Eight- to twelve-week-old male C57BL/6J (The Jackson Laboratory, Bar Harbor, ME, USA) and VISTA knockout mice (15) were maintained in our animal care facility. All experiments were conducted in accordance with NIH Guidelines for Animal Use and Care and as approved by the Rhode Island Hospital Institutional Animal Care and Use Committee (Providence, RI; AWC#0040-16). Of note, male mice were used in this study because our laboratory has previously observed sex-based differences in immune responses to hemorrhage (Hem) and/or cecal ligation and puncture (CLP), with males being significantly more sensitive than pro-estrus females (16, 17). In other words, to avoid overlooking the potential effects of Hem/CLP due to the known protective influence of the female sex, we selected male mice, which are more susceptible to developing experimental ARDS.

Indirect ARDS (iARDS) was induced using a two-insult mouse model involving an initial hemorrhagic shock and a septic challenge (18). Mice were anesthetized in the atmosphere of servofluorane during the procedure. For the Hem protocol, polyethylene tubing (PE-10, 0.28 mm inner diameter × 0.64 mm outer diameter) was inserted into the femoral artery under an optical microscope. One side was used for drawing blood, and the other side for monitoring blood pressure. The volume of drawn blood was recorded and stopped once the blood pressure dropped to 35 mmHg ± 5 mmHg and was maintained at that level for 90 min. Following this, four times lactated Ringer’s solution (relative to the volume of blood drawn) was administered through the tube for resuscitation. The femoral arteries were then tied and sutured after resuscitation. In the sham group, the femoral arteries were tied and sutured without blood withdrawal or resuscitation. Mice were housed in the animal care facility with free access to food. At –24 h post-Hem, a secondary insult was induced by polymicrobial sepsis using the CLP method, which creates a consistent and reproducible septic focus. In the sham group, the abdomen was incised and sutured without CLP. Mice were euthanized at –24 h post-CLP, and lung tissues and blood samples were collected for further analysis.

WT mice in the treatment group were subcutaneously injected on the back with a hamster monoclonal antimouse VISTA antibody (13F3) (10 μg/g mouse body weight, 45 μL; clone 13F3, Bio X Cell Inc, Hanover, NH, USA.) after Hem and CLP, respectively. The 13F3 was diluted in saline. Mice in the control group received the same volume of nonspecific hamster IgG diluted in saline.

The expression levels of VISTA on various cell types—including neutrophils, monocytes/macrophages, and endothelial and epithelial cells—were assessed 24 h after CLP surgery. Flow cytometry was used to analyze VISTA expression in both whole blood samples and single-cell suspensions derived from lung tissues. The lung tissues were processed using an enzymatic dissociation buffer to generate single-cell suspensions, following these steps: (1) Prepare 10 mL of dissociation medium by adding 1 mL of collagenase/hyaluronidase and 1.5 mL of DNase I solution (1 mg/mL) to 7.5 mL of RPMI 1640 medium. Warm the solution to room temperature (15°C–25°C). (2) Harvest lung tissue into a tube containing PBS with 2% FBS. (3) Transfer the lung tissue to a dish without medium and mince it into a homogenous paste (< 1 mm in size) using a razor blade. (4) Transfer the minced lung tissue and the dissociation medium to a sterile 50-mL conical tube. Rinse the dish with the remaining 5 mL of dissociation medium and add it to the tube containing the minced tissue. Incubate at 37°C for 20 min using a Miltenyi^© GentleMACS Dissociator. Following digestion, the cell suspension was carefully filtered through a 70-mm cell strainer to obtain a homogenous single-cell mixture suitable for accurate flow cytometric analysis. The resultant cells were diligently counted using a microscope, and the concentration was adjusted to achieve a final cell density of 1 to 10 million cells per milliliter. The single-cell suspension was then stained, and fluorochrome expression was analyzed using a Miltenyi^© MACS Quant 10 flow cytometer (Miltenyi Biotec Inc., Auburn, CA, USA), as previously described in our laboratory (19, 20).

