The aryl hydrocarbon receptor (AhR) is a ligand-activated transcription factor that mediates immune modulation following exposure of animals to many environmental xenobiotics. However, its role in innate immune responses during viral infection is not fully understood, especially in invertebrates. In this study, a cDNA encoding an AhR homolog was cloned from an arthropod Litopenaeus vannamei (LvAhR). The expression of LvAhR was strongly upregulated in response to the challenge of white spot syndrome virus, a pathogen of highly contagious and fatal infectious disease of shrimp. The relevance of LvAhR to host defense was underlined by heightened susceptibility and elevated virus loads after AhR-silenced shrimp exposure to white spot syndrome virus. LvAhR could induce an apoptosis response through regulating the expression of L. vannamei caspase-1 (homologous to human caspase-3) by directly targeting its promoter that was required to couple with AhR nuclear translocator. Additionally, knockdown of L. vannamei caspase-1 resulted in elevated virus titers and a lower cell apoptotic rate. Thus, we demonstrate that an AhR–caspase axis restrains virus replication by promoting antiviral apoptosis, supporting a previously unidentified direct link between AhR signaling and caspase-mediated apoptosis signaling and, furthermore, suggests that the AhR–caspase axis could be a potential therapeutic target for enhancing antiviral responses in arthropods.

The aryl hydrocarbon receptor (AhR) is a ligand-dependent transcription factor that integrates environmental, dietary, metabolic, and microbial cues to control complex transcriptional programs (1). The protein structure of AhR consists of a basic helix-loop-helix (bHLH) domain, a Per–Arnt–Sim (PAS) A domain, a PAS B domain, and a transcriptional activation domain (TAD), in which the PAS domains can sense both endogenous factors (such as oxygen tensions or redox potentials) and exogenous factors (such as polyaromatic hydrocarbons and environmental toxins) (2). Under normal physiological conditions, AhR is kept in an inactive state in the cytoplasm as part of a protein complex that consists of a dimer of the 90-kDa heat shock protein (HSP90), the AhR interacting protein (AIP), the co-chaperone p23, and the protein kinase SRC (1). This chaperone complex keeps AhR inactive, prevents its proteasomal degradation, and keeps it in a high-affinity state for its ligands. Upon ligand (agonist) binding, the AhR–ligand complex translocates to the nucleus, where AhR dissociates from the complex and pairs with its binding partner, AhR nuclear translocator (ARNT, also known as HIF-1β). The AhR–ARNT ligand complex then is recruited to the xenobiotic response element (XRE; also known as the dioxin response element) that displays a common DNA consensus motif (5′-TNGCGTG-3′) to control gene expression (1, 2). These target genes include xenobiotic metabolizing enzymes, the most prominent of which are the microsomal cytochrome P450-dependent monooxygenases CYP1A1 and CYP1A2 (3). These enzymes are induced in an attempt to metabolize and inactivate toxic pollutants that bind AhR, such as 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) (3). In addition, AhR also controls the expression of genes that do not contain XREs by interacting with other transcription factors. For example, the AhR–ARNT ligand complex has been shown to interact with the estrogen receptor, retinoic acid receptor, and retinoblastoma protein or Krüppel-like factor 6 (KLF6) to inspire the expression of genes that do not bear XREs in their regulatory regions (47).

Growing evidence shows that the AhR plays an important role in a wide range of biological progresses, including growth, development, immunity, tryptophan metabolism, gut microbiota, and tumor therapy (811). Apoptosis is one of the basic cellular activities of multicellular organisms, and AhR is shown to be involved in the regulation of apoptosis. An early study reported that TCDD induced apoptosis in the CNS of zebrafish during development by activating AhR (12). In mouse cortical neurons, plasticizer dibutyl phthalate (DBP) can be mediated by AhR at micromolar concentrations to activate caspase-3, reduce cell survival rates, and induce the production of apoptotic bodies (13). In dioxin-like polychlorinated biphenyls–treated mouse spleen cells, the expressions of Bax, caspase-3, caspase-8, and caspase-9 were increased, which are related to the activation of AhR (14). Recently, research has suggested that in the early stage of SARS-CoV-2 infection, the activated AhR increases the expression of CYP1 family genes such as CYP1A1, CYP1B1, and CYP1A2, resulting in the activation of the caspase-dependent apoptosis pathway (15).

In multicellular organisms suffering from pathogenic infection or adverse stimulation, apoptosis commonly functions as a protective mechanism and a defensive strategy, which has positive survival significance (16, 17). In the early stages of viral infection, this kind of autonomous programmed cell death (apoptosis) can disintegrate a variety of proteins and destroy the “base” needed for virus replication, thereby inhibiting viral proliferation (16, 17). The central components of the apoptotic response are the caspase proteins, which comprise two distinct classes: the initiators and the effectors (18). Generally, the activation of an effector caspase is carried out by an initiator caspase, whereas the initiator caspases are auto-activated. The effector (executioner) caspases, which include caspase-3, caspase-6, and caspase-7, are able to perform the cleavage of the proteins in host cells (1821), as well as mediate the cleavage of viral proteins directly (22). Therefore, caspases are regarded as important antiviral factors in host immunity. In clinical antiviral therapy, it was found that patients with low caspase activity cannot clear viral infection during antiviral therapy and show viral relapse after antiviral treatment (23).

