The primary HUVEC cell line was purchased from the American Type Culture Collection (ATCC, Manassas, VA) and cultured in a cell incubator at a temperature of 37°C and a CO₂ concentration of 5%. Microgravity conditions were simulated using a 2-D clinostat (Supplementary Figure S1). The direction changes through continuous rotation deprive the cells of the capacity to perceive gravity. Similar changes in the gene expression and cell morphology have been observed under clinorotation as compared to real microgravity (Herranz et al., 2013), suggesting the validity of this system for simulating the effects of microgravity. In this study, HUVECs were seeded on a 2.55 × 2.15 cm coverslip at a density of 1 × 10⁵ cells. Each coverslip was then placed in each well of 6-well plates. When the cells reached a confluence of 60%, the coverslips were placed into the fixture of the chambers (Astronaut Research and Training Center, Beijing, China), which were subsequently filled with a complete cell culture medium, removing air bubbles to avoid the effects of shear stress. Then, the chambers were rotated around the horizontal axis at a speed of 24 r/min before the cells were harvested (Supplementary Figure S1D). The control group was rotated around the vertical axis at the same speed (Supplementary Figure S1E).

Special reagents used in this study were purchased as indicated: mdivi-1(338967-87-6, TargetMol, Boston, Massachusetts, USA), MG132 (133407-82-6, Apexbio, Suzhou, China), NAC (616-91-1, Apexbio), human IL-1β neutralizing antibody (mabg-hil1b-3, InvivoGen, San Diego, CA, United State), MitoTEMPO (ab144644, Abcam, Cambridge, United Kingdom), and 2-APB (ab120124, Abcam).

Cell mitochondrial extraction kits (C3601, Beyotime, Shanghai, China) were used for mitochondrial isolation of HUVECs according to the manufacturer’s instructions. The mitochondria and cytoplasm were isolated through differential centrifugation and kept in a storage solution with PMSF (1mM, ST506, and Beyotime) for Western blot analysis.

The cells were digested and centrifuged after clinorotation for 48 h. Then, the cell precipitate was fixed in 2.5% glutaraldehyde for two hours at 4°C and post-fixed with osmic acid, dehydrated, embedded, and cut. After double staining with lead citrate and uranyl acetate, the images were acquired using a TEM system (HT7800, Hitachi, Japan).

The cells on the coverslips after clinorotation were incubated in media containing 250 nmol/L MitoTracker Deep Red FM (40743ES50, Yeasen, Shanghai, China) for 30 min at 37°C and washed with pre-warmed PBS. The cells were then incubated in media containing 50 nmol/L LysoTracker Green DND-26(40738ES50, Yeasen) for 1 h in the cell incubator and observed under a confocal microscope (LSM 800, Zeiss, Oberkochen, Germany) to determine the red and green fluorescence.

Knockdown of PINK1, MMP1, NLRP3, and HSF1 was performed by reverse transfection of PINK1 siRNA (GenePharma, Shanghai, China), MMP1 siRNA (GenePharma), NLRP3 siRNA (GenePharma), and HSF1 siRNA (GenePharma). RBM3-overexpression plasmid pGV141-RBM3 was purchased from GeneChem (Shanghai, China). The cells were placed into the chambers of a 2-D clinostat 24 h after being transfected with the abovementioned siRNA or plasmid by using Lipofectamine 2000 (11668-019, Invitrogen, Waltham, Massachusetts, United States).