For flow cytometric analysis, debris (low FSC/SSC) and doublets (identified using FSC-H vs. FSC-W for singlet discrimination) were first excluded. Intact cells (moderate-to-high FSC/SSC) were then gated, followed by the exclusion of dead cells using a viability dye (e.g., PI). The harvested cells were stained with the following fluorochrome-conjugated antibody panel: anti-Ly6G (clone 1A8), anti-VISTA (clone MH5A), anti-CD11b (clone M1/70), anti-PD-1 (clone 135209), anti-CD31 (clone W18222B), anti-CD115 (clone T38-320), and anti-programmed cell death-ligand 1 (PD-L1; clone 124336). The cells were identified and quantified based on their expression of the following markers: CD11b⁺Ly6G⁺ (neutrophils), CD11b⁺F4/80⁺ (macrophages), CD31⁺ (endothelial cells), and CD115⁺ (epithelial cells).

Lung tissues were processed using hematoxylin and eosin (H&E) staining to assess morphological changes in these animals, as we previously described (20). Specimens from the lungs of experimental mice were harvested and fixed in formalin to preserve cellular details and prevent degradation. The fixed tissues were embedded in paraffin wax to provide support for sectioning. Thin sections (3–5 µm) were cut using a microtome and mounted on glass slides. The slides were then placed in xylene to dissolve the paraffin. The tissues were then rehydrated by passing the slides through a series of graded alcohol solutions (100%, 95%, 70%) and finally into distilled water. The slides were immersed in hematoxylin solution for 5–10 min. They were then rinsed in running tap water to remove excess stain. Bluing was performed by immersing the slides in ammonia water or lithium carbonate solution for 30 s to 1 min. The slides were subsequently in eosin solution for 1–3 min. Finally, the slides were passed through graded alcohol solutions (70%, 95%, 100%) to remove water. The tissue sections were cleared by immersing them in xylene, which rendered the tissue transparent. A few drops of mounting medium were applied to the tissue section. A cover slip was carefully placed over the section, ensuring that air bubbles were avoided. The slides were examined under a microscope to confirm optimal staining quality. As previously described by us, the extent of morphological changes in the lung specimens was assessed using a pathological scoring system. Specimens were scored from 0 to 4: 0 (normal), 1 (very mild impairment, < 25% of the field area), 2 (mild impairment, 25% to 50% of the visual field area), 3 (moderate impairment, 50% to 75% of the visual field area), and 4 (severe impairment, > 75% of the view area). Scoring was performed by two blinded pathologists based on alveolar wall thickening, interstitial edema, infiltration of inflammatory cells (such as neutrophils and macrophages), and the presence of hyaline membranes. Additionally, areas of atelectasis, Hem, and cellular debris may be observed, reflecting the pathological changes associated with the progression of ARDS (20, 21).

Survival analysis was conducted to assess the impact of VISTA on the survival of mice in the iARDS model. Mice were monitored daily for survival outcomes, and a Kaplan–Meier survival curve was generated to compare survival rates between wild-type and VISTA knockout mice. This analysis was used to evaluate the efficacy of VISTA in improving survival outcomes in the context of iARDS.

To quantify the levels of cytokines, including interleukin (IL)-6), IL-10, monocyte chemoattractant protein 1 (MCP-1), and tumor necrosis factor-alpha (TNF-α) (BD Biosciences, San Diego, CA, USA), as well as chemokines such as keratinocyte chemoattractant (KC) and macrophage inflammatory protein 2 (MIP-2) (R&D Systems, Minneapolis, MN, USA), enzyme-linked immunosorbent assays (ELISAs) were conducted according to the manufacturer’s instructions using lung tissue homogenates and plasma samples obtained from the experimental animals, as recently described by our laboratory (20). In brief, following the initial coating of the ELISA plate with the capture antibody, any excess unbound antibody was washed from the plate. Next, the unknown samples (lung tissue homogenates or plasma), along with diluted standard cytokine, were added. The samples were added in duplicate and at varying concentrations to ensure that their values fell within the detection range of the assay/standard cytokine curve. After sample incubation, excess material was again washed from the plate, and a biotinylated detection antibody was added. Finally, a biotin-binding chromogenic substrate was added to the plate. The antigen concentrations in the unknown samples were determined by comparing the optical density (OD) values to a standard curve generated using cytokines/chemokines of known concentrations, in accordance with the manufacturer’s instructions.

