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. Author manuscript; available in PMC: 2023 Feb 1.
Published in final edited form as: Prostaglandins Other Lipid Mediat. 2021 Dec 15;158:106604. doi: 10.1016/j.prostaglandins.2021.106604

The soluble epoxide hydrolase inhibitor GSK2256294 decreases the proportion of adipose pro-inflammatory T cells

Mona Mashayekhi a, Celestine N Wanjalla b, Christian M Warren b, Joshua D Simmons b, Kakali Ghoshal c, Mark Pilkinton d, Samuel S Bailin b, Curtis L Gabriel e, Ambra Pozzi c,f, John R Koethe b,f, Nancy J Brown g, Spyros A Kalams b,*, J Matthew Luther h,*
PMCID: PMC8742790  NIHMSID: NIHMS1765562  PMID: 34922004

Abstract

Adipose tissue contains a complex immune environment and is a central contributor to heightened systemic inflammation in obese persons. Epoxyeicosatrienoic acids (EETs) are lipid signaling molecules that decrease inflammation in obese animals, but their effect on inflammation in humans is unknown. The enzyme soluble epoxide hydrolase (sEH) hydrolyzes EETs to less active diols, and we hypothesized that pharmacologic sEH inhibition would decrease adipose inflammation in obese individuals. We treated obese prediabetic adults with the sEH inhibitor GSK2256294 versus placebo in a crossover design, collected subcutaneous abdominal adipose tissue via lipoaspiration and characterized the tissue T cell profile. Treatment with GSK2256294 decreased the percentage of pro-inflammatory T cells producing interferon-gamma (IFNγ), but not interleukin (IL)-17A, and decreased the amount of secreted tumor necrosis factor-alpha (TNFα). Understanding the contribution of the EET/sEH pathway to inflammation in obesity could lead to new strategies to modulate adipose and systemic inflammation.

Keywords: epoxyeicosatrienoic acids, soluble epoxide hydrolase, inflammation, obesity, adipose

Introduction

Obesity is increasing in prevalence globally and is associated with increased risk of cardiometabolic diseases including type 2 diabetes and atherosclerotic cardiovascular disease. Expansion of adipose tissue in obesity is accompanied by changes in the immune cell profile that contribute to chronic systemic inflammation and cardiometabolic diseases. With obesity, there is hypoxia associated with adipocyte hypertrophy and decreased blood supply leading to formation of necrotic crown-like structures, and infiltration and differentiation of pro-inflammatory CD8+ T cells. In addition, there is a decrease in homeostatic cell types such as gamma-delta T cells (γδT) and regulatory T cells (Tregs), and an increase in pro-inflammatory interferon-gamma (IFNγ)-producing T helper (Th) type 1 and interleukin (IL)-17-producing Th17 cells.[1–3] These changes not only affect local adipose inflammation, but also contribute to adipocyte dysfunction, impaired lipid storage and handling, dysregulated adipokine production, and systemic inflammation and metabolic disorders,[4] representing a rich potential target for interventions.

Epoxyeicosatrienoic acids (EETs) are autocrine/paracrine lipid signaling molecules produced by the action of CYP epoxygenases on arachidonic acid. EETs are rapidly hydrolyzed to the more stable, but less active, dihydroxyeicosatrienoic acids (DHETs) by the enzyme soluble epoxide hydrolase (sEH). EETs promote vasodilation, sodium excretion, and improve insulin signaling.[5, 6] Increasing the action of EETs attenuates hypertension, endothelial cell dysfunction, renal injury, insulin resistance and cardiovascular remodeling in animal models.[7] In humans, higher circulating EETs are associated with greater insulin sensitivity, and the level of EETs is significantly lower in individuals with obesity and metabolic syndrome.[8, 9] Genetic variants that result in decreased EETs are more common in individuals with cardiovascular disease.[10]

