Introduction

Salmonella is a prevalent zoonotic food-borne pathogen that can lead to foodborne illness1. The consumption of contaminated food or water can lead to the entry of the pathogen into the body and cause ST infection2. Salmonella-induced food poisoning poses a significant public health concern worldwide2. Millions of individuals contract Salmonella from consuming contaminated food and experience symptoms such as enteric fever, diarrhea, fever, and vomiting each year3. Young chickens, piglets, children, elderly individuals and individuals with compromised immune systems are particularly vulnerable to Salmonella infection2.

The overuse and misuse of antibiotics over a long period of time has caused many bacteria to develop antibiotics resistance4. This has made treatment for bacterial infections more difficult and has increased the time and economic cost required for patients to recover4. In view of drug resistance, countries around the world have formulated corresponding measures to reduce the harm caused by drug-resistant bacteria5. Antibiotics should be judiciously utilized in medical practice and not overused6. The use of antibiotics in agriculture should also be carefully regulated, and the safety of livestock and poultry products should be guaranteed7. Additionally, there is a need to intensify research and development efforts for new antibiotics. However, the process of developing new antibacterial drugs is time-consuming and economically burdensome, while the emergence of drug-resistant bacteria has led to the development of effective antibacterial medications8. Hence, there is an urgent need to develop novel antibacterial agents.

New antibacterial drugs that target key virulence factors in bacterial disease processes, known as antivirulence drugs9, offer distinct advantages over traditional antibiotics. By not directly affecting essential bacterial growth elements, they present a lower risk of inducing bacterial resistance9. This makes them a promising alternative to antibiotic therapy for treating infectious diseases caused by bacteria9. The T3SS encoded by the Salmonella pathogenicity island, which is divided into T3SS-1 and T3SS-2, plays key roles in the invasion and intracellular proliferation processes of Salmonella10,11. Salmonella can use its SPI-1-encoded T3SS-1 to inject effector proteins into host cells, thereby achieving invasion and replication in host cells and leading to infection and disease10,11. During the infection process, Salmonella releases various effector proteins through T3SS-1, which can interfere with host cell signal transduction, promote endocytosis, and inhibit immune responses, ultimately causing damage to host cells and triggering severe intestinal infections10,11. T3SS-1 is crucial for bacterial invasion of cells and their intracellular replication.

The important role of Salmonella T3SS-1 in the infection process and the elucidation of its pathogenic mechanism have provided a theoretical basis and ideal targets for the development of novel antibacterial drugs and vaccines. Therefore, inhibitors targeting T3SS-1 can exert antibacterial effects without exerting bacteria-selective pressure. This approach is expected to be one of the solutions for the development of new antibiotic replacement strategies9. Many studies have investigated antibacterial drugs that target bacterial virulence factors. Flavonoids with antibacterial properties obtained from medicinal plants chemically inactivate substrates of type III protein secretion12. Prazosin has strong inhibitory effects on quorum sensing, the T3SS, and bacterial communication, indicating its potential as a promising antivirulence treatment13. However, this still cannot meet the treatment needs for clinically resistant bacterial infections. Therefore, in the preliminary stage of this study, a cell model of ST infection was used to screen for inhibitors of ST invasion, and it was found that PPR can significantly inhibit the invasion of ST into host cells. This study investigated the molecular mechanism by which PPR inhibits invasion and the therapeutic effects of PPR in a mouse model of ST infection. This study provides a hit compound for the research of therapeutic agents for treating ST infection and provides a theoretical basis for the formulation of future treatment strategies.

Results

PPR has the potential to prevent ST from invading HeLa cells

To identify compounds that can inhibit ST invasion of host cells, a screening based on the gentamicin protection assay was carried out (Fig. 1A). The concentration used for screening assay was 100 µg/mL. The relationship between dose and inhibition efficiency was further detected, and the results revealed that, as the concentration of PPR increased, its inhibitory effect gradually increased (Fig. 1B). At a concentration of 16 µg/mL, PPR inhibited the ST invasion of HeLa cells by 90% (Fig. 1B). The immunostaining results also confirmed these findings (Fig. 1C). These results suggest that PPR can inhibit the invasion of HeLa cells by ST.

