Introduction
Sphingosine-1-phosphate receptors belong to G-protein-coupled receptors that participate in different cellular responses depending on cell types and availabilities of downstream effectors [
1]. Sphingosine-1-phosphate receptor 2 (S1PR2) is predominately expressed in vascular tissues [
2], where its activity is tightly associated with the function of vasculature, especially the function of endothelial cells [
3]. JTE-013, a potent and selective S1PR2 antagonist, could reverses the inhibitory effects of S1PR2 on vascular endothelial cells [
4]. Windh and colleagues have revealed that S1PR2 couples to Gi and G12/13 along with the Gq family members [
5], thus activating multiple small GTPases (Rho, Rac, and Ras) [
6‐
8] to exerts distinct effects via mediating diverse signaling pathways [
9,
10]. Recently, S1PR2 has been found to be an endothelial barrier-unstabilizing mediator in the activation of Rho-associated coiled coil-containing protein kinase (ROCK) pathway [
11]. Human renal glomerular endothelial cells (HRGECs) are highly specialized primary endothelial cells packed with fenestrae, which contribute to the glomerular filtration barrier [
12,
13]. However, the specific role of S1PR2 in the regulatory function of HRGECs remains unclear.
As a major energy source of cells, mitochondria have an important role in the regulation of endothelial function [
14]. In renal glomerular endothelial cells, mitochondria are the most important effectors of cellular metabolism and oxidative stress, and their changes in morphology and function are closely related to the varying cellular microenvironment and functions [
15]. Numerous studies have shown that sphingosine-1-phosphate receptors affect the morphology and function of mitochondria in various cell types [
16‐
19]. However, the role of S1PR2 activity in the morphology and function of mitochondria of HRGECs and the downstream signaling molecules of S1PR2 coupling associated with mitochondrial and endothelial dysfunctions have not been identified.
Endothelial dysfunction induced by high glucose (HG) is a classical in vitro model to mimic the development of diabetes. HG induces endothelial cell apoptosis, oxidative stress, and inflammation [
20]. Also, HG increases permeability [
21] and enhances endothelial-to-mesenchymal transition in HRGECs [
22]. Moreover, increased mitochondrial fragmentation and superoxide production [
23] and lowered mitochondrial bioenergetics [
24] have been observed in HG-treated endothelial cells. Although mitochondrial impairment is indicative of HG-induced endothelial dysfunction, the changes of mitochondrial morphology and functions induced by HG in HRGECs and the molecular mechanisms underlying these pathological events are still unclear. In the present study, we aim to investigate whether S1PR2 modulates the morphology and function of mitochondria in HRGECs via regulating ROCK1, thus affecting the dysfunction of glomerular endothelium mediated by HG.
Materials and methods
Cell culture and treatment
Primary HRGECs were purchased from Sciencell (Carlsbad, CA, USA) and cultured in Endothelial Cell Medium (ECM) (Carlsbad, CA, USA) containing 5% fetal bovine serum at 37 °C in a 5% CO
2 incubator. Cells at passages 2–4 were treated with 30 mM HG for 72 h to induce cell dysfunction [
25] or treated in parallel with 5.5 mM normal glucose (NG) as a control group or 24.5 mM mannitol (Mnt) plus 5.5 mM glucose as an osmotic control. As indicated elsewhere [
26], cells were pretreated with 10 μM JTE-013 (Cayman, Ann Arbor, MI, USA), a specific S1PR2 inhibitor, or 10 μM Y27632 (Selleck, Houston, TX, USA), a selective ROCK1 inhibitor, for 30 min to assess the regulatory effects of SIPR2 and ROCK1 in HRGECs.