To assess select changes in gene transcription, a concise version of the quantitative RT-PCR protocol was employed to detect mRNA expression of VISTA in lung tissue. RNA was isolated from preserved lung tissue using a suitable extraction kit (Takara Bio USA, Inc., San Jose, CA, USA). The extracted RNA was then converted to cDNA using a reverse transcription kit. The VISTA gene region was amplified from the cDNA using specific primers and a PCR master mix. cDNA synthesis was performed using a PrimeScript™ RT Reagent Kit (Takara Bio USA, Inc., San Jose, CA, USA). qRT-PCR was then performed using the ABIQ3 system (Applied Biosystems Corp., Waltham, MA, USA) and the SYBER PrimeScript™ RT Reagent Kit (Takara Bio USA, Inc., San Jose, CA, USA) according to the manufacturer’s recommendations. The following primer sequences (synthesized by Life Technologies Inc., Carlsbad, CA, USA) were used: murine VISTA 5′-CTC CTT GCT ATT TTC CTG GCT G-3′ and 5′-AGG TGA GGG TGG CAT TCT GT-3′; IL-6 5′-ATG GAT GCT ACC AAA CTG GAT-3′ and 5′-TGA AGG ACT CTG GCT TTG TCT-3′ (sense/antisense); TNF-α 5′-AGG CTC ATC CTT GCC TTT GTC TCT-3′ and 5′-TCA GCA GCT ACC CAC ACT TCA CTT-3′ (sense/antisense); IL-10 5′-CCA GTT TTA CCT GGT AGA AGT GAT G-3′ and 5′-TGT CTA GGT CCT GGA GTC CAG CAG ACT C-3′ (sense/antisense); MCP-1 5′-TCA CCT GCT GCT ACT CAT TCA CCA-3′ and 5′-AAA GGT GCT GAA GAC CTT AGG q′ (sense/antisense); MIP-2 5′-AAAGTTTGCCTTGACCCTGAA-3′ and 5′-TCTTTGGTTCTTCCGTTGAGG-3′ (sense/antisense); KC 5′-ACTGCACCCAAACCGAAGTC-3′ and 5′-TGGGGACACCTTTTAGCATCTT-3′ (sense/antisense); and 18s 5′-GGA CAC GGA CAG GAT TGA CAG ATT-3′ and 5′-AAT CGC TCC ACC AAC TAA GAA CGG-3′ (sense/antisense). Amplified products were quantified using real-time PCR analysis, employing either a fluorescent probe or SYBR Green. Duplicate CT values were analyzed using the comparative CT (ΔΔCt) method. The amount of target (2−ΔΔCt) was obtained by normalizing to GAPDH relative to a control (nonstimulated cells or sham mice). VISTA expression data were normalized to 18s, and relative expression levels were calculated (22).

Data were presented as mean values with standard error of the mean (SEM). As the group sizes were typically fewer than 10 animals/group, a Kruskal–Wallis test was applied to determine whether statistically significant differences existed between group means. Statistical significance for survival analysis was assessed using log-rank tests. A p-value of less than 0.05 was considered indicative of a statistically significant difference between group means.

To investigate the development of iARDS, we employed a “double-hit” mouse model that simulates septic shock (23). This approach involves HEM followed by sepsis induced by CLP—a sequence referred to as Hem/CLP. As depicted schematically in Figure 1 , C57BL/6J mice first undergo Hem (day 1), followed by CLP (day 2).

To investigate the role of VISTA in iARDS, we established an experimental model using C57BL/6J (wild-type [WT]) and VISTA^−/− mice.

