CRISPR/Cas9-Induced Knockout of Sting Increases Susceptibility of Zebrafish to Bacterial Infection

Stimulator of interferon genes (STING) is an adapter protein that is activated when cyclic dinucleotides (CDNs) are present. CDNs originate from the cytosolic DNA of both pathogens and hosts. STING activation promotes efficient immune responses against viral infections; however, its impact in bacterial infections is unclear. In this study, we investigated the role of Sting in bacterial infections by successfully creating a sting-deficient (sting(−/−) with a 4-bp deletion) knockout zebrafish model using CRISPR/Cas9. The transcriptional modulation of genes downstream of cGAS (cyclic GMP-AMP synthase)-Sting pathway-related genes was analyzed in seven-day-old wild-type (WT) and sting(−/−) embryos, as well as in four-day-old LPS-stimulated embryos. The expression of downstream genes was higher in sting(−/−) than in healthy WT fish. The late response was observed in sting(−/−) larvae following LPS treatment, demonstrating the importance of Sting-induced immunity during bacterial infection by activating the cGAS–STING pathway. Furthermore, adult sting(−/−) fish had a high mortality rate and significantly downregulated cGAS–STING pathway-related genes during Edwardsiella piscicida (E. piscicida) infection. In addition, we assessed NF-κB pathway genes following E. piscicida infection. Our results show fluctuating patterns of interleukin-6 (il6) and tumor necrosis factor-α (tnfα) expression, which is likely due to the influence of other NF-κB pathway-related immune genes. In summary, this study demonstrates the important role of Sting against bacterial infection.


Introduction
The innate immune system recognizes invading pathogens through pattern recognition receptors (PRRs) [1]. PRRs include retinoic-acid-inducible gene 1 (RIG-1) receptors (RLRs), nucleotide-binding oligomerization domain-like receptors (NLRs), toll-like receptors (TLRs), AIM2-like receptors (ALRs), and C-type lectin receptors (CLRs). Cell surface PRRs (TLRs and CLRs) are responsible for sensing pathogens from the extracellular environment, cytoplasmic PRRs (NLRs, RLRs, and ALRs) are responsible for detecting intracellular pathogens, and endosomal PRRs (TLRs) are used to detect microbes that have entered phagolysosomes [2]. PRRs regulate the expression of inflammatory mediators through intracellular signaling cascades to eliminate pathogens from infected cells [3]. The RLR family plays an important role in recognizing non-self-signatures (of viral RNA) in the cytoplasm, and their signaling controls viral infection. RLRs modulate the host intracellular immune response by regulating the expression of interferons (IFNs) and antiviral genes and activating downstream transcription factors [4,5]. from our study may encourage future studies on the antibacterial activity of zebrafish Sting, and the sting (−/−) zebrafish model can be an important tool for further studies on the role of Sting in the zebrafish immunity.

Animals
Wild-type (WT; AB strain) zebrafish were maintained as previously described [25]. Laboratory water circulation and filtration systems for the zebrafish cultures were maintained at a constant temperature of 28 ± 0.5 ℃ in a 14:10 h light-dark cycle. All animal experiments were approved by the Animal Care and Use Committee of Jeju National University.

CRISPR/Cas9-Mediated Generation of Sting (−/−) Zebrafish
The sting (−/−) zebrafish was generated using CRISPR/Cas9 gene-editing technology [15]. The CRISPR/Cas9 sting-targeting site (5′ AGAGCGCGCAGCAGGCTGCC 3′) were designed using an online tool [http://zifit.partners.org/ZiFiT/ (accessed on 3 September 2019)]. Single-guide RNA (sgRNA) synthesis, microinjection, and mutant confirmation were performed as previously described [25]. The sgRNA target site was chosen to be 20bp before the protospacer adjacent motif (PAM) sequence in exon 3 ( Figure 1A). To confirm target site mutations in the genomic DNA, polymerase chain reaction (PCR) was carried out using T7E1 primers designed from intron 2 to exon 3 of the sting sequence after thatT7E1 assaywas performed. (Table 1).   To evaluate the tissue-specific sting transcriptional patterns, healthy male and female zebrafish (5 per sex) were dissected after anesthesia, and tissues (including the brain, gill, heart, intestine, kidney, liver, muscle, ovary, spleen, and testis) were collected. Internal organs (including the intestine, kidney, liver, and spleen) and muscle were collected from fish challenged with bacteria. Immediately after isolation, all tissues were snap-frozen in liquid nitrogen and stored at −80 • C until RNA extraction.