Fig. 1
figure 1

The results of the gentamicin protection assay revealed that PPR has the ability to prevent the invasion of HeLa cells by ST. (A) Screening process for inhibitors of ST invasion of HeLa cells. The concentration used in the screening assay was 100 µg/mL. (B) The ability of PPR to inhibit ST invasion of HeLa cells increased in a dose-dependent manner with increasing PPR concentration (n = 3). (C) Immunofluorescence analysis was used to detect the effects of PPR on ST invasion of HeLa cells (scale bar, 50 μm). The data are presented as the means ± standard deviations and were subjected to analysis via one-way ANOVA. NS, not significant; **p < 0.01.

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PPR does not impact the natural growth or mobility of ST cells or the viability of HeLa cells

Both changes at the level of the pathogen and the host cell may be factors affecting the invasion of ST. To determine whether the inhibition of ST invasion by PPR is due to its inhibition of bacterial growth, we determined the MIC of PPR. The results revealed that the MIC of PPR was > 1024 µg/mL (Fig. 2A). When different concentrations of PPR were applied to SL1344, the results revealed that ≤ 64 µg/mL PPR had no effect on the natural growth of ST (Fig. 2B). The mobility of bacteria is an important factor that affects their ability to invade host cells. To determine whether PPR affects the ability of ST to move and thus its ability to invade HeLa cells, we conducted swimming and swarming assays. The results revealed that PPR had no effect on the swimming ability (Fig. 2C) or swarming ability (Fig. 2D) of ST. To investigate the potential impact of PPR on host cells via the inhibition of ST invasion, we conducted a cell counting kit-8 (CCK8) assay to evaluate its cytotoxicity to HeLa cells. The findings revealed that PPR, at concentrations ≤ 128 µg/mL, did not exhibit toxicity toward HeLa cells (Fig. 2E). These results suggest that PPR has the ability to hinder ST invasion of HeLa cells without compromising their natural bacterial growth or motility or causing cytotoxic effects on the HeLa cells.

Fig. 2
figure 2

PPR has no effect on the natural growth or mobility of ST cells or on the viability of HeLa cells. (A) Measurement of the MIC for PPR against ST. (B) Influence of PPR on the in vitro growth curve of ST. (C-D) The impact of PPR on ST swimming (n = 3). (C) and swarming (D). (E) PPR did not exhibit cytotoxic effects on HeLa cells (n = 3). The data are presented as the means ± standard deviations and were subjected to analysis via one-way ANOVA. NS, not significant.

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PPR inhibits the secretion of T3SS-1 effector proteins

T3SS-1 is an important virulence apparatus for the invasion of host cells by ST10. It can secrete various effector proteins, including SipA, SipB, SipC, and SopB, to enable the invasion of host cells. Therefore, we aimed to determine whether PPR inhibition of the invasive ability of ST is achieved by suppressing the function of the ST T3SS-1. We detected the secretion function of T3SS-1 using the β-lactamase (TEM) fluorescence reporting system (SipA-TEM). The results revealed that PPR inhibited SipA-TEM translocation to HeLa cells (Fig. 3A), and the ability of PPR to inhibit the translocation of SipA-TEM increased as the concentration of PPR increased (Fig. 3A).

To determine the effect of PPR on the protein secretion capacity of the T3SS effector protein, we analyzed the secretion of T3SS effector proteins in ST with PPR via trichloroacetic acid (TCA) precipitation. The results showed that PPR could also inhibit the secretion of T3SS-1 effector proteins, such as SipA, SipB, SipC, SopB, and SipD (Fig. 3B). In addition, the secretion of FliC was inhibited by PPR (Fig. 3B). To determine whether the inhibition of SipA secretion was due to its suppression of effector protein expression, we assayed the expression of SipA via Western blotting, and the results revealed that PPR had no effect on the expression of the effector protein SipA (Fig. 3C).