An anti-inflammatory effect of EETs has been described in animals. In a mouse sepsis model, sEH inhibition prevents mortality, improves blood pressure, reduces liver and kidney damage, and decreases pro-inflammatory cytokines such as IL-6.[11] In rodent models of diet-induced obesity, increasing EETs through genetic or pharmacologic manipulation with sEH inhibition decreases adipose tissue inflammation, including decreased adipose necrotic crown-like structures, decreased adipose tissue infiltration by pro-inflammatory macrophages and CD8+ T cells, decreased adipose tissue gene expression of pro-inflammatory cytokines including tumor necrosis factor-alpha (TNFα), and decreased circulating inflammatory cytokines IL-1β, IL-6, TNFα and monocyte chemoattractant protein-1 (MCP-1).[12, 13] The effects of modulating the EET/sEH axis on inflammation in humans has only been investigated in vitro. TNFα activated human endothelial cells have decreased pro-inflammatory vascular cell adhesion molecule-1 (VCAM-1) transcription and NF-κB nuclear translocation in the presence of EETs, thus blocking mononuclear cell adhesion to the endothelium.[14]

The anti-inflammatory effect of EETs on adipose could represent a common mechanism underlying many of the observed benefits of EETs, yet it is unknown if EETs affect inflammation in humans. We hypothesized that blocking the degradation of EETs via sEH inhibition in obese individuals would improve adipose tissue inflammation. We used the sEH inhibitor GSK2256294 to block sEH enzyme activity and evaluated adipose tissue T cell profiles.

Materials and Methods

Clinical Study Design

The clinical study design and cohort has been previously reported (NCT03486223).[15] In brief, we enrolled obese (BMI ≥ 30 kg/m2) and pre-diabetic volunteers, age 21 through 60 years. Pre-diabetes was defined as a fasting plasma glucose of 100–125 mg/dL, a two-hour plasma glucose of 140–199 mg/dL during a 75-gram oral glucose tolerance test, or a hemoglobin A1c of 5.7–6.4%. All individuals gave written informed consent. Participants were randomized to the sEH inhibitor GSK2256294 (10 mg once daily) or matching placebo using a permuted-block algorithm for one week in a double-blind, crossover design. Participants underwent a seven-week washout period between drugs. On the seventh day of each drug treatment, participants underwent subcutaneous adipose biopsy by lipoaspiration. Study drug was provided by GlaxoSmithKline.

We enrolled and randomized 16 individuals. One participant dropped out prior to receiving drug, and 15 completed both study days. Eight individuals opted out of one or both of the adipose tissue biopsies during their study days. Thus, we had paired adipose tissue from both placebo and GSK2256294 study days in seven individuals, for a total of 14 adipose tissue samples which were used for the analyses in this manuscript. All comparisons are from within-individual paired samples. Four individuals were randomized to placebo treatment first and GSK2256294 treatment second, while the remaining three received GSK2256294 treatment first and placebo treatment second. There was no significant carryover effect detected between the treatments.

Adipose harvest

Adipose tissue was obtained from the periumbilical area using a Tulip™ Medical closed syringe system for lipoaspiration. Under aseptic conditions and local lidocaine anesthesia, a small incision was made in the skin. The GEMS Johnnie Snap lock was placed into a 60-cc syringe, the Tulip liposuction cannula with 60-cc syringe attached was inserted at an angle through the incision to below Scarpa’s fascia, and suction was applied until the syringe activates the clicker lock. The needle was oscillated at a rate of approximately 1 Hz without breaking suction with a twisting motion. The sampling continued until approximately 5 g of tissue was removed, and the sample was immediately processed for cryopreservation.

Stromal Vascular Fraction Isolation and Cryopreservation

Harvested adipose tissue was rinsed with media and weighed, then digested in gentleMACS C tubes with a Dissociator (Miltenyi Biotec). Collagenase D (Roche) was added at 2mg/mL and the sample was shaken at 37 degrees at 150 rpm for 40 minutes. The digested tissue was rinsed, filtered through a 70 micron filter, and the floating adipocyte fraction removed. The remaining immune-rich stromal vascular fraction was pelleted and washed, resuspended in cryopreservation solution (9 parts FBS, 1 part DMSO) at 1 million cells per vial, and slow frozen in a controlled-rate freezing container at −80 degrees overnight, then transferred to liquid nitrogen for long term storage. Adipose tissue macrophages do not survive cryopreservation and were not analyzed in this study.

In Vitro Characterization of Adipose Tissue Inflammatory Profile

To characterize the adipose immune milieu, the cryopreserved stromal vascular fraction was thawed and split into different treatment conditions in vitro: 1) media as negative control with DMSO carrier; 2) phorbol 12-myristate 13-acetate (PMA) (50ng/mL) and ionomycin (1μg/mL) stimulation for 6 hours to elicit cytokine responses. After 2 hours of culture, the supernatant was collected for measurement of secreted cytokines (Meso Scale Discovery), and Brefeldin A (BD Biosciences) was added to the cells at 1 μg/mL to trap intracellular cytokines for flow cytometry.