Immunoblotting
Total protein extracts (30 μg) was separated on 8–12% Bis-Tris gels, followed by transferring onto polyvinylidene fluoride membranes. After blocking with 5% skimmed milk for 2 h at room temperature (RT), the blots were incubated with a primary antibody overnight at 4 °C and then with horseradish peroxidase-conjugated secondary antibody for 1 h at RT. The primary antibodies included ROCK1 (Abcam, Cambridge, UK; 1:1000), RhoA (Abcam; 1:1000), and dynmin-related protein-1(Drp1) (Abcam; 1:1000). The intensities of protein bands were visualized using chemiluminescence with an ImageQuant 350 (GE Healthcare, Woburn, MA, USA). GAPDH was used as a loading control.
Mitochondrial morphology assay
After treated with glucose at different levels, 1 × 106 cells were fixed in 3% glutaraldehyde for 48 h at 4 °C and post-fixed in 1% osmium tetroxide for 2 h at RT. After pre-stained with acetatebarbitone for 10 min, dehydrated with acetone, and embedded in Epon 812, cells were isolated on nickel fitters, stained with 2% uranyl acetate for 10 min, and then with Reynold’s lead citrate for 5 min. Finally, mitochondrial morphology of HRGECs was evaluated by transmission electron microscopy (Hitachi-7650, Tokyo, Japan) at 60 kV.
ATP assay
Intracellular ATP levels of HRGECs were quantified by using an ATPlite assay kit according to the manufacturer’s instructions (PerkinElmer, Akron, OH, USA). Briefly, following cell exposure to HG with or without pretreatment with an inhibitor, mammalian cell lysis solution was added to cell-containing 96-well microplates. After shaking for 5 min at RT, equal amount of substrate was added and mixed for another 5 min at RT. The luminescent intensities were monitored 10 min later to indicate the ATP levels and mitochondrial activities.
Intracellular ROS assay
Intracellular ROS production of living HRGECs was detected by using 2′,7′- dichlorofluorescein diacetate (DCF-DA) (Beyotime, Jiangsu, China), which is oxidized to a fluorescent dye when ROS are present. After cultured with HG, cells were stained with 10 μM DCF-DA dye in the dark for 30 min at 37 °C and then washed twice. Intracellular fluorescence generated was subsequently imaged using a fluorescent microscope with a FITC filter (Ex/EM = 485/535 nm; Nikon, Tokyo, Japan) and quantified by a flow cytometry (Becton-Dickinson, San Jose, CA, USA).
Intracellular calcium assay
Intracellular Ca2+ concentrations were detected by the Rhod-2 AM assay (Thermo Fisher, Waltham, MA, USA). After cultured with HG, cells were stained with 10 μM Rhod-2 AM dye in the dark for 30 min at 37 °C and then washed twice, and mitochondrial Ca2+ levels were measured by using a fluorescent microscope with filter (Ex/Em = 552/581 nm) and a flow cytometry.
Permeability assay
Endothelial cell permeability was detected by using an In Vitro Vascular Permeability Assay kit following the manufacturer’s protocol (Millipore, Burlington, MA, USA). After cells formed a tighter monolayer and exposed to HG, they were stained with FITC-labeled BSA for 20 min, and the amount of FITC-BSA diffusion across endothelial monolayer was measured with a fluorescent microscope with a FITC filter and quantified by a fluorescence plate reader.
Apoptosis assay
To quantify the apoptosis of HRGECs, cells were stained with Annexin V and propidium iodide (PI) using an Annexin V- FITC apoptosis detection kit (BD Biosciences, Franklin Lakes, NJ, USA) and analyzed by flow cytometry.
Transwell migration assay
To determine the migration of HRGECs, Matrigel transwell culture plates with 4 μm pore size inserts were used. After cultured with HG, 2 × 105 cells were loaded onto the upper chambers of transwell culture plates with FBS-free ECM, which were suspended over the lower wells with 5% FBS-containing ECM. After 24 h incubation, migrated cells in the upper chambers were stained with 1% crystal violet and then counted.
