ATF3 -/- mice were reported by us previously (14). Sex- and age-matched (8–10 weeks) WT and ATF3 KO mice were used during the whole experiments. The mice were bred and kept on a 12 h reverse light/dark cycle and were provided with adequate water and food under specific pathogen-free conditions. The animals were transferred and housed in an animal facility for 3 d of stable housing prior to any experiments. The experiments were conducted in strict accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals, approved by the Ethics Committee of Ruijin Hospital, Shanghai Jiao Tong University School of Medicine.

RAW 264.7 murine macrophages were maintained in Dulbecco’s modified Eagle’s medium (DMEM) containing 10% fetal calf serum at 37°C in a 5% CO₂-humidified incubator. For bone marrow-derived macrophages (BMDMs), isolation and differentiation were performed in line with our published procedure with minor changes (15). Briefly, we flushed the tibias and femurs with precooled phosphate-buffered saline (PBS) using a 25-gauge needle and a 5 mL syringe. Then, the collected bone marrow was gently resuspended into single-cell suspension, and sometimes it was necessary to use red blood cell lysis solution (Sangon Biotech, Shanghai, China) to lyse the red blood cells in it. The cells were centrifuged at 700g for 4 min at room temperature. Then, they were resuspended using a 10 mL conditioned medium (DMEM + 10% fetal bovine serum (FBS) 1% penicillin/streptomycin, and 30% L929) and transferred into a 10-cm cell culture dish. Three days later, 10 mL of fresh conditioned medium was used to replace the old medium in the Petri dishes, and most of the cells were found to be stuck to the Petri dishes. After 7 d of cultivation, fresh medium was replaced, and the cells were used for experiments.

The MRSA strain (USA300) reported by our previous study (16) was cultured in Luria Bertani broth at 37°C until its stable growth phase, and then collected by centrifugation at 8,000 rpm for 5 min. The bacteria were washed three times with PBS and resuspended to an optical density of 0.8 at 600 nm. Then, 500 µL of the bacterial suspension was incubated with an equal volume of 5.0 μM solution of 5(-and 6) carboxyfluorescein diacetate succinimidyl ester (CFDA/SE, Selleck) for 30 min in the dark at 37°C. Subsequently, the stained bacteria were washed three times with PBS and the resuspended bacteria were used for later experiments.

For lung bacterial infections, a total volume of 40 μL of PBS containing S. aureus (5 × 10⁶ CFU/mouse) was injected into the trachea of mice. For survival experiments, we used 2 × 10⁸ CFU/mouse of S. aureus and observed their survival 48 h after the infection. The survival rate of the mice was monitored every 3 h.

BALF collection was performed in line with our published procedure with minor modifications (16). Briefly, we sacrificed the mice by cervical dislocation and exposed the trachea. A 20-gauge catheter was used, and 0.8 mL of PBS was dripped into the lungs and gathered into clean tubes. This process was repeated four times to collect about 3 mL of BALF from each mouse. We used a cell counting plate to count the total number of cells in the alveolar lavage fluid and a flow cytometer to count the number of macrophages and neutrophils in BALF. The remaining BALF was then centrifuged at 700 g for 5 min, and supernatants were stored at −80°C until cytokine analysis.

To evaluate the phagocytic capability, macrophages were incubated with CFSE-labeled S. aureus (MOI 10) for the indicated time periods. Then, the infected cells were washed using PBS, and extracellular bacteria were eliminated by treatment with lysostaphin (20 μg/mL) for 30 min. Macrophages were then collected, and intracellular bacterial loads were quantified by flow cytometry. To determine the bactericidal capability of macrophages, the cells were seeded on coverslips for 24 h and incubated with CFSE-labeled S. aureus (MOI 10) for 2 h. The cells were then washed and further cultured in a fresh medium containing lysostaphin (2 μg/mL) for the indicated time periods. Thereafter, the cells were fixed, and the nuclei were counterstained with DAPI, and then they were observed by fluorescence microscopy.