T cell Flow Cytometry

Sequential gating strategy was performed as follows for all: lymphocyte gate → exclusion of doublets → exclusion of dead cells using LIVE/DEAD Fixable Aqua, B cells using CD19, monocytes and macrophages using CD14 → selection of immune cells by CD45. CD3+ T cells have additional gating as follows: CD3+ T cells → exclusion of invariant natural killer T cells using CD1d tetramer (NIH tetramer core facility) and 6B11 anti-human TCR → exclusion of γδT cells using 11F2 anti-human γδTCR → exclusion of regulatory T cells using CD25 and CD127 → exclusion of IL-4-producing cells.

Reagents used for flow cytometry include: ThermoFisher LIVE/DEAD Aqua Stain Kit L34957. BD anti-CD14 antibody V500 561391; anti-CD19 antibody V500 561121; anti-IFNγ antibody PE-Cy7 557643; anti-CD4 antibody PerCP-Cy5.5 560650; anti-CD25 antibody BB515 565096; anti-CD8 antibody BUV395 563795; anti-γδ TCR antibody BUV737 748533. Biolegend anti-iNKT TCR BV611 342922; anti-IL-4 antibody BV421 500826; anti-IL-13 antibody BV421 501916; anti-CD127 antibody BV786 351330; anti-CD56 antibody BV605 318334; anti-IL-17A antibody PE-Dazzle 512336; anti-CRTH2 antibody AF647 350104; anti-CD3 antibody APC-Fire750 344840; anti-CD45 antibody AF700 304024.

Statistical analysis

Data are presented as mean ± standard deviation unless otherwise indicated. A Wilcoxon-signed rank test was used for paired within-subject comparison of GSK2256294 versus placebo. Correlation between different outcomes was assessed by Spearman’s correlation coefficient.

Results

Sixteen participants were randomized to treatment order. One individual dropped out prior to receiving study drug, and the remaining 15 completed both study visits. As reported previously, seven days of treatment with GSK2256294 decreased sEH activity as measured by conversion rate of 14,15-EET to 14,15-DHET in plasma (Placebo 4.08 ± 4.53, GSK2256294 1.93 ± 2.62 pmol/mL/hr, P=0.005) and in adipose tissue (Placebo 2,176 ± 965, GSK2256294 1,280 ± 675 pmol/mg/hr, P=0.0005).[15] There were no significant changes in measured plasma or adipose EET levels, suggesting possible compensatory changes in EET production in the setting of sEH inhibition in humans, which will require further investigation. GSK2256294 also decreased serum F2-isoprostanes, which are markers of oxidative damage and inflammation (Placebo 50.7 ± 15.8, GSK2256294 37.2 ± 17.3 pg/mL, P=0.03), but did not change plasma IL-6 levels.[15] Paired adipose after treatment with placebo versus GSK2256294 was available from seven individuals for the following exploratory immune analyses. These seven individuals were all female, with average age 41.9 ± 10.1.

We first treated the immune-rich stromal vascular fraction from adipose with PMA and ionomycin to elicit cytokine responses, then evaluated T cells using flow cytometry. We identified immune cells within the adipose using the marker CD45. T cells were identified as CD3+ lymphocytes and comprised 68.9 ± 9.6% of the CD45+ cells. CD3+ T cells were further subdivided into CD8+ cytotoxic cells and CD8− cells. Nearly all CD8− T cells express CD4 and are designated T helper (Th) cells (Supplementary Figure 1). The CD4 marker is known to be downregulated with PMA and ionomycin stimulation, thus we elected to show results based on CD8 positivity or negativity rather than gating on CD4. CD3− lymphocytes, the majority of which are natural killer (NK) cells, comprised 31.1 ± 9.5% of the CD45+ cells. Concatenated flow cytometry plots to illustrate the gating strategy are shown in Figure 1 (combined plot of seven fat samples in each treatment group). Figure 1A illustrates IFNγ-producing Th1 (CD8−), CD8+ cytotoxic, and NK cells. Figure 1B illustrates IL-17A-production from the same cell types. Cytokine production in the absence of PMA and ionomycin stimulation was low as expected (media control), and is subtracted as background cytokine response from the remaining graphs.

