USP7-mediated stabilization of CXCL3 aggravates inflammation in models of acute lung injury

Introduction

Sepsis, a deadly condition arising from an overwhelming host response to infections, poses a significant burden on intensive care units and global healthcare, with 1.7 million fatalities (1,2). This disorder progresses to multi-organ dysfunction in nearly 30% of cases, and the lungs are particularly susceptible, accounting for approximately 40% of sepsis-induced acute lung injury (ALI) cases (3,4). Sepsis-induced ALI has been attributed to various mechanisms, such as inflammation, immune reactions, cellular hypoxia, apoptosis, and oxidative stress (5). However, the exact mechanisms that lead to and exacerbate sepsis-induced ALI remain unclear. Currently, patients with sepsis-induced ALI rely primarily on antibiotics and supportive care, which show limited success in decreasing mortality, emphasizing the pressing need for novel and efficacious therapies.

CXCL3, a member of the CXC chemokine family, acts as a neutrophil attractant and undergoes alternative splicing to generate multiple transcript variants (6). CXCL3 affects the development and progression of diverse diseases, including inflammatory and autoimmune conditions and cancer. Its impact stems from modulating genes in the ERK pathway and G-protein activity, as well as in engaging with signaling cascades, such as β-arrestin and MAPK/ERK pathways (7,8). Emerging evidence has implicated CXCL3 in the progression of lung disease. It plays a pivotal role in pulmonary fibrosis by directing vascular restructuring and fibroblast migration and spread (9). Furthermore, CXCL3 acts as a key attractor for polymorphonuclear leukocytes during lung damage and stimulates p21-activated kinase phosphorylation (10). CXCL3 attracts CXCR2-expressing neutrophils and exacerbates rhinovirus-induced inflammation in asthma (11). It has also been identified as a differentially expressed gene in both mechanical ventilation and lipopolysaccharide (LPS)-induced ALI mouse models (12). However, its specific function and mechanism in sepsis-related ALI are yet to be elucidated.

Ubiquitination, a crucial post-translational modification, ensures cellular health and balance by managing protein degradation and is integral to gene expression and DNA repair (13). The ubiquitin-proteasome system, which targets key proteins, is central to this process. USP7, composed of 1,102 amino acids, is a deubiquitinating enzyme of the USP family (14). It regulates the stability of proteins involved in immunity, DNA damage repair, transcription, and epigenetic processes, thereby influencing a wide spectrum of cellular functions (15,16). USP7 not only stabilizes substrates but also removes monoubiquitination from transcription factors and Lys-63-linked polyubiquitin chains (17). However, the relationship between USP7 and CXCL3 in sepsis-induced ALI remains unclear.

Endothelial cells, specifically human lung microvascular endothelial cells (HLMVECs), play a crucial role in the pathophysiology of sepsis-induced ALI. They form the interface between the bloodstream and the lung parenchyma and are among the first cells to be exposed to circulating endotoxins such as LPS during sepsis. Activation of HLMVECs by LPS leads to a series of events, including the upregulation of adhesion molecules, release of pro-inflammatory cytokines, and promotion of leukocyte recruitment, all of which contribute to the development of ALI (18). Therefore, studying the inflammatory response of HLMVECs is essential for understanding the early events in sepsis-induced ALI. Macrophages, on the other hand, are key immune cells in the lung’s defense mechanism. In the context of sepsis, macrophages can polarize into different phenotypes. The M1-polarized macrophages are pro-inflammatory and produce high levels of cytokines such as interleukin-1β (IL-1β) and tumor necrosis factor-α (TNF-α), which can exacerbate lung injury. THP-1 cells, which can be differentiated into macrophages, provide a valuable in-vitro model to study macrophage polarization and its role in the inflammatory cascade (19). By assessing the inflammatory response of both HLMVECs and macrophages, we can gain a more comprehensive understanding of the complex network of events that lead to sepsis-induced ALI and the potential role of CXCL3 and USP7 in modulating this process. Therefore, this study examined the regulatory function of CXCL3 in LPS-induced HLMVECs and macrophages, and a sepsis-induced ALI mouse model, with the aim of establishing a link between CXCL3 and USP7 in this context. Overall, our findings identify potential therapeutic targets for sepsis-related ALI. We present this article in accordance with the ARRIVE and MDAR reporting checklists (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2024-2026/rc).

Methods

Participants and clinical samples

A total of 25 healthy individuals and 27 patients with sepsis-related ALI aged 40–65 years were included in this study. Participants contributed 4 mL fasting blood, which was refrigerated at 4 ℃ for plasma extraction. These patients who were being treated for bacterial infections were recruited from The Second Affiliated Hospital of Soochow University with the approval of the Ethics Committee of The Second Affiliated Hospital of Soochow University (approval number: JD-LK-2022-105-01). Patients with severe comorbidities or those treated prior to admission were excluded. The study was conducted in accordance with the Declaration of Helsinki and its subsequent amendments. All participants provided written informed consent.