Litopenaeus vannamei, also known as the Pacific white shrimp, has become the largest cultured crustacean species in the world, attributable to its ability to survive in a wide range of environments and its fast growth at high densities (24). However, emerging diseases such as white spot syndrome (WSS), acute hepatopancreatic necrosis disease, decapod iridescent virus 1–caused disease, and the microsporidian parasite Enterocytozoon hepatopenaei–caused disease have resulted in substantial production losses and sustained economic costs to the shrimp industry (25). Among these diseases, WSS, caused by a dsDNA virus of WSS virus (WSSV), frequently causes up to 100% mortality within 3–10 d, and leads to the most serious economic losses of about USD $1 billion per year (26). WSSV is a bacilliform, non-occluded enveloped virus belonging to the family Nimaviridae, genus Whispovirus (27). In the nucleus of infected cells, the WSSV genome is replicated and new virus particles are assembled, followed by the mature virus particles being formed in the cytoplasm (28). WSSV has a replication cycle of ∼22–24 h (29). The WSSV genome is ∼300 kbp and contains at least 181 open reading frames (ORFs) (30). Based on temporal expression profiles, WSSV genes can be classified as immediate-early, early, or late genes. Most of these encode proteins that show no homology to known proteins (31). Apoptosis has been shown to be vital for shrimp defense against pathogenic infections, including that of WSSV. Previous studies have found that WSSV infection can induce apoptosis in shrimp (32). In shrimp L. vannamei, several potential apoptosis-related genes such as apoptosis inducing factor and cytochrome c showed strong antiviral effects during WSSV infection (33, 34).

Although there is significant evidence indicating that AhR can affect caspase activity and promote apoptosis, how AhR modulates caspase-mediated apoptosis remains, to a wide extent, enigmatic, especially in invertebrates. In the current study, we identified an AhR homolog from an arthropod (shrimp) L. vannamei (LvAhR). We uncovered that AhR–caspase axis-mediated apoptosis is essential for host defense against WSSV infection in shrimp. In particular, LvAhR activated the expression of L. vannamei caspase (LvCaspase-1) by targeting an XRE motif in its promoter, which in turn induced an antiviral apoptosis. Our findings thus demonstrate a link between AhR signaling and apoptosis signaling in an arthropod (shrimp), and understanding which factors affect the LvAhR–caspase axis may help in designing therapies to prevent and treat shrimp suffering from viral infections such as WSSV.

An expressed sequence tag encoding a putative AhR protein was retrieved from L. vannamei transcriptome data analyzed by our laboratory (35). The 3′ and 5′ ends of LvAhR were obtained with gene-specific primers using the rapid amplification of cDNA ends (RACE) method as previously described (Supplemental Table I) (36). The cDNA template for RACE-PCR was prepared with the SMARTer PCR cDNA synthesis kit (Clontech, Shiga, Japan). 5′- and 3′-RACE PCR amplifications were performed with universal primer A mix and LvAhR-specific reverse primer 5RACE1 or 3RACE1. The products from the first round of PCR were diluted 50-fold as templates for the second round of PCR. Primers of nested universal primer A and LvAhR-5RACE2 or 3RACE2 were used for the second round of 5′- and 3′-RACE PCR, respectively. The final products were cloned into pMD-20T cloning vector (Takara Bio, Shiga, Japan), and 12 positive clones were selected and sequenced.

Protein domains of AhRs were predicted using the SMART program (http://smart.embl-heidelberg.de/) (37). Protein sequences of AhR homologs from other species were found in the National Center for Biotechnology Information (http://www.ncbi.nlm.nih.gov/) databases. Sequences of LvAhR and AhR homologs from other species were aligned by the Clustal X v2.0 program (38) and visualized by GeneDoc software (http://www.nrbsc.org/gfx/genedoc/) (39). Phylogenetic trees were constructed by MEGA 5.0 software with the neighbor-joining method (40).

The pAC5.1A-hemagglutinin (HA) plasmid was reconstructed by inserting a 3× HA sequence at the N terminal of the multiple cloning site of pAC5.1A-V5/His A (36). The ORFs of LvAhR, L. vannamei ARNT (LvARNT) (GenBank no. ACU30155.1, http://www.ncbi.nlm.nih.gov), or their domain or fragment deletion mutants were cloned into pAc5.1A-HA to generate pAc5.1-LvAhR-HA, pAc5.1-LvARNT-HA, pAc5.1-LvAhR-ΔbHLH-HA, pAc5.1-LvAhR-ΔPAS A-HA, pAc5.1-LvAhR-ΔPAS B-HA, pAc5.1-LvAhR-ΔTAD-HA, pAc5.1-LvARNT-ΔbHLH-HA, pAc5.1-LvARNT-ΔPAS A-HA, pAc5.1-LvARNT-ΔPAS B-HA, or pAc5.1-LvARNT-ΔTAD-HA plasmids for expressing HA-tagged proteins. The ORFs without stop codons of LvAhR and LvARNT were cloned into pAc5.1A-V5/His A to generate pAc5.1-LvAhR-V5 and pAc5.1-LvARNT-V5 for expressing V5-tagged proteins. The ORFs without stop codons of LvAhR were cloned into pAc5.1A-GFP to generate pAc5.1-LvAhR-GFP for expressing GFP-tagged proteins. The promoter sequence of LvCaspase-1 was retrieved from the L. vannamei genome (41) and constructed into the pGL3-Basic vector (Promega) for luciferase reporter gene analysis.

Healthy shrimp were provided by a shrimp farm in Maoming, Guangdong Province, China. Tissues of hemocytes, gills, hepatopancreas, and intestines were sampled and pooled from 15 shrimp for the tissue expression distribution. For the immune challenge assay, each shrimp was injected with 50 μl of viral suspension containing ∼1 × 105 particles of WSSV at the second abdominal segment. Gills, hemocytes, and intestines of challenged shrimp were sampled at 0, 4, 8, 12, 24, 36, 48, and 72 h postinjection, and each sample was collected and pooled from 15 shrimp. Total RNA extraction and a quantitative RT-PCR (qRT-PCR) assay were performed according to former descriptions (35).

qRT-PCR assays were also performed to assess the mRNA levels of genes in the in vivo RNA interference (RNAi) experiments. Tissues of gill were sampled and pooled from nine shrimp. Expression levels of LvAhR, LvARNT, LvCaspase-1–LvCaspase-1, VP28, VP15, wsv056, and wsv069 were calculated using the Livak (2−ΔΔCt) method after normalization to LvEF-1a (GenBank no. GU136229, http://www.ncbi.nlm.nih.gov), whereas the expression levels of Drosophila melanogaster Dcp-1, Grim, and Reaper (DmDcp-1, DmGrim, and DmReaper, respectively) were normalized by Rp49 (GenBank no. Y13939.1, http://www.ncbi.nlm.nih.gov). Primer sequences are listed in Supplemental Table I. Each experiment was done at least three times.

