines are not synthesized by cells in the absence of an inflammatory reaction or immune response [14]. We observed elevated levels of GRO-KC, a neutrophil chemoattractant, and of MIP-3α, which is strongly chemotactic for lymphocytes and mildly for neutrophils [14,38]. We also observed an increase in levels of TNFα in the stretched myometrium, which is produced by activated macrophages [14,38]. Interestingly, TNFα has been found to be present in the myometrium only during active labor [39]. During pregnancy in humans and animals such as cows, macrophages and neutrophils are recruited to the placenta as part of the mild inflammatory response [40,41].
Mechanical stretch has been found to enhance the expression of HIF-1α, the molecular precursor to VEGF, which is associated with an increase in the expression of VEGF through transcriptional regulation [14]. The >900-fold elevation in VEGF concentrations within the exudate relative to circulating levels was both striking and unexpected. VEGF plays a role in many important processes in the human body including endothelial cell proliferation, promotion of cellular migration, inhibition of apoptosis, induction of angiogenesis and vascular permeability, and regulation of vasculogenesis [42]. It has been implicated in the pathogenesis of preeclampsia, as reduced tissue availability secondary to elevated sFlt-1 levels is well documented [43]. It is possible that the high levels of VEGF in the luminal exudate may stimulate vascular remodeling, particularly axial growth. Its ability to increase uterine venous permeability >10-fold, as shown in an earlier study from our laboratory [44], may also augment the transfer of vasoactive and/or growth-promoting signals to the arteries, which run in parallel with, and in close apposition to, the veins. Venoarterial signaling has been established as a mechanism of luteolysis in a number of species, providing a physiological precedent, although whether it plays an active role in maternal gestational uterine vascular remodeling is not known at this time [45]. Further studies will need to be completed to elucidate the particular roles of VEGF in our model.
PDGF, which was observed to be elevated in the exudate, has been found to play an important role in wound healing, particularly in the process of smooth muscle cell proliferation, as well as in angiogenesis, and the pathophysiology of pregnancy-induced hypertension [46,47]. An elevation of PDGF in the serum of hypertensive women can cause atherosclerotic changes in the spiral arteries of placentas and an increase in the proliferation of smooth muscle cells [47]. Additionally, PDGF has been found to contribute to smooth muscle hyperplasia in chronic pulmonary hypertension, as well as endothelial cell hyperplasia [48]. The presence of PDGF in the exudate may have facilitated the increase in vascular smooth muscle mitosis in the radial arteries, as noted in our preliminary study completed in the laboratory [10], and in the increase in the rate of the endothelial cells of the distended myometrium.
In summary, this study has further validated this novel surgical model of myometrial distension for studying the effects of mechanical stretch on the myometrium without the confounding factors of pregnancy. Uterine hypertrophy was present as evidenced by the increased axial length of the uterine corpus combined with the significantly increased smooth muscle mitotic index and increased matrix elastin content. Distension was accompanied by a mild inflammatory response, confirmed by cytokine, growth factor, and cellular analyses of the luminal exudate. The origin of the exudate is thought to be caused by a combination of factors, including the introduction of a foreign substance (silicone) and subsequent mechanical perturbation (distension) of the uterine wall. Of particular note were the large increases in exudate VEGF and PDGF concentrations, which support their proangiogenic actions, and suggest their involvement in the observed arterial remodeling, particularly axial growth, following uterine distension.

Author Contributions

M.M., Conceptualization, Methodology, Validation, Investigation, Data Curation, and Writing—Original Draft; M.E.P., Methodology, Validation, Formal Analysis, Resources, and Data Curation; B.T.S., Methodology, Validation, Formal Analysis, and Resources; G.O., Conceptualization, Validation, Formal Analysis, Resources, Data Curation, and Writing—Review and Editing). All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded to G.O. by NIH, grant number R21HL112216.

Institutional Review Board Statement

The animal study protocol was approved by the Institutional Review Board of University of Vermont, (protocol code A3301-01, 19 August 2015).

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in the study are included in the article, further inquiries can be directed to the corresponding author.

Acknowledgments

We would like to thank Shannon Kostin, Rebecca Nelson, and Darren Clas for their technical assistance, and Ruth Blauwiekel, DVM, for her invaluable surgical training and support. The authors acknowledge R21 support from the National Institutes of Health (HL112216).

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Views of the vasculature in a non-stretched (A) vs. stretched (B) uterine horn from the same animal at equivalent magnification. Scale bar = 0.5 mm in both photographs.
Figure 1. Views of the vasculature in a non-stretched (A) vs. stretched (B) uterine horn from the same animal at equivalent magnification. Scale bar = 0.5 mm in both photographs.
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Figure 2. H&E-stained myometrial cross-sections of a non-stretched uterine horn (A) and a stretched uterine horn (B). Scale bar = 200 µm in both photographs.
Figure 2. H&E-stained myometrial cross-sections of a non-stretched uterine horn (A) and a stretched uterine horn (B). Scale bar = 200 µm in both photographs.
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Figure 3. Image of cytospin stained with Hema 3 showing activated (two nuclei) vs. unactivated macrophages (one nucleus) and neutrophils, 40×.
Figure 3. Image of cytospin stained with Hema 3 showing activated (two nuclei) vs. unactivated macrophages (one nucleus) and neutrophils, 40×.
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Table 1. Vascular measurements. * p < 0.05.
Table 1. Vascular measurements. * p < 0.05.
Non-Stretched Horn
(n = 9)		Stretched Horn
(n = 9)
MUA diameter (µm)43 ± 4.144.9 ± 4.5
MUV diameter (µm)155 ± 15.3184 ± 20.1 *
SA length (mm)2.9 ± 0.094.3 ± 0.5 *
MUA/MUV length (cm)3.2 ± 0.184.5 ± 0.4 *
Table 2. Cytokine and growth factor protein levels. * p < 0.05.
Table 2. Cytokine and growth factor protein levels. * p < 0.05.
CytokinesMyometriumPlasma
(pg/mL)		Exudate
(pg/mL)
Non-Stretched
(pg/mg)		Stretched
(pg/mg)
GRO-KC--10.7 ± 2.9101 ± 20.9 *
MIP-3α--0.403 ± 0.130.646 ± 0.313
RANTES5.6 ± 2.225.6 ± 7.5 *22.6 ± 11.6-
TNFα4.8 ± 0.974.15 ± 0.66--
VEGF0.144 ± 0.0440.505 ± 0.102 *0.561 ± 0.36514 ± 62 *
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Mandalà, M.; Poynter, M.E.; Suratt, B.T.; Osol, G. Uterine Myometrial Distension Augments the Production of Angiogenic and Proinflammatory Factors. Targets 2025, 3, 3. https://doi.org/10.3390/targets3010003

AMA Style

Mandalà M, Poynter ME, Suratt BT, Osol G. Uterine Myometrial Distension Augments the Production of Angiogenic and Proinflammatory Factors. Targets. 2025; 3(1):3. https://doi.org/10.3390/targets3010003

Chicago/Turabian Style

Mandalà, Maurizio, Matthew E. Poynter, Benjamin T. Suratt, and George Osol. 2025. "Uterine Myometrial Distension Augments the Production of Angiogenic and Proinflammatory Factors" Targets 3, no. 1: 3. https://doi.org/10.3390/targets3010003

APA Style

Mandalà, M., Poynter, M. E., Suratt, B. T., & Osol, G. (2025). Uterine Myometrial Distension Augments the Production of Angiogenic and Proinflammatory Factors. Targets, 3(1), 3. https://doi.org/10.3390/targets3010003

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