ABSTRACT
Cronobacter sakazakii poses a significant threat, particularly to neonates and infants. Despite its strong pathogenicity, understanding of C. sakazakii biofilms and their role in infections remains limited. This study investigates the roles of HmsP and c-di-GMP in biofilm formation and identifies key genetic and proteomic elements involved. Gene knockout experiments reveal that HmsP and c-di-GMP are linked to biofilm formation in C. sakazakii. Comparative proteomic profiling identifies the lysozyme inhibitor protein LprI, which is downregulated in hmsP knockouts and upregulated in c-di-GMP knockouts, as a potential biofilm formation factor. Further investigation of the lprI knockout strain shows significantly reduced biofilm formation and decreased virulence in a rat infection model. Additionally, LprI is demonstrated to bind extracellular DNA, suggesting a role in anchoring C. sakazakii within the biofilm matrix. These findings enhance our understanding of the molecular mechanisms underlying biofilm formation and virulence in C. sakazakii, offering potential targets for therapeutic intervention and food production settings.
IMPORTANCE
Cronobacter sakazakii is a bacterium that poses a severe threat to neonates and infants. This research elucidates the role of the lysozyme inhibitor LprI, modulated by HmsP and c-di-GMP, and uncovers a key factor in biofilm formation and virulence. The findings offer crucial insights into the molecular interactions that enable C. sakazakii to form resilient biofilms and persist in hostile environments, such as those found in food production facilities. These insights not only enhance our understanding of C. sakazakii pathogenesis but also identify potential targets for novel therapeutic interventions to prevent or mitigate infections. This work is particularly relevant to public health and the food industry, where controlling C. sakazakii contamination in powdered infant formula is vital for safeguarding vulnerable populations.
KEYWORDS: proteomics, biofilm forming, virulence, lysozyme inhibitor, Cronobacter sakazakii
INTRODUCTION
Cronobacter species are recognized as opportunistic pathogens capable of causing severe infections with elevated mortality rates, particularly among neonates and infants who are preterm, low birth weight, or immunocompromised (1, 2). Clinical presentations commonly associated with Cronobacter infections include necrotizing enterocolitis, septicemia, and neonatal meningitis (3–5). The mortality rate among neonates afflicted with Cronobacter infections varies between 27% and 80%, with 20% of survivors experiencing subsequent neurological complications (6, 7).
Cronobacter species have been isolated from a variety of inert and bioactive surfaces and are prevalent in the food industry (8, 9). These bacteria can persist in production environments, with certain strains reported to survive in factories for months or even years (10). Infant infections are often associated with the consumption of contaminated powdered infant formula (PIF) (11, 12). Currently, the genus Cronobacter includes seven species: C. sakazakii, C. malonaticus, C. turicensis, C. muytjensii, C. dublinensis, C. universalis, and C. condiment (4, 13). Of these, C. sakazakii is the most frequently isolated from patients (14). Given the widespread use of powdered infant formula (PIF) by infants, preventing C. sakazakii contamination in food has become a crucial strategy in reducing the risk of infant infections.
During the manufacturing process of PIF, the utilization of high-temperature and drying procedures is common practice, leading to a reduction in bacterial levels (15). Nevertheless, the presence of Cronobacter contamination may arise during post-processing stages, particularly in storage and packaging areas, as well as on various equipment surfaces, such as drum dryers, packaging machines, drying towers, and air filters (8, 16). Pathogens that adhere to these surfaces have the capability to establish biofilms, predominantly consisting of extracellular polymeric substances (EPSs) (17). The formation of biofilms serves as a critical mechanism for bacteria to resist external stresses within the production environment (18).
Numerous studies have demonstrated that bacterial cells within biofilms display heightened resistance to detergents and disinfectants in comparison to planktonic cells (19–21). Additionally, investigations on bacterial pathogens have revealed that biofilm formation promotes recurrent and enduring infections in vivo (22, 23). Enhanced comprehension of the mechanisms underlying biofilm formation and maturation is essential for the advancement of control measures and interventions. Recent studies utilizing random transposon mutagenesis have revealed the genes associated with cellulose and flagellar biosyntheses, as well as virulence in Cronobacter species (9, 24). Nevertheless, the current understanding of Cronobacter biofilms is relatively limited in comparison to other pathogenic organisms renowned for their biofilm-forming abilities, such as Pseudomonas aeruginosa (25), Salmonella (26), Escherichia coli O157 (27), and Staphylococcus aureus (28).
Recent developments in omics technologies have provided new opportunities for identifying key factors implicated in biofilm formation, presenting potential targets for novel antimicrobial treatments (26, 29). Given that the proteins of C. sakazakii BAA-894 have been clearly annotated (30, 31), this strain has been selected as the starting organism. Our research employs comparative proteomics to study biofilm formation in wild-type and gene knockout strains, aiming to uncover genes involved in C. sakazakii biofilm formation. Based on a thorough examination of the current literature, we have pinpointed nine candidate genes that may play a role in environmental stress tolerance, as well as the endogenous plasmid pESA3. Our study involved extensive gene knockout experiments and subsequent evaluation of phenotypic characteristics, which demonstrated that the deletion of LprI significantly hinders both biofilm formation and virulence in C. sakazakii.