Embryo Sample Collection
To analyze sting expression during early development, zebrafish embryos at each developmental stage were collected based on morphological criteria [26]. A total of 50 embryos each of WT and sting (−/−) zebrafish were collected seven days post-fertilization (dpf) for downstream transcriptional studies and mutation analysis. Embryos were washed with RNA-grade 1 × phosphate-buffered saline (PBS) and stored at −80 • C for RNA extraction.

LPS Stimulation of Sting (−/−) and WT Zebrafish Larvae
WT and sting (−/−) zebrafish larvae (3 dpf) were divided into six groups, each containing 20 larvae. Three groups were treated with 100 µg/mL LPS (Escherichia coli 0111:B4) and three were maintained as a PBS-treated control. Embryos were collected 6, 12, and 24 h after LPS treatment. RNA extraction and cDNA synthesis were performed as described in Section 2.6. Using quantitative real-time PCR (qPCR), the expression of downstream genes, such as interferon phi 1 (ifnphi1), caspase b, tnfα, and interleukin 6 (il6), in WT and sting (−/−) zebrafish was examined at various time points (qPCR). The same experiment was repeated, and larval survival was monitored for three days following LPS treatment.

Bacterial Challenge against Sting (−/−) and WT Zebrafish
Two-month-old juvenile WT and sting (−/−) zebrafish were challenged with E. piscicida by the immersion method as previously described, with some modifications [27]. To examine the percentage mortality of sting (−/−) and WT zebrafish following E. piscicida infection, the zebrafish were divided into three groups of 20 individuals each. Two groups were wounded by removing ten scales before bacterial or PBS exposure, and the third group was used as a non-injured control. The first group was exposed to E. piscicida at a final concentration of 10 8 CFU/mL, and the control groups were treated with the same volume of PBS in a total volume of 300 mL. After a 5 h immersion bath, the zebrafish were transferred to a new tank with 2 L of water, and half of the water was replaced with new water every day. Zebrafish were maintained within an incubator at 28°C for 12 days to observe mortality. Feeding was paused one day before infection, and resumed two days post-infection (dpi).
For the challenge experiment, WT and sting (−/−) zebrafish were divided into three groups, each containing 24 zebrafish, and a wound was generated in two groups before bacterial treatment. These two groups (one from both WT and sting (−/−) ) were exposed to E. piscicida at the final concentration of 10 8 CFU/mL in a total volume of 300 mL for 5 h, and the other was maintained as a PBS-treated control without injury. The bath immersion and maintenance of zebrafish were performed as described in the mortality experiment. In addition, the zebrafish were not fed after bath immersion. Tissue samples, including the internal organs and muscles, at the infected site of six zebrafish from each group were collected at 6, 24, 48, and 72 h post-infection (hpi), as shown in Supplementary Figure S2A. Immediately after collection, tissue samples were snap-frozen in liquid nitrogen. RNA extraction and cDNA synthesis were performed as described in Section 2.6. The expression of downstream genes, including tbk1, irf3, irf7, nf-κb, tnfα, ifnphi1, il6, and an internal control gene elongation factor-1α (ef1α) in WT and sting (−/−) zebrafish were analyzed by qPCR at different time points post-E. piscicida challenge. Relative expression of the target genes was calculated with respect to ef1α expression using the Livak method [28,29]. The expression of target genes in E. piscicida-infected samples were normalized with respect to the PBS-treated controls.
To confirm that E. piscicida infection occurred, six randomly selected zebrafish from the E. piscicida-infected WT and sting (−/−) groups were sampled at 6 hpi. The genomic DNA was extracted, and the presence of E. piscicida was confirmed by PCR using E. piscicida and gapdh detection primers following agarose gel electrophoresis.