Cell culture and transfection

HLMVECs (EK-Bioscience, Shanghai, China) were cultured in Dulbecco’s Modified Eagle’s Medium (DMEM; HuanKai Microbial, Guangzhou, China) enriched with endothelial growth factors and 5% fetal bovine serum (FBS; HuanKai Microbial) at 37 ℃ with 5% CO₂. Meanwhile, THP-1 macrophages (EK-Bioscience) were maintained in RPMI-1640 medium (Amyjet, Wuhan, China) supplemented with 10% FBS and 1% antibiotics (Sunncell, Wuhan, China) under standard conditions (37 ℃, 95% air, 5% CO₂). Both cell lines were passaged tri-weekly. For experiments, HLMVECs were treated with varying concentrations of LPS (0, 5, 10, and 15 µg/mL) (MedChemExpress, Princeton, NJ, USA), and THP-1 cells stimulated with 100 ng/mL LPS for 24 h. Heyuan Biotechnology (Shanghai, China) supplied small interfering RNAs (siRNAs) targeting CXCL3, STAT1-STAT4, USP7, and USP14, along with overexpression plasmids for STAT3, USP7, and CXCL3. The control materials included si-NC (negative control siRNA) and the pcDNA 3.1(+) vector. The specific target sequences used are listed in Table 1.

Transfection of HLMVECs and THP-1 cells at 70–80% confluency was performed using siRNAs, plasmids, and LipoFiter (Hanbio, Shanghai). After a 48-h transfection period, the cells were prepared for functional assays through stimulation with LPS. Following transfection with si-NC, si-USP7, vectors, or USP7 overexpression constructs, HLMVECs were treated with actinomycin D (cat. A9415; Sigma-Aldrich, St. Louis, MO, USA) and cycloheximide (CHX; MedChemExpress) to assess USP7 messenger RNA (mRNA) and protein stability, following established protocols (20). A proteasome inhibitor (MG132; MedChemExpress) was used to analyze USP7 ubiquitination following transfection with si-NC and si-USP7, as previously described (20).

Cell viability analysis

HLMVECs in logarithmic growth were seeded onto 96-well plates and allowed to adhere. Following the treatments detailed in the previous transfection and/or LPS-stimulation procedures, each well received fresh DMEM supplemented with Cell Counting Kit-8 (CCK-8; Beyotime, Shanghai, China). The plates were then incubated for 4 h in the dark, after which the optical density was determined using a microplate reader.

5-ethynyl-2'-deoxyuridine (EdU) assay

HLMVECs were plated into 96-well plates and cultivated at 37 ℃ under standard culture conditions (5% CO₂, 95% atmospheric air) until they were ready for the EdU incorporation assay. The EdU solution was prepared according to the instructions provided in the Cell Proliferation Detection Kit (RiboBio, Guangzhou, China). The cells were then exposed to freshly prepared EdU-containing medium, fixed with paraformaldehyde, and subjected to the click reaction in the dark. Thereafter, the cells were stained with DAPI. Images were captured from five distinct areas per well under a fluorescence microscope to visualize proliferating cells.

Flow cytometry analysis

Apoptosis in HLMVECs was assessed using an Annexin V-FITC/PI (propidium iodide) Staining Kit (BioVision, Milpitas, CA, USA). Briefly, HLMVECs in logarithmic growth were harvested through trypsinization in the absence of EDTA. Following the manufacturer’s protocol, both Annexin V-FITC and PI were introduced into the cell suspension. Following a 15-min incubation period in the dark, each sample was analyzed using flow cytometry to quantify the number of apoptotic cells.

To induce the differentiation of human monocyte cells (THP-1) into macrophages, 100 ng/mL phorbol 12-myristate 13-acetate (Sigma-Aldrich) was administered for 24 h, following the protocol outlined in a previous study (21). After differentiation, the cells were subjected to individual treatments and subsequently stimulated with 100 ng/mL LPS for an additional 24-h period. Thereafter, the cells were rinsed with ice-cold phosphate-buffered saline and probed with primary antibodies specific to the CD86 and CD11b markers. Finally, the antibody-labeled cells were analyzed via flow cytometry to assess macrophage phenotype expression.

Superoxide dismutase (SOD) and malondialdehyde (MDA) measurements

Following the specified transfection and/or LPS-stimulation procedures, HLMVECs and frozen lung tissue samples were lysed in a suitable lysis buffer. The lysates were then centrifuged at 8,000 ×g for 10 min to separate the soluble fractions. The resultant supernatants were carefully collected to assess SOD activity and MDA content. These biochemical analyses were performed in strict adherence to the protocols provided by the SOD Activity Assay Kit (Solarbio, Beijing, China) and MDA Content Assay Kit (Solarbio).