Over a 14-day follow-up period, we observed a significantly lower survival rate in VISTA^−/− mice compared to their WT counterparts. This striking difference in mortality underscores the protective role of VISTA in mitigating the severity of iARDS. Specifically, VISTA deficiency appears to exacerbate disease progression, leading to increased lethality. These findings are visually summarized in Figure 2 , which depicts the survival curves for both groups.

To assess the impact of VISTA deficiency on lung tissue pathology, we performed histopathological analysis using optical microscopy. VISTA^−/− mice exhibited a marked increase in immune cell infiltration within lung tissue compared to WT mice. This infiltration was accompanied by loss of epithelial integrity—a hallmark of tissue damage—and compromised barrier function. These findings suggest that VISTA deficiency exacerbates lung injury by promoting immune cell recruitment and disrupting the structural organization of the lung epithelium.

Following Hem/CLP, lung tissue sections from VISTA^−/− mice revealed abundant infiltrating cells, including neutrophils, macrophages, and lymphocytes, indicative of an intense inflammatory response. In contrast, WT mice exhibited a more controlled and less severe immune cell influx. The severity score, quantifying the extent of tissue damage and inflammation, was significantly higher in VISTA^−/− mice compared to WT controls (p < 0.01), as illustrated in Figure 3 . This elevated severity score underscores VISTA’s critical role in modulating inflammatory responses and maintaining tissue homeostasis.

To elucidate the impact of VISTA on the pathogenesis of inflammatory iARDS, we measured plasma levels of proinflammatory cytokines and chemokines using ELISA. In VISTA^−/− mice, we observed a postprocedural surge in the cytokines MCP-1 (p < 0.01) and TNF-α (p < 0.01), as well as the chemokine MIP-2 (p < 0.01), compared to WT mice. Concurrently, the anti-inflammatory cytokine IL-10 (p < 0.01) was significantly suppressed in VISTA^−/− mice, as depicted in Figure 4A . This heightened proinflammatory state in the plasma suggests that VISTA deficiency exacerbates systemic inflammation, potentially contributing to the onset of iARDS.

Expanding our analysis to lung tissues, we found that levels of MCP-1 (p < 0.01), TNF-α (p < 0.01), MIP-2 (p < 0.01), IL-6 (p < 0.01), and KC (p < 0.01) increased significantly following Hem/CLP in WT mice. However, in VISTA^−/− mice, these mediators did not exhibit significant changes compared to WT controls, as illustrated in Figure 4B . This divergence between plasma and lung tissue responses underscores the compartment-specific effects of VISTA deficiency, with systemic inflammation predominating in plasma but a muted response in the lung microenvironment.

In peritoneal fluid, most cytokines and chemokines were reduced in VISTA^−/− mice, except for MCP-1, IL-6, IL-10, MIP-2, and KC, which remained unchanged compared to WT mice ( Figure 4C ). These findings further underscore the tissue-specific nature of VISTA-mediated immune regulation, with distinct inflammatory profiles observed across different biological compartments.

Intriguingly, analysis of mRNA expression revealed significant suppression of MCP-1 (p < 0.01), MIP-2 (p < 0.01), IL-6 (p < 0.01), IL-10 (p < 0.01), and KC (p < 0.01) in VISTA^−/− mice, indicating a dynamic molecular response at the transcriptional level. In contrast, TNF-α mRNA expression remained unchanged, suggesting a more nuanced regulatory mechanism potentially involving posttranscriptional or posttranslational modifications. These findings, visually depicted in Figure 4D , provide further insight into the molecular underpinnings of the observed biological response.

To investigate the impact of 13F3 on VISTA, a critical immune checkpoint molecule, we measured its expression on various cell populations using flow cytometry. In peripheral blood, VISTA expression was assessed on monocytes (p < 0.05) and neutrophils (p < 0.01), two key innate immune cell types. As shown in Figure 5A , 13F3 treatment significantly suppressed VISTA expression on both monocytes and neutrophils. This downregulation suggests that 13F3 may modulate immune checkpoint activity in circulating immune cells, potentially enhancing their responsiveness to inflammation.