The protein from lysates of HUVECs was separated using 12% sodium dodecyl sulphate polyacrylamide gel electrophoresis and transferred onto PVDF membranes (IPVH00010, Millipore, Billerica, MA, United States). After blocking in 4% milk, the membranes were incubated with primary antibodies overnight at 4°C. The primary antibodies used are as follows: Tom20 (1:2000 dilution, 11802-1-AP, Proteintech, Rosemont, USA), Tim23 (1:1000 dilution, 11123-1-AP, Proteintech), PINK1(1:1000 dilution, ab137361, Abcam), Parkin (1:1000 dilution, 14060-1-AP, Proteintech), Drp1 (1:1000 dilution, 12957-1-AP, Proteintech), Mfn2 (1:1000 dilution, 12186-1-AP, Proteintech), p62 (1:1000 dilution, 18420-1-AP, Proteintech), caspase 1(1:1000 dilution, ab179515, Abcam), ZO-1 (1:1000 dilution, 20742-1-AP, Proteintech), occludin (1:1000 dilution, 13409-1-AP, Proteintech), NLRP3 (1:1000 dilution, 19771-1-AP, Proteintech), IL-1β (1:1000 dilution, ab2105, Abcam), MMP1 (1:1000 dilution, 10371-2-AP, Proteintech), RBM3 (1:1000 dilution, 14363-1-AP, Proteintech), Hsp70 (1:1000 dilution, 10995-1-AP, Proteintech), Hsp90 (1:5000 dilution, 60318-1-Ig, Proteintech), clusterin (1:1000 dilution, 12289-1-AP, Proteintech), COXIV (1:1000 dilution, 11242-1-AP, Proteintech), and GAPDH (1:2000 dilution, EK020, Zhuangzhi Biotech, Xi’an, China). The membranes were subsequently washed and incubated with HRP-conjugated secondary antibody (1:5000 dilution, EK020, Zhuangzhi Biotech) for 1 h at room temperature. The proteins were detected using an ECL Western blot detection reagent (WBKLS0500, Millipore). The gray value was quantified using ImageJ software (1.31v, Bethesda, USA).

After clinorotation, the cells growing in coverslips were fixed in 4% paraformaldehyde for 15 min at room temperature and rinsed several times with PBS. Then, the cells on coverslips were permeabilized in PBS with 0.1% Triton X‐100 (9036-19-5, Sigma-Aldrich, St. Louis, MO, United States) for 15 min and blocked with goat serum for 1 h. The coverslips were incubated in Tom20 (1:100 dilution, 11802-1-AP, Proteintech), LC3 (1:100 dilution, ab192890, Abcam), PINK1 (1:100 dilution), and Hsp60 (1:100 dilution, 15282-1-AP, Proteintech) primary antibodies in blocking solution at 4°C overnight. Fluor 488 (1:500 dilution, EK011, Zhuangzhi Biotech) or Cy3 (1:500 dilution, EK022, Zhuangzhi Biotech) secondary antibodies were used for incubation for 1 h in the dark at room temperature. The cells were then treated with DAPI solution (1:1000 dilution) for 5 min in the dark, washed with PBS, and kept at 4°C until imaged. The cells were visualized using laser confocal microscopy (LSM 800, Zeiss). The quantification of colocalization was performed using ImageJ software.

RNA was extracted using TRIzol reagent (10296010, Thermo Fisher Scientific, Waltham, Massachusetts, United States), according to the manufacturer’s instructions. cDNA was generated using the PrimeScript II First-Strand cDNA Synthesis Kit (RR036A, Takara, Shiga, Japan) as described in the manufacturer’s protocol. Then, we performed quantitative PCR using SYBR Green reagent (RR420L, Takara) on a CFX96 Real‐Time System (Bio‐Rad, Hercules, CA). The sequences are listed in Table 1. The comparative CT method was used to calculate the relative amounts of mRNA.

Proteasome activity was analyzed using a proteasome activity assay kit (ab107921, Abcam), which included fluorescent substrates that were Succ-LLVY-AMC specific for the chymotrypsin-like activity. The cell lysate was prepared using 0.5% NP-40 in dH₂O. The supernatant was collected after centrifugation. Samples, standards, and positive control were seeded to a 96-well plate. The proteasome inhibitor MG132, positive control, and proteasome substrate were added to wells according to the manufacturer’s instructions and incubated for 10 min at 37°C. Fluorescence intensity was analyzed using a microplate reader (51119670DP, Thermo Fisher Scientific). The substrates were incubated for a further 30 min in the dark at 37°C and fluorescence intensity was analyzed again. The excitation and emission wavelengths were 350 and 440 nm, respectively. The standard curve was constructed, and the proteasome activity was calculated according to the manufacturer’s instructions.

SUnSET is a nonradioactive method to monitor protein synthesis (Schmidt et al., 2009). Briefly, the extent of puromycin incorporation into the nascent peptides is assessed using Western blot after the treatment with antibiotic puromycin, a tyrosyl-tRNA analog. The cells were removed from the wells of 6-well plates after clinorotation and incubated with fresh media containing puromycin (10 μg/ml, ab141453, Abcam) for 30 min. Subsequently, cellular protein lysates were obtained for Western blot analysis. The primary antibody was anti-puromycin (12D10, Millipore) at the dilution of 1:10000.

JC-1 dye formed aggregates inside the healthy mitochondrial matrix with red fluorescence while effluxes from the depolarized mitochondria to the cytoplasm acting as monomers with green fluorescence. After clinorotation, the cells on the coverslips were moved to wells of 6-well plates loaded with JC-1 dye (c2006, Beyotime) in the dark at 37°C for 20 min. The cells were washed and incubated in the assay buffer. The images were captured using laser confocal microscopy (LSM 800, Zeiss).

After clinorotation, the cells were washed with PBS and incubated in PBS containing 1 µM Rhod-2 AM (T19052, TargetMol) for 15 min at 37°C. Then, the cells were washed with PBS three times prior to incubation with PBS for 20 min at 37°C. The fluorescence images were captured using laser confocal microscopy (LSM 800, Zeiss). The excitation and emission wavelengths were 540 and 590 nm, respectively.

Human IL-1β in the cell culture medium was quantified using a human IL-1β ELISA kit (JL13662-48T, J&L Biological, Shanghai, China) according to the manufacturer’s instructions.

HUVECs seeded on transwell inserts with a filter of 3-μm pore (3492, Corning, New York, United States) were exposed to clinorotation for 48 h. The transwell inserts were moved into 6-well plates with 1.5 ml fresh media. 100μL FITC–dextran (70kDa, 60842-46-8, Seebio, Shanghai, China) solution was added to each insert, and the plate was incubated at 37°C for 2 h to allow fluorescein molecules to flow through the endothelial cell monolayer. The media below the inserts were added to 96-well plates. The fluorescent intensity in each well was determined at excitation/emission of 485/530 nm using a microplate reader (51119670DP, Thermo Fisher Scientific). The arbitrary fluorescence intensity was used to calculate the relative permeability.

Cellular ROS levels were detected using a DCFH-DA probe (ab113851, Abcam). Briefly, cells on the coverslips after clinorotation were washed with PBS three times before incubating with 5 μM DCFH-DA for 30 min at 37°C. After washing twice with a serum-free cell culture medium, we measured fluorescence using laser confocal microscopy (LSM 800, Zeiss). MitoSOX (40778ES50, Yeasen) was used to detect the mitochondrial ROS level in HUVECs. The cells were incubated in 5 μM MitoSOX for 30 min at 37°C, and positive staining was subsequently detected using fluorescence microscopy for imaging.

Transwell migration assay was performed using transwell inserts (3422, Corning) with a 8 μm pore filter. Cells exposed to clinorotation for 48 h were seeded into the upper chamber of the insert, and 600 μl of complete medium was added to the lower chamber. After 24 h incubation at 37°C, cells on the lower surface were fixed with methanol and stained with 0.2% crystal violet for 10 min. After washing with PBS three times, the cells were observed using an Olympus IX73 microscope.

Briefly, the coverslips on which the HUVECs grew after clinorotation were removed into a 6-well plate and grown to nearly 100% confluence. Linear scratches were made on confluent cell monolayers using a sterile 200-μl pipette tip. The cells were then incubated with serum-free medium. Images were taken at 0 and 24 h with an Olympus IX73 microscope, and the rate of wound healing was measured using ImageJ software.

Data derived from three independent experiments were shown as mean ± standard deviation. SPSS software (version 19.0, IL, United States) was used for one-way ANOVA or Student’s t-test whenever appropriate. In all experiments, the differences were considered significant when p < 0.05.