The dsRNAs specifically targeting LvAhR, LvCaspase-1 (GenBank no. DQ988351.1, http://www.ncbi.nlm.nih.gov), and GFP (as a control) were synthesized by in vitro transcription using the kit with the gene-specific primers in Supplemental Table I. The lengths of LvAhR dsRNA #1, LvAhR dsRNA #2, LvCaspase-1 dsRNA, and GFP dsRNA are 418, 545, 489, and 504 bp, respectively. Shrimp in the experimental group were injected with LvAhR dsRNA or LvCaspase-1 dsRNA (2 μg/g shrimp), whereas the control groups were injected with GFP dsRNA (2 μg/g shrimp). The qRT-PCR was used to investigate the RNAi efficiency. Each experiment was done at least three times.

Healthy shrimp (5 ± 0.5 g) were injected at the second abdominal segment with 10 μg (2 μg/g shrimp in 50 μl of PBS) of dsRNA (LvAhR dsRNA, LvCaspase-1 dsRNA, or GFP dsRNA). Forty-eight hours later, the shrimp in LvAhR-knockdown experiment were injected with WSSV suspension (1 × 105 copies/shrimp, n = 40) and mock challenged with PBS as a control (n = 40), respectively. In the LvCaspase-1–knockdown experiment, the shrimp were injected with WSSV suspension (∼1 × 105 copies/shrimp, n = 15) and mock challenged with PBS as a control (n = 15), respectively. Shrimp were kept in culture flasks for ∼7 d following infection. Cumulative mortality was recorded every 4 h. Differences in mortality levels between treatments were tested for statistical significance using the Kaplan–Meier plot (log-rank χ2 test) using GraphPad Prism software.

Absolute quantitative PCR was performed to monitor WSSV replication in shrimp in vivo. Briefly, we collected gills from shrimp at 48 and 168 h after WSSV infection (12 shrimp/treatment in the dsRNA-LvAhR experiment; 8 shrimp/treatment in the dsRNA-LvCaspase-1 experiment). Gill DNA was extracted with a Tiangen marine animals DNA kit (Tiangen Biotech, Beijing, China) according to the user’s introduction. The copies of WSSV genome were measured by absolute quantitative PCR using WSSV32678-F/WSSV32753-R primers and a TaqMan fluorogenic probe as described previously (Supplemental Table I) (42). The WSSV genome copy numbers in 0.1 μg of shrimp DNA were then calculated. All of the experiments were repeated three times.

In the dsRNA-LvAhR experiment, the hemocytes of shrimp were collected at 0, 2, 4, and 6 h after WSSV infection, respectively. In the dsRNA-LvCaspase-1 experiment, the hemocytes of shrimp were collected at 6 h after WSSV infection. Drosophila S2 cells were collected at 48 h after plasmid transfection. After being washed twice in cold PBS, ∼1 × 105 hemocytes and S2 cells were resuspended in 500 μl of 1× annexin binding buffer, followed by the addition of 10 μl of annexin V–allophycocyanin and 5 μl of propidium iodide (PI) (Keygen, catalog no. KGA1030-100), and then the cells were incubated for 15 min in the dark. Finally, the fluorescence emission of the stained samples was examined by flow cytometry (BD Biosciences, San Jose, CA). All of the experiments were repeated three times.

The gating strategy was set by the following steps: unstained hemocytes and S2 cells were used to measure the forward scatter (FSC) and side scatter (SSC) signals to remove debris and dead cells. Cellular debris were considered to be low FSC signals, whereas dead cells were considered to be low FSC signals and high SSC signals. Debris and dead cells were excluded by setting polygons that include events with high FSC signals and low SSC signals (viable cells). Then, the other gate was set to exclude doublets from viable cells. After the gate was set, unstained cells, annexin V–allophycocyanin-stained cells (no PI), and PI-stained cells (no annexin V–allophycocyanin) were used to set up compensation and divide the plots into four quadrants.

The hemocytes of dsRNA-treated shrimp were collected at 0, 2, 4, and 6 h after WSSV infection, respectively, and S2 cells were collected at 48 h after plasmid transfection. The caspase-3/7 activity of L. vannamei hemocytes was detected using a Caspase-Glo 3/7 assay (Promega, Madison, WI) in line with the manufacturer’s instructions. After being washed twice in cold PBS, 100 μl of Caspase-Glo 3/7 reagent was mixed with 2 × 104 shrimp hemocytes and incubated at 25°C for 1.5 h. The mixture was assayed using a luminometer to detect the activity of caspase-3/7. Each experiment was done at least three times.

Drosophila S2 cells were seeded onto glass slides in 24-well plates. After culturing for 24 h, S2 cells were transfected with 0.2 μg of firefly luciferase reporter gene plasmids, 0.04 μg of pRL-TK Renilla luciferase plasmid (internal control), 0.2 μg of protein expression plasmids, or 0.2 μg of pAc5.1a-V5 plasmid (as a control). Forty-eight hours later, the cells were lysed, 60% of the lysate was used for Dual-Luciferase reporter assays, and the remaining 40% of the cell lysate was used to analyze the expression level of proteins by Western blotting. All of experiments were repeated three times.