RNA Extraction and Complementary DNA Synthesis
RNA was extracted from zebrafish tissues and embryos using TRIzol reagent (Sigma-Aldrich, St. Louis, MO, USA). RNA concentrations were assessed using a Multiskan™ GO Microplate Spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA) at 260 nm. The quality of the RNA samples was confirmed by agarose gel electrophoresis. First-strand cDNA was synthesized from 3 µg RNA using a Prime Script™ 1st strand cDNA synthesis kit (Takara, Shimogyo-ku, Kyoto, Japan). The cDNA samples were diluted 30-fold and stored at −20 • C for PCR and qPCR analysis.

Transcriptional Analysis Using qPCR
The qPCR reaction mixture (10 µL) was prepared using 5 µL 2× TaKaRa Ex Tag SYBR premix, 0.5 µL each of the forward and reverse primers (Table 1), and 4 µL cDNA. qPCR reactions were performed using a Thermal Cycler Dice Real-Time System III (TaKaRa, Japan) under the following thermal conditions: initial denaturation at 95 • C for 10 s; 45 PCR cycles, each consisting of denaturation at 95 • C for 5 s, annealing at 58 • C for 10 s, and extension at 72 • C for 20 s, and finally, one melting cycle at 95 • C for 15 s, 60 • C for 30 s, and 95 • C for 15 s. Relative mRNA expression was calculated using the 2 −∆∆Ct method.

Statistical Analyses
All experiments were carried out in triplicate, and data expressed as the means ± standard deviation (SD). Student's t-tests were used for statistical analysis, and p < 0.01 (indicated by *) and <0.001 (indicated by **) were considered statistically significant.

Expression Analysis of Sting
Sting and β-actin transcription during the embryonic developmental stages from the 2-cell stage to 7 dpf was analyzed using PCR, and the gel electrophoresis images are shown in Supplementary Figure S1B. Moderate sting expression is observed starting from the shield stage, and the sting expression pattern in our study is similar to that observed in previous studies [17].
Sting expression is observed in all analyzed tissues of the adult zebrafish, and the results are plotted as fold-values of the expression in muscle (Supplementary Figure S1A). High expression is observed in the kidney, gill, and testis, followed by in the spleen and brain. A previous study found that sting expression was highest in the kidney, followed by the gill, heart, and spleen [17]. We also examined the transcription of stings in the ovary and testis. Our results, in accordance with previous reports, show high expression of sting in important immune organs in fish, such as the kidney, gill, and spleen, which may be due to the role of Sting in autoimmunity [8]. However, they are slightly different regarding sting expression in the heart, muscle, and intestine. Adaptations occur in organisms depending on environmental and nutritional changes, and this may explain the different gene expression patterns observed in different studies [30][31][32]. Furthermore, moderate sting expression is observed in the brain, possibly due to Sting's role(s) in controlling neuronal gene expression [33]. The transcription of sting in mandarin fish was highest in the gill, followed by the kidney, spleen, and blood [34].

Generation of Sting (−/−) Zebrafish by CRISPR/Cas9 Gene Editing
The target site mutation with a 4-bp deletion was selected using nucleotide sequence analysis, as it produced a prematurely truncated protein ( Figure 1B,C). The selected mutation was verified by PCR in WT and sting (−/−) larvae at 7 dpf using the original target site sequence as a primer ( Figure 1D). The PCR-amplified band was observed in WT fish only, confirming the absence of sting in the sting (−/−) group. The sting (−/−) zebrafish were normal in terms of survival, morphology, and fertility. Sting knockout mice have also been found to be viable and fertile [35].