Figure 1. GSK2256294 Decreases the Percentage of Adipose Tissue IFNγ-Producing, but not IL-17A-Producing Cells.

Figure 1.

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Shown are concatenated contour plots from seven individuals after treatment with placebo or GSK2256294 for seven days. Adipose cells were treated in vitro with media control or PMA and ionomycin to elicit cytokine production. (A+B) Top row is gated on CD3+ T cells, and further separated into CD8− and CD8+ positive cytotoxic cells based on expression of CD8 on the x-axis. Bottom row is gated on CD3− lymphocytes. (A) IFNγ-producing Th1 (CD8−), CD8+ cytotoxic, and CD3− NK cells are shown in boxes. (B) IL-17A-producing Th17 (CD8−), CD8+ cytotoxic, and CD3− cells are shown in boxes. CD3− IL-17A-producing cells are not NK cells as they are negative for CD56.

Figure 2 illustrates the percentage of IFNγ-producing cell types for each individual during treatment with placebo or GSK2256294, with the denominator of total CD3+ or CD3− cells. Treatment with GSK2256294 decreased the percentage of adipose IFNγ-producing Th1 (CD8−) cells (median difference −4.69%, P=0.016). The effect of GSK2256294 treatment on IFNγ-producing CD8+ cytotoxic cells and NK cells was more variable and not significant (CD8+ median difference −0.73%, P=0.297; NK median difference −6.50%, P=0.297). Treatment with GSK2256294 did not significantly change the percentage of IL-4/13-producing Th2 cells (Supplementary Figure 2), IFNγ-producing γδT cells, IL-17A-producing Th17 cells, or regulatory T cells (Tregs) (data not shown). IFNγ production on a per-cell basis as measured by median fluorescence intensity (MFI) was highest in the Th1 (CD8−) cells, and not altered by treatment with GSK2256294 (Supplementary Figure 3).

Figure 2. GSK2256294 Decreases the Percentage of IFNγ-Producing Th1 (CD8−) Cells in Adipose Tissue.

Figure 2.

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Each line represents one individual. Graphed are the percentage of IFNγ-producing cells after negative control subtraction (PMA/ionomycin minus media condition). P-values generated using Wilcoxon signed-rank test. Summary data as median and inter-quartile range shown as diamonds with error bars.

As we could only capture a few cytokines using flow cytometry, we additionally measured secreted cytokines in the supernatant after media or PMA/ionomycin treatment. Several cytokines tested were not detectable in our culture conditions (IL-2, IL-10, IL-13, IL-17, IL-4, IL-5, IL-9). Treatment with GSK2256294 decreased the amount of secreted pro-inflammatory TNFα (median difference −15.79 pg/mL, P=0.016), and decreased secreted IFNγ in most individuals, although the difference did not reach significance (median difference −80.07 pg/mL, P=0.109) (Figure 3A). The percentage of Th1 (CD8−) and CD8+ IFNγ-producing cells was significantly correlated with the amount of secreted IFNγ and TNFα (Figure 3B). We did not measure intracellular TNFα by flow cytometry and could not quantify those cells numbers, although our data suggest that most cells co-produce both pro-inflammatory cytokines. Finally, IFNγ and TNFα levels were significantly correlated with plasma sEH activity as measured by conversion of EETs to DHETs (Supplementary Table 1). There was no correlation between the adipose tissue inflammatory measures and plasma F2 isoprostanes or IL-6 (data not shown).

Figure 3.

Figure 3.

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(A) GSK2256294 Decreases Secreted Pro-Inflammatory Cytokines from Adipose Tissue. Each line represents one individual. Graphed are the concentrations of each cytokine after negative control subtraction (PMA/ionomycin minus media condition). P-values generated using Wilcoxon signed-rank test. Summary data as median and inter-quartile range shown as diamonds with error bars. (B) Relationship Between Secreted Pro-Inflammatory Cytokines and the Proportion of IFNγ-Producing Cells. Sub- populations of IFNγ-producing cells are plotted on the x-axis and the secreted IFNγ and TNFα on the y-axis, with the linear regression line to visualize the correlation. Spearman’s rho (rs) and P-values generated using Spearman’s rank-order correlation.