RNA preparation and quantitation analysis

For the extraction of RNA from patient plasma, 4 mL of fasting blood samples were first centrifuged at 2,000 ×g for 10 min at 4 ℃. This step was crucial to separate the plasma from the cellular components. The supernatant, which was the plasma, was carefully collected. RNA was then isolated from the plasma using RNAzol (IGene Biotech, Guangzhou, China) following the manufacturer’s protocol. Genomic DNA contamination was removed with DNase I treatment prior to cDNA synthesis using the First-Strand cDNA Synthesis Kit (GeneCopoeia, Guangzhou, China). Quantitative real-time polymerase chain reaction (qRT-PCR) was subsequently performed using SYBR Green RT-qPCR Master Mix (GeneCopoeia) to analyze gene expression levels. For the differential expression analysis, RNA was isolated from LPS-treated and control HLMVECs using RNAzol (IGene Biotech) as described above. After ensuring RNA quality and quantity, the samples were subjected to microarray analysis. The microarray platform used was Affymetrix Human Genome U133 Plus 2.0 Array. The raw data obtained from the microarray were pre-processed using the Robust Multi-array Average algorithm in Bioconductor. Normalization was carried out to account for differences in hybridization and signal intensity across arrays. Differentially expressed genes were identified using the Limma package in R with a significance threshold set at P<0.001 and an absolute log-fold change (|logFC|) >4. This approach allowed us to detect genes that were significantly up-or down-regulated in LPS-treated cells compared to the controls. The top five dysregulated genes, SELE, CSF3, CSF2, CXCL3, and CXCL2, were further validated via qRT-PCR. The threshold cycle (Ct) values for gene expression were recorded, and normalization of the qRT-PCR data achieved using the 2^−∆∆Ct calculation method. The primer sequences used are listed in Table 2.

mRNA and protein stability

HLMVECs were treated with actinomycin D at a final concentration of 5 µg/mL for durations of 0, 3, 6, or 9 h. Following this, total RNA was extracted using an RNA isolation kit and subjected to qRT-PCR analysis. The half-life of target mRNAs was determined following established methodologies (22), with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) serving as an internal control for normalization. CHX, a known inhibitor of protein synthesis, was used to evaluate protein stability. HLMVECs, seeded at a density of 3×10⁵ cells/well in a 6-well plate, were allowed to adhere for 24 h prior to exposure to 20 µg/mL CHX for specified time intervals (0, 3, 6, 9, or 12 h).

Chromatin immunoprecipitation (ChIP) assay

The interaction between STAT3 and the CXCL3 promoter region was examined, following the instructions provided in the ChIP Assay Kit (Wanleibio, Shenyang, China). To initiate the cross-linking process, cells were exposed to formaldehyde, and the reaction halted with glycine. Subsequently, chromatin was fragmented to a size range of 200–1,000 base pairs through sonication. The lysates were then incubated overnight with the STAT3 primary antibody (#ab32500; Abcam, Cambridge, UK) or control IgG antibody (ab109489; Abcam) as a negative control. Magnetic bead-mediated immunoprecipitation was used to isolate DNA-protein complexes, followed by DNA purification. The recovered DNA was quantified through qRT-PCR to assess enrichment of the target DNA sequence.

Dual-luciferase reporter assay

The CXCL3 promoter sequence harboring STAT3-binding motifs was amplified via PCR and subsequently cloned into the pGL3 vector to establish a wild-type CXCL3 reporter construct (WT-CXCL3). To create a mutant version, STAT3-binding sites within the CXCL3 promoter were altered by GenScript Biotech (Nanjing, China), leading to the generation of a mutant CXCL3 reporter plasmid (MUT-CXCL3). HLMVECs were then transfected with a mixture of WT-CXCL3 or MUT-CXCL3, along with either si-NC or si-STAT3, using the LipoFiter transfection reagent. After a 48-h incubation period, transcriptional activity was assessed using the Dual-Luciferase Reporter Assay System (Solarbio), which enabled quantification of fluorescence signals emanating from the transfected cells.

Co-immunoprecipitation

HLMVEC lysates were prepared using RIPA buffer. For immunoprecipitation, the lysates were incubated overnight at 4 ℃ with specific antibodies against USP7 and CXCL3 (Abcam). Protein A Agarose Beads (BioVision) were then added to capture the immune complexes. Following centrifugation to pellet the agarose beads, the precipitated proteins were analyzed using Western blotting. Anti-CXCL3 (Affinity, Nanjing, China) and anti-USP7 (Affinity) antibodies were used to detect CXCL3 and USP7 protein levels, respectively. Immunoprecipitation with an anti-IgG antibody served as a negative control to ensure specificity.

Generation of adenoviral vectors

An effective small hairpin RNA (shRNA) targeting CXCL3 was designed and subsequently integrated into an adenoviral vector (Life Technologies, Shanghai, China) using the Gateway LR Clonase II Enzyme (Livning, Beijing, China), giving rise to the recombinant adenovirus, Ad-sh-CXCL3. As a control, another adenovirus was constructed by introducing a non-targeting “universal control” shRNA sequence into the same vector, designated as Ad-sh-NC. Both recombinant adenoviruses were amplified and purified, according to the manufacturer’s instructions. The following sequences were used: 5'-CCACTCTCAAGGATGGTCAAGAAGT-3' for Ad-sh-CXCL3 and 5'-CCATCACGAAGGTTGACGAATCAGT-3' for Ad-sh-NC.