Downstream Gene Expression Analysis in Sting (−/−) Zebrafish
Sting-deficiency-mediated variations in the expression of downstream genes were analyzed using cDNA from 7 dpf WT and sting (−/−) zebrafish. The expression of downstream genes was normalized to that of β-actin (Figure 2), and we found that tbk1, irf3, irf7, and nf-κb were significantly upregulated in sting (−/−) zebrafish when compared to WT individuals. In addition to its role in innate immunity, STING regulates genotoxic stress homeostasis following pathway activation [36,37]. Knockdown of cGAS, STING, TBK1, and IRF3 in HeLa cells results in increased levels of micronuclei formation and chromosomal mis-segregation [38]. RLR, cGAS-STING, and non-RLR DExD/H-box RNA helicase pathways detect cytoplasmic nucleic acids and activate type 1 IFN and pro-inflammatory cytokines [5]. Deficiency in Sting can lead to chromosomal instability through micronuclei formation, which may explain the increased levels of tbk1, irf3, irf7, and nf-κb expression in sting (−/−) zebrafish. These observed transcription patterns support the involvement of Sting in regulating downstream gene transcription in zebrafish [39,40].
irf3, irf7, and nf-κb were significantly upregulated in sting (−/−) zebrafish when compa WT individuals. In addition to its role in innate immunity, STING regulates geno stress homeostasis following pathway activation [36,37]. Knockdown of cGAS, ST TBK1, and IRF3 in HeLa cells results in increased levels of micronuclei formatio chromosomal mis-segregation [38]. RLR, cGAS-STING, and non-RLR DExD/H-box helicase pathways detect cytoplasmic nucleic acids and activate type 1 IFN and inflammatory cytokines [5]. Deficiency in Sting can lead to chromosomal insta through micronuclei formation, which may explain the increased levels of tbk1, irf3 and nf-κb expression in sting (−/−) zebrafish. These observed transcription patterns su the involvement of Sting in regulating downstream gene transcription in zebrafish [3 Figure 2. Comparison of transcription of downstream genes in 7 dpf larvae of sting (−/−) an zebrafish. The internal control gene, ꞵ-actin, was used to analyze the relative mRNA express genes downstream of sting using the Livak method. Experiments were performed in triplicat the error bar represents the standard deviation (SD). Student's t-tests were used for sta analysis, and p < 0.01 (indicated by *) and < 0.001 (indicated by **) were considered statis significant.

LPS-Induced Expression Modulation in Sting (−/−) and WT Larvae
LPS is a major component of the outer membranes of Gram-negative bacteri initiates the innate immune response of host organisms (and activates downs pathways) following its recognition. Normally, LPS recognition and signal initiatio carried out by the TLR4 receptor [41]. Interestingly, zebrafish TLR4 has been identif a negative regulator of TLR signaling and causes sequestration of TLR adaptors to i the activation of NF-κB by MyD88 [42]. Recent studies observed LPS-induced activ of the cGAS-STING pathway and promotion of endometritis [43]. To investigate th of zebrafish Sting in LPS recognition and innate immune response activ transcriptional modulation of downstream genes (including ifnphi1, caspb, tnfα, an was compared between WT and sting (−/−) zebrafish larvae at 6, 12, and 24 h pos treatment (Figure 3). At 6 h post-LPS treatment, ifnphi1 expression is signific Figure 2. Comparison of transcription of downstream genes in 7 dpf larvae of sting (−/−) and WT zebrafish. The internal control gene, β-actin, was used to analyze the relative mRNA expression of genes downstream of sting using the Livak method. Experiments were performed in triplicate, and the error bar represents the standard deviation (SD). Student's t-tests were used for statistical analysis, and p < 0.01 (indicated by *) and <0.001 (indicated by **) were considered statistically significant.