Protein samples were separated in SDS-PAGE gels, transferred to polyvinylidene difluoride membranes, and incubated with the appropriate Abs. The primary Abs used in Western blotting included a rabbit anti-V5 Ab (Merck Millipore, Burlington, MA, catalog no. AB3792), rabbit anti-HA Ab (Sigma-Aldrich, cat. no. H6908-100UL), rabbit anti-GFP Ab (Sigma-Aldrich, catalog no. G1544-100UG), mouse anti-LvAhR Ab (customized in Abmart), mouse anti–α-tubulin Ab (Cell Signaling Technology, catalog no. 3873S), or mouse anti-actin (C4) Ab (Cell Signaling Technology, catalog no. 41185S). The secondary Abs were the anti-rabbit IgG-HRP conjugate (Promega, catalog no. W401B) and anti-mouse IgG-HRP conjugate (Promega, catalog no. W402B). Both primary and secondary Abs were incubated in TBST (0.05 M Tris [pH 7.6] with 0.9% NaCl and 0.1% Triton X-100). Membranes were developed with the ECL blotting substrate (Thermo Scientific), and chemiluminescence was detected using the 5200 chemiluminescence imaging system (Tanon).

All reagents and experimental data are available from corresponding author upon reasonable request.

The complete transcript of LvAhR was 2957 bp long, consisting of a 296-bp 5′-untranslated region, a 333-bp 3′-untranslated region including a poly(A) tail, and a 2328-bp ORF encoding a 775-aa polypeptide with a calculated molecular mass of 85.8 kDa (GenBank accession no. OM046573, https://www.ncbi.nlm.nih.gov/nuccore/OM046573). The LvAhR protein sequence contained a bHLH domain, a PAS A domain, and a PAS B domain at 43–97, 123–189, and 264–377 residues in the N-terminal, respectively (Supplemental Fig. 1).

The multiple sequences analysis showed that LvAhR resembled the Homo sapiens AhR (21% sequence identity), Mus musculus AhR (21% identity), Gallus gallus AhR (22% identity), Xenopus laevis AhR (23% identity), D. melanogaster AhR (46% identity), Crassostrea gigas AhR (31% identity), Mya arenaria AhR (30% identity), Caenorhabditis elegans AhR (23% identity), and Danio rerio AhR (22% identity) (Supplemental Fig. 1A). Among the regions of bHLH domains, PAS A domains, and PAS B domains, these domains were highly conserved in both invertebrates and vertebrates (Supplemental Fig. 1A). In addition, LvAhR and AhR homologs of several other species were phylogenetically analyzed by the neighbor-joining method. The AhR proteins used in this study were clustered to three groups, including vertebrates, chordates, and invertebrates, and LvAhR was mostly clustered to the subgroup of Arthropoda AhR proteins, including D. melanogaster AhR, Apis mellifera AhR, and Daphnia magna AhR (Supplemental Fig. 1B). Taken together, these results strongly suggest that LvAhR is a homolog of the AhR family and displays high sequence conservation in regions of the bHLH domain, PAS A domain, and PAS B domain.

Tissue distribution of LvAhR mRNA was analyzed by qRT-PCR. The mRNA of LvAhR could be detected in four immune-related tissues including hemocytes, the hepatopancreas, gills, and intestines (Fig. 1A). The expressional levels of LvAhR in the hepatopancreas, gills, and intestines were ∼4.4-, ∼5.3-, and ∼5.7-fold higher than those in hemocytes (normalization to 1.0), respectively. (Fig. 1A). Shrimp hemocytes are leukocyte-like blood cells with the function of eliminating pathogens, whereas gills and intestines are the crucial organs involved in immune defense against virus infection (43). Next, we explored the expression pattern of LvAhR after WSSV infection in the three tissues of hemocytes, gills, and intestine. In the hemocytes of WSSV-infected shrimp, the expression of LvAhR was significantly upregulated at 4 h with an ∼6.93-fold increase, and at 8 h with a peak of ∼15.65-fold increase. After that, the expression of LvAhR remained high during 12–36 h with ∼5.23-, ∼2.7-, and ∼8.42-fold increases at 12, 24, and 36 h after WSSV infection, respectively (Fig. 1B). In gills, LvAhR expression quickly reached its maximum with an ∼6.57-fold increase at 4 h, followed by the increased expression levels declining at 8–24 h postinfection. Notably, the expression of LvAhR was downregulated with an ∼0.57-fold decrease at 36 h, whereas its expression was significantly upregulated again at 72 h with an ∼3.50-fold increase (Fig. 1C). In intestines, LvAhR expression was sharply upregulated during the early stage of infection with ∼5.60-, ∼5.47-, and ∼3.67-fold increases at 4, 8, and 12 h, respectively (Fig. 1D). Subsequently, the levels of LvAhR expression in the intestines of shrimp infected with WSSV resembled those of shrimp in the PBS group (Fig. 1D). The control group injected with PBS did not show clear changes in LvAhR expression (Fig. 1B–D). Taken together, these results suggest that LvAhR could respond to WSSV infection.

FIGURE 1.

Expression of LvAhR in healthy and immune challenged shrimp detected by qRT-PCR. Expression of LvAhR was normalized to that of LvEF-1α using the Livak (2−ΔΔCt) method, and the data were provided as the means ± SD of triplicate assays with at least nine shrimp for each sample. (A) Tissue distribution of LvAhR in healthy L. vannamei. Expression level in the hemocytes was used as control and set to 1.0. (BD) Expression profiles of LvAhR after WSSV infection in hemocytes (B), gills (C), and intestines (D). The expression level at each time point was normalized to the PBS-injected group. *p < 0.05, **p < 0.01.

FIGURE 1.

Expression of LvAhR in healthy and immune challenged shrimp detected by qRT-PCR. Expression of LvAhR was normalized to that of LvEF-1α using the Livak (2−ΔΔCt) method, and the data were provided as the means ± SD of triplicate assays with at least nine shrimp for each sample. (A) Tissue distribution of LvAhR in healthy L. vannamei. Expression level in the hemocytes was used as control and set to 1.0. (BD) Expression profiles of LvAhR after WSSV infection in hemocytes (B), gills (C), and intestines (D). The expression level at each time point was normalized to the PBS-injected group. *p < 0.05, **p < 0.01.