RESULTS
Biofilm-forming abilities of C. sakazakii mutants
Nine homologous genes, including kdpE, recA, rpfB, envZ/ompR, glrK, flgK, treR, hmsP, and ESA_01588 (which encodes c-di-GMP phosphodiesterase), associated with biofilm formation in other bacteria were selected for study in C. sakazakii. Additionally, an endogenous plasmid in C. sakazakii was selected to examine its specific role in biofilm formation within this organism. The biofilm formation capabilities of the resultant C. sakazakii mutants were subsequently assessed, as illustrated in Fig. 1. The kdpE, recA, and rpfB knockout strains displayed diminished biofilm formation coupled with decreased growth rates. Conversely, the hmsP knockout strain exhibited reduced biofilm formation without a notable alteration in growth rate. In contrast, the knockout of envZ/ompR, glrK, pESA3, flgK, and treR had no discernible impact on biofilm formation. Additionally, the knockout of ESA_01588 did not affect growth rate but did enhance biofilm formation in C. sakazakii. These findings highlight the crucial roles of hmsP and c-di-GMP in biofilm formation in C. sakazakii.
Fig 1.
Biofilm formation and growth curves of wild-type (WT) and knockout strains. (A) Biofilm formation of WT and knockout strains for kdpE, recA, rpfB, envZ/ompR, glrK, flgK, treR, hmsP, ESA_01588 (encoding c-di-GMP phosphodiesterase), and pESA3. Biofilm formation was quantified after a total incubation period of 72 h using crystal violet staining, with the absorbance measured at 570 nm. Significant differences from the WT are marked with an asterisk (*), determined by t-test with three biological replicates. (B to F) Growth curves of WT and the knockout strains that showed significant differences in biofilm formation (A).
Comparative proteomic profiling to identify protein related to biofilm forming
The hypothesis posited that the disruption of hmsP and ESA_01588 would impact bacterial biofilm formation through the regulation of other biofilm-associated genes. To elucidate the potential regulatory mechanisms of biofilm formation modulated by HmsP and c-di-GMP, a comparative proteomic analysis was undertaken, resulting in the identification of 1,447 proteins. Notably, the protein abundances exhibited significant variability across samples: the deletion of hmsP was associated with a notable increase in 90 proteins and a decrease in 146 proteins (Fig. 2A), whereas the deletion of c-di-GMP resulted in a significant increase in 56 proteins and a decrease in 83 proteins (Fig. 2B). Analysis of the Venn diagram indicated that the lipoprotein lysozyme inhibitor N-terminal domain-containing protein encoded by lprI (ESA_00658) was the sole protein downregulated in the hmsP knockout and upregulated in the c-di-GMP knockout (Fig. 2C). This suggests that lprI may serve as a potential regulator of biofilm formation, potentially being co-regulated by HmsP and c-di-GMP.
Fig 2.
Proteomic analysis of hmsP and ESA_01588 deletion mutants. (A-B) Volcano plots illustrating the impact of hmsP knockout (A) and ESA_01588 knockout (B) on protein abundance. Significant changes, marked in red (upregulated) and green (downregulated), show fold changes greater than two with P < 0.05. (C) Venn diagram showing the overlap of proteins downregulated in ΔhmsP (blue) and upregulated in ΔESA_01588 (yellow) strains, with the intersecting proteins listed.
Impaired biofilm forming and virulence of knockout strain encoding lipoprotein LprI
To investigate the function of lprI in C. sakazakii, a knockout strain of lprI was generated through the utilization of a suicide plasmid technique. The absence of lprI did not impact the growth rate of the bacteria; however, it did significantly hinder the formation of biofilms. The formation of biofilms is often associated with the ability of bacterial pathogens to establish persistent infections. This research utilized a rat oral infection model to assess the role of lprI in the virulence of C. sakazakii. Three-day-old rats were randomly assigned to three groups and orally exposed to wild-type (WT), ΔlprI, and ΔlprI complemented strains of C. sakazakii. The survival of the rats was observed for more than 5 days. The findings indicated that rats infected with either the WT or ΔlprI complemented strains died within 24 h, with an 85% fatality rate observed within 72 h. Conversely, rats infected with the ΔlprI strain began to succumb after 48 h, with 80% of them surviving for at least 5 days following infection. The survival curves depicted in Fig. 3A unequivocally illustrated the notable distinction between the ΔlprI and WT strains. Furthermore, the study examined the influence of LprI on the colonization of C. sakazakii in mice. It was observed that rats infected with the ΔlprI strain exhibited markedly reduced bacterial counts in their blood, liver, and spleen compared with those infected with the WT strain, as depicted in Fig. 3B, C and D. These findings underscore the pivotal involvement of lprI in the formation of C. sakazakii biofilms and its virulence.
Fig 3.