LPS-Induced Expression Modulation in Sting (−/−) and WT Larvae
LPS is a major component of the outer membranes of Gram-negative bacteria and initiates the innate immune response of host organisms (and activates downstream pathways) following its recognition. Normally, LPS recognition and signal initiation are carried out by the TLR4 receptor [41]. Interestingly, zebrafish TLR4 has been identified as a negative regulator of TLR signaling and causes sequestration of TLR adaptors to inhibit the activation of NF-κB by MyD88 [42]. Recent studies observed LPS-induced activation of the cGAS-STING pathway and promotion of endometritis [43]. To investigate the role of zebrafish Sting in LPS recognition and innate immune response activation, transcriptional modulation of downstream genes (including ifnphi1, caspb, tnfα, and il6) was compared between WT and sting (−/−) zebrafish larvae at 6, 12, and 24 h post-LPS treatment (Figure 3). At 6 h post-LPS treatment, ifnphi1 expression is significantly upregulated in both WT and sting (−/−) fish, but is more prominent in WT fish. At 12 h, the transcription levels of all the analyzed downstream genes are significantly upregulated in WT, while only il6 is upregulated in sting (−/−) larvae. At 24 h, ifnphi1 is downregulated in WT, while all the analyzed genes are upregulated in sting (−/−) larvae. Studies suggested the crucial role of STING upon LPS stimulation in inducing IFN production via cGAMP-primed enhancement [44]. LPS challenge in Oplegnathus fasciatus also caused a gradual increase in ifn1 transcription at early time points, which reduced at later time points. Peak transcript levels were observed at 12 h in the blood and 24 h in the head kidney [45]. Our results (upregulated ifnphi1 expression at 6 and 12 h, and downregulated expression at 24 h in WT) agree with this previous study. Furthermore, the observed fold reduction in ifnphi1 upregulation in sting (−/−) compared to WT fish at early time points (6 and 12 h) indicates the involvement of a STING-mediated pathway in Ifnphi1 activation upon LPS recognition. In endothelial cells, mitochondrial DNA (mtDNA) is released from the mitochondria into the cytosol via mitochondrial pores induced by activation of the pore-forming protein gasdermin D. Gasdermin D is activated by LPS [46]. Subsequently, cytosolic mtDNA stimulates the cGAS-STING pathway. Another critical factor of Gram-negative bacterial sensing is the noncanonical inflammasome. In zebrafish fibroblasts, Caspase b binds to LPS directly via its N-terminal pyrin death domain, and its oligomerization is critical for pyroptosis [47]. In a previous study, caspase expression patterns were examined after LPS treatment to analyze the activation of noncanonical inflammasomes in zebrafish. It was revealed that Caspase a promotes pyroptosis canonically, similar to mammalian Caspase 1, while Caspase b induces non-canonical pyroptosis [48]. Moreover, the upregulated expression of downstream genes in sting (−/−) (but not in WT) fish at 24 h indicates that WT larvae recover from LPS-induced stress earlier than sting (−/−) larvae. Altogether, the lack of transcriptional stimulation of downstream genes in sting (−/−) zebrafish compared to WT zebrafish at early time points (6 and 12 h) after LPS treatment, and the late (24 h) response in sting (−/−) fish indicate the essential role of zebrafish Sting in conferring antibacterial immunity. Furthermore, LPS-treated WT and sting (−/−) larvae show 100% survival for three days. The results from the LPS tolerance study in zebrafish larvae 2, 5, and 10 dpf also show that mortality starts to occur at LPS concentrations of 150 µg/mL [49]. Therefore, the LPS concentration (100 µg/mL) used in this study was not lethal to WT or sting (−/−) zebrafish larvae.