Close modal

To accurately determine the function of LvAhR during WSSV infection, we designed and synthesized two pieces of dsRNA-LvAhR, named dsRNA-LvAhR #1 (targeting 807–1224 bp) and dsRNA-LvAhR #2 (targeting 1398–1942 bp). We then performed two separate knockdown experiments using RNAi strategy in vivo. The silencing efficiency of LvAhR was checked by qRT-PCR at 48 and 168 h after dsRNA injection, respectively. We observed that both the transcription levels and protein levels of LvAhR can be downregulated by dsRNA-LvAhR #1 or dsRNA-LvAhR #2 (Fig. 2A) at 48 h after dsRNA injection; meanwhile, the inhibitory effect of dsRNA on LvAhR still works at 168 h (Fig. 2A). In addition, at 48 h after dsRNA injection, the shrimp were challenged with WSSV or PBS (as a control), and their surviving numbers were counted every 4 h. Survival rate showed that both shrimp in the dsRNA-GFP group and in the dsRNA-LvAhR group started to die at 36 h after WSSV infection. However, the overall survival rates of shrimp in the dsRNA-LvAhR #1 group (χ2 of 4.181, p = 0.0409) and the dsRNA-LvAhR #2 group (χ2 of 7.385, p = 0.0066) were significantly lower than those in the GFP-knockdown group. As negative controls, both the GFP and LvAhR knockdown did not result in death in PBS-treated shrimp (Fig. 2B). We subsequently investigated whether the differences in survival rates were associated with changes of viral copies and viral gene expression levels in vivo. We performed the RNAi experiment again during WSSV infection in parallel to survival rate and probed the viral copies and several viral genes, including two immediate early genes, wsv056 and wsv069, and two late structural genes, VP28 and VP15 (4446). As expected, absolute quantitative PCR results revealed that the WSSV genomic copies (viral loads) were significantly increased in the dsRNA-LvAhR #1 and dsRNA-LvAhR #2 groups compared with the dsRNA-GFP group (Fig. 2C). Accordingly, the expression levels of four viral genes, including wsv056, wsv069, VP28, and VP15, in the dsRNA-LvAhR+WSSV group were higher than those of the dsRNA-GFP+WSSV group at 48 and 168 h after WSSV infection (Fig. 2D, 2E). Taken together, these results strongly suggest that LvAhR is critical for host defense against WSSV.

FIGURE 2.

Functional analysis of LvAhR in WSSV infection by dsRNA-mediated RNAi. (A) The silencing efficiency of two dsRNA-LvAhRs (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) was checked by qRT-PCR (mRNA levels) and Western blotting (protein levels) at 48 and 168 h after WSSV infection. (B) Survival of WSSV-challenged LvAhR-silenced shrimp and GFP dsRNA–treated shrimp. Experiments were performed three times with similar results and analyzed statistically by the Kaplan–Meier plot (log-rank χ2 test). (C) The copies of WSSV in dsRNA-LvAhR–treated (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) or dsRNA-GFP–treated shrimp at 48 and 168 h after WSSV infection. The data were provided as the means ± SD of copies from 12 shrimp and analyzed statistically by a Student t test. **p < 0.01. (D and E) Expression level of VP28, VP15, wsv056, and wsv069 in dsRNA-GFP–treated or dsRNA-LvAhR–treated (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) shrimp at 48 and 168 h after WSSV infection were detected by qRT-PCR; the internal control was LvEF-1a, and the Livak (2−ΔΔCt) method was used. All data were provided as the means ± SD of triplicate assays with nine shrimp for each sample (A, D, E) and analyzed statistically by a Student t test. **p < 0.01.

FIGURE 2.

Functional analysis of LvAhR in WSSV infection by dsRNA-mediated RNAi. (A) The silencing efficiency of two dsRNA-LvAhRs (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) was checked by qRT-PCR (mRNA levels) and Western blotting (protein levels) at 48 and 168 h after WSSV infection. (B) Survival of WSSV-challenged LvAhR-silenced shrimp and GFP dsRNA–treated shrimp. Experiments were performed three times with similar results and analyzed statistically by the Kaplan–Meier plot (log-rank χ2 test). (C) The copies of WSSV in dsRNA-LvAhR–treated (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) or dsRNA-GFP–treated shrimp at 48 and 168 h after WSSV infection. The data were provided as the means ± SD of copies from 12 shrimp and analyzed statistically by a Student t test. **p < 0.01. (D and E) Expression level of VP28, VP15, wsv056, and wsv069 in dsRNA-GFP–treated or dsRNA-LvAhR–treated (dsRNA-LvAhR #1 and dsRNA-LvAhR #2) shrimp at 48 and 168 h after WSSV infection were detected by qRT-PCR; the internal control was LvEF-1a, and the Livak (2−ΔΔCt) method was used. All data were provided as the means ± SD of triplicate assays with nine shrimp for each sample (A, D, E) and analyzed statistically by a Student t test. **p < 0.01.

Close modal

As evidenced above, LvAhR is critical for shrimp defense against WSSV; however, its exact role and the underlying mechanism are still elusive. In vertebrates, emerging evidence shows that AhR is implicated in the process of apoptosis, and that apoptosis commonly functions as a protective mechanism and defensive strategy during virus infection in evolution (43). Thus, we hypothesize that LvAhR could regulate the apoptotic response during WSSV infection. To verify the idea, we first explored the effect of the ectopic expression of LvAhR on apoptosis in Drosophila cells, because there is no available shrimp cell line. The pAc-LvAhR-GFP plasmid and empty plasmid (as a control) were transfected into Drosophila S2 cells (Fig. 3A). Forty-eight hours after transfection, the S2 cells were collected and apoptotic cells of the two groups were detected by flow cytometry assay. The gating strategy was set as shown in Supplemental Fig. 2. After the GFP-positive cells were selected, ∼8.5% of the control group cells were apoptotic, whereas ∼51.8% of cells in the LvAhR overexpressed group were apoptotic (Fig. 3B). Accordingly, the caspase-3/7 activity of S2 cells transfected with pAc-LvAhR-GFP was significantly increased compared with that of S2 cells transfected with pAc-GFP empty plasmid (Fig. 3C). We next investigated whether the caspase cascade was activated after LvAhR was overexpressed in S2 cells. In Drosophila, the occurrence of the caspase-mediated apoptotic response requires proapoptotic proteins such as DmReaper and DmGrim to drive the caspase cascade, and finally activating the downstream effector DmDcp-1 (caspase-3–like protein) (47). We observed that the ectopic expression of LvAhR was able to induce the expression levels of DmDcp-1, DmGrim, and DmReaper with ∼2.45-, ∼2.31-, and ∼2.23-fold increases, respectively (Fig. 3D). In summary, LvAhR could induce apoptosis and activate the caspase cascade in Drosophila S2 cells.