Impact of lprI knockout on bacterial phenotypes. (A) Effect of lprI knockout on biofilm formation. (B) Survival of rat pups during the 120-h period following oral infection with WT, ΔlprI, and ΔlprI complemented strains. Combined data from three independent experiments are shown (log-rank test; *, P < 0.05). (C to E) CFUs present in the blood, liver, and spleen of infected rats at 24-h post-infection. Blood was collected from the facial vein, and the liver and spleen were harvested and homogenized in ice-cold PBS for colony enumeration.
LprI binds to DNA during biofilm formation
Prior research has confirmed the existence of extracellular DNA (eDNA) within bacterial biofilm matrices, as well as the presence of DNA-binding proteins that aid in biofilm formation. Due to its classification as an outer membrane lipoprotein, the potential eDNA-binding ability of LprI was examined. Cell-free protein expression of LprI was carried out, followed by electrophoretic mobility shift assay (EMSA) to evaluate its interaction with DNA. Upon co-incubation of purified LprI protein with E. coli plasmid DNA pUC57 or PCR products of the dnaK gene from C. sakazakii DNA, alterations in DNA migration were observed, showing a decrease in migration rates with increasing LprI concentration (Fig. 4A and B). Conversely, no significant changes in DNA fragment migration were observed when LprI was heat-inactivated or when the highest concentration of bovine serum albumin (BSA) was utilized as a control. To assess the role of LprI in mediating bacterial adhesion to extracellular DNA (eDNA), DNA-binding assays were performed utilizing intact C. sakazakii cells. The PCR products utilized in the EMSA were immobilized onto microarray plates, enabling a comparison of the binding capacities of wild-type and ΔlprI mutant strains. The findings revealed that the wild-type bacteria exhibited a greater propensity to adhere to DNA within the wells, whereas the ΔlprI mutant bacteria displayed significantly diminished attachment (Fig. 4C). The addition of DNAse I to wells resulted in a significant decrease in the attachment of wild-type bacteria, while having a minimal effect on ΔlprI mutant attachment. Consequently, these experiments, along with findings from EMSA analysis, suggest that LprI may have the ability to bind to eDNA in a non-specific manner, potentially facilitating the anchoring of C. sakazakii onto eDNA within the biofilm.
Fig 4.
Interaction of DNA and LprI on bacterial biofilm formation. (A-B) Impact of DNA and LprI interaction on DNA electrophoretic mobility. DNA samples include linearized pUC57 plasmid DNA (A) and PCR products of the dnaK gene (B). Electrophoretic migration rates were compared between DNA alone, DNA mixed with 10 ng/mL LprI protein [LprI (L)], DNA mixed with 100 ng/mL LprI protein [LprI (H)], DNA mixed with heat-inactivated high-concentration LprI protein, and DNA mixed with BSA. (C) Relative retention of C. sakazakii cells to extracellular DNA (eDNA). The attachment of wild-type (WT) C. sakazakii cells was compared to that of the ΔlprI mutant with respect to eDNA. Additionally, the effect of DNase I on the binding of C. sakazakii cells to eDNA was investigated.
DISCUSSION
Microorganisms residing in biofilms are surrounded by a matrix primarily composed of extracellular polymeric substances (EPSs), which include extracellular polysaccharides, proteins, eDNA, and their substrates (32, 33). This matrix facilitates surface adhesion and cell aggregation, providing mechanical cohesion and stability between bacteria (34). The resulting stability contributes to the high survival and persistence potential of microorganisms within biofilms. Initial research on LprI revealed its function as a lysozyme inhibitor in Mycobacterium tuberculosis. Our study demonstrates the role of LprI in the regulation of biofilm formation and its interaction with eDNA in C. sakazakii. This interaction is believed to strengthen the mechanical cohesion among bacteria, ultimately facilitating biofilm formation. These results indicate that targeting the lipoprotein LprI could be a promising approach for developing anti-biofilm strategies.
Lipoproteins located in the outer membrane have been identified as key players in the formation of biofilms and other phenotypes in a variety of bacterial species (35, 36). For instance, the outer membrane lipoprotein VacJ is indispensable for maintaining membrane integrity, resisting serum, and facilitating biofilm formation in Actinobacillus pleuropneumoniae (37, 38). Likewise, the type II secretion system and its widely distributed lipoprotein substrate, SslE, are essential for both biofilm formation and virulence in enteropathogenic E. coli (39, 40). Furthermore, the OmpA family lipoprotein present in P. aeruginosa plays a crucial role in the development of outer membrane vesicles, biofilms, and periplasmic structure, emphasizing the significance of outer membrane lipoproteins in the process of biofilm formation (35, 41). Additionally, the eDNA serves as a fundamental element in the matrix of most biofilms (33). Research has demonstrated that the enzymatic breakdown of eDNA can impede biofilm formation, disperse established biofilms, or increase the susceptibility of biofilms to antibiotics, underscoring the pivotal role of eDNA in the dynamics of biofilm formation (42). In alignment with our study, the identification of a lipoprotein (CD1687) on the surface of Clostridioides difficile, which, when overexpressed, facilitated biofilm formation potentially through its capacity for nonspecific binding to extracellular DNA, indicates a potential conserved mechanism involving lipoproteins located on the bacterial outer membrane and their interaction with extracellular DNA in the formation of bacterial biofilms (43, 44).