Effects of Sting Deficiency on Susceptibility to E. piscicida Infection in Zebrafish
To study the effects of sting deficiency during bacterial infection, percent mortality was analyzed in sting (−/−) and WT zebrafish following E. piscicida infection ( Figure 4A). Mortality begins in sting (−/−) and WT zebrafish at 4 and 5 dpi, respectively. Mortality in the sting (−/−) group reaches 55% at 8 dpi and remains unaltered thereafter, whereas it reaches 10% in WT fish at 8 dpi and remains unaltered. CDN-mediated activation of STING leads to the activation of genes that control pathogen replication and boost host adaptive immunity [8,[50][51][52]. The involvement of the cGAS-STING pathway in Gram-positive and Gram-negative bacterial infection has been reported previously. However, the role of the cGAS pathway in bacterial infection is complex compared to its role in antiviral responses [6]. The importance of Sting in the antibacterial immunity of zebrafish is clearly demonstrated by our observed percent mortality values. Biomolecules 2023, 13, x FOR PEER REVIEW 9 of 15  (Figure 3B, 3D, and 3F respectively) zebrafish larvae. Ef1∝ was used as an internal control to analyze the relative mRNA expression of genes downstream of sting using the Livak method. Transcription levels of target genes in the PBS-treated group were considered as 1, and expression levels in the LPS-treated groups were normalized to those in the PBS-treated group and are represented as fold values. Standard deviation (SD; n = 3) is indicated by the error bars. Significantly differentially transcribed genes (when compared to the respective PBStreated control) are marked with an asterisk (*: p < 0.05). 10% in WT fish at 8 dpi and remains unaltered. CDN-mediated activation of STING leads to the activation of genes that control pathogen replication and boost host adaptive immunity [8,[50][51][52]. The involvement of the cGAS-STING pathway in Gram-positive and Gram-negative bacterial infection has been reported previously. However, the role of the cGAS pathway in bacterial infection is complex compared to its role in antiviral responses [6]. The importance of Sting in the antibacterial immunity of zebrafish is clearly demonstrated by our observed percent mortality values. ef-1α expression was used as an internal control to analyze relative mRNA expression using the Livak method, and gene expression was normalized to that of the respective PBS control group. The error bars and the asterisk (* p < 0.01, ** p < 0.001) indicate the standard deviation and significantly different transcription levels compared to the respective control, respectively.