Discussion

Our study found that pharmacologic inhibition of sEH decreases adipose IFNγ-producing Th1 cells and secreted TNFα in obese pre-diabetic individuals, suggesting a reduction in adipose tissue inflammation compared to placebo. Adipose tissue inflammation in obesity is a major contributor to chronic systemic inflammation, which promotes metabolic disease by increasing insulin resistance[16] and cardiovascular disease by increasing endothelial cell dysfunction and activation.[17–19] Thus, adipose inflammation may be a critical link between obesity and cardiometabolic disease development.

Our understanding of adipose tissue inflammation in humans is limited due to difficulty obtaining samples in clinical trials. Furthermore, systemic measures of inflammation do not adequately reflect tissue-level physiology. Importantly, while adipose tissue immune populations have been extensively characterized in obese animals, there are critical differences in adipose immune cells in humans that require further investigation. For example, while adipose macrophages in mice can be broadly classified into pro-inflammatory M1 and homeostatic M2 phenotypes based on markers such as CD11c and CD206, in humans there are mixed inflammatory phenotypes with markers of both M1 and M2.[20] Thus, discoveries in animals do not consistently translate to humans.

Based on animal studies, the mechanisms by which EETs modulate adipose inflammation may involve “browning” of white adipose tissue by increasing expression of thermogenic genes such as uncoupling protein-1 (UCP-1), reducing reactive oxygen species and inducing antioxidant genes such as heme oxygenase-1 (HO-1), and reducing NF-κB activation.[12–14, 21] Deletion of sEH results in reprogramming of white adipose to express thermogenic genes including UCP-1.[22] Human adipose tissue metabolomics also reveal that 5,6-EET is associated with the abundance of brown adipocytes in human brown adipose tissue.[23] We did not measure markers of thermogenesis in the current study, and future studies will need to determine if the same mechanism is involved in humans after sEH inhibition.

This study is the first to our knowledge to examine the effect of sEH inhibition on inflammation in humans in vivo. Weaknesses of this study include the small sample size, and our inability to evaluate adipose macrophages in cryopreserved samples. In addition, there were no significant changes in plasma and adipose EET levels with sEH inhibition, suggesting the observed effects may be mediated by another mechanism. Strengths of the study include the crossover design, allowing paired analysis within participants, and the placebo-controlled randomized order of treatment. Furthermore, by performing lipoaspiration to obtain our clinical samples, we aimed to understand tissue-level immune cell effects in adipose directly. Thus, despite our small sample size, we demonstrated the feasibility of evaluating the immune effects of interventions in humans in vivo, with direct measurements of adipose tissue T cell numbers and function. Future studies will compare the effects of our interventions on T cells in adipose tissue and peripheral blood simultaneously, to better understand if any circulating markers may be used as a surrogate for adipose inflammation. In addition, we are developing new protocols to study adipose macrophages obtained from lipoaspiration.

In conclusion, the present study suggests that sEH inhibition decreases adipose inflammation in humans, and future studies will determine if reducing adipose inflammation via sEH inhibition improves cardiometabolic health. Thus the anti-inflammatory potential of the EET/sEH axis in humans warrants further study.

Supplementary Material

1

Highlights:

  • sEH inhibition decreases adipose tissue IFNγ-producing T helper cells and secreted TNFα

  • sEH inhibition does not affect adipose tissue IL-17A production

  • Modulating the sEH/EET pathway has anti-inflammatory potential in humans

Sources of Funding:

This project was supported by NIH grant DK117875 (JML, NJB), the National Center for Research Resources Grant UL1 RR024975-01, now at the National Center for Advancing Translational Sciences Grant 2 UL1 TR000445-06. This work utilized the core(s) of the Vanderbilt Diabetes Research and Training Center funded by grant DK020593 from the National Institute of Diabetes and Digestive and Kidney Disease. GlaxoSmithKline generously provided drug and matching placebo for this study. MM supported by T32DK007061 and KL2TR002245.

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.

Disclosures:

MM: none

CNW: none

CMW: none

JDS: none

KG: none

MP: none

SSB: none

CLG: none

AP: none

JRK: none

NJB: none

SAK: none

JML: reports consultant relationship with Selenity Therapeutics and Mineralys.

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Supplementary Materials

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NIHMS1765562-supplement-1.docx (396.9KB, docx)

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