The analysis was extended to lung tissue, where VISTA expression was evaluated on a broader range of cell types, including macrophages (p < 0.05), neutrophils (p < 0.05), epithelial cells (p < 0.05), and endothelial cells (p < 0.05). As shown in Figure 5B , VISTA expression was significantly suppressed across all these cell populations following 13F3 treatment. This widespread downregulation highlights the systemic effects of 13F3 on immune checkpoint regulation within the lung microenvironment.

The suppression of VISTA on macrophages and neutrophils in lung tissue aligns with findings in peripheral blood, reinforcing the consistency of 13F3’s effects on innate immune cells. Additionally, the reduced VISTA expression on epithelial and endothelial cells suggests that 13F3 may also influence the immune-regulatory functions of structural cells, which are critical for maintaining tissue homeostasis and modulating immune responses.

These findings collectively underscore the broad and potent ability of 13F3 to suppress VISTA expression across diverse cell types, both in circulation and within tissues. Given VISTA’s role in inhibiting T-cell activation and promoting immune tolerance, its downregulation by 13F3 may have significant implications for modulating anti-inflammatory actions.

The immunomodulatory effects of 13F3 were investigated by analyzing the expression of key cytokines and chemokines in different biological compartments. As demonstrated in Figure 6A , treatment with 13F3 led to significant suppression of the cytokines MCP-1 (p < 0.05), TNF-α (p < 0.05), and IL-10 (p < 0.01), as well as the chemokine MIP-2 (p < 0.01). These molecules are central to inflammatory and immune responses, and their downregulation suggests a potent anti-inflammatory effect of 13F3 in this context.

In lung tissue, as shown in Figure 6B , 13F3 treatment resulted in suppression of IL-6 (p < 0.05) and MIP-2 (p < 0.01), while TNF-α (p < 0.05) expression was notably elevated. This tissue-specific response highlights the complexity of 13F3’s effects, potentially reflecting differences in local immune microenvironments or signaling pathways.

Interestingly, in peritoneal fluid, the cytokines TNF-α (p < 0.05) and IL-10 (p < 0.05) remained suppressed following 13F3 treatment, consistent with the findings in Figure 6A . However, IL-6, MCP-1, and MIP-2 exhibited no significant changes in expression. In contrast, the chemokine KC (p < 0.05) was significantly suppressed, as illustrated in Figure 6C . This differential regulation suggests that 13F3 exerts distinct effects depending on the tissue or fluid compartment, potentially reflecting variations in cellular composition or microenvironmental factors.

These findings collectively emphasize the tissue-specific and context-dependent nature of 13F3’s immunomodulatory activity. The selective suppression or enhancement of cytokine and chemokine expression by 13F3 across different compartments highlights its potential as a targeted therapeutic agent for modulating immune responses in inflammatory or disease settings.

As illustrated in Figure 7A , treatment with 13F3 suppressed mRNA expression of IL-6 (p < 0.05), IL-10 (p < 0.01), KC (p < 0.05), and MIP-2 (p < 0.01), all of which play pivotal roles in inflammation and immune regulation. In contrast, the expression of MCP-1 (p < 0.05), a key chemokine involved in monocyte recruitment, was notably amplified by 13F3, suggesting selective modulation of chemokine signaling pathways.

A particularly intriguing observation was the significant suppression of PD-L1 (p < 0.05) mRNA expression by 13F3, as depicted in Figure 7B . PD-L1 is a critical immune checkpoint molecule that inhibits T-cell activation, and its downregulation could potentially enhance antitumor immune responses. In contrast, VISTA mRNA expression remained unchanged following 13F3 treatment. This differential regulation highlights the specificity of 13F3 in targeting certain immune checkpoint pathways while sparing others.

Under an optical microscope, iARDS lung injury was characterized by alveolar wall thickening, alveolar edema, hyaline membrane formation, macrophage cell infiltration, alveolar collapse, hemorrhage, and epithelial disruption. These features were collectively demonstrated in the diffuse alveolar damage, which was attenuated by 13F3 (p < 0.05, Figure 8 ).