Fig. 3
figure 3

PPR suppresses the secretion of T3SS effector proteins. (A) The β-lactamase reporting system was used to detect the effect of PPR on the translocation of the ST T3SS (scale bar, 100 μm). (B) TCA precipitation assay combined with Coomassie Brilliant Blue (CBB) staining to detect the effects of PPR on T3SS effector protein secretion. (C) Detection of the impact of PPR on the expression of the effector protein SipA via WB.

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HilA is an important regulatory molecule for the expression and secretion of T3SS effector proteins, and it is a key node in the regulatory network of the T3SS10 (Fig. 4A). To investigate whether HilA mediates the inhibitory effect of PPR on T3SS secretion, we overexpressed the HilA protein and detected the influence of HilA overexpression on the inhibitory effect of PPR on T3SS function. The WB results revealed that PPR had no effect on the expression of T3SS effector proteins when successful overexpression of HilA was achieved (Fig. 4B). The TCA precipitation CBB results revealed that the inhibitory effect of PPR on T3SS-1 secretion remained after the overexpression of HilA (Fig. 4C). The WB results also revealed the inhibitory effect of PPR on the secretion of SipA (Fig. 4D). Therefore, the inhibitory effect of PPR on T3SS secretion is independent of HilA.

Fig. 4
figure 4

The ability of the PPR to suppress T3SS secretion is unrelated to HilA. (A) The simplified regulatory network for the T3SS. (B) Western blotting was used to assess the impact of HilA overexpression on the production of T3SS effector proteins. (C) A CBB assay was used to investigate the effect of HilA overexpression on the secretion of T3SS effector proteins. (D) Western blotting was used to examine the effect of overexpression of HilA on the secretion of the T3SS effector protein SipA.

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InvC ATPase is one of the core components of the T3SS of Salmonella and can hydrolyze ATP to provide energy for the secretion/translocation of T3SS effector proteins14. In this study, we established an InvC ATPase reaction system using the InvC recombinant protein construct developed in our laboratory (Fig. 5A) to determine the effect of PPR on the activity of InvC ATPase. The results revealed that PPR had no effect on the ATPase activity of InvC (Fig. 5B). Thus, PPR does not target InvC to inhibit the secretion function of T3SS-1.

Fig. 5
figure 5

PPR does not impact InvC ATPase activity. (A) ATPase reaction system based on InvC (n = 3). (B) PPR had no effect on the ATPase activity of InvC (n = 3). The data are presented as the means ± standard deviations and were subjected to analysis via one-way ANOVA. NS, not significant; **p < 0.01.

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PPR provides effective protection for mice from ST infection

These findings suggest that PPR has the potential to inhibit ST invasion of host cells by suppressing the secretory function of T3SS-1. To investigate the efficacy of PPR in treating ST infections, a mouse enteritis model of ST infection was established. The therapeutic impact of PPR on the model was assessed. The results from the mortality test revealed that all the mice in the ST infection group succumbed after 10 days of infection, while the survival rate of the mice in the PPR group was 50% (Fig. 6A). Analysis of colony colonization in target organs revealed a significantly lower level of ST colonization in the spleen of the mice in the PPR treatment group than in those in the ST infection group (Fig. 6B). Anatomical examination revealed no significant changes in liver pathology before or after PPR treatment (Fig. 6C). In the ST infection group, intestinal obstruction was observed, with thinning of the intestinal wall, increase ed fragility, and a shortened length. Conversely, the stool in the PPR treatment group was similar to that in the PBS blank group (Fig. 6C). HE staining revealed shedding and thinning of intestinal epithelial cells in the ST infection group, which was notably alleviated in the PPR treatment group (Fig. 6D). Liver inflammatory cell infiltration increased in the mice in the ST infection group but was significantly reduced following PPR treatment (Fig. 6D). Additionally, splenic inflammatory cell infiltration was markedly weaker in the PPR treatment group than in the ST infection group (Fig. 6D). The inflammation scores for the intestine, liver, and spleen revealed a significant reduction in inflammation due to ST infection following PPR treatment (Fig. 6E). These findings suggest that PPR is effective in treating enteritis caused by ST infection in a mouse model.