Sepsis-induced ALI mouse model

Twenty male C57BL/6 mice (Hunan Slyke Jingda Experimental Animal Co., Ltd., Changsha, China) were housed in a controlled environment with ad libitum access to food and water. These mice were then randomly assigned to four groups, each consisting of five animals: the sham, cecal ligation and puncture (CLP), CLP + Ad-sh-NC, and CLP + Ad-sh-CXCL3 groups. Following a one-week period of environmental adaptation, all mice underwent intravenous injections via their caudal veins with 20 µL recombinant adenoviral vectors—either Ad-sh-CXCL3, designed to suppress CXCL3 expression, or Ad-sh-NC, a nonspecific control adenovirus. One week after viral administration, the mice underwent CLP, which was performed using a 22-gauge needle to induce sepsis. In summary, mice were anesthetized using pentobarbital sodium (40 mg/kg; Sigma), and their abdominal hair removed. The procedure involved exposing the cecum, which was then ligated, punctured, and replaced after gently expressing a small amount of its content. The abdominal incision was then closed, all of which were performed within a 10-min timeframe. Mice that underwent identical procedures without CLP served as the sham control group. Mice underwent fluid resuscitation every 6 h postoperatively (23). One week after CLP treatment, the mice were humanely euthanized, and serum along with lung tissues harvested for subsequent analyses. All animal experiments in this study were conducted in accordance with the experimental protocols approved by the Animal Ethics Committee of Soochow University (Approval No. 202209A0301), strictly complying with China’s Regulations on the Administration of Laboratory Animals and national guidelines for the care and use of animals.

Measurement of inflammatory factors

The concentrations of inflammatory cytokines, specifically IL-1β, using kits PI305 for humans and PI301 for mice, and TNF-α, with kits PT518 for humans and PT512 for mice (Beyotime), were quantified in cell culture supernatants and mouse serum through enzyme-linked immunosorbent assays (ELISAs). The samples were incubated with biotin-labeled antibodies, and optical density was measured using a microplate reader.

Western blotting assay

Proteins were extracted from HLMVECs, THP-1 cells, and frozen lung tissues at specified time points using RIPA buffer. The samples were homogenized in ice-cold buffer containing proteinase inhibitors. Protein concentrations were standardized using bicinchoninic acid assays before separation via SurePAGE gels and subsequently transferred onto methanol-pretreated polyvinylidene difluoride membranes. Membranes were blocked and then incubated with primary antibodies against inducible nitric oxide synthase (iNOS) (#AF0199; Affinity), COX-2 (#AF7003; Affinity), CXCL3 (#DF8554; Affinity), STAT3 (#AF6294; Affinity), USP7 (#DF6931; Affinity), p-P65 (#AF2006; Affinity), P65 (#BF8005; Affinity), p-IκBα (#AF2002; Affinity), IκBα (#AF5002; Affinity), Bax (#K001593P; Solarbio), Bcl-2 (#K003505P; Solarbio), and GAPDH (#AF7021; Affinity). After washing, the cells were incubated with the corresponding secondary antibodies (Solarbio). Detection was performed using the Beyotime EyoECL Plus reagent.

Histopathological analysis

Mouse lung tissues underwent a series of preparatory steps, beginning with fixation in formaldehyde and dehydration using alcohol. For the fixation of mouse lung tissues, after euthanizing the mice, the lungs were carefully exposed. The trachea was cannulated, and the lungs were inflated with 10% formaldehyde at a constant pressure of 20 cmH₂O for 15 min to ensure uniform fixation. This was followed by immersion in formaldehyde for an additional 24 hours for complete fixation. They were subsequently embedded in paraffin, sectioned, and stained with hematoxylin and eosin staining solution (Solarbio). Next, the slides were covered with neutral balsam. Pathological alterations were meticulously assessed under a microscope, adhering to established lung injury scoring criteria (24,25).

Measurement of lung wet-to-dry ratios

Wet-to-dry weight (W/D) ratios of the lungs were calculated to assess pulmonary edema. Mice were euthanized, and excess fluid absorbed using filter paper. The right upper lobes of the lungs were excised and weighed (wet weight). These lobes were subsequently dried in an oven at 180 ℃ for 24 h to obtain their dry weights. Finally, the W/D ratio was calculated to provide a quantitative measure of lung water accumulation.

Statistical analysis

Data analysis was conducted using GraphPad Prism (GraphPad Software, La Jolla, CA, USA), and measurement data were expressed as the mean ± standard deviation. Statistical comparisons were performed using the Wilcoxon signed-rank test, two-tailed Student’s t-test, or one-way analysis of variance, as appropriate. A probability value of P<0.05 was considered statistically significant.

Results

LPS treatment induced human lung microvascular endothelial cell apoptosis, inflammation, oxidative stress, and M1 macrophage polarization

As expected based on previous literature, LPS treatment of HLMVECs led to a series of inflammatory responses, including decreased cell viability, increased apoptosis, and elevated cytokine production. The detailed data regarding these effects can be found in Figure S1. HLMVECs were subjected to LPS to induce an ALI-mimicking injury. LPS reduced the viability and proliferation of HLMVECs in a dose-dependent manner (Figure S1A,S1B). Furthermore, LPS exposure led to a concentration-dependent increase in cell apoptosis and elevated IL-1β and TNF-α levels (Figure S1C-S1F). LPS administration resulted in concentration-dependent suppression of SOD activity and elevation of MDA levels (Figure S1G-S1H). It also enhanced expression of the M1 macrophage markers, iNOS and COX-2 (Figure S1I), and increased the M1 macrophage population (Figure S1J). Consequently, a concentration of 10 µg/mL LPS was selected for subsequent HLMVEC experiments.