FIGURE 3.

Proapoptosis function of LvAhR in S2 cells. (A) pAc-GFP and pAc-LvAhR-GFP plasmids were transfected into S2 cells, respectively, and the overexpression level of LvAhR was detected by Western blotting at 48 h posttransfection. (B) Left, pAc-GFP and pAc-LvAhR-GFP plasmids were transfected into S2 cells, respectively. At 48 h post transfection, the S2 cells were stained with annexin V–allophycocyanin and PI. GFP-positive cells were distinguished to analyze apoptosis. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) Caspase-3/7 activity in the S2 cells overexpressing pAc-GFP and pAc-LvAhR-GFP. (D) The expression of proapoptotic genes of DmDcp-1, DmGrim, and DmReaper from D. melanogaster in S2 cells overexpressing pAc-GFP and pAc-LvAhR-GFP were detected by qRT-PCR; the internal control was DmRp49, and the Livak (2−ΔΔCt) method was used. All data were provided as the means ± SD of triplicate assays (B–D) and analyzed statistically by a Student t test. **p < 0.01.

FIGURE 3.

Proapoptosis function of LvAhR in S2 cells. (A) pAc-GFP and pAc-LvAhR-GFP plasmids were transfected into S2 cells, respectively, and the overexpression level of LvAhR was detected by Western blotting at 48 h posttransfection. (B) Left, pAc-GFP and pAc-LvAhR-GFP plasmids were transfected into S2 cells, respectively. At 48 h post transfection, the S2 cells were stained with annexin V–allophycocyanin and PI. GFP-positive cells were distinguished to analyze apoptosis. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) Caspase-3/7 activity in the S2 cells overexpressing pAc-GFP and pAc-LvAhR-GFP. (D) The expression of proapoptotic genes of DmDcp-1, DmGrim, and DmReaper from D. melanogaster in S2 cells overexpressing pAc-GFP and pAc-LvAhR-GFP were detected by qRT-PCR; the internal control was DmRp49, and the Livak (2−ΔΔCt) method was used. All data were provided as the means ± SD of triplicate assays (B–D) and analyzed statistically by a Student t test. **p < 0.01.

Close modal

Next, we explored whether LvAhR could induce apoptosis in response to WSSV infection in shrimp in vivo. We used dsRNA-LvAhR #2 to knock down LvAhR in vivo and found that both the transcript levels and protein levels of LvAhR were substantively suppressed at 0, 2, 4, and 6 h after WSSV infection (Fig. 4A). A flow cytometry assay was then performed to evaluate the apoptotic cell rates of hemocytes in LvAhR-silenced shrimp and GFP-silenced shrimp (as controls). The gating strategy was set as shown in Supplemental Fig. 2. We observed that the total numbers of apoptotic hemocyte cells, including early apoptotic cells and late apoptotic cells, were evidently lessened in LvAhR-silenced shrimp during the early stage of WSSV infection (0, 2, 4, and 6 h), compared with those of the control group (Fig. 4B). Meanwhile, WSSV infection could induce hemocyte cell apoptosis, and a gradual increase in the total number of apoptotic hemocyte cells was observed in both LvAhR-silenced shrimp and GFP-silenced shrimp (Fig. 4B).

FIGURE 4.

Proapoptosis function of LvAhR in shrimp in the early stage of WSSV infection. (A) The gills from nine shrimp were sampled and pooled at 0, 2, 4, and 6 h after WSSV infection was checked by qRT-PCR (mRNA levels) and Western blotting (protein levels) for the silencing efficiency of LvAhR. (B) Left, he shrimp hemocytes in dsRNA-LvAhR–treated (n = 9) and dsRNA-GFP–treated (n = 9) shrimp at 0, 2, 4, and 6 h after WSSV infection were stained with annexin V–allophycocyanin and PI and then detected by flow cytometry. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) Caspase-3/7 activity in dsRNA-GFP–treated or dsRNA-LvAhR–treated shrimp at 0, 2, 4, and 6 h after WSSV infection. (D) Expression level of LvCapsase-1 in dsRNA-GFP–treated or dsRNA-LvAhR–treated shrimp at 0, 2, 4, and 6 h after WSSV infection was detected by qRT-PCR; the internal control was LvEF-1a. The data were provided as the means ± SD of triplicate assays with nine shrimp for each sample (A–D) and analyzed statistically by a Student t test. **p < 0.01.

FIGURE 4.

Proapoptosis function of LvAhR in shrimp in the early stage of WSSV infection. (A) The gills from nine shrimp were sampled and pooled at 0, 2, 4, and 6 h after WSSV infection was checked by qRT-PCR (mRNA levels) and Western blotting (protein levels) for the silencing efficiency of LvAhR. (B) Left, he shrimp hemocytes in dsRNA-LvAhR–treated (n = 9) and dsRNA-GFP–treated (n = 9) shrimp at 0, 2, 4, and 6 h after WSSV infection were stained with annexin V–allophycocyanin and PI and then detected by flow cytometry. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) Caspase-3/7 activity in dsRNA-GFP–treated or dsRNA-LvAhR–treated shrimp at 0, 2, 4, and 6 h after WSSV infection. (D) Expression level of LvCapsase-1 in dsRNA-GFP–treated or dsRNA-LvAhR–treated shrimp at 0, 2, 4, and 6 h after WSSV infection was detected by qRT-PCR; the internal control was LvEF-1a. The data were provided as the means ± SD of triplicate assays with nine shrimp for each sample (A–D) and analyzed statistically by a Student t test. **p < 0.01.