Temporal Expression Analysis in Zebrafish upon E. piscicida Infection
To understand the role of Sting in controlling immune-related gene expression during bacterial infection, we compared the modulation of downstream gene expression in sting (−/−) and WT zebrafish following E. piscicida infection. The relative mRNA expression of tbk1 is significantly downregulated in the sting (−/−) group at 24 and 72 hpi, while no significant changes are observed in the WT control group ( Figure 4B). The mRNA expression of nf-κb is significantly upregulated at 6 and 48 hpi in the WT control group, and significantly downregulated at 6 and 72 hpi in the sting (−/−) group ( Figure 4C). The transcription of irf3 is significantly upregulated at 48 hpi in the WT control group; however, no significant change is observed in the sting (−/−) group ( Figure 4D). The transcription of irf7 is upregulated at 6 hpi in the WT control group, but is significantly downregulated at 6 and 24 hpi, and upregulated at 72 hpi in the sting (−/−) group ( Figure 4E). The transcription of ifnphi1 is significantly upregulated at 6 hpi in the WT control group and significantly downregulated at 48 hpi in both the WT and sting (−/−) groups ( Figure 4F). tnfα transcription is significantly upregulated at 6 hpi in the WT control group, whereas it fluctuates in the sting (−/−) group ( Figure 4G). The transcriptional pattern of il6 is similar to that of tnfα in both groups ( Figure 4H), which suggests that these two genes may be regulated by the same mechanism.
Studies using murine models elucidated a STING-mediated mechanism that contributes to NF-κB activation [9]. STING-mediated activation of IRF3 and NF-κB is speciesdependent. Observations using human and mouse primary immune cells indicate strong IRF3 and weak NF-κB responses to STING alleles (reporter signaling, approximately 40-60-fold and 15-fold, respectively) [8,51,53]. The activation of sting alleles from Danio rerio (zebrafish) and Salmo salar (salmon) in human cells demonstrated a stronger response by NF-κB compared to that by IRF3 (>100-fold stimulation). Furthermore, the CTT motif of zebrafish Sting dramatically enhances NF-κB signaling by recruiting Traf6 (Tnf receptor associated factor 6) [39]. The observed changes in nf-κb and irf3 expression post E. piscicida challenge in the current study supports these previous results, as sting-deficient zebrafish show decreased expression of nf-κb and irf3. Since cytokines play a critical role in host immune defense and repair mechanisms, we analyzed the transcriptional modulation of cytokines il6 and tnfα upon E. piscicida challenge [54]. Transcription modulation of il6 and tnfα at an early time point (6 hpi) is similar to that of nf-κb and irf7. The expression patterns at 6 hpi clearly indicate that sting deficiency inhibits ifnphi1, il6, and tnfα expression through the nf-κb pathway. However, different cellular signaling pathways may be involved in modulating il6 and tnfα expression at later time points (24, 48, and 72 hpi). Previous studies reported that deficiencies in TBK1 induced TNFα-mediated cell death [55,56]. This may explain the upregulation of tnfa and il6 in sting (−/−) zebrafish at 24 and 72 hpi, as tbk1 is downregulated at these time points. Further, mortality starts in sting (−/−) fish at 4 dpi, while it begins at 5 dpi in WT fish. Therefore, tnfα and il6 upregulation at 72 hpi may result from early death signaling in sting (−/−) zebrafish. LPS-stimulated ROS generation in cardiomyocytes induces the translocation of NLRP3 from the nucleus to the cytoplasm via a STING-independent pathway [57]. The presence of NLRP3 inflammasomes is critical in triggering the expression of proinflammatory cytokines [58]. ROS levels in the immune organs of fish increase upon bacterial infection, which may trigger translocation of NLRP3 to the cytoplasm in a Sting-independent manner, inducing inflammation and apoptosis [59,60]. This may also explain the upregulation of tnfα and il6 at later time points in the sting (−/−) group. STING activation by L. monocytogenes triggers ifn1 to downregulate cell-mediated immunity [61]. IFN-1 is mostly related to antiviral immunity; however, it plays a role in antibacterial responses in mice [62,63]. A study on zebrafish Ifnphi1 and Ifnphi3 activation by Irf1, Irf3, and Irf7 shows that Irf3 acts both as a positive and negative regulator of ifn genes, depending on Irf1, Irf3, and Irf7 load in the cell [64]. This might explain the observed differential expression patterns of irf3, irf7, and ifnphi1.

Conclusions
In summary, we established a sting (−/−) zebrafish model to analyze the role of zebrafish Sting against bacterial infection. The transcription of genes downstream of sting (tbk1, nf-κb, irf3, and, irf7) was compared in WT and sting (−/−) zebrafish. The elevated expression of downstream genes in the sting (−/−) zebrafish illustrates the role of zebrafish Sting in controlling downstream gene expression. Furthermore, the role of Sting in survival and downstream gene modulation during bacterial infection were studied via E. piscicida challenge in sting (−/−) and WT zebrafish. The observed premature mortality in sting (−/−) zebrafish compared to WT zebrafish, and the differential expression patterns of downstream genes observed during E. piscicida and LPS stimulation, suggest that zebrafish Sting plays an essential role against bacterial infection by controlling downstream gene expression. The results of our study provide a reference for future studies on the antibacterial and other immune responses of zebrafish. Further, the understanding of the molecular pathways behind human diseases is being advanced by disease modeling in zebrafish. The sting (−/−) zebrafish model should be a key component of future research on antibacterial activity in humans based on these findings.

Supplementary Materials:
The following supporting information can be downloaded at: https: //www.mdpi.com/article/10.3390/biom13020324/s1, Figure S1A: The relative transcription of sting in different tissues of healthy adult zebrafish; Figure S1B: Transcription of sting through the embryonic developmental stages. Figure S2A: The experimental flowchart of E. piscicida infection; Figure S2B: PCR confirmation of E. piscicida infection at 6 hpi.