CXCL3 expression was upregulated in the plasma of patients with sepsis-induced ALI

Using the Gene Expression Omnibus (GEO) database (website: https://www.ncbi.nlm.nih.gov/geo/; accession number GSE5883), we identified differentially expressed genes in LPS-treated versus control HLMVECs (P<0.001 and |logFC| >4; Figure 1A). The top five dysregulated genes, SELE, CSF3, CSF2, CXCL3, and CXCL2, were further validated via qRT-PCR in both LPS- and DMSO-treated cells. LPS notably upregulated SELE, CSF3, CXCL3, and CXCL2 mRNA levels, with CXCL3 showing the most prominent increase (Figure 1B). Consequently, CXCL3 was selected for subsequent experiments. Figure 1C presents the online database analysis of the differences in CXCL3 expression between LPS-exposed HLMVECs and untreated controls. Notably, CXCL3 mRNA levels were significantly higher in the plasma of patients with sepsis-induced ALI than in that of healthy volunteers (Figure 1D). Furthermore, LPS augmented CXCL3 mRNA and protein expression in HLMVECs in a dose-dependent manner (Figure 1E,1F).

Figure 1 CXCL3 expression in patient plasma and LPS-stimulated endothelial cells. (A) Differential gene expression analysis between LPS-treated HLMVECs and controls was conducted using the GEO database (GSE5883). (B) LPS’s impact on SELE, CSF3, CSF2, CXCL3, and CXCL2 mRNA levels was quantified by qRT-PCR. (C) Online database (GEO GSE5883) was utilized to assess CXCL3 expression changes in LPS-exposed HLMVECs versus controls. (D) qRT-PCR measured CXCL3 mRNA in sepsis-induced ALI patient plasma (N=27) compared to healthy controls (N=25). (E,F) HLMVEC CXCL3 mRNA and protein responses to LPS were examined via qRT-PCR and Western blot respectively. **, P<0.01; ****, P<0.0001. ALI, acute lung injury; GEO, Gene Expression Omnibus; HLMVEC, human lung microvascular endothelial cell; LPS, lipopolysaccharide; mRNA, messenger RNA; qRT-PCR, quantitative real-time polymerase chain reaction.

CXCL3 depletion protected against LPS-induced human lung microvascular endothelial cell apoptosis, inflammation, oxidative stress, and M1 macrophage polarization

LPS-induced CXCL3 protein upregulation was effectively reversed upon CXCL3 siRNA transfection (Figure 2A). Silencing CXCL3 mitigated the decline observed in HLMVEC viability and proliferation and increase in apoptosis induced by LPS (Figure 2B-2E). Moreover, the LPS-induced increase in IL-1β, TNF-α, and MDA levels, along with SOD activity reduction, were alleviated after CXCL3 knockdown (Figure 2F-2I). Additionally, LPS-enhanced iNOS and COX-2 expression, and the proportion of M1 macrophages, were restored to baseline levels upon CXCL3 suppression (Figure 2J,2K).

Figure 2 CXCL3 depletion protected against LPS-induced HLMVEC apoptosis, inflammation, oxidative stress and M1 macrophage polarization. The HLMVECs divided into Control, LPS, LPS + si-NC, and LPS + si-CXCL3. Protein expression of CXCL3 was assessed via Western blotting (A), cell viability through CCK-8 assay (B), proliferation with EdU test (C), apoptosis by flow cytometry (D,E), cytokines IL-1β and TNF-α levels via ELISAs (F,G), SOD activity using a specific kit (H), and MDA content by another kit (I). Similarly, THP-1 cells were grouped identically, and their iNOS and COX-2 protein expressions were measured by Western blotting (J), with M1 macrophage proportion determined by flow cytometry (K). **, P<0.01; ***, P<0.001; ****, P<0.0001. CCK-8, Cell Counting Kit-8; EdU, 5-ethynyl-2'-deoxyuridine; ELISAs, enzyme-linked immunosorbent assays; HLMVEC, human lung microvascular endothelial cell; IL, interleukin; iNOS, inducible nitric oxide synthase; LPS, lipopolysaccharide; MDA, malondialdehyde; NC, negative control; SOD, superoxide dismutase; TNF, tumor necrosis factor.

STAT3 induced activation of CXCL3 transcription

We examined the impact that silencing of the four STAT transcription factors has on CXCL3 mRNA levels in HLMVECs via qRT-PCR. STAT3 knockdown alone reduced CXCL3 mRNA expression (Figure 3A), guiding our focus to STAT3 for subsequent investigations. Using the JASPAR database (website: https://jaspar.elixir.no/), we predicted STAT3-binding sites within the CXCL3 promoter region. Figure 3B illustrates three potential STAT3-binding sequences. A subsequent ChIP assay confirmed the direct interaction of STAT3 with sites 1 and 3 in the CXCL3 promoter (Figure 3C). Our findings also demonstrated that si-STAT3 transfection reduced luciferase activity in cells containing WT-CXCL3 but not in those with MUT-CXCL3 (Figure 3D). The efficacy of STAT3 silencing and overexpression was validated through Western blot (Figure 3E). STAT3 knockdown consistently decreased CXCL3 protein levels, whereas STAT3 overexpression exhibited the opposite effect (Figure 3F).

Figure 3 STAT3 induced activation of CXCL3 transcription. (A) qRT-PCR assessed CXCL3 mRNA changes after reducing STAT1, 2, 3, and 4 in HLMVECs. (B) A diagram depicted STAT3’s binding areas on the CXCL3 promoter. (C) ChIP assay confirmed STAT3’s interaction with the CXCL3 gene. (D) Dual-luciferase testing verified the STAT3-CXCL3 connection. (E) Western blots gauged STAT3 knockdown and overexpression efficiency in HLMVECs. (F) STAT3 modulation’s impact on CXCL3 protein was measured via Western blot. **, P<0.01; ****, P<0.0001. ChIP, chromatin immunoprecipitation; HLMVECs, human lung microvascular endothelial cells; MUT, mutant; mRNA, messenger RNA; NC, negative control; qRT-PCR, quantitative real-time polymerase chain reaction; WT, wild type.

USP7 regulated CXCL3 via deubiquitination

LPS administration notably elevated USP7 and USP14 mRNA expression levels in HLMVECs, with USP7 showing a more substantial increase (Figure 4A). Remarkably, the silencing of USP7 led to a substantial decrease in CXCL3 protein levels, whereas USP14 knockdown had no such effect (Figure 4B), guiding our focus towards USP7 for further study. Figure 4C illustrates that USP7 knockdown did not affect CXCL3 mRNA levels in HLMVECs. The Co-IP assay confirmed the interaction between CXCL3 and USP7 (Figure 4D). The effectiveness of USP7 silencing and overexpression is depicted in Figure 4E. Our results indicated that the mRNA half-life of CXCL3 was shortened by USP7 knockdown and prolonged by USP7 overexpression (Figure 4F,4G). A CHX-chase experiment demonstrated the role of USP7 in stabilizing the CXCL3 protein (Figure 4H). Importantly, although USP7 depletion reduced CXCL3 protein levels, this effect was mitigated upon treatment with the proteasome inhibitor, MG132 (Figure 4I).

Figure 4 USP7 regulated CXCL3 via deubiquitination. (A) qRT-PCR analyzed LPS-induced changes in USP7 and USP14 mRNA levels. (B) Western blots showed how USP7 and USP14 knockdown altered CXCL3 protein expression. (C) qRT-PCR probed USP7 depletion’s impact on CXCL3 mRNA. (D) Co-IP assays illuminated interactions between CXCL3 and USP7. (E) Western blots validated USP7 knockdown and overexpression in HLMVECs. (F,G) Actinomycin D tests assessed USP7’s role in CXCL3 mRNA stability. (H) After CHX exposure, USP7 overexpression’s effect on CXCL3 protein was measured by Western blot. (I) Western blots, post-treatment with si-NC, si-USP7, or si-USP7 + MG132, evaluated CXCL3 protein levels. ns, no statistically significant difference; **, P<0.01, ***, P<0.001; ****, P<0.0001. HLMVECs, human lung microvascular endothelial cells; LPS, lipopolysaccharide; mRNA, messenger RNA; NC, negative control; qRT-PCR, quantitative real-time polymerase chain reaction.

USP7 knockdown ameliorated LPS-induced human lung microvascular endothelial cell apoptosis, inflammation, oxidative stress, and M1 macrophage polarization through the regulation of CXCL3

LPS-induced CXCL3 protein upregulation was reversed upon si-USP7 transfection and partially offset by CXCL3 overexpression (Figure 5A). USP7 silencing alleviated the LPS-induced decline in HLMVEC viability and proliferation, as well as the increase in apoptosis rates, whereas these improvements were hindered upon CXCL3 elevation (Figure 5B-5E). Furthermore, the LPS-induced elevation in IL-1β, TNF-α, and MDA concentrations, alongside SOD activity reduction, were rectified by USP7 silencing. These effects, however, were dampened upon CXCL3 overexpression (Figure 5F-5I). Likewise, the LPS-stimulated increases in iNOS and COX-2 protein expression levels and the M1 macrophage population were partially mitigated by USP7 knockdown but reinstated upon CXCL3 upregulation (Figure 5J,5K).

Figure 5 USP7 knockdown ameliorated LPS-induced HLMVEC apoptosis, inflammation, oxidative stress and M1 macrophage polarization through the regulation of CXCL3. HLMVECs were grouped into Control, LPS, LPS + si-NC, LPS + si-USP7, LPS + si-USP7 + pcDNA, LPS + si-USP7 + CXCL3 for assessing CXCL3 protein (Western blot) (A), cell viability (CCK-8) (B), proliferation (EdU) (C), apoptosis (flow cytometry) (D,E), cytokine levels (IL-1β, TNF-α via ELISAs) (F,G), SOD activity (kit assay) (H), and MDA content (kit assay) (I). Similarly, THP-1 cells were grouped likewise, examining iNOS and COX-2 proteins (Western blot) (J) and M1 macrophage ratio (flow cytometry) (K). *, P<0.05, **, P<0.01, ***, P<0.001; ****, P<0.0001. CCK-8, Cell Counting Kit-8; EdU, 5-ethynyl-2'-deoxyuridine; ELISAs, enzyme-linked immunosorbent assays; HLMVECs, human lung microvascular endothelial cells; IL, interleukin; iNOS, inducible nitric oxide synthase; LPS, lipopolysaccharide; MDA, malondialdehyde; NC, negative control; SOD, superoxide dismutase; TNF, tumor necrosis factor.

USP7 depletion inactivated the NF-κB pathway by regulating CXCL3

USP7 silencing decreased the ratios of p-P65 to P65 and p-IκBα to IκBα in LPS-induced HLMVECs (Figure 6). However, these reductions were partially negated when cells were co-transfected with si-USP7 and the CXCL3 overexpression vector.

Figure 6 USP7 depletion inactivated the NF-κB pathway by regulating CXCL3. HLMVECs were categorized into Control, LPS, LPS + si-NC, LPS + si-USP7, LPS + si-USP7 + pcDNA, and LPS + si-USP7 + CXCL3. Western blot analysis was conducted to determine the protein levels of p-P65, P65, p-IκBα, and IκBα. ****, P<0.0001. HLMVECs, human lung microvascular endothelial cells; LPS, lipopolysaccharide; NC, negative control.

CXCL3 knockdown ameliorated cecal ligation and puncture-induced lung injury in C57BL/6 mice

CLP-induced increases in lung injury scores and W/D ratios were mitigated by CXCL3 knockdown (Figure 7A,7B). Moreover, CLP stimulated CXCL3 and Bax protein expression while suppressing that of Bcl-2 in mouse lung tissue; these effects were alleviated upon CXCL3 depletion (Figure 7C,7D). Reducing CXCL3 expression counteracted the CLP-elevated serum IL-1β and TNF-α levels (Figure 7E,7F). Furthermore, CLP increased the ratios of p-P65 to P65 and p-IκBα to IκBα in mouse lung tissue; these effects were partially reversed by CXCL3 silencing (Figure 7G).

Figure 7 CXCL3 knockdown ameliorated CLP-induced lung injury of C57BL/6 mice. Mice were allocated to Sham, CLP, CLP + Ad-sh-NC, and CLP + Ad-sh-CXCL3 groups. Prior to surgery, CLP + adenovirus groups received intravenous injections of adenovirus vectors. Post-CLP for a week, mice were euthanized for serum and lung sample collection, followed by lung injury scoring. The lung tissues of mice were first fixed with formaldehyde and dehydrated with alcohol, respectively. Then, the tissues were embedded in paraffin and stained with hematoxylin and eosin solution. Magnification: 200× (A). W/D assessment (B). CXCL3, Bax, Bcl-2 protein quantification in lung tissue via Western blot (C,D). ELISA measurement of serum IL-1β and TNF-α levels (E,F). Western blot analysis of p-P65, P65, p-IκBα, and IκBα protein expression in lung tissue (G). **, P<0.01, ***, P<0.001; ****, P<0.0001. CLP, cecal ligation and puncture; ELISA, enzyme-linked immunosorbent assay; IL, interleukin; NC, negative control; TNF, tumor necrosis factor; W/D, wet-to-dry weight ratio.

Discussion

Sepsis-related ALI significantly influences patient outcomes; however, despite extensive research, the precise mechanisms underlying its initiation and progression remain unclear (26,27). LPS, a pivotal mediator of sepsis, activates the CD14/TLR4 receptor complex in certain cell types, triggering the production of inflammatory mediators (28). This study explored the function and mechanism of action of CXCL3 in sepsis-driven ALI using LPS-induced HLMVECs and a CLP mouse model. We observed elevated CXCL3 expression in sepsis-related ALI. We further elucidated that STAT3 activates CXCL3 transcription, whereas USP7 modulates CXCL3 via deubiquitination. Silencing USP7 alleviated LPS-induced HLMVEC injury and M1 macrophage polarization, as well as suppressed NF-κB pathway activation, through CXCL3 regulation. In-vivo experiments showed that CXCL3 depletion markedly diminished CLP-induced lung injuries in mice, reinforcing the central role CXCL3 plays in sepsis-mediated ALI.

STAT proteins, comprising STAT1-STAT6, are a group of transcription factors integral to various cellular signaling pathways. Database analysis using GeneCards suggested the potential involvement of STAT1-STAT4 in the regulation of CXCL3 transcription. Our qRT-PCR analysis of HLMVECs with siRNAs targeting the four transcription factors indicated that STAT3 suppression alone reduced CXCL3 mRNA levels. STAT3, which is activated by cytokines and growth factors, such as oncostatin M upon stimulation, typically remains inactive in the cytoplasm until triggered (29). LPS rapidly activates STAT3 in the lungs, peaking at 1–2 h (30). STAT3 inhibition has been proven to be protective against ALI, potentially by controlling the transcription of certain molecules, such as lncRNA XIST (31,32). Our findings support that STAT3 stimulates CXCL3 transcription in HLMVECs.

A previous study reported that CXCL3 expression was upregulated in LPS-exposed C57BL/6 mice (12). Moreover, activating endothelial A2b adenosine receptors, which have low affinity for adenosine compared to other adenosine receptor subtypes, alleviates LPS-induced lung microvascular inflammation through CXCL3 downregulation (33), highlighting the pro-inflammatory function of CXCL3 in LPS-triggered pulmonary responses. Consistently, our findings demonstrated elevated CXCL3 levels in the plasma of patients with sepsis-induced ALI and in LPS-treated HLMVECs. LPS exposure led to a decline in HLMVEC proliferation and an increase in apoptosis and THP-1 cell polarization that favors M1 macrophages. It also enhanced IL-1β, TNF-α, and MDA secretion while suppressing SOD activity. In vivo, CLP increased lung injury scores, lung W/D ratios, and Bax levels while decreasing Bcl-2 levels. These changes were attenuated after CXCL3 silencing. A previous study implicated CXCL3 in LPS-induced pulmonary endothelial injury via MAPK activation (34). Our study reinforces the hypothesis that CXCL3 suppression mitigates sepsis-induced ALI.

USP7 stabilizes proteins vital for immunity, transcription, and epigenetic processes in disease, affecting processes such as Axin deubiquitination via direct TRAF domain interactions (35). USP7 has been shown to play a significant role in the regulation of the immune response. For instance, in a study on viral infections, USP7 was found to deubiquitinate and stabilize key antiviral proteins. This stabilization enhanced the cell’s ability to mount an effective antiviral response, highlighting its importance in the innate immune defense mechanism (36). In the context of autoimmune diseases, USP7 has been associated with the regulation of T-cell activation. Aberrant expression of USP7 in T-cells can lead to abnormal activation and cytokine production, contributing to the pathogenesis of autoimmune conditions such as rheumatoid arthritis (37). Moreover, in inflammatory bowel disease models, USP7 has been implicated in modulating the inflammatory response in intestinal epithelial cells. Its dysregulation can lead to increased production of pro-inflammatory cytokines and disruption of the epithelial barrier function (38). Our findings revealed that USP7 could deubiquitinate CXCL3 and thereby enhance its stability. Additionally, reducing USP7 levels alleviated LPS-induced cellular damage through its interaction with CXCL3. Macrophages use pattern recognition receptors, such as TLRs, to detect invading pathogens. Notably, LPS functions as a ligand for TLR4, and activation of TLR4-mediated pathways initiates a cascade of intracellular signals, leading to the nuclear translocation of transcription factors, including NF-κB (37). Furthermore, P22077, a specific inhibitor of USP7, effectively suppresses activation of the NF-κB pathway in LPS-stimulated peritoneal macrophages (39). Building upon this, our study delved deeper into the relationship between USP7 and CXCL3 in modulating the NF-κB signaling cascade, revealing that downregulation of USP7 leads to inactivation of this pathway through the mediation of CXCL3.

LPS- and CLP-induced endotoxemia and tissue injury do not comprehensively reflect the entire spectrum of pathophysiological processes observed in all instances of sepsis-associated ALI. Moreover, the impact of anesthesia on the inflammatory response remains inadequately understood and may introduce confounding variables in the interpretation of results derived from murine model experiments. Consequently, additional studies are warranted to validate this novel mechanism in alternative animal models.

Conclusions

CXCL3 expression was significantly increased in sepsis-induced ALI. Knockdown of CXCL3 ameliorated sepsis-induced ALI. In in-vitro models, USP7 modulated CXCL3 through deubiquitination. Silencing USP7 alleviated LPS-induced HLMVEC injury and inhibited M1 macrophage polarization by regulating CXCL3. Although the direct role of USP7 in the in-vivo CLP model was not experimentally demonstrated, our in-vitro results suggest a potential mechanism by which CXCL3, in relation to USP7, contributes to the pathogenesis of sepsis-induced ALI. This study contributes to a deeper understanding of the molecular pathways driving the development and progression of sepsis-related ALI, highlighting CXCL3 as a promising therapeutic target for future interventions. In addition, although our study focused on the relationship between USP7 and CXCL3, we noted SELE and CXCL2 are also highly upregulated in endothelial cells stimulated with LPS. Future studies could explore this aspect, as determining that USP7 specifically modulates CXCL3, while not affecting SELE and CXCL2, would significantly strengthen the significance of the link between USP7 and CXCL3.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE and MDAR reporting checklists. Available at https://jtd.amegroups.com/article/view/10.21037/jtd-2024-2026/rc

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Funding: None.

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://jtd.amegroups.com/article/view/10.21037/jtd-2024-2026/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. The study was conducted in accordance with the Declaration of Helsinki and its subsequent amendments, with the approval of the Ethics Committee of The Second Affiliated Hospital of Soochow University (approval number: JD-LK-2022-105-01). All participants provided written informed consent. All animal experiments in this study were conducted in accordance with the experimental protocols approved by the Animal Ethics Committee of Soochow University (Approval No. 202209A0301), strictly complying with China’s Regulations on the Administration of Laboratory Animals and national guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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