Close modal

The caspase-cascade system plays vital roles in the induction, transduction, and amplification of intracellular apoptotic signals (48). We thus explored the potential link between LvAhR and the caspase cascade under WSSV infection. Until now, there have been a total of five caspases identified in shrimp L. vannamei, including LvCaspase-1–LvCaspase-5, which were named based on the cloning orders rather than the corresponding orthologs of caspase-1–caspase-5 in mammals. LvCaspase-1, LvCaspase-2, and LvCaspase-5 are regarded as the executioner caspases, whereas LvCaspase-3 and LvCaspase-4 are the initiator caspases (49, 50). It is noteworthy that LvCaspase-1 is considered the ortholog of the caspase-3 family, and thus it is also referred to as caspase-3 or cap-3 in some literature (34, 51). Considering that the activation of caspase-3 and caspase-7 are landmark events in apoptosis, we thus detected the activity of caspase-3/7 of hemocytes in LvAhR-silenced shrimp. The results showed that the activity of caspase-3/7 in hemocytes from LvAhR-silenced shrimp was significantly lower than that in the control group in the early stage of WSSV infection (Fig. 4C), which is consistent with the rates of apoptotic hemocyte cells (Fig. 4B). We also investigated the effect of LvAhR knockdown on the expression levels of the five caspases (LvCaspase-1–LvCaspase-5). We found that the expression levels of LvCaspase-1–LvCaspase-5 in hemocytes from LvAhR-silenced shrimp were significantly downregulated compared with those of control groups at multiple or all time points in the early stage of WSSV infection (Fig. 4D, Supplemental Fig. 3). Taken together, these findings suggest that WSSV infection inducing the apoptosis of hemocytes could partly depend on LvAhR to affect the expression levels and activities of caspases.

It is generally accepted that AhR functions as a transcription factor by direct or functional interaction with other transcription factors, among which the most common partner is the ARNT, also known as HIF-1β (52). In the present study, we found that the expression of LvARNT (LvHIF-1β) in hemocytes was not induced by WSSV (Fig. 5), which was consistent with previous work (53). We next explored whether LvAhR and LvARNT could interact with each other and sought to map which domain of LvAhR mediates its association with LvARNT. We thus generated several deletion constructs of LvAhR (Fig. 5A) for coimmunoprecipitation assays. The results showed that the full length of LvAhR and LvAhR-ΔTAD (the C terminus of LvAhR was truncated) could bind to LvARNT, whereas LvAhR-ΔbHLH, LvAhR-ΔPAS A, and LvAhR-ΔPAS B abrogated the association with LvARNT (Fig. 5B). Therefore, these results suggest that the complete N-terminal region of LvAhR—consisting of the bHLH domain, PAS A domain, and PAS B domain—is indispensable for its interaction with LvARNT.

FIGURE 5.

LvCasapse-1 was regulated by the heterodimer of LvAhR and LvARNT. (A) Schematic diagram of the LvAhR deletion mutation constructs. (B) Coimmunoprecipitation analysis mapped the domains of LvAhR that were required to bind LvARNT. (C) Schematic diagram of the LvCaspase-1 promoter regions in the luciferase reporter gene constructs. (D) Expression profiles of LvARNT after WSSV infection in hemocytes. The expression level at each time point was normalized to the PBS-injected group. Each sample from 15 shrimp is shown. (E) Schematic diagram of the LvARNT deletion constructs. (F and G) The heterodimer composed of the full-length LvAhR and the full-length LvARNT efficiently activated the promoter activity of LvCaspase-1. All data (D, F, G) were provided as the means ± SD of at least three assays and analyzed statistically by a Student t test. **p < 0.01.

FIGURE 5.

LvCasapse-1 was regulated by the heterodimer of LvAhR and LvARNT. (A) Schematic diagram of the LvAhR deletion mutation constructs. (B) Coimmunoprecipitation analysis mapped the domains of LvAhR that were required to bind LvARNT. (C) Schematic diagram of the LvCaspase-1 promoter regions in the luciferase reporter gene constructs. (D) Expression profiles of LvARNT after WSSV infection in hemocytes. The expression level at each time point was normalized to the PBS-injected group. Each sample from 15 shrimp is shown. (E) Schematic diagram of the LvARNT deletion constructs. (F and G) The heterodimer composed of the full-length LvAhR and the full-length LvARNT efficiently activated the promoter activity of LvCaspase-1. All data (D, F, G) were provided as the means ± SD of at least three assays and analyzed statistically by a Student t test. **p < 0.01.