Our study demonstrates that LprI acts both as a biofilm-promoting factor and a virulence factor, suggesting a possible link between bacterial infection and biofilm formation. The identification of genes that coordinate both biofilm formation and virulence is well-documented in the literature. For example, luxS, a quorum-sensing system in Staphylococci, plays a crucial role in regulating both biofilm development and virulence (28). Similarly, two quorum-sensing systems are responsible for controlling biofilm formation and virulence in Burkholderia cepacia (45). The ClpP protease, HD-GYP domain proteins, and SarZ have been identified as key regulators of biofilm development and virulence in Staphylococcus epidermidis and P. aeruginosa (46, 47). These findings indicate that biofilm formation serves as both an adhesive foundation and a defensive barrier, protecting bacteria from detachment and host immune defenses, ultimately enhancing bacterial persistence within the host. In addition, in M. tuberculosis, the lprI gene co-transcribes with the glbN gene, and both are simultaneously upregulated during macrophage infection (48). The expression of the lprI gene protects bacterial growth from lysozyme inhibition in vitro and enhances phagocytosis and survival during intracellular infection of peritoneal and monocyte-derived macrophages (48). The co-occurrence of lprI in both C. sakazakii and M. tuberculosis suggests that it is a functionally conserved gene. However, the activity of the lysozyme inhibitor encoded by lprI in C. sakazakii requires further investigation.
Certain compounds or materials have demonstrated the ability to hinder bacterial biofilm formation and virulence. For instance, ZnO nanoparticles have been found to impede both biofilm formation and virulence factor production in P. aeruginosa (49, 50). Quercetin has also shown efficacy in inhibiting biofilm formation and virulence factors in the same bacterium (51, 52). Additionally, streptomycin has been observed to facilitate biofilm inhibition and suppress virulence traits in P. aeruginosa PAO1 (25). Andrographolide exhibits inhibitory effects on biofilm formation and virulence in Listeria monocytogenes (53), whereas an oxazole derivative exhibits inhibitory effects on biofilm formation, extracellular polysaccharide production, and virulence in Streptococcus mutans (54). These findings offer valuable insights into potential strategies for the control of C. sakazakii. Several studies have identified compounds that demonstrate inhibitory effects on the virulence and biofilm formation of C. sakazakii, such as Thymoquinone and trans-cinnamaldehyde (55, 56). Thymoquinone has been shown to inhibit virulence-related traits and possess anti-biofilm formation potential in C. sakazakii ATCC 29544, whereas trans-cinnamaldehyde has been found to inhibit and deactivate C. sakazakii biofilm on non-living surfaces (12, 56). Nevertheless, further investigation is required to ascertain the impact of these chemicals on the interaction between LprI and eDNA in C. sakazakii.
In this research, knockouts were performed on nine gene cassettes, including envZ/ompR, glrK, flgK, treR, kdpE, recA, rpfB, hmsP, and c-di-GMP, as well as the endogenous plasmid pESA3 in C. sakazakii. The findings of this study suggest that the deletion of the c-di-GMP gene promotes biofilm formation in C. sakazakii, whereas the deletion of hmsP inhibits biofilm formation, underscoring the crucial roles of c-di-GMP and hmsP in the regulation of biofilm formation. The role of hmsP in influencing biofilm formation has been thoroughly investigated in Yersinia pestis, where it modulates Hms-dependent biofilm formation (57). The phosphodiesterase activity of the HmsP EAL domain has been found to have a negative impact on biofilm formation (58). Similarly, in Pseudomonas aeruginosa, specific residues within the HmsP domain of SagS have been shown to independently regulate both biofilm formation and biofilm drug tolerance (59). These findings suggest that hmsP is a conserved gene that plays a key role in the regulation of bacterial biofilms (60). Furthermore, c-di-GMP has been identified as a key regulator of biofilm formation in various bacterial species, including Vibrio cholerae (61, 62). The effects of c-di-GMP on biofilm formation can exhibit significant variability, as evidenced by the synergistic support of biofilm maintenance through direct interaction of their effectors in Shewanella putrefaciens (63). Surprisingly, our discovery that the knockout of c-di-GMP enhances biofilm formation in C. sakazakii suggests a compensatory response triggered by the absence of c-di-GMP. The upregulation of LprI following c-di-GMP knockout may serve as an indicator of this compensatory mechanism.