Fig. 6
figure 6

PPR protects mice from ST infection. (A) The impact of PPR on the survival of ST-infected mice (n = 10). The survival rate was analyzed via log-rank analysis (n = 5). (B) The impact of PPR on ST colony formation in the liver and spleen of infected mice. (C) Autopsy image of the effect of PPR on the ST-infected mice. (D) The influence of PPR treatment on pathological changes in mice infected with ST (scale bar, 100 μm). (E) Inflammation scores based on HE staining results (D) (n = 3). The data are presented as the means ± standard deviations and were subjected to analysis via one-way ANOVA. **p < 0.01.

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Discussion

The issue of drug resistance is increasingly severe, necessitating the urgent development of new antibacterial drugs4,15. Traditional antibacterial medications primarily exert selective pressure on bacteria by targeting essential components of bacterial growth or vital life processes15. In this study, PPR effectively inhibited the ST invasion of host cells by suppressing T3SS-1 secretion without affecting bacterial growth or virulence. These findings suggest that PPR not only has an anti-infective function but also does not induce bacterial resistance, making it a promising candidate for antivirulence strategies with potential minimal impact on normal gut microorganisms.

Previous studies have identified compounds such as tannic acid, harmine, and fraxetin as inhibitors of T3SS-1 secretion, primarily by targeting the regulatory network of T3SS-1 to suppress the expression of effector proteins16,17,18. Fisetin was found to specifically target HilD to inhibit T3SS-1 effector protein expression19. Our study revealed that PPR selectively inhibits T3SS-1 secretion without affecting effector protein expression. Mechanistic exploration revealed that PPR does not impact the regulatory network centered on HilA or the activity of the T3SS ATPase InvC. However, further investigations are needed to elucidate the detailed molecular mechanism underlying the inhibitory effect of PPR on T3SS-1 secretion. Nevertheless, the discovery of PPR has significant implications for the development of specific inhibitors of T3SS-1.

Flagellin C (FliC) is a flagellin protein of Salmonella20. When TCA precipitation was used to detect the effect of PPR on the secretion of the T3SS-1 effector protein, it was unexpectedly discovered that PPR also significantly suppressed the expression of FliC. However, the results from the swimming and swarming assays indicated that PPR did not impact ST motility. This remains a key area for future exploration.

PPR is a natural anthraquinone with multifaceted pharmacological activities that is isolated from the roots of Rubia cordifolia2122. This study revealed that PPR can target T3SS-1 and inhibit the invasion of host cells by ST from a pathogenic perspective. However, this study detected only the impact of PPR on cell viability at the cellular level and revealed that PPR alleviates the inflammatory damage caused by ST infection; however, substantial exploration of the impact of PPR on host physiological functions has not been conducted. Many previous studies have shown that PPR has a wide range of pharmacological characteristics, including antioxidant23, neuroprotective, antibacterial24,25, neuroprotective26and anticancer27 effects. Therefore, we hypothesize that the protective effects of PPR against ST infection may not only be due to its ability to inhibit ST T3SS-1 secretion and suppress inflammation caused by infection but also involve other host physiological functions that are altered during the course of infection. This represents a potential area for future investigations. Furthermore, the pharmacokinetics and toxicology of PPR are also key research directions that we must explore in the future for the application of PPR treatment for ST infections.