Close modal

The silencing of LvAhR affects the expression of caspases (Fig. 4D, Supplemental Fig. 3), which implies that LvAhR could be able to regulate the transcription of caspase directly. To address this, we focused on LvCaspase-1, an ortholog of caspase-3 in mammals (34, 51). We found that the promoter region of LvCaspase-1 contains a putative XRE motif located between 522 and 517 bp upstream of the transcriptional start site (Fig. 5C). The Dual-Luciferase reporter assays showed that LvARNT could strongly induce the promoter activity of LvCapsase-1 when coexpressed with the full length of LvAhR (∼5.42-fold), whereas overexpression of LvARNT alone or with LvAhR-ΔbHLH (∼0.97-fold) and LvAhR-ΔPAS A (∼1.15-fold) could not efficiently induce LvCaspase-1 promoter activity. Interestingly, the coexpression of LvARNT with LvAhR-ΔPAS B or LvAhR-ΔTAD showed low induction activity toward the LvCaspase-1 promoter with ∼2.06- and ∼3.40-fold increases, respectively (Fig. 5F). In comparison, LvARNT is also composed of a bHLH domain, a PAS A domain, a PAS B domain, and a C-terminal TAD domain (54) (Fig. 5E). To explore which domains are important for cooperation with LvAhR to induce the promoter activity of LvCaspase-1, several deletion mutations of LvARNT domains were used to perform the reporter assays. As shown in (Fig. 5G, LvAhR could efficiently induce LvCapsase-1 promoter activity only with the full length of LvARNT (∼5.39-fold), whereas the expression of LvAhR alone (∼1.07-fold) or alongside LvARNT-ΔbHLH (∼1.62-fold), LvARNT-ΔPAS A (∼1.57-fold), LvARNT-ΔPAS B (∼1.62-fold), and LvARNT-ΔTAD (∼2.85-fold) had no or very low induced LvCaspase-1 promoter activity. Taken together, these data indicate that the effective induction of LvCaspase-1 promoter activity in vitro by LvAhR requires the formation of dimer with LvARNT.

Although caspase-3 is a definite proapoptotic factor in mammals, the apoptotic function and antiviral role of LvCaspase-1, an ortholog of caspase-3, is still unknown. We have revealed that LvCaspase-1 was regulated by LvAhR, and that the latter played a positive role in promoting apoptosis and shrimp resistance against WSSV (Fig. 4). We thus speculated that LvCaspase-1 might resist WSSV by promoting apoptosis. As expected, we found that the expression of LvCaspase-1 in the gills of shrimp injected with dsRNA-LvCaspase-1 was only 23.0% of that of shrimp injected with dsRNA-GFP at 6 h after WSSV infection (Fig. 6A). The data showed that silencing of LvCaspase-1 remarkably restrained the apoptosis rate of hemocytes (4.3%), compared with that of the dsRNA-GFP group (17.1%, as a control) (Fig. 6B). In summary, this result suggests that LvCaspase-1 could promote apoptosis at the early stage of WSSV infection (6 h).

FIGURE 6.

The role of LvCaspase-1 in promoting virus-induced apoptosis and inhibiting virus replication. (A) The silencing efficiency of LvCaspase-1 was checked by qRT-PCR. The gills from nine shrimp were sampled and pooled at 6 h postinfection with WSSV. (B) Left, The hemocytes of shrimp in dsRNA-LvCaspase-1 group (n = 9) and dsRNA-GFP group (n = 9) at 6 h after WSSV infection were stained with annexin V–allophycocyanin and PI, and then detected by flow cytometry. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. The graphs are representative of three independent assays. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) The silencing efficiency of LvCaspase-1 was checked by qRT-PCR. The gills from nine shrimp were sampled and pooled at 48 h postinfection with WSSV. (D) Survival of LvCaspase-1–silenced shrimp and GFP dsRNA–treated shrimp during WSSV infection. Experiments were performed three times with similar results and analyzed statistically by the Kaplan–Meier plot (log-rank χ2 test). (E) The copies of WSSV in dsRNA-LvCaspase-1–treated and dsRNA-GFP–treated shrimp at 48 h after WSSV infection. The data were provided as the means ± SD of copies from eight shrimp and analyzed statistically by A Student t test. All data (A, B Right, C, E) were provided as the means ± SD of triplicate assays and analyzed statistically by a Student t test. **p < 0.01.

FIGURE 6.

The role of LvCaspase-1 in promoting virus-induced apoptosis and inhibiting virus replication. (A) The silencing efficiency of LvCaspase-1 was checked by qRT-PCR. The gills from nine shrimp were sampled and pooled at 6 h postinfection with WSSV. (B) Left, The hemocytes of shrimp in dsRNA-LvCaspase-1 group (n = 9) and dsRNA-GFP group (n = 9) at 6 h after WSSV infection were stained with annexin V–allophycocyanin and PI, and then detected by flow cytometry. Q1 represents viable cells, Q2 denotes early apoptotic cells, Q3 represents late apoptotic cells, and Q4 includes necrotic cells and nonviable and nonapoptotic cells. The graphs are representative of three independent assays. Right, Percentage of apoptotic cells indicated by histogram (percentage of apoptotic cells = percentage of apoptotic cells in Q3 + percentage of apoptotic cells in Q2). (C) The silencing efficiency of LvCaspase-1 was checked by qRT-PCR. The gills from nine shrimp were sampled and pooled at 48 h postinfection with WSSV. (D) Survival of LvCaspase-1–silenced shrimp and GFP dsRNA–treated shrimp during WSSV infection. Experiments were performed three times with similar results and analyzed statistically by the Kaplan–Meier plot (log-rank χ2 test). (E) The copies of WSSV in dsRNA-LvCaspase-1–treated and dsRNA-GFP–treated shrimp at 48 h after WSSV infection. The data were provided as the means ± SD of copies from eight shrimp and analyzed statistically by A Student t test. All data (A, B Right, C, E) were provided as the means ± SD of triplicate assays and analyzed statistically by a Student t test. **p < 0.01.

Close modal

To determine the function of LvCaspase-1 on antiviral roles, we suppressed LvCaspase-1 expression via a RNAi strategy during WSSV infection. The silencing efficiency of LvCaspase-1 was checked by qRT-PCR at 48 h after WSSV infection. The injection of LvCaspase-1 dsRNA resulted in a significant decrease in LvCaspase-1 transcription levels, which was downregulated to ∼0.23-fold of the GFP dsRNA injection groups (as a control) (Fig. 6C). Interestingly, we observed no significant difference in shrimp survival rates between the LvCapsase-1–knockdown group and the GFP-knockdown group (χ2 of 0.5093, p = 0.4754) (Fig. 6D). Importantly, we found that the WSSV genomic copies in shrimp gills of the dsRNA-LvCaspase-1 groups were ∼97.49-fold that of the GFP-dsRNA group (Fig. 6E) