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Bortezomib induces Rho-dependent hyperpermeability of endothelial cells synergistically with inflammatory mediators

BMC Pulmonary Medicine volume 24, Article number: 617 (2024) Cite this article

Abstract

Background

Bortezomib (BTZ), a selective 26 S proteasome inhibitor, is clinically useful in treating multiple myeloma and mantle cell lymphoma. BTZ exerts its antitumor effect by suppressing nuclear factor-B in myeloma cells, promoting endothelial cell apoptosis, and inhibiting angiogenesis. Despite its success, pulmonary complications, such as capillary leak syndrome of the vascular hyperpermeability type, were reported prior to its approval. Although the incidence of these complications has decreased with the use of steroids, the underlying mechanism remains unclear. This study aims to investigate how BTZ influences endothelial cell permeability.

Methods

We examined the impact of BTZ on vascular endothelial cells, focusing on its effects on RhoA and RhoC proteins. Stress fiber formation, a known indicator of increased permeability, was assessed through the Rho/ROCK pathway.

Results

BTZ was found to elevate the protein levels of RhoA and RhoC in vascular endothelial cells, leading to stress fiber formation via the Rho/ROCK pathway. This process resulted in enhanced vascular permeability in a Rho-dependent manner. Furthermore, the stress fiber formation induced by BTZ had synergistic effects with the inflammatory mediator histamine.

Conclusions

Our findings suggest that BTZ accumulates RhoA and RhoC proteins in endothelial cells, amplifying the inflammatory mediator-induced increase in the active GTP-bound state of Rho, thereby exaggerating vascular permeability during pulmonary inflammation. This study provides novel insights into the molecular mechanism underlying the pulmonary complications of BTZ, suggesting that BTZ may enhance inflammatory responses in pulmonary endothelial cells by increasing RhoA and RhoC protein levels.

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Background

Bortezomib (BTZ) is a reversible proteasome inhibitor that selectively blocks 26 S chymotrypsin-like activity of the proteasome, causing functional derangement and apoptosis in malignant cells. The Food and Drug Administration has approved BTZ usage for treating multiple myeloma and mantle cell lymphomas. The antitumor effect of BTZ strongly inhibits nuclear factor-kB (NF-κB) activity in myeloma cells and suppresses tumor cell proliferation and apoptosis [1, 2]. Further, BTZ acts on endothelial cells, inducing endothelial cell apoptosis, inhibiting angiogenesis, and suppressing endothelial–tumor cell adhesion, thereby enhancing its antitumor effects [3,4,5].

Dose-limiting adverse events, such as decreased platelet counts and peripheral neuropathy, occur with BTZ administration. Adverse effects of BTZ also include fever and pneumonitis. Therefore, elucidating the mechanisms underlying these adverse effects and overcoming them are important clinical issues. BTZ remains the first-line drug for multiple myeloma treatment and is used in combination with lenalidomide and dexamethasone. In recent years, pulmonary complications of BTZ have rarely been reported compared to hematologic and neurotoxic side effects. However, prior to treatment approval, BTZ had caused numerous pulmonary complications, including fatalities [6,7,8,9]. Although the reason for the recent decrease in the frequency of BTZ-induced pneumonitis remains unclear, concomitant use of steroids reportedly reduces the frequency of BTZ-induced pneumonitis [10]. Therefore, steroid recommendations after treatment approval may have decreased the incidence of BTZ-induced pneumonitis. BTZ-induced pneumonitis exhibits different patterns of pathogenesis from conventional drug-induced interstitial pneumonitis [6, 11]. Pulmonary involvement, characterized by fever and systemic inflammation, is observed in BTZ-induced pneumonitis, accompanied by capillary leak syndrome-like hypoxemia without concomitant imaging abnormalities. However, the mechanism underlying BTZ-induced pneumonitis remains unclear.

Vascular permeability is dynamically regulated by the intercellular adhesion of endothelial cells [12, 13]. Vascular endothelial (VE)-cadherin, a cell adhesion molecule specifically expressed on endothelial cells, restricts vascular permeability by constituting endothelial adherence junctions (AJs). VE-cadherin forms cis-dimers on the cell surface and contributes to intercellular adhesion by trans-linking the cis-dimers of neighboring cells [13]. In normal tissues, endothelial cells strengthen the VE-cadherin-mediated intercellular adhesion, thereby maintaining low vascular permeability. However, once inflammation is induced, inflammatory mediators and cytokines instantly weaken the VE-cadherin-mediated endothelial cell adhesion, increasing vascular permeability [14, 15]. Notably, the lungs are a highly vulnerable organ in which pulmonary inflammation induces vascular hyperpermeability, leading to severe pathological conditions, such as acute respiratory distress syndrome, pulmonary edema, and capillary leak syndrome.

VE-cadherin forms two types of AJs: stable linear and focal AJs. Linear AJs are supported by circumferential actin bundles aligned along endothelial cell–cell adhesions, stabilizing junctional VE-cadherin to restrict endothelial permeability [16]. However, during inflammation, inflammatory mediators induce the formation of focal AJs, where actin stress fibers perpendicularly bind VE-cadherin-based cell–cell adhesions and exert a traction force to generate zipper-like structures, thereby increasing vascular permeability. The Rho subfamily members of Rho GTPases, RhoA, RhoB, and RhoC, play key roles in forming actin stress fibers connected to focal AJs [17]. Rho cycles between the active GTP-bound and the inactive GDP-bound forms through processes mediated by guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs), respectively. Inflammatory mediators stimulate the exchange of GTP for GDP on Rho to generate the activated form, thereby inducing actin stress fiber formation through the activation of Rho-associated coiled-coil-forming protein kinase (ROCK) to increase vascular permeability [18, 19].

Here, we investigated the molecular mechanisms underlying BTZ-induced pneumonitis by focusing on its effect on endothelial cell permeability. We found that BTZ induces the accumulation of RhoA and RhoC proteins, stimulating ROCK-dependent formation of actin stress fibers to disrupt VE-cadherin-mediated endothelial cell–cell junctions, thereby promoting vascular permeability. We further showed that the BTZ-induced accumulation of RhoA and RhoC enhanced the disruption of endothelial cell–cell junctions induced by inflammatory mediators.

Methods

Cell culture

Human umbilical vein endothelial cells (HUVECs), purchased from KURABO (KE-4109), were cultured in collagen-coated dishes at 37 °C in a 5% CO2 atmosphere, as described previously [20]. Humedia-EG2 medium (KURABO, KE-2170 S) was used to culture HUVECs. The cells were used for experiments before passage eight.

Materials

In this study, various primary antibodies were used, including mouse anti-vinculin (Sigma Aldrich V913, 1:400), rabbit anti-VE-cadherin (Cell Signaling #2500, 1:300), mouse anti-β-actin (Sigma Aldrich A2228, 1:3000), rabbit anti-RhoA (Cell Signaling #2117, 1:1000), mouse anti-RhoB (Santa Cruz sc-8048, 1:100), and anti-rabbit RhoC (Cell Signaling #3430, 1:1000) antibodies.

For the secondary antibody applications, we used Alexa Fluor 633 goat anti-mouse IgG (Thermo Fisher Scientific A21052, 1:500), Alexa Fluor 488 goat anti-rabbit IgG (Thermo Fisher Scientific A11008, 1:500), goat anti-mouse IgG-HRP (Southern Biotech 1030-05, 1:5000), and goat anti-rabbit IgG-HRP (Southern Biotech 4030-05, 1:5000).

We purchased other materials for this study from various sources. These include BTZ (Cayman CHEMICAL 10008822), Exoenzyme C3 transferase (cytoskeleton #CT03), Y-27,632 (FUJIFILM WAKO 253–00513), histamine (FUJIFILM WAKO 081-03551), tumor necrosis factor-α (TNF-α) (PeproTech, Inc. #300–01 A), FITC-dextran (R&D Systems 3475-096-0, 1:100), and Rhodamine Phalloidin (Thermo Fisher Scientific R415, 1:500).

Transwell permeability assay

HUVECs (2 × 105 cells/well) were seeded onto type-I collagen-coated upper inserts of Transwell chambers (pore size of polycarbonate filter: 3 μm, purchased from Corning) and cultured for 72 h. After 2 h of serum starvation in Medium 199 (Thermo Fisher Scientific) containing 0.5% AlbuMAX II (Thermo Fisher Scientific), the cells were pretreated with DMSO or 25 µM Y-27,632 for 0.5 h and subsequently exposed to 100 nM BTZ in the presence of DMSO or Y-27,632 for 6 h. After the incubation, permeability was measured by adding 1 µL of FITC-labeled dextran (1 mg/mL; molecular weight, 42,000) to the upper inserts containing 100 µL media. After incubation for 0.5 h, 500 µL of media containing FITC-labeled dextran passed through the HUVEC monolayers was collected from the lower compartment of the transwell chamber. The amount of FITC-labeled dextran was measured by measuring the fluorescence at 520 nm when excited at 492 nm using an Infinite 200 Pro microplate reader (Tecan).