The mitochondria dynamic has been tied to mitochondrial quality control. Damaged or dysfunctional mitochondria undergo fragmentation from the healthy mitochondrial network. The fission of mitochondria is the requisite step for mitochondria to be accommodated within autophagosomes. Mitochondrial fission protein Drp1 and fusion protein Mfn1/Mfn2 mediate the structure alteration of mitochondria. We found that clinorotation for 24 and 48 h increased Drp1 protein expression and reduced the protein level of Mfn2 (Supplementary Figure S2A). To further investigate the altered mitochondrial dynamics, HUVECs were stained for the mitochondria marker Hsp60 before mitochondrial morphology was analyzed using confocal microscopy. As shown in Supplementary Figure S2B, confocal imaging revealed that mitochondria appeared fragmented after clinorotation for 48 h. Depolarization of mitochondria is known to induce their fragmentation (Legros et al., 2002). To determine whether mitochondrial depolarization occurred under clinorotation in HUVECs, the mitochondrial membrane potential in living cells was measured using the JC-1 probe and observed using confocal imaging. As shown in Supplementary Figure S2C, exposure to simulated microgravity for 48 h resulted in decreased membrane potential of mitochondria, which was reflected by the decreased ratio of red to green fluorescence.

Next, we explored the effect of simulated microgravity on the initiation of mitophagy. Western blot confirmed that the mitochondrial proteins Tom20, Tim23, and COXIV were decreased after 24 and 48 h clinorotation (Figure 1A). Furthermore, exposure to clinorotation upregulated the expression of the mitophagy marker PINK1 at 12 h, with a sustained increase up to 48 h (Figure 1A). LC3 colocalized with Tom20 in HUVECs under clinorotation for 48 h, and the colocalization of MitoTracker and LysoTracker significantly increased in HUVECs under clinorotation for 48 h (Figures 1B,C). To further validate the mitophagy in HUVECs under clinorotation, we explored the formation of mitophagosomes using TEM and found increased mitophagosomes and phagolysosomes in HUVECs after clinorotation for 48 h (Figure 1D).

PINK1 is degraded in the healthy mitochondrial matrix. The depolarization of mitochondria stabilizes PINK1 on the outer membrane of mitochondria, leading to Parkin recruitment. The mitochondrial outer membrane proteins such as Mfn1 and Mfn2 are ubiquitination substrates of the cytosolic ubiquitin ligase Parkin. The polyubiquitinated cargo recruits p62, ultimately leading to autophagosome formation coupled with mitochondria via LC3-interacting domain. Given that the expression of PINK1 was increased in response to clinorotation, we hypothesized that clinorotation triggers PINK1/Parkin-dependent mitophagy. To examine PINK1, Parkin, and p62 accumulation on mitochondria in HUVECs under clinorotation for 48 h, we investigated their expression in mitochondrial extracts. We found that PINK1, Parkin, and p62 were recruited to mitochondria under clinorotation (Figure 2A), demonstrating that the PINK1/Parkin pathway was activated. As expected, PINK1 notably colocalized with Tom20 in HUVECs under clinorotation for 48 h (Figure 2B), thereby confirming the recruitment of PINK1 to mitochondria. To further confirm the role of PINK1 in the mitophagy induction process, the mitophagy level was evaluated in cells that were transfected with PINK1 siRNA and then exposed to clinorotation for 48 h. As expected, gene silencing of PINK1 inhibited the clinorotation-induced downregulation of mitochondrial proteins Tom20, Tim23, and COXIV (Figure 2C), thus underscoring the role of PINK1 in the induction of mitophagy in HUVECs under clinorotation.

We have discovered that microgravity largely affected mitochondrial dynamics and the level of mitophagy. However, a detailed mechanism through which clinorotation induces mitophagy remains unclear. The ER has a great influence on mitochondrial morphology and function. Alterations in ER–mitochondria crosstalk and ER stress are reported to be closely related to the impairment of mitochondrial function, fission, and mitophagy (Wang et al., 2021). Ca²⁺ transfer is the vital crosstalk between the ER and mitochondria. To explore the influence of ER–mitochondria Ca²⁺ transfer on the level of mitophagy, we detected the level of Ca²⁺ in the mitochondria after clinorotation using the Rhod-2 AM probe, a mitochondrion-specific chemical Ca²⁺ indicator. The fluorescence signal of Rhod-2 AM rose after clinorotation, and the presence of 2-APB, which is classically used to inhibit IP₃R, dramatically decreased mitochondrial Ca²⁺ levels upon stimulation with simulated microgravity (Figure 3A), supposedly by blocking Ca²⁺ transfer and mitochondrial Ca²⁺ uptake.