A previous study has demonstrated that rats fed with a basal diet containing 1% purpurin over a long period (520 days) presented widespread renal lesions and a certain extent of bladder tumor28. In the V79-HGPRT mammalian cell mutagenicity assay, 5 µg/ml of purpurin exhibited mutagenic effects, while in comparison with the strong mutagens 1,5-DHA and emodin in primary rat hepatocytes, the mutagenic effect of purpurin was relatively weaker, and 1 µg/ml of purpurin had significant transforming activity on C3H/M2 mouse fibroblasts29,30. In this study, the results of the cytotoxicity assay indicated that no more than 128 µg/ml of purpurin showed no obvious cytotoxicity by the CCK8 assay. However, the carcinogenicity, teratogenicity and mutagenicity still need to be systematically evaluated in accordance with the existing drug evaluation criteria to determine whether purpurin can be used in clinical practice at all.

In summary, this study revealed that PPR can inhibit the secretion of the T3SS and thus suppress the invasion of ST into host cells, effectively protecting mice from ST infection. These findings can provide a hit compound for the development of anti-Salmonella virulence drugs and serve as a reference for the development of alternative drugs based on antivirulence strategies.

Materials and methods

Bacterial strains, cell lines, culture conditions and compounds

The strains and primers used in this study are listed in Tables 1 and 2 respectively. The ST strain SL1344 was subsequently grown in Luria–Bertani (LB) broth supplemented with antibiotics at 37 °C. SPI-1 gene expression in ST was induced by adding 0.3 M NaCl to the medium. HeLa cells were cultured in DMEM (Sigma) supplemented with 10% fetal bovine serum (FBS). The incubation of HeLa cells occurred in a CO2-controlled environment at 37 °C. The natural compounds containing purpurin (PPR) used for the screening assay were purchased from Chengdu Herbpurify Co., Ltd (Chengdu, China) (https://www.herbpurify.com/).

The sipA and bla(M) genes were fused and cloned into the pEX233 vector. The recombinant plasmid was transformed into SL1344 WT and SL1344 ΔinvA respectively. The obtained strains were respectively named SL1344 WT (SipA-TEM) and SL1344 ΔinvA (SipA-TEM). The pJL03 vector and pJL03-hilA recombinant plasmid were transformed into SL1344 WT. The obtained strains were named SL1344 pJL03-vector and SL1344 pJL03-hilA respectively. The SL1344 ΔflhCwas constructed using a λ-red-based method31.

Table 1 Bacterial strains used in this study.
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Table 2 The primers used in this study.
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Bacterial invasion assays

The ST culture was diluted in 3 mL of 0.3 M NaCl LB at a 1:20 ratio. Then, increasing concentrations of PPR were added to the aliquoted cultures, which were subsequently grown at 37 °C with consistent agitation at 220 rpm/min for 4 h. In a 24-well plate, 4 × 105 HeLa cells were challenged with SL1344 at an MOI of 100. After infection for 1 h, the cells were washed with prewarmed PBS three times. The plates were replaced with fresh medium containing 100 µg/mL gentamicin and incubated in the incubator for 40 min to kill the extracellular bacteria. The cells were washed with PBS and lysed with 0.02% saponin. The lysates were spread onto LB plates and grown at 37 °C for 18 h before the CFUs were enumerated.

Determination of the minimum inhibitory concentration (MIC) and in vitro growth curve

The MIC was determined via the CLSI broth microdilution method16. Resazurin was added to the 96-well plate of SL1344, which was subsequently incubated overnight at 37 °C. The color change in the 96-well plate was observed. The MIC was determined on the basis of the color change (a pink color indicating the reduction of fuchsin from the oxidized blue state indicates bacterial growth in the well). The in vitro growth curve was generated as follows: A bacterial culture that had been cultured overnight was diluted with fresh LB to an OD600nm value of 0.1. After the various concentrations of PPR were added, the mixture was activated for 5 h at 37 °C and 220 rpm/min, and the OD600nm was measured every 30 min and plotted as a curve.

Swimming and swarming assay

A gradient concentration of PPR was added to the swimming medium (LB medium with 0.3% (w/v) agar) and swarming medium (0.5% (w/v) agar, 25 g/L LB, 5 g/L glucose, and drying at 25 °C for 1 h). For the swimming assay, 5 µL of bacterial suspension (1 × 105 CFUs) was added to the liquid surface of the swimming medium, which was subsequently incubated upright at 37 °C for 7 h. The radius of the bacterial movement range was calculated. For the swarming assay, swarming medium was inoculated with 5 µL of bacterial suspension (1 × 105 CFUs) and incubated at 37 °C for 7 h. The radius of the bacterial movement range was calculated.

Cell counting Kit-8 (CCK-8) assay

The CCK-8 test was performed with a CCK-8 kit (#C0039, Beyotime, China) following the manufacturer’s instructions. Briefly, HeLa cells (1 × 104/well) were seeded into 96-well plates and cultured for 12 h at 37 °C. After three washes, the medium was replaced with media containing varying concentrations of PPR, and the mixture was incubated for 6 h at 37 °C. Then, CCK-8 solution (10 µL/well) was added to each well, and the samples were incubated for 1 h before analysis. Finally, the optical densities of the wells were measured at 450 nm via a microplate reader.

β-Lactamase (TEM) reporting system

The translocation of T3SS effector proteins in ST was determined via a transmission electron microscopy (TEM) reporting system32. SipA-TEM was subsequently transferred into wild-type SL1344. The T3SS-1-defective strain (ΔinvA) was transformed into a negative control strain. ST cultured overnight was diluted in fresh LB containing 0.3 M NaCl at a ratio of 1:30. After various concentrations of PPR were added, the culture was incubated at 37 °C and 220 rpm/min for 4 h. HeLa cells were inoculated into 96-well plates at a concentration of 1.2 × 104 per well, and SL1344 cells were infected with an MOI of 20 for 2 h. The cells were washed with Hank’s balanced salt solution (HBSS) twice to remove extracellular bacteria. A reaction mixture of 6×CCF4/AM (K1095, Thermo Fisher) was added to each well. The transport of effector proteins was observed under a fluorescence microscope after incubation at room temperature for 1 h in the dark.

Trichloroacetic acid (TCA) precipitation

The secretion of T3SS effector proteins was determined via the TCA precipitation method18. SL1344 was cultured in high-salt LB with various concentrations of PPR for 5 h. Two milliliters of the culture were collected and centrifuged at 12,000x g for 5 min. The bacterial precipitates were resuspended in 100 µL of 1x SDS loading buffer. TCA was added to the supernatant at a ratio of 1/9 volume and precipitated overnight at 4 °C. After centrifugation at 12,000×g for 20 min, the bacterial pellet was resuspended in 20 µL of 1xSDS loading buffer. Effector protein secretion was detected by SDS‒PAGE combined with Coomassie brilliant blue (CBB) staining or Western blot (WB) analysis with specific antibodies.

ATPase assay

(1) ATPase reaction: Referring to existing methods33 and making necessary improvements, the reaction system consists of 45 µL, including 4.5 µL 10× reaction buffer (500 mM HEPEs, 300 mM KCl, 300 mM ammonium acetate, 10 mM DTT, 50 mM MgCl2), 4.5 µL 10× BSA (5 mg/mL), 1 µL ATP (10 mmol/L, pH = 7.0), 1 µL purified InvC (0.8 µM), and different concentrations of PPR. Finally, H2O was added to bring the reaction system to a total volume of 45 µL, and the mixture was incubated at room temperature (25 °C) for 20 min. (2) Color development: 40 µL of developer solution (containing 30 µL of 0.045% cochineal solution, 10 µL of 4.2% ammonium molybdate tetrahydrate, and 0.8 µL of 1% Triton X-100) was added. Color was developed at room temperature for 1 min. (3) The reaction was stopped: 5 µL of 34% sodium citrate was added, and the OD660nm was measured within 2 h.