The intracellular killing assay was conducted following a previously described procedure with slight modifications (17). BMDM cells from WT and ATF3 KO mice were infected with S. aureus (MOI 10) for 2 h. The cells were then washed and treated with a medium containing gentamicin (300 µg/mL) for 30 min to kill the extracellular bacteria. Then, the cells were further cultured in a fresh medium containing gentamicin (100 µg/mL) for the indicated time periods (2 h, 6 h, 12 h, and 18 h). Subsequently, the cells were washed several times with PBS and then lysed with 0.1% Triton X-100 to release the bacteria inside the cells. To estimate the number of bacteria, the lysate was serially diluted with sterile PBS, applied to LB plates, and incubated overnight in a dedicated bacterial incubator.

Total RNA was extracted using TRIzol reagent (Thermo Fisher Scientific, Waltham, MA, USA) in accordance with the manufacturer’s guidelines. One microgram of total RNA was reverse-transcribed to cDNA using SuperScript II (Invitrogen, Carlsbad, CA), and qRT-PCR was performed using SYBR Green technology (Takara, Tokyo, Japan). β-actin was used for normalization, and data were analyzed through the ΔΔCt method. The primers used for the detection in the study were synthesized by GENEWIZ (Suzhou, China), and sequences are shown in Supplementary Table 1 .

MPO activity was assessed as an indicator of neutrophil accumulation in lung tissues by using Myeloperoxidase (MPO) Activity Fluorometric Assay Kit (#K745-100, BioVision, California, USA) in accordance with its instruction.

WT and ATF3 KO BMDM cells were seeded on glass slides and stimulated with S. aureus for the indicated time. The cells were washed twice with prewarmed PBS and fixed with 4% paraformaldehyde solution for 15 min at room temperature. We washed the samples and permeabilized them in 0.1% Triton X-100 for 20 min. After washing the samples three times with PBS, we added the fluorescent phalloidin staining solution to each coverslip and incubated them in a covered container for 50 min at room temperature. We again washed the samples three times with PBS, and recorded immunofluorescence images by laser scanning confocal microscopy.

For histological analysis, mouse lung samples were thoroughly washed in PBS, fixed in 4% (wt/vol) paraformaldehyde for 24 h, embedded in paraffin, and sliced into 5-μm-thick sections. Hematoxylin and eosin staining was performed in line with standard procedures. For immunostaining, the lung sections were deparaffinized, hydrated, and blocked in Dulbecco’s phosphate-buffered saline (DPBS)containing 2% normal goat serum. The slides were then stained with the indicated primary antibody and biotin-conjugated secondary antibody and then incubated with streptavidin-conjugated horseradish peroxidase (HRP). Finally, the slides were incubated with DAB reagent and counterstained with hematoxylin for observation.

The levels of TNFα, IL-6, IL-1β, and IL-12 in the cell culture supernatants and BALF were measured by ELISA kits (R&D Systems, Minneapolis, MN) in accordance with the manufacturers’ guidelines.

BMDMs (1 × 10⁵ cells) were seeded onto the top chamber of an 8-μm pore transwell insert (Corning, NY, USA) in a 24-well plate with media containing 2% FBS. After 36 h, the media within the transwell inserts were carefully removed, and the cells were fixed with 2% paraformaldehyde and stained with 0.2% crystal violet. Cells that did not migrate across the transwell membrane were wiped from the top of the chamber. The migrated cells were viewed using a Nikon DS-F2 microscope. The inserts were then incubated with 30% acetic acid, and the absorbance was read at 570 nm. The experiment was repeated three times.

The cell lysate was prepared with SDS-lysis buffer (Beyotime, Shanghai, China). Total protein concentration was measured by Nano-100 Micro-Spectrophotometer. Protein was then separated by 10% SDS-polyacrylamide microgel and transferred to 0.45-μm polyvinylidene difluoride membranes (Invitrogen). The PVDF membranes were then blocked in Tris-buffered saline containing 5% nonfat dry milk (w/v) (Sangon Biotech) in Tween-20 (TBST) for 1 h at room temperature. After that, the membranes were incubated with the indicated primary antibodies overnight. Then, they were incubated with a secondary antibody conjugated with HRP. We used Tanton™ Chemistar High-sig ECL Western Blotting Substrate (ECL) to display the signal. Full blots of images cropped for presentation are presented in Supplementary Material .

The ChIP assay was performed using the ChIP-IT Express Magnetic Chromatin Immunoprecipitation kit (53008, Active Motif, Carlsbad, CA, USA) according to the manufacturer’s instructions. The chromatin solution was immunoprecipitated using either an anti-ATF3 antibody (CST, USA) or normal anti-IgG antibody (CST, USA) and then incubated with magnetic beads overnight at 4°C with rotation. Next, the DNA-Antibody complexes were washed sequentially by CHIP buffer1 and CHIP buffer2 and then eluted with elution buffer AM2. Then, RT-PCR was performed using purified DNA fragments and primers directed to the specific area spanning the putative ATF3-binding motif. Primer pairs used for ChIP are shown in Supplementary Table 2 .

The plasmids pLVX-flag-REG3β-IRES-Puro and pLVX-flag-REG3γ-IRES-Puro were purchesed from Shanghai Xitubio biotechnology Co., Ltd. To generate stable cell lines expressing REG3β or REG3γ, pLVX-flag-REG3β-IRES-Puro or pLVX-flag-REG3γ-IRES-Puro plasmid together with the packaging plasmids (psPAX2+pMD2.G) were transfected into human embryonic kidney 293T (HEK293T) cells using EZ Trans transfection resgents (Life iLab Bio-Technology, China). The supernatants containing the virus were harvested after 48 h post–transfection and filtered using a 0.45-μm filter. Cells were incubated with viral supernatants plus equal complete medium in the presence of 5 μg/mL polybrene (Sigma) for 24 h. After infection, positive clones were selected by puromycin selection and infection efficacy were validated by immunoblotting assays.

Unless otherwise stated, all data are expressed as the mean ± SD of three independent experiments. Student’s t test or one-way analysis of variance was used for comparison between groups. P values < 0.05 were considered statistically significant. Kaplan–Meier survival analysis with log-rank test was used to evaluate the survival curve. All calculations were performed using the Prism 8 software program (GraphPad Software).

To explore the potential role of ATF3 in the host’s lung defense against S. aureus infection, we used a lethal dose of S. aureus (US 300) (2 × 10⁸ CFUs/mouse) to infect WT and ATF3 KO mice intratracheally and observed survival patterns for 48 h. Although all ATF3 KO mice died within 24 h, 60% of WT mice survived more than two days after infection ( Figure 1A ). To determine whether the difference in survival was due to the difference in bacterial load in various organs, we measured the bacterial load in the lungs, BALF, and extrapulmonary organs after infecting mice with a sublethal inoculum (5 × 10⁷ CFU) of S. aureus. Compared with WT mice, ATF3 KO mice had a higher bacterial load in the lungs and BALF, while there was no significant difference in spleen and blood bacterial load at 6 and 18 h after infection ( Figures 1B – E ). Meanwhile, the total protein (a measure of lung leakage) in the BALF of ATF3 KO mice was higher than that in the WT mice ( Figure 1F ). This suggests that ATF3 has a protective function against lethal S. aureus pneumonia and limits the host bacterial load.