Fluorescent immunocytochemistry

HUVECs were plated on a 35-mm glass-based dish (Iwaki, ASAHI GLASS Company, Ltd.) coated with Cellmatrix type I-C (Nitta Gelatin) at a density of 5 × 104 cells/dish. The following day, cells were treated with vehicle or varying concentrations of BTZ for the periods indicated in the Figure legends. To examine the effect of Y-27,632 and C3 exoenzyme on BTZ-induced stress fiber formation and disruption of endothelial cell–cell junctions, we pretreated the cells with 25 µM Y-27,632 and 1 µg/mL C3 exoenzyme for 0.5 h and 4 h, respectively, and subsequently exposed them to 100 nM BTZ for 6 h. To assess the synergistic effect of histamine with BTZ, we treated the cells first with 10 nM BTZ for 6 h and then stimulated with 10 µM histamine for 5 min.

After incubation, cells were fixed with phosphate-buffered saline (PBS) containing 2% formaldehyde for 15 min at room temperature (RT), permeabilized with 0.05% Triton X-100 in PBS for 30 min at RT, and blocked with 4% BSA in PBS for 1 h at RT. Then, the cells were stained with mouse anti-vinculin and rabbit anti-VE-cadherin antibodies at 4 °C overnight. After extensive washing with PBS, the proteins that reacted with the antibody were visualized using Alexa Fluor 633 goat anti-mouse IgG and Alexa Fluor 488 goat anti-rabbit IgG antibodies for 1 h at RT. To visualize filamentous actin, we simultaneously stained the cells with rhodamine–phalloidin for 1 h at RT. Fluorescence images of Alexa Fluor 488, Alexa Fluor 633, and rhodamine were recorded with an inverted fluorescence microscope (IX-83; Olympus) equipped with UPlanSApo 60x / 1.35 Oil ∞ / 0.17 / FN 26.5 objective lenses (Olympus) and a Zyla 4.2 PLUS sCMOS camera (Andor). The microscope and image acquisition were controlled by MetaMorph 7.8 software (Molecular Devices).

Quantification of cytoplasmic actin stress fiber and vinculin-labeled focal adhesions

To quantify the number of cytoplasmic actin stress fibers in HUVECs stained with rhodamine–phalloidin, anti-VE-cadherin, and anti-vinculin antibodies, we identified cell areas using the threshold methods of ImageJ. The areas of cell–cell adhesion were also identified by analyzing the VE-cadherin fluorescence signal using threshold methods. Non-cell and cell–cell junction areas were excluded from the background-subtracted rhodamine–phalloidin images to identify the cytoplasmic regions. The average pixel intensity of rhodamine–phalloidin fluorescence in the cytoplasmic region was defined as the number of cytoplasmic actin stress fibers. At least five images were analyzed per experiment, and the experiments were repeated at least four times.

To quantify the number of vinculin-labeled focal adhesions, fluorescence images of Alexa Fluor 633-labeled vinculin were analyzed using the ImageJ software according to a previously described protocol [21].

Immunoblotting

HUVECs were starved with Medium 199 containing 0.5% AlbuMAX II for 2 h and treated with 100 nM BTZ for 6 h. Cell lysates for immunoblotting were collected by lysing cells in sample buffer (240 mM Tris-HCl (pH 6.8) containing 8% SDS, 40% glycerol, and 0.01% BPB), separated by SDS-PAGE on 12% polyacrylamide gels, and then transferred onto PVDF membranes (Millipore). Membranes were blocked in blocking buffer [5% skim milk in Tris-buffered saline with Tween 20 (TBS-T)] for 1 h, probed with primary antibodies diluted in the blocking buffer overnight at 4 °C, and then reacted with HRP-labeled secondary antibodies at RT for 1 h. Immunocomplexes reacted with Immobilon Forte Western HRP substrate (Millipore) were visualized under a chemiluminescence detection system, Amersham™ ImageQuant™ 800 (IQ800). Densitometric analysis of band intensities was performed using ImageJ software.

qPCR analysis of inflammatory gene expression induced by BTZ

HUVECs (3 × 105 cells/well) were seeded on type-I collagen-coated 6-well dishes (Iwaki, ASAHI GLASS Company, Ltd.), cultured overnight, and serum-starved in Medium 199 containing 0.5% AlbuMAX II for 2 h., The cells were then treated with DMSO, 100 nM BTZ or 50 ng/ml TNF-α in the Medium 199 containing 0.5% AlbuMAX II for 6–12 h. After washing with ice-cold PBS, total RNA was extracted from the cells by TRIzol reagent (Thermo Fisher Scientific) in combination with Direct-zol RNA Microprep (Zymo Research). For each reaction, 0.5 µg of total RNA was reverse-transcribed to first-strand cDNA with DNase treatment using ReverTra Ace qPCR RT Master Mix with gDNA Remover (TOYOBO). Real-time quantitative PCR (qPCR) was performed using THUNDERBIRD Next SYBR™ qPCR Mix (TOYOBO) on a qPCR 96-well plate in a CFX96 machine (Bio-Rad) following the manufacturer’s instructions. The primers used for amplification were as follows: human 18 S-rRNA, 5-TGCGCCGCTAGAGGTGAAATTCC-3 and 5-CGCCGGTCGGCATCGTTTATG-3; human IL-1β, 5-CAGCCAATCTTCATTGCTCA-3 and 5-GAACCAGCATCTTCCTCAGC-3; human IL-6, 5-AGGAGACTTGCCTGGTGAAA-3 and 5-AAAGCTGCGCAGAATGAGAT-3; human IL-8, 5-tctggaccccaaggaaaac-3 and 5-ttctcagccctcttcaaaaact-3; human ICAM-1, 5-CCCTGTCAGTCCGGAAATAA-3 and 5-GATGACTTTTGAGGGGGACA-3. The obtained data were analyzed by ⊿⊿Ct method. For normalization, expression of human 18 S-rRNA was determined in parallel as an endogenous control.

Statistical analysis

All data shown in the figures represent means ± s.d. Statistical analyses were performed using GraphPad Prism 9 software (GraphPad Software Inc.) software. Statistical significance was determined by applying a two-tailed Student’s t-test for comparing two groups and a one-way ANOVA followed by Tukey test for comparing more than two groups. Data were considered statistically significant if the p-value was less than 0.05. p < 0.05, p < 0.01, and p < 0.001 are shown as *, **, and ***, respectively.

Results

BTZ increases vascular endothelial cell permeability by disrupting endothelial cell–cell junctions

We first performed an in vitro endothelial permeability assay using FITC-labeled dextran in HUVECs to evaluate the effect of BTZ on endothelial cell permeability. The permeability of FITC-labeled dextran across HUVEC monolayers significantly increased by treatment with 100 nM BTZ for 6 h (Fig. 1A), suggesting that BTZ disrupted endothelial cell–cell junctions. Therefore, we tested the effect of BTZ on VE-cadherin-mediated endothelial cell–cell adhesion and actin cytoskeleton organization using immunofluorescence staining. Treatment of confluent HUVECs with BTZ for 6 h increased cytoplasmic actin stress fibers and formed vinculin-labeled focal AJs in a concentration-dependent manner (Fig. 1B, C). Consistently, the number of punctate spots of vinculin localized in focal AJs significantly increased following BTZ treatment (Fig. 1B, D). We also treated HUVECs with 100 nM BTZ for different periods and found that the BTZ-induced formation of focal AJs was evident after 4 h of treatment (Supplementary Figure S1 in Additional File 1). These results indicate that BTZ promotes vascular permeability by disrupting VE-cadherin-mediated endothelial cell–cell junctions.

Fig. 1
figure 1

Bortezomib (BTZ) increases endothelial cell permeability by inducing actin stress fiber to disrupt cell–cell adhesions. (A) Endothelial cell permeability was assessed using a Transwell permeability assay with FITC-labeled dextran. HUVECs grown on Transwell filters were exposed to either vehicle or BTZ at a concentration of 100 nM for 6 h. Endothelial cell permeability was expressed as a fold increase compared to vehicle-treated cells. Data are means ± s.d. (n = 9). (B) HUVECs plated on collagen-coated dishes were treated with vehicle or varying concentrations of BTZ (10, 30, 100, and 300 nM) for 6 h, immunostained with anti-VE-cadherin and anti-vinculin antibodies, and then stained with rhodamine–phalloidin. Representative fluorescence images of F-actin (left panels, red), VE-cadherin (left middle panels, green), and vinculin (right middle panels, white) and their merged images (right panels) are shown at the top. The boxed regions are magnified on the right side. Scale bars, 200 μm. (C, D) The number of actin stress fibers (C) and vinculin-labeled focal adhesions (D), as observed in A, were quantified as described in the Materials and Methods. Data are means ± s.d (n = 9), expressed as fold increases compared to the vehicle-treated cells. In A, C, and D, *, p < 0.05; **, p < 0.01; ***, p < 0.001

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BTZ increases endothelial cell permeability via the Rho/ROCK pathway

Next, we aimed to elucidate the molecular mechanisms by which BTZ increases endothelial cell permeability. Inflammatory mediators increase endothelial cell permeability via the Rho/ROCK pathway. Hence, we investigated whether the Rho/ROCK pathway was involved in the BTZ-induced increase in endothelial cell permeability. Treatment with Y-27,632, an inhibitor of ROCK, completely abolished the BTZ-induced increase in endothelial cell permeability (Fig. 2A). BTZ-induced formation of actin stress fibers and increase in vinculin-labeled focal AJs were suppressed by treatment with Y-27,632 (Fig. 2B–D). We also examined the effect of the C3 exoenzyme, which inhibits Rho through ADP-ribosylation, on BTZ-induced disruption of endothelial cell–cell junctions. Inhibiting Rho by the C3 exoenzyme prevented the BTZ-induced formation of vinculin-labeled focal AJs (Fig. 3A–C). The C3 exoenzyme also tended to suppress the BTZ-induced formation of actin stress fibers. These findings suggest that the BTZ-induced increase in endothelial cell permeability occurs through the Rho/ROCK pathway-mediated disruption of endothelial cell–cell junctions.

Fig. 2
figure 2

ROCK inhibitor, Y27632, prevents BTZ-induced hyperpermeability by inhibiting stress fiber, disrupting endothelial cell–cell junctions. (A) Endothelial cell permeability was assessed as described in Fig. 1A. HUVECs were treated with vehicle or 100 nM BTZ in the absence or presence of 25 µM Y-27,632 for 6 h. Data are means ± s.d (n = 6). (B) HUVECs treated with vehicle or 100 nM BTZ in the absence or presence of 25 µM Y-27,632 for 6 h were immunostained with anti-VE-cadherin and anti-vinculin antibodies, followed by staining with rhodamine–phalloidin. Representative fluorescence images of F-actin (left panels, red), VE-cadherin (left middle panels, green), and vinculin (right middle panels, white) and their merged images (right panels) are shown at the top. The boxed regions are magnified on the right side. Scale bars, 200 μm. (C, D) The number of actin stress fibers (C) and vinculin-labeled focal adhesions (D), as observed in A, were quantified as described in the Materials and Methods. Data are means ± s.d. (n = 4), expressed as fold increases compared to the vehicle-treated cells. In A, C, and D, *, p < 0.05; **, p < 0.01; ***, p < 0.001

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Fig. 3
figure 3

C3 exoenzyme inhibits stress fiber formation to prevent disruption of endothelial cell–cell junctions. (A) HUVECs pretreated with the C3 exoenzyme for 4 h were exposed to vehicle or 100 nM BTZ for 6 h, immunostained with anti-VE-cadherin and anti-vinculin antibodies, and stained with rhodamine–phalloidin. Representative fluorescence images of F-actin (left panels, red), VE-cadherin (left middle panels, green), and vinculin (right middle panels, white) and their merged images (right panels) are shown at the top. The boxed regions are magnified on the right side. Scale bars, 200 μm. (B, C) The number of actin stress fibers (B) and vinculin-labeled focal adhesions (C), as observed in A, were quantified as described in the Materials and Methods. Data are means ± s.d. (n = 4), expressed as fold increases compared to the vehicle-treated cells. *, p < 0.05; **, p < 0.01; ***, p < 0.001

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BTZ causes accumulation of RhoA and RhoC proteins in endothelial cells

A previous study showed that BTZ induces the accumulation of RhoA protein through inhibition of the proteasomal degradation pathway in megakaryocytes, leading to impaired thrombosis [22]. Therefore, we investigated whether BTZ increased the levels of Rho proteins in endothelial cells by performing western blot analysis. The levels of RhoA and RhoC proteins in HUVECs significantly increased following BTZ treatment for 6 h (Fig. 4A, C). However, the protein level of RhoB in HUVECs was very low, even after treatment with BTZ (Fig. 4B). This result is consistent with a previous report showing that RhoB continuously undergoes lysosomal degradation, thereby maintaining low levels of RhoB protein in quiescent endothelial cells [23]. These data suggest that BTZ induces the accumulation of RhoA and RhoC proteins, likely disrupting VE-cadherin-mediated endothelial cell–cell junctions and increasing endothelial cell permeability.

Fig. 4
figure 4

BTZ increases protein levels of RhoA and RhoC in endothelial cells. (A–F) HUVECs were treated with vehicle or 100 nM BTZ for 6 h. Protein levels of RhoA (A), RhoB (B), and RhoC (C) were assessed using western blot analysis with anti-RhoA, anti-RhoB anti-RhoC, and anti-β-actin antibodies. Representative western blot images are shown at the top of the panel. Relative expression levels of RhoA, RhoB, and RhoC were quantified by normalizing their expression levels to those of β-actin. Data are expressed as fold increases compared to the vehicle-treated cells and shown as means ± s.d. (RhoA, n = 4; RhoB, n = 4; RhoC, n = 3). *, p < 0.05

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BTZ induces the disruption of endothelial cell–cell junctions independently of inflammatory gene expression

BTZ is known to suppress inflammation by preventing the proteasome-mediated degradation of the NF-κB inhibitor [1]. However, other studies have also reported that BTZ exerts a pro-inflammatory effect by inducing the expression of pro-inflammatory cytokines [11, 24, 25]. Therefore, we investigated whether BTZ induces the disruption of endothelial cell-cell junctions not only through the activation of the Rho-ROCK pathway but also by inducing the expression of inflammatory genes. To this end, we performed qPCR analysis on HUVECs treated with or without BTZ, using TNF-α as a positive control. Treatment with BTZ for 6–12 h did not significantly upregulate the expression of interleukin-1β (IL-1β), interleukin-6 (IL-6), interleukin-8 (IL-8), or intercellular adhesion molecule-1 (ICAM-1), whereas their expression was dramatically increased by stimulation with TNF-α (Supplementary Figure S2). These results indicate that inflammatory gene expression is not involved in the BTZ-induced disruption of endothelial cell-cell junctions.

BTZ exaggerates histamine-induced disruption of endothelial cell–cell junctions

Patients with BTZ-induced pneumonitis exhibit elevated levels of blood inflammatory markers [11, 24]. Thus, we hypothesized that BTZ might enhance inflammatory mediator-induced disruption of endothelial cell–cell junctions to increase vascular permeability, contributing to the development of pneumonitis. To test this hypothesis, we examined the effect of BTZ on histamine-induced disruption of endothelial cell–cell junctions as histamine induces vascular leakage through the Rho/ROCK pathway-mediated formation of focal AJs [18]. Stimulation with 10 µM histamine and treatment with 10 nM BTZ tended to induce actin stress fiber formation (Fig. 5A, B). However, we did not observe a significant synergistic effect of histamine and BTZ (Fig. 5A, B). Because the histamine-induced formation of vinculin-labeled focal AJs was more prominent than histamine-induced stress fiber formation, we examined the effect of BTZ on histamine-induced disruption of endothelial cell–cell junctions by measuring the number of vinculin-labeled focal AJs. Stimulation of HUVECs with 10 µM histamine for 5 min resulted in the formation of vinculin-labeled focal AJs (Fig. 5A, C). Treatment with 10 nM BTZ for 6 h also increased the number of vinculin-labeled focal AJs (Fig. 5A, C). However, a greater increase in vinculin-labeled focal AJs was observed when HUVECs pretreated with 10 nM BTZ for 6 h were subsequently stimulated with 10 µM histamine, revealing the synergistic effect of BTZ and histamine (Fig. 5A, C). These results suggest that BTZ potentiates the histamine-induced disruption of endothelial cell–cell junctions.

Fig. 5
figure 5

BTZ potentiates histamine-induced formation of stress fiber and disruption of endothelial cell–cell adhesions. (A) HUVECs treated with vehicle or 10 nM BTZ for 6 h were stimulated with 10 µM histamine for 5 min, immunostained with anti-VE-cadherin and anti-vinculin antibodies, and stained with rhodamine–phalloidin. Representative fluorescence images of F-actin (left panels, red), VE-cadherin (left middle panels, green), and vinculin (right middle panels, white) and their merged images (right panels) are shown at the top. The boxed regions are magnified on the right side. Scale bars, 200 μm. (B, C) The number of actin stress fibers (B) and vinculin-labeled focal adhesions (C), as observed in A, were quantified as described in the Materials and Methods. Data are means ± s.d.(n = 5), expressed as fold increases compared to the vehicle-treated control cells. *, p < 0.05; **, p < 0.01

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Discussion

In this study, we investigated the molecular mechanisms underlying BTZ-induced pneumonitis, focusing on its effects on endothelial cell permeability. Our data revealed that BTZ disrupted VE-cadherin-mediated endothelial cell–cell junctions, thereby promoting endothelial cell permeability. As the underlying mechanism, we showed that BTZ induces the accumulation of RhoA and RhoC proteins, which, in turn, promotes the ROCK-dependent formation of actin stress fibers connected to focal AJs to disrupt endothelial cell–cell junctions. Furthermore, our study indicates that BTZ exaggerates histamine-induced disruption of endothelial cell–cell junctions, suggesting that pulmonary inflammation may increase the susceptibility to the adverse effects of BTZ.

BTZ disrupts endothelial cell–cell junctions by accumulating RhoA and RhoC proteins. Given that BTZ is a proteasome inhibitor, our data suggest that it prevents the degradation of RhoA and RhoC, leading to a significant increase in their protein levels. How does the accumulation of RhoA and RhoC disrupt endothelial cell–cell junctions? The strength of G protein signaling is controlled by the balance between the GTP-bound active form and GDP-bound inactive form [15, 17]. GEFs increase the GTP-bound active forms of G proteins by stimulating the exchange of GDP for GTP. Conversely, GAPs promote the intrinsic GTPase activity of G proteins, leading to GTP hydrolysis and conversion of the GTP-bound active form to the GDP-bound inactive form. Hence, GEF and GAP activities determine the equilibrium between the GTP-bound active form and GDP-bound inactive form within cells. Therefore, a small population of G proteins possibly exists in the GTP-bound active form that moderately stimulates downstream signaling, even in unstimulated cells. This notion is supported by evidence that overexpression of wild-type RhoA can exert biological functions [26, 27]. Therefore, BTZ-induced accumulation of RhoA and RhoC proteins may increase the levels of GTP-bound active forms, thereby inducing downstream signaling that disrupts endothelial cell–cell junctions. However, further investigations are required to test this hypothesis.

BTZ may enhance inflammatory mediator-induced vascular permeability by potentiating the Rho-mediated disruption of endothelial cell–cell junctions. Our data showed that pretreatment with BTZ exaggerated the histamine-induced disruption of endothelial cell–cell junctions. Inflammatory mediators, such as histamine, thromboxane A2, and bradykinin, promote endothelial cell permeability by inducing Rho-mediated actin stress fiber formation [18, 19, 28]. These inflammatory mediators increase the amount of GTP-bound active form of Rho by stimulating the Rho GEFs through their cognate G protein-coupled receptors. Because BTZ treatment led to an increase in the protein levels of RhoA and RhoC, inflammatory mediators may strongly increase their GTP-bound active forms in BTZ-treated endothelial cells compared with those in untreated cells, thereby promoting vascular hyperpermeability through the disruption of endothelial cell–cell junctions. Therefore, pulmonary inflammation could potentially enhance susceptibility to BTZ-induced pneumonitis. In recent years, pulmonary complications caused by BTZ treatment have rarely been reported, although they frequently occur before drug treatment [6,7,8,9]. Because the combined use of BTZ and steroids has been recommended for the treatment of multiple myeloma, steroid-induced suppression of inflammation might decrease the incidence of BTZ-induced pneumonitis by preventing an increase in endothelial permeability.

BTZ-induced vascular permeability does not depend on the expression of inflammatory factors. BTZ suppresses NF‑κB signaling by preventing proteasome-dependent degradation of NF‑κB inhibitor, thereby inhibiting inflammation. Indeed, Stellari et al. reported that BTZ attenuates lipopolysaccharide-induced lung inflammation through inhibition of NF‑κB [29]. BTZ ameliorates inflammatory diseases, such as ulcerative colitis and periodontitis, in animal model [30, 31]. However, some clinical case reports showed that BTZ administration in patients with multiple myeloma caused vasculitis and fever through the production of pro-inflammatory cytokines, such as IL-6, TNF-α, and C-reactive protein [11, 24, 25]. Furthermore, Maruyama et al. showed that bone marrow stromal cells produce pro-inflammatory cytokines upon BTZ administration [25]. These results indicate that BTZ exerts both anti-inflammatory and pro-inflammatory effects in a context dependent manner. Nevertheless, BTZ induces vascular permeability independently of its pro-inflammatory effect, as it did not upregulate the expression of inflammatory genes in endothelial cells.

BTZ has been shown to directly act on endothelial cells in vivo. Roccaro et al. reported that BTZ inhibits angiogenesis in a chick embryo chorioallantoic membrane model [4]. Additionally, BTZ has been shown to downregulate ICAM-1 expression in both T-cell lymphoma and endothelial cells, thereby inhibiting the interaction between tumor cells and endothelial cells in vivo [32]. Furthermore, BTZ upregulates endothelial nitric oxide synthase in pulmonary endothelial cells in an animal model of pulmonary arterial hypertension, potentially alleviating its pathogenesi [33]. However, whether BTZ acts on alveolar endothelial cells to induce vascular permeability in vivo remains unexplored. Therefore, further studies are needed to confirm that BTZ induces Rho-dependent hyperpermeability of alveolar endothelial cells in vivo.

Conclusions

In conclusion, we demonstrated that BTZ induces the accumulation of RhoA and RhoC, leading to the disruption of endothelial cell–cell junctions to promote vascular permeability. We also showed that BTZ exaggerates inflammatory mediator-promoted vascular permeability by potentiating Rho-mediated disruption of endothelial cell–cell junctions. Thus, the inhibition of Rho signaling could reduce the risk of adverse effects of BTZ.

Data availability

All data generated or analyzed during this study are included in this published article and its supplementary information file.

Abbreviations

AJs:

Adherence junctions

BTZ:

Bortezomib

GAPs:

GTPase-activating proteins

GEFs:

Guanine nucleotide exchange factors

HUVECs:

Human umbilical vein endothelial cells

NF-κB:

Nuclear factor-kB

PBS:

Phosphate-buffered saline

RT:

Room temperature

ROCK:

Rho-associated coiled-coil-forming protein kinase

TBS-T:

Tris-buffered saline with Tween

VE-cadherin:

Vascular endothelial-cadherin

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Acknowledgements

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Funding

This study was supported by JSPS KAKENHI (grant number: JP19K17652 and 23K07661 to Takeru Kashiwada). The funder had no role in the conceptualization, design, data collection, analysis, decision to publish, or preparation of the manuscript.

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Authors and Affiliations

  1. Department of Pulmonary Medicine and Oncology, Graduate School of Medicine, Nippon Medical School, Tokyo, Japan

    Shunichi Nishima, Takeru Kashiwada, Yoshinobu Saito, Kuniko Matsuda, Koichiro Kamio, Masahiro Seike & Akihiko Gemma

  2. Department of Molecular Pathophysiology, Institute for Advanced Medical Sciences, Nippon Medical School, Tokyo, Japan

    Shinya Yuge, Tomohiro Ishii & Shigetomo Fukuhara

Authors
  1. Shunichi Nishima
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  2. Takeru Kashiwada
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Contributions

SN, TK, SY, TI, KM, and SF performed the experiments; SN, TK, YS, KM, KK, MS, SF, and AG conceived and designed the study; and SN and SF analyzed the data. TK and SF drafted the manuscript. SN, YS, SY, TI, KK, MS, and AG edited and revised the manuscript. All authors read and approved the final manuscript.

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Correspondence to Takeru Kashiwada or Shigetomo Fukuhara.

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Supplementary Material 1:

Figure S1. BTZ-induced disruption of endothelial cell–cell junctions after treatment for 4 h. HUVECs plated on collagen-coated dishes were treated with vehicle or 100 nM BTZ for 1, 2, 4, or 6 h, immunostained with anti-VE-cadherin and anti-vinculin antibodies, and then stained with rhodamine–phalloidin. Representative fluorescence images of F-actin (left panels, red), VE-cadherin (left middle panels, green), and vinculin (right middle panels, white) and their merged images (right panels) are shown at the top. The boxed regions are magnified on the right side. Scale bars, 200 μm

Supplementary Material 2:

Figure S2. BTZ does not induce the expression of inflammatory genes. HUVECs plated on collagen-coated plates were serum-starved and treated with DMSO (Vehicle), 100 nM BTZ, or 50 ng/ml TNF-α for 6 or 12 h. After stimulation, total RNA was extracted and analyzed by qPCR to determine the expression levels of IL-1β, IL-6, IL-8, and ICAM-1 as described in Methods. Bar graphs show relative mRNA levels normalized to that of 18S-rRNA. Each dot represents an individual sample. Data are means ± s.d. (n=3) and are expressed as fold increases compared to vehicle-treated cells. *, p < 0.05; **, p < 0.01; ***, p < 0.001

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Nishima, S., Kashiwada, T., Saito, Y. et al. Bortezomib induces Rho-dependent hyperpermeability of endothelial cells synergistically with inflammatory mediators. BMC Pulm Med 24, 617 (2024). https://doiorg.publicaciones.saludcastillayleon.es/10.1186/s12890-024-03387-x

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Keywords

BMC Pulmonary Medicine

ISSN: 1471-2466

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