Western blotting

An overnight culture of SL1344 cells was diluted at a ratio of 1:30 in fresh LB medium containing 0.3 M NaCl, and a gradient concentration of PPR was subsequently added, after which the mixture was incubated for 4 h. The cells were then collected, 40 µL of 1× SDS loading buffer was added, and the mixture was boiled for 5 min. The lysates were subjected to SDS‒PAGE and transferred onto a nitrocellulose membrane. The membrane was blocked with 5% skim milk (w/v) in TBST for 1 h at room temperature before incubation with primary antibodies. The following antibodies were used: homemade rabbit anti-SipA antibody (1:1000), rabbit anti-HilA antibody (1:1000), rabbit anti-FilC antibody (1:1000, ab93713, Abcam), and rabbit anti-ICDH antibody (Sigma, abs2090, 1:5000). After washing with TBST three times, the membrane was incubated with the corresponding secondary antibodies (ab175775 and ab175781; Abcam) at room temperature for 1 h. Protein abundance was analyzed via the Odyssey CLx Imaging System (LI-COR) and Tanon 4600 imaging system (Biotanon), all original gels/blots images were shown in supplementary information.

Immunostaining

To observe the invasion of ST, HeLa cells were placed on coverslips in 24-well plates. The ST cultures were treated with PPR and then used for infections following the methods described earlier. The immunostaining methods were as follows. The samples were fixed with 4% (w/v) paraformaldehyde at room temperature for 30 min, incubated with 0.02% (v/v) Triton X-100 for 5 min, blocked with 4% goat serum at 37 °C for 30 min, and incubated with primary antibody at room temperature for 1 h. The following antibodies were used: rabbit anti-Salmonella antibody (Abcam, ab69238, 1:1000) and secondary antibody (Alexa Fluor 488) (Thermo Fisher Scientific). The nuclei were stained with Hoechst 33,342. An Olympus IX83 microscope was used to observe and collect images.

Animal experiments

The animal experiments described in this report were carried out in accordance with the guidelines of the National Institutes of Health and approved by the Institutional Animal Care and Use Committee of Jilin University (permit number: SY202405300).

BALB/c mice (6–8 weeks old, 18–20 g) were purchased from Liaoning Changsheng Biotechnology Co., Ltd. The mice were randomly divided into 3 groups: the SL1344 infection group, the PPR treatment group and the PBS control group (survival rate, n = 10; target organ colony colonization, n = 5). All the mice were given streptomycin (5 g/L) in drinking water for 3 days. On the 4th day postinfection, the mice in the infection group and the PPR treatment group were infected with SL1344 by intragastric administration (the survival rate test was 1 × 107 CFUs; the attack dose of the target organ colony colonization test was 5 × 106 CFUs). The mice in the treatment group were administered PPR at a dose of 50 mg/kg every 12 h by gavage, with a volume of 100 µL. The ST group and PBS control group mice were administered with 100 µL DMSO (DMSO is the solvent of PPR). On the 8th day postadministration, the mice were euthanized using an overdose of carbon dioxide (CO2) in accordance with approved ethical protocols. After euthanasia, the mice were dissected, and the spleen and liver tissues were collected aseptically, homogenized, diluted, coated with LB plates and counted to determine their CFUs. Mouse tissues were collected, fixed with 4% (w/v) paraformaldehyde for 48 h, and stained with hematoxylin‒eosin (HE) for histopathological analysis. The inflammation score was based on the number of suppurative foci in the hepatic lobules, the number and size of the germinal centers in the spleen, the integrity of the intestinal epithelial cell layer and the infiltration of inflammatory cells in the lamina propria. The scoring criteria were as follows: mild inflammation, 0–4 points; moderate inflammation, 5–8 points; and severe inflammation, 9–12 points.

Statistical analysis

The data were analyzed via GraphPad Prism 10 software (GraphPad Software, La Jolla, CA, USA; available at https://www.graphpad.com/). A t test was employed to compare the two groups of samples. One-way analysis of variance was used to analyze the data of 3 groups and more than 3 groups. The survival rate of the mice was assessed through log-rank analysis. The p value in the figure is indicated as follows: NS indicates no significant difference. ***p < 0.001; **p < 0.01; *p < 0.05.