Statistical analysis
All experiments were conducted at least thrice. Data are expressed as the mean ± standard deviation (SD). Differences between the treatment groups were analyzed by one-way ANOVA followed by Tukey’s post hoc test and Student’s t test using Prism (version 5, GraphPad). A value P < 0.05 was considered statistically significant.
Discussion
In this study, we explored the role of S1PR2 and its downstream signal molecules in the morphology and function of mitochondria in primary HRGECs. Our results demonstrated that the S1PR2 antagonist (JTE-013) almost completely blocked endothelial dysfunction and associated fission and dysfunction of mitochondria under HG conditions along with down-regulated HG-promoted expression of RhoA/ROCK1. Our data also indicate that ROCK1 is a critical step that links S1PR2 and mitochondrial fission and dysfunction through regulating Drp1. Therefore, we collected direct evidence for the mechanistic association between S1PR2 and cell dysfunction via the activation of the RhoA/ROCK1/Drp1 signaling pathway in HRGECs.
Vascular complications are the initial factor of mortality associated with type 2 diabetes mellitus [
27]. Diabetic nephropathy, a microvascular complication,, is the most common cause of end-stage renal disease in developed countries [
28]. Extensive studies have confirmed that vascular diseases usually start with endothelial cell damage [
29]. We used the primary HRGECs for the first time to explore the potentially pathological role of S1PR2 in the dysfunction of primary renal endothelium cells. We had previously observed that S1PR2 displays a positive regulation of human umbilical vein endothelial cell dysfunctions induced by hyperglycemia [
30]. Here, we further demonstrated that S1PR2 is important for regulating HG-induced dysfunction in HRGECs, as indicated by the use of JTE-013, which reversed the inducing effects of HG on the barrier function, migration, and cell death of HRGECs. Small GTPases (Rho, Rac and Ras) are known downstream signaling effectors of S1PR2 [
6‐
8]. We confirmed that S1PR2 inhibition blocks HG-induced upregulation of the expression of the small GTPase RhoA and its downstream ROCK1, indicating that S1PR2 regulates the dysfunctions of primary HRGECs through activating the RhoA/ROCK1 pathway.
Although mitochondria represent only 5–10% of cytoplasmic volume of endothelial cells [
31], they are the most important organelles for cell survival [
32]. Balanced mitochondria dynamics, ATP production, ROS synthesis, and local Ca
2+ concentrations are all indispensable for healthy endothelial function [
14,
33]. The present study shows the reversal of HG-mediated mitochondria fission and dysfunction by S1PR2 inhibition, suggesting that S1PR2 is responsible for the regulation of endothelial mitochondria morphology and function. Previous work has shown that the balance between mitochondrial fission and fusion, in which Drp1is a crucial player, holds the key to maintain mitochondrial morphology and function [
34,
35]. As the RhoA/ROCK1 is the downstream pathway of S1PR2 in HRGECs and Drp1 a direct substrate of ROCK1 in podocytes contributing to the enhanced mitochondrial fission [
36], we hypothesized that ROCK1 is the key regulator linking S1PR2 and mitochondrial fission in HRGECs. As expected, inhibition of ROCK1 downregulated the expression of Drp1 and consequently improved HG-induced mitochondrial morphology and biochemical changes in HRGECs.
Conclusions
Overall, our study provides the first evidence that S1PR2 exerts multiple effects on the fission and dysfunction of mitochondria induced by HG in HRGECs, which in turn contribute to endothelial dysfunction. Additionally, ROCK1 is a key regulator that functionally links S1PR2 and mitochondrial fission and dysfunction through activating Drp1. Although the exact mechanisms behind the ROCK1/Drp1 regulation by S1PR2 remind further investigation, we support the concept of S1PR2-mediated morphology and function of mitochondria of HRGECs induced by HG through the RhoA/ROCK1/Drp1 signaling pathway, suggesting that targeting this S1PR2-associated signaling pathway might be a potential therapeutic target for renal vascular disease treatment.
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