It is reported that mitochondria exposed to Ca²⁺ overload exhibit reduced mitochondrial membrane potential, increased fission, and ROS production (Buhlman et al., 2014; Park et al., 2017). Less functional mitochondria activate the elimination of defective mitochondria by mitophagy. We determined whether excessive mitochondrial Ca²⁺ accumulation impacted the morphology, the membrane potential of mitochondria, and the level of mitophagy under clinorotation. As shown in Figures 3B and C, 2-APB ameliorated the decreased mitochondrial membrane potential and the fission of mitochondria in HUVECs under simulated microgravity. Next, we showed that 2-APB reversed the increased protein levels of p62, PINK1, and COXIV in mitochondrial fraction and the decreased protein level of Tom20 and Tim23 in the cell lysate under clinorotation (Figure 3D). These findings confirmed that ER–mitochondria Ca²⁺ transfer through the IP₃R channel is the leading cause of mitophagy induction.

Cellular stress, especially ER stress, has been shown to modulate the interaction between the ER and mitochondria, thus causing Ca²⁺ in ER to flow to mitochondria (Bravo et al., 2011). Our previous study has confirmed that clinorotation induces ER stress and UPR (Li et al., 2019). As the protein aggregate has been shown to induce ER stress and influence the ER–mitochondria Ca²⁺ crosstalk (Hedskog et al., 2013), we analyzed the impact of simulated microgravity on proteostasis in HUVECs. The accumulation of unfolding proteins due to cytoplasmic proteotoxic stimuli triggers transcriptional activation of HSF1, which promotes the expression of genes encoding chaperone proteins such as Hsp70, Hsp90, and clusterin. Chaperone proteins maintain protein homeostasis by stabilizing protein structures and assisting in the correct folding and unfolding of proteins (Neef et al., 2011). Exposure to clinorotation for 12, 24, and 48 h increased the expression of Hsp70, Hsp90, and clusterin at both mRNA and protein levels in HUVECs (Figures 4A–D). The effect of clinorotation on the expression of Hsp70, Hsp90, and clusterin was abolished by HSF1 knockdown (Figure 4E). The heat shock response suggested proteotoxic stress under clinorotation.

Apart from the activation of chaperones, eukaryotic cells have several other systems to maintain proteostasis, such as ER-associated degradation, autophagy, and impaired protein translation (Su et al., 2016; Baumgartner et al., 2021). Our previous study found that clinorotation-induced autophagy promotes the clearance of ubiquitinated protein aggregates, thus protecting cells against apoptosis (Li et al., 2019). The results in this study indicated that clinorotation for 12, 24, and 48 h increased proteasome activity (Figure 5A). The restraint of the protein synthesis rate after clinorotation for 12, 24, and 48 h was verified by fewer proteins labeled with puromycin using the SUnSET assay (Figure 5B). The cold stress-induced protein RBM3 enhances global levels of protein synthesis at colder temperatures (Dresios et al., 2005). We found that HUVECs exposed to clinorotation for 12, 24, and 48 h displayed lower expression of RBM3 (Figure 5B). To shed light on the role of RBM3 in protein synthesis under simulated microgravity, we used RBM3 overexpression plasmid and found that the slower protein synthesis rate was rescued by RBM3 overexpression under clinorotation (Figure 5C). Moreover, we treated HUVECs with MG132, a proteasome inhibitor that is known to induce intracellular accumulation of misfolded proteins. MG132 significantly reduced the level of RBM3 protein and protein synthesis rate (Figure 5D). These results uncovered the RBM3-mediated restraint of protein synthesis rate under clinorotation. Collectively, coordinated regulation of protein synthesis and degradation along with enhanced autophagy level revealed the proteotoxic stress after clinostat-simulated microgravity.

A few studies have indicated that PINK1 and Parkin are also activated upon accumulation of misfolded proteins in the mitochondrial lumen (Burbulla et al., 2014; Fiesel et al., 2017). Here, we used mitochondrial chaperone Hsp60 as the mtUPR biomarker (Keerthiga et al., 2021; Zhou et al., 2021) and found that the simulated microgravity did not show a much significant effect on the mRNA expression of Hsp60 (Supplementary Figure S3). The result presented here supported that clinorotation activated PINK1 and induced mitophagy independently of mitochondrial-unfolded protein response.

Overall, we concluded that ER stress and UPR due to the imbalance of protein homeostasis upregulated the transfer of Ca²⁺ from the ER to mitochondria through the IP₃R under clinorotation. The augmented ER-to-mitochondria Ca²⁺ transfer caused a collapse in mitochondrial membrane potential, fission of mitochondria, and mitophagy.

Mitochondrial ROS are the byproduct of the electron transport chain, and the depolarized and damaged mitochondria lead to the overproduction of ROS (Ma et al., 2018). Based on the strict connection between altered mitochondrial function and mitochondrial ROS, we explored whether ROS production was increased under clinorotation in HUVECs using the DCFH-DA assay. HUVECs under clinorotation exhibited a significant increase in ROS, rescued by NAC, a specific scavenger of ROS (Figure 6A). Mitochondria are an important source of ROS, so HUVECs were incubated in MitoSOX to label mitochondrial superoxide. Treatment with MitoTEMPO, the mitochondria-targeted antioxidant, reduced ROS and mtROS levels under simulated microgravity (Figures 6B,C), signifying mtROS as the main source of ROS under clinorotation. In addition, 2-APB reversed the elevated mtROS level under clinorotation (Figure 6D), suggesting that ER–mitochondria Ca²⁺ transfer was the leading cause of mtROS production.

ROS are well-known promoters of the NLRP3 inflammasome in various cells. Recently, it was reported that simulated microgravity induces NLRP3 inflammasome activation (Jiang et al., 2020). To determine whether mitochondrial ROS are the trigger of the NLRP3 inflammasome in HUVECs after simulated microgravity, the protein levels of the markers of inflammasome such as NLRP3, mature IL-1β, and cleaved caspase 1 were determined. We found increased expression of the inflammatory factors NLRP3, mature IL-1β, and cleaved caspase 1, and the increased release of IL-1β was confirmed by ELISA analysis in HUVECs under clinorotation for 48 h (Figures 6E,F). Treatment with NAC or MitoTEMPO obviously diminished the activation of the NLRP3 inflammasome, indicating that the NLRP3 inflammasome was mtROS-dependent (Figures 6E,G).

Mitochondrial quality control such as mitophagy is protective against oxidative injury in many diseases (Haslip et al., 2015; Zhou et al., 2021). Evidence is accruing that the removal of mitochondria via mitophagy prevents apoptosis by reducing mitochondria-derived ROS and subsequent activation of the NLRP3 inflammasome (Chen et al., 2019; Lin et al., 2019). We aimed at assessing whether mitophagy induced by clinorotation ameliorated NLRP3 inflammasome activation via controlling mtROS production. The mdivi-1 was used as the specific inhibitor of the mitochondrial fission and mitophagy. The effects of mdivi-1 and PINK1 knockdown on the NLRP3 inflammasome were explored using Western blot and ELISA assays. The presence of mdivi-1 under clinorotation made the expression of NLRP3, IL-1β p17, and caspase 1 p20 higher than that of the MG group (Figure 7A). Similarly, PINK1 knockdown exacerbated the activation of the NLRP3 inflammasome (Figure 7B).

To demonstrate that the inhibition of NLRP3 inflammasome by mitophagy is linked to mitochondrial ROS production, we investigated the effect of MitoTEMPO treatment on the NLRP3 inflammasome. As shown in Supplementary Figure S4A, the mitochondrial ROS level was markedly increased when PINK1 was knocked down by siRNA under simulated microgravity. As expected, the increased mtROS level and activation of the NLRP3 inflammasome induced by clinorotation were abolished by the presence of MitoTEMPO (Supplementary Figure S4B). Taken together, these results suggested that mitophagy inhibited NLRP3 inflammasome activation by reducing mitochondria-derived ROS production.

The vascular endothelium barrier contributes to vascular homeostasis. Endothelium alteration by the reduced level of tight-junction proteins induces endothelial hyperpermeability, which is the pathogenesis of many human diseases. Many stimuli induce changes in endothelial permeability via ROS and the NLRP3 inflammasome (Boini et al., 2017; Zhang et al., 2019; Zhou et al., 2019). To investigate whether ROS and the NLRP3 inflammasome induced by clinorotation cause the disassembly of tight-junction proteins, we examined the expression of the representative tight-junction proteins ZO-1 and occludin. As shown in Figure 8A, simulated microgravity decreased the expression of ZO-1 and occludin. Moreover, dextran flux significantly increased in HUVECs under clinorotation for 48 h (Figure 8B). Consistently, our confocal analysis showed that clinorotation markedly decreased the expression of the tight-junction protein ZO-1 on endothelial cell monolayers (Figure 8C). This clinorotation-induced increase in EC permeability was markedly reduced in the presence of NAC or NLRP3 siRNA transfection (Figure 8D). Next, we detected the role of mitophagy in EC permeability under clinorotation, given that mitophagy ameliorated ROS production and NLRP3 inflammation activation. It was found that PINK1 knockdown aggravated the inter-endothelial junction disruption (Figures 8D–F). Therefore, our results indicated that mitophagy attenuated endothelial hyperpermeability induced by clinorotation.

The integrity of endothelial cells is regulated by MMPs, which are destructive to adhesion and tight-junction structures by degrading tight-junction proteins (Pan et al., 2021). To determine the influence of the MMP family on the permeability of HUVECs, the expressions of MMP1, MMP2, and MMP9 were explored. Clinorotation increased the mRNA and protein expression of MMP1 but had no effect on mRNA levels of MMP2 and MMP9 (Figures 9A–C). In addition, HUVECs treated with PINK1 siRNA had significantly increased MMP1 compared to that of the control group (Figure 9D), indicating that mitophagy inhibited MMP1 expression. Pro-inflammatory cytokine IL-1β is known to promote MMP expression (Kaden et al., 2003; Chen et al., 2018). We hypothesized that mitophagy could suppress MMP1 expression by inhibiting the activation of the NLRP3 inflammasome and IL-1β. To verify our conjecture, HUVECs were transfected with NLRP3 siRNA or subjected to IL-1β neutralizing antibody under clinorotation. As shown in Figure 9E, the presence of IL-1β neutralizing antibody or NLRP3 gene silencing prevented the changes in protein levels of MMP1, ZO-1, and occludin induced by clinorotation, suggesting that increased IL-1β release after NLRP3 inflammasome formation contributed to increased expression of MMP1 and disassembly of tight-junction proteins ZO-1 and occludin. To provide direct evidence for the involvement of MMP1 in the disrupted endothelial tight junction, we examined the effect of MMP1 siRNA on the protein level of ZO-1 and occludin. It is interesting to note that MMP1 knockdown mitigated the decrease of occludin expression under clinorotation, whereas it failed to rescue downregulated ZO-1 expression (Figure 9F). These data suggested that the effects of IL-1β on tight-junction proteins ZO-1 and occludin may be mediated by other unknown ways apart from MMP1.

In addition to endothelium barrier function, cell migration is a key aspect of endothelial cells which could be regulated by MMP1 (Li et al., 2018b). Transwell migration assay and wound healing assay were performed to assess the cellular ability of migration in HUVECs transfected with siRNA-NLRP3, siRNA-MMP1, or siRNA-PINK1 under simulated microgravity. Genetic deletion of MMP1 or NLRP3 reversed the increased cellular migration, whereas PINK1 knockdown under clinorotation enhanced the cellular migration compared to the MG group (Figures 9G,H). These results confirmed that mitophagy negatively regulated cellular migration via the NLRP3-MMP1 pathway.