Macrophages and neutrophils are necessary to control S. aureus infection in the lungs (18). Since macrophages and neutrophils are critical to the survival of pneumonia, we investigated whether the loss of ATF3 affects the recruitment of macrophages or neutrophils to the alveolar space during S. aureus pneumonia. ATF3 KO mice displayed profound lung pathology during the infection ( Figures 2A, B ). To further analyze the neutrophil accumulation in the lung parenchyma, we performed MPO assay and showed that the MPO activity was similar between the two genotypes ( Figure 2C ). We also found that WT and ATF3 KO mice had a comparable neutrophil number as reflected by s100a9 staining and Ly6G+/CD11b+ population in the FACS analysis in post-infection lung tissue sections ( Figures 2D, E ). In addition, the total number of infiltrating cells was very similar between the WT and ATF3 KO mice from BALF samples ( Figure 2F ). Interestingly, we noticed that, compared to WT mice, ATF3 KO mice had fewer macrophages recruited into alveolar spaces, as revealed by F/80 staining for the macrophages ( Figure 2G ). We further confirmed ATF3 controls F4/80 macrophage cell recruitment to the lung with histologic observations, the increased F4/80 macrophage cell in the WT mice lung through time-dependent manner during the S. aureus infection (added Figure 1 Supplementary Data ). ATF3 KO mice also had a lower macrophage infiltration from BALF samples the FACS analysis and absolute cell counts ( Figures 2H, I ), which indicated that ATF3 had a host protective effect on survival through macrophage-mediated anti-S. aureus immunity. Collectively, in S. aureus infection, ATF3 exerts host protection by increasing the number of macrophages in the alveolar lavage fluid.

We next examined the effect of ATF3 on the ability of macrophages to clear bacteria. The phagocytic activity of macrophages was initially assessed by carboxyfluorescein succinimidyl ester (CFSE)-labeled S. aureus. There was no significant difference in the number of internalized bacteria between the WT and ATF3 KO macrophages ( Figures 3A, B ), indicating that ATF3 had no significant effect on macrophage phagocytosis. However, the count of bacterial colony-forming units (CFU) showed that the bacterial load was significantly reduced in WT, while ATF3 KO had less reduction during the 6-18 h infection process ( Figure 3C ). To further evaluate the bactericidal ability of macrophages, we conducted a lysostaphin protection test, in which the survival of internalized bacteria was observed with a fluorescence microscope (19). Remarkably, the number of living bacteria was increased in ATF3 KO macrophages compared to WT macrophages and lungs ( Figures 3D, E ), indicating that ATF3 enhanced the ability of macrophages to eliminate invading bacteria.

Given the importance of macrophages in host defense against bacterial infections (5, 20), we next evaluated the effect of ATF3 on macrophages during staphylococcal infection. First, we noticed that ATF3 was induced in BMDM cells and RAW264.7 macrophages in a time-dependent manner after S. aureus infection ( Figures 4A, B ). Our previous study has reported that ATF3 is a crucial response gene in the lipid/cholesterol mediated inflammatory responses (14). The other study also showed that the transcriptional response suggests that the cholesterol de novo synthesis increases considerably in RAW264.7 cells to compare other macrophage cells, by those ATF3 was the top list of upregulated genes (21). It is indicated that ATF3 appears to have differing effects in immune response and is related to the metabolic regulation. Such an altered expression was also observed in the lungs from staphylococcal infection mice ( Figure 4C ). Indeed, the similar regulatory effect between WT and ATF3 KO significantly increased the expression of M1 proinflammatory cytokines, such as IL-1β, TNF-α ( Figures 4D, E ), with the exception of IL-6 and IL-12p40 being more significantly elevated in BMDM of WT ( Figures 4F, G ), while there were no differences in BALF among the major proinflammatory agents involved in the antimicrobial response in vivo ( Figures 4H – K ). In conclusion, our data suggest that ATF3 promotes the inflammatory response of macrophages in the early stages of staphylococcal infection regardness of lung released pro inflammatory cytokines.

There is increasing evidence that antimicrobial peptides (AMP) play a central role in restricting bacterial replication and preventing tissue damage (22). The AMP such as Regenerating islet-derived protein type 3 [Reg3] was regulated by the IL-17, IL-22/23 through stat3 signaling (23). First, we experimentally found no significant difference in STAT3 signaling between WT and ATF3 KO cells with respect to the expression of phosphorylated STAT3 and its target gene SOCS3 ( Figures 5A, B ). Given that IL-22 can mediate the expression of many AMPs, including Reg3 family and S100A8,9 (24), we examined whether ATF3 has an effect on AMP and analyzed the changes in IL-22 levels. Notably, increased levels of IL-22 and the related cytokines IL-17 and IL-23 were observed in macrophages infected by S. aureus, but no differences in induction of IL-17, IL-22, and IL-23 were observed between WT and ATF3 KO cells ( Figure 5C ). Interestingly, the expression levels of the molecules with antibacterial properties, such as S100A8 and Reg3 family genes were increased in WT macrophages but decreased in ATF3-null cells ( Figures 5D, E ). Further we found that WT and ATF3 KO BMDM cells had a comparable induced expression of cytokines, chemokines after S. aureus infection ( Figures 5F, G ). Collectively, our data suggest that ATF3 enhances the production of its antimicrobial effector molecules, thereby potentially enhancing the ability of macrophages to clear bacteria.

We next investigated the functional relevance of ATF3 induction by S. aureus infection with respect to AMP expression. We studied the regulatory role of the transcription factor ATF3 in AMP transcription. To explore whether ATF3 could regulate AMP expression, CiiiDER, a tool for predicting and analyzing transcription factor binding sites was used to analyze the potential ATF3 binding site in these AMP genes (25). In our computational prediction results, those AMP genes had at least one predicted ATF3-binding site ( Figure 6A ). Consistent with Reg3β and Reg3γ as a direct target, chromatin immunoprecipitation analysis revealed that endogenous ATF3 bound to different regions of Reg3β and Reg3γ regulatory elements containing ATF3 or ATF/CREB binding sites; such binding was abolished by ATF3 KO BMDM ( Figures 6B, C ). Hence, ATF3 seems to regulate the levels of Reg3β and Reg3γ via direct transcription control. To evaluate a role for antimicrobial protein Reg3, we overexpressed Reg3β and Reg3γ in macrophages. Immunoblotting of BMDM cell lysates showed the exogenous Reg3β and Reg3γ proteins ( Figures 6D, E ). Reg3β and Reg3γ have potent growth-inhibitory activity against S. aureus ( Figures 6F, G ). These data are consistent with the finding that Reg3 has selective bactericidal activity against pulmonary S. aureus infections (9).

Figure 2H shows that ATF3 KO mice had a lower macrophage infiltration after S. aureus infection. To further investigate how ATF3 controls cell recruitment through motility, we studied the effect of ATF3 on the actin cytoskeleton. The polymerization and depolymerization of filamentous (F) actin have been reported to control the reorganization of the cytoskeleton, which is essential for cell movement through morphological changes (26). We next investigated whether ATF3 may regulate cellular recruitment by altering the actin cytoskeleton. We performed staining for F-actin in WT and ATF3 KO BMDM, revealing that there was no significant difference in actin stress fibers between WT and KO cells. Interestingly, in the case of S. aureus infection, we found that WT cells had significantly more actin stress fibers( Figures 7A, B ).In addition, this impacted F-actin correlated with actin-modifying protein gelsolin (GSN) expression, which was regulated by ATF3, as described in our previous study (27). Positive or negative regulation of GSN by ATF3 is dependent on cofactors, such as different histone deacetylases (HDACs) in a context-dependent manner. Results showed that ATF3 negatively regulated the GSN mRNA and proteins expression in BMDM and GSN expression was higher in ATF3 KO cells compared to WT cells after S. aureus infection ( Figure 7C, D ). We further examined the role of ATF3 in cell motility and found no significant difference in the degree of migration of WT BMDM and ATF3 KO BMDM in the unstimulated conditions; however, WT BMDM were significantly more migratory than ATF3 KO cells in S. aureus-stimulated conditions ( Figures 7E, F ). These data show that ATF3 regulates cell motility in vitro and may drive macrophage recruitment in lung tissues.