MATERIALS AND METHODS
Strains and plasmids
The study utilized bacterial strains and plasmids (Table 1) along with corresponding primers (Table 2). Bacterial samples were stored at −80°C in cryovials containing LB medium with 15% glycerol. Before the experiments, overnight cultures of bacterial strains in LB medium were prepared. Target gene deletion mutants were generated using the pCVD442 suicide plasmid (Miaolingbio, China). The plasmid was linearized via PCR with primers pCVD442-fwd and pCVD442-rev. Upstream and downstream fragments (800–900 bp) of the gene to be deleted were amplified using primers listed in Table 2. A 5-µL mixture containing 100-µg linearized pCVD442, 10-µg upstream, and 10-µg downstream fragments was combined with 5 µL of 2× recombinase (Vazyme, China) and incubated at 45°C for 15 min. The reaction was transformed into 100 µL of thawed E. coli S17 lambda pir cells (Weidi, China). After ice incubation, heat shock, and shaking at 37°C, the cells were centrifuged, and the pellet was spread onto an ampicillin-resistant plate. In situ deletion in C. sakazakii was achieved via conjugation with E. coli S17 lambda pir, followed by antibiotic and sucrose selection. Gene deletions were confirmed by PCR and sequencing. E. coli mixed with C. sakazakii (kanamycin-resistant) was spread onto double antibiotic plates, and colonies were verified. Correct clones underwent sucrose selection, and single colonies were screened for loss of resistance. PCR and sequencing confirmed gene deletion by checking the absence of the target gene.
TABLE 1.
Bacterial strains and plasmids used in this study
| Strain or plasmid | Description | Source |
|---|---|---|
| C. sakazakii strains | ||
| WT | Wild-type C. sakazakii BAA-894 | ATCC |
| ΔenvZ/ompR | Markerless deletion mutant ΔenvZ/ompR | This study |
| ΔrecA | Markerless deletion mutant ΔrecA | This study |
| ΔkdpE | Markerless deletion mutant ΔkdpE | This study |
| ΔglrK | Markerless deletion mutant ΔglrK | This study |
| ΔrpfB | Markerless deletion mutant ΔrpfB | This study |
| ΔpESA3 | Markerless deletion mutant ΔpESA3 | (64) |
| ΔhmsP | Markerless deletion mutant ΔhmsP | This study |
| ΔflgK | Markerless deletion mutant ΔflgK | This study |
| ΔtreR | Markerless deletion mutant ΔtreR | This study |
| ΔESA_01588 | Markerless deletion mutant ΔESA_01588 | This study |
| ΔlprI | Markerless deletion mutant ΔlprI | This study |
| ΔenvZ/ompR-C | envZ/ompR complementation in ΔenvZ/ompR | This study |
| ΔrecA-C | recA complementation in ΔrecA | This study |
| ΔkdpE-C | kdpE complementation in ΔkdpE | This study |
| ΔglrK-C | glrK complementation in ΔglrK | This study |
| ΔrpfB-C | rpfB complementation in ΔrpfB | This study |
| ΔpESA3-C | pESA3 complementation in ΔpESA3 | This study |
| ΔhmsP-C | hmsP complementation in ΔhmsP | This study |
| ΔflgK-C | flgK complementation in ΔflgK | This study |
| ΔtreR-C | treR complementation in ΔtreR | This study |
| ΔESA_01588-C | ESA_01588 complementation in ΔESA_01588 | This study |
| ΔlprI-C | lprI complementation in ΔlprI | This study |
| Escherichia coli strains | ||
| S17 λ pir | Strain for construction harboring λ pir | (65) |
| S17 λ pir-ΔenvZ/ompR | S17 λ pir harboring pCVD442-ΔenvZ/ompR | This study |
| S17 λ pir-ΔrecA | S17 λ pir harboring pCVD442-ΔrecA | This study |
| S17 λ pir-ΔkdpE | S17 λ pir harboring pCVD442-ΔkdpE | This study |
| S17 λ pir-ΔglrK | S17 λ pir harboring pCVD442-ΔglrK | This study |
| S17 λ pir-ΔrpfB | S17 λ pir harboring pCVD442-ΔrpfB | This study |
| S17 λ pir-ΔpESA3 | S17 λ pir harboring pCVD442-ΔpESA3 | This study |
| S17 λ pir-ΔhmsP | S17 λ pir harboring pCVD442-ΔhmsP | This study |
| S17 λ pir-ΔflgK | S17 λ pir harboring pCVD442-ΔflgK | This study |
| S17 λ pir-ΔtreR | S17 λ pir harboring pCVD442-ΔtreR | This study |
| S17 λ pir-ΔESA_01588 | S17 λ pir harboring pCVD442-ΔESA_01588 | This study |
| S17 λ pir-ΔlprI | S17 λ pir harboring pCVD442-ΔlprI | This study |
| Plasmids | ||
| pACYC184 | Low-copy plasmid | (65, 66) |
| pACYC184-envZ/ompR | envZ/ompR complementation vector | This study |
| pACYC184-recA | recA complementation vector | This study |
| pACYC184-kdpE | kdpE complementation vector | This study |
| pACYC184-glrK | glrK complementation vector | This study |
| pACYC184- rpfB | rpfB complementation vector | This study |
| pCVD442-hybrid-pESA3 | pESA3 complementation vector | This study |
| pACYC184- hmsP | hmsP complementation vector | This study |
| pACYC184- flgK | flgK complementation vector | This study |
| pACYC184- treR | treR complementation vector | This study |
| pACYC184- ESA_01588 | ESA_01588 complementation vector | This study |
| pACYC184- lprI | lprI complementation vector | This study |
TABLE 2.
Primers used in this study
| Primer | Sequence (5′−3′) |
|---|---|
| For construction | |
| pCVD442-fwd | GGCTGTCAGACCAAGTTTACTCATATATACTTTAGATTG |
| pCVD442-rev | GCAGATACTCTTCCTTTTTCAATATTATTGAAGCATTTATCAGGGTTATTG |
| ΔpESA3-F | AAAAAGGAAGAGTATCTGCGGTACGGTACGGCCATACTG |
| ΔpESA3-R | CGATTAACCCATCTAAACGTCTCCACTAAAAAATCG |
| ΔrecA-A | GAAAAAGGAAGAGTATCTGCGGATAACCATAGTACGCACTATG |
| ΔrecA-B | CTGCATCAGCAGCCCTTGAGATTACATTTTTACTCCTGTCATGCAGG |
| ΔrecA-C | TAATCTCAAGGGCTGCTGATGCAG |
| ΔrecA-D | GATTAATTGTCAAGGCTAGCGATGATTATTCGGGACCAGAGAGCTACTG |
| ΔenvZ/ompR-A | GAAAAAGGAAGAGTATCTGCGGGTGAAGCGCCTGGCGCGCGAAAACG |
| ΔenvZ/ompR-B | ATTGGCTAAGGAGCAGTGAAATGCATGCTTTAGCGCCGTCCGGCAC |
| ΔenvZ/ompR-C | CATTTCACTGCTCCTTAGCCAAT |
| ΔenvZ/ompR-D | GATTAATTGTCAAGGCTAGCGCGAGAAAACAGCAGCGCGATAA |
| ΔkdpE-A | GAAAAAGGAAGAGTATCTGCGGGACACCCTGCCCGGCGTGCCGTATC |
| ΔkdpE-B | CAGGCGTGACGCGGCGTCAGCGTTACATGACCTCGTGGATATCCAGT |
| ΔkdpE-C | CGCTGACGCCGCGTCACGCCTG |
| ΔkdpE-D | GATTAATTGTCAAGGCTAGCGATGATTCGAAGACGCCCATTGCGACGTGGC |
| ΔglrK-A | GAAAAAGGAAGAGTATCTGCGGCCTGGTGGAAGTGACGGCCAGCCCGT |
| ΔglrK-B | CCATGCGCTTACCAGACGTGAATTCACATGACTAGTTGGCGAAGCGAGCGGGG |
| ΔglrK-C | TTCACGTCTGGTAAGCGCATGG |
| ΔglrK-D | GATTAATTGTCAAGGCTAGCGATGACTGGAGCTTCTGGATCTCGGCGA |
| ΔrpfB-A | GAAAAAGGAAGAGTATCTGCGGCCGAGTACCAGCGCGATGAGCA |
| ΔrpfB-B | TCAGGGCGACGTGATTGCAGCTCACATCACACCAAAAACAAATAAACAGC |
| ΔrpfB-C | GAGCTGCAATCACGTCGCCCTGA |
| ΔrpfB-D | GATTAATTGTCAAGGCTAGCGATGAGCTGATTAATTCCGGCAATTGAT |
| ΔhmsP-A | GAAAAAGGAAGAGTATCTGCGGCTGGGGCACCTGCGCCGAAACG |
| ΔhmsP-B | CTTACACCTTTTGCAGCGACGTCACATCTGTTTTATCGTGAGGGAAACGACTG |
| ΔhmsP-C | TGTGACGTCGCTGCAAAAGGTGTAAG |
| ΔhmsP-D | GATTAATTGTCAAGGCTAGCGATGACCATCGTCAGGTAATGGAAAGTGC |
| ΔflgK-A | GAAAAAGGAAGAGTATCTGCGGCCAAAGGCTTAGGCCTCGCTGAC |
| ΔflgK-B | CGTTCGGTTCCCTGTTAGCGGGACATGGAGGTTCCTTTTGAA |
| ΔflgK-C | CCCGCTAACAGGGAACCGAACG |
| ΔflgK-D | GATTAATTGTCAAGGCTAGCGATGATCTGCTGAGTACCGACTTCC |
| ΔtreR-A | GAAAAAGGAAGAGTATCTGCGGCGAAACACGCCTGCGCCGAG |
| ΔtreR-B | TTAGCCGAGACGGGCGGGGATCCATCGTCCATTCCTCTGGC |
| ΔtreR-C | CCCCGCCCGTCTCGGCTAA |
| ΔtreR-D | GATTAATTGTCAAGGCTAGCGATGAGATGGTAAAGAGCCCAAAATCC |
| ΔESA_01588-A | GAAAAAGGAAGAGTATCTGCGGCGCACCGAGCCAGGCCCGCACTTCGGC |
| ΔESA_01588-B | CGTTTACGGCGTCTGTTTCCCGTTATTCTCCAGGTCGTCTTTCATGCA |
| ΔESA_01588-C | GGGAAACAGACGCCGTAAACG |
| ΔESA_01588-D | GATTAATTGTCAAGGCTAGCGATGATTATCTGGATGATGTTGCG |
| ΔlprI-A | GAAAAAGGAAGAGTATCTGCGCTTCATGAGAGATTCCTTAATAAG |
| ΔlprI-B | GCTTTTTTTATGGCGTTACATAACAATCCTTTTTACGTA |
| ΔlprI-C | TAACGCCATAAAAAAAGC |
| ΔlprI-D | GATTAATTGTCAAGGCTAGCGATGACGTCATGACGCGCATGAAAT |
| For sequencing confirmation | |
| Check2-pESA3-F | GAGCGGCAGTGTTGCCTGGC |
| Check2-pESA3-R | TCCAGCGTTGCGCTTTTTCA |
| ΔenvZ/ompR-E | GTGAAGCGCCTGGCGCGCGAA |
| ΔenvZ/ompR-F | AGCGCGAGCAAGACCAGCGCC |
| ΔrecA-E | CCCGCCAACGGCCAGGCGCCG |
| ΔrecA-F | TGAAGTTGCCCAAGCATTTCGAAGAAGGT |
| ΔflhD-E | TGCCGTTAACGGGTTTAACGATA |
| ΔflhD-F | GCAGCGCCTTCAGCGTGCCTTTG |
| ΔkdpE-E | CGCAGCCGCTCGCGGAAATCACC |
| ΔkdpE-F | TTCTGTGCGGCGGCCTCAAGCTGG |
| ΔglrK-E | GCGAAGTCTATTCTGTTCAACG |
| ΔglrK-F | TGACAATCGTCTCGCGCCACTGTT |
| ΔrpfB-E | GGCGCAGGGCGCATGGCGCGTCA |
| ΔrpfB-F | TCACGCCGTTAATAATTTTCCAG |
| ΔhmsP-E | CTGCGCGACGGGCGGCTGTTCCT |
| ΔhmsP-F | CAGCAACAGCACTACCAGCAGCGT |
| ΔESA_01588-E | TACAACGTCCACGCGGGCTTTGACCCGGC |
| ΔESA_01588-F | CAGATAAATAATGGTTATAAAAC |
| ΔlprI-E | TAATTGTTCATCCAGCGCGCTGATTTT |
| ΔlprI-F | CATTTCGGCCAAAGCCTGATCTA |
| ΔtreR-E | TGAGTAACCTTACTCGATAGTAACATAAC |
| ΔtreR-F | CGATAAACGCGTGCGCCAGGA |
Curing of pESA3 and complementation
A DNA segment from the pESA3 plasmid was amplified using primers ΔpESA3-F and ΔpESA3-R (30). The amplified segment was then ligated to linearized pCVD442 plasmid using a recombinant kit (Vazyme, China). The resulting circularized plasmid was transformed into C. sakazakii BAA-894 using S17-1 λ pir competent cells (Weidi, China) and electroporation. Transformants were selected with antibiotics, and plasmid-curing strains were selected on 20% sucrose LB agar plates. The resulting hybrid plasmid from recombination was isolated and introduced into a plasmid-curing strain to generate a complementation strain lacking the pESA3 plasmid.
Rat virulence assay
The C. sakazakii infection in rats was conducted following established protocols (67). Animal survival over a 5-day period post-infection was monitored at specific intervals. To evaluate bacterial infection, the rats were euthanized 24 h post-infection. The organs were retrieved, homogenized in pre-cooled PBS buffer with steel beads, and then serially diluted in PBS. The diluted homogenates were plated on LB agar plates, and bacterial colonies were counted after 24 h of incubation to determine bacterial load.
Biofilm formation
Biofilms were cultivated over a 72-h incubation period to allow for maturation (68). Biomass attachment was assessed using the crystal violet assay. Glass slides were stained with 0.1% crystal violet solution for 10 min, rinsed twice with double-distilled water, and then solubilized with 50% (v/v) ethanol/water solution for 10 min. The optical density of the solubilized crystal violet solution was measured at 570 nm to quantify biomass attached to the glass substrates at regular intervals.
Protein synthesis
E. coli lysate served as the enzyme system for cell-free protein synthesis (69). The full-length gene, including the promoter, was amplified by PCR, with point mutations incorporated into the amplification primers when necessary. Fragment fusion PCR was utilized to merge multiple segments into a complete segment. The resulting PCR products were purified, sequenced for confirmation, and added to the cell-free protein synthesis system according to manufacturer’s instructions. After the reaction, proteins were purified using ultrafiltration tubes and quantified using a BCA assay.
Electromobility shift assays
Electromobility shift assays (EMSAs) are conducted according to the method described previously (43). Freshly purified LprI, synthesized via a cell-free protein synthesis kit, was exclusively used in these assays. LprI concentrations ranged from 10 to 100 ng/mL, incubated with DNA (pUC57 or PCR product) in 10 µl of sodium phosphate buffer (50 mM, pH 8.0) at room temperature for 30 min. The samples were loaded onto TAE-buffered agarose gels (1% w/v) and electrophoresed for 90 min at 100 V. Denatured LprI controls were treated at 100°C for 15 min before the assay. Gels were stained with ethidium bromide and imaged using a ChemiDoc gel imaging system (Bio-Rad). The pUC57 plasmid was extracted from E. coli using a plasmid extraction kit (Axygen, USA), and the PCR amplicon was generated using C. sakazakii DNA template and primers targeting the dnaK region.
Protein extraction and digestion
Protein extraction and digestion are conducted according to the method described previously (67). The bacterial suspension was subjected to centrifugation and subsequently washed with phosphate-buffered saline (PBS) to eliminate the culture medium. The bacterial cells were then lysed using ultrasonic waves until the solution became clear. The resulting supernatant was purified by filtration through a 0.22-µm membrane following centrifugation at 21,000g for 15 min at 4°C. For proteomic analysis, the protein concentration was quantified using a bicinchoninic acid (BCA) assay. Fifty micrograms of protein was denatured in 100 µL of 8 M urea (Aladdin, China). Disulfide bonds were reduced by adding 1 µL of 200 mM dithiothreitol (DTT, Aladdin, China) and incubating for 30 min at 60°C. Following cooling, 1 µL of 500 mM iodoacetamide (Aladdin, China) was introduced, and the mixture was agitated for 30 minutes in the dark to facilitate the alkylation of cysteine residues. The sample subsequently underwent overnight digestion at 37°C with 2 µg of recombinant trypsin (Thermo, USA). The digestion process was terminated by the addition of 10% trifluoroacetic acid (Aladdin, China) to achieve a final concentration of 0.4%. The samples were then desalted using a C18 solid-phase extraction (SPE) column (Millipore, USA) and vacuum-dried before mass spectrometry analysis.
Bacteria-DNA binding assay
A 500 bp segment of the dnaK gene was amplified using a primer modified with C6 amine. The DNA was immobilized on the bottom of a 96-well plate using a 50 mM carbonate buffer at pH 9.6. After incubating at room temperature for 3 h, the plate was washed with PBST to remove unbound DNA. Subsequently, 200 µL of bacterial suspension was added to the DNA-coated wells of the 96-well plate. The plate was incubated at 37°C for 20 min, followed by three washes with PBS to remove unbound bacteria. Each well was then treated with 20 µg of DNase I and incubated at 37°C for 30 min. Bacteria from the wells were serially diluted and plated to quantify viable cells.
LC-MS/MS and statistical analysis
Peptide samples were analyzed with a Q Exactive Plus mass spectrometer (70). Data processing used Maxquant software against the UniProt C. sakazakii database (71). The specific experimental parameters for the machine setup were followed according to the previously described method (30, 31, 72). Peptide compounds were dissolved in buffer A (0.1% formic acid) and loaded onto a reversed-phase trapping column (Thermo Fisher Scientific, PepMap100, 100 µm × 2 cm, nano Viper C18). This trapping column was connected to a reversed-phase analytical column (Thermo Fisher Scientific, 75 µm × 10 cm, 3 µm resin). Maxquant parameters included fixed carbamidomethylation, variable methionine oxidation, and trypsin specificity allowing up to two missed cleavages. Identifications were filtered to a 1.0% false discovery rate, and protein quantification was based on the median of unique peptides per protein.
All experiments were conducted in triplicate for robustness. Statistical analysis was conducted using GraphPad Prism software (version 8.3). Significance of differences was assessed using one-way ANOVA, with statistical significance defined as P < 0.05.
ACKNOWLEDGMENTS
This work was financially supported by the National Key Research and Development Program of China (grant number 2022YFF1100704) and the Fundamental Research Funds for the Central Universities of Nankai University (grant numbers 63241341, 63241583, and 63241584).
X.J.: Conceived and designed research. C.D., J.W.，B.Z., Y.H., H.L., J.W., Y.S., J.L.: Writing – review and editing, Data curation. Y.Z.: Writing – review and editing, Supervision. S.W.: Writing – review and editing, Supervision, Conceptualization.
Contributor Information
Shuo Wang, Email: wangshuo@nankai.edu.cn.
Edward G. Dudley, The Pennsylvania State University, University Park, Pennsylvania, USA
ETHICS APPROVAL
The study was approved by the Ethics Committee of our department, and written informed consent was obtained from all participants before the study.
DATA AVAILABILITY
The MS proteomics data have been deposited in iProX and can be accessed using the accession number IPX0005984000.
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Associated Data
This section collects any data citations, data availability statements, or supplementary materials included in this article.
Data Availability Statement
The MS proteomics data have been deposited in iProX and can be accessed using the accession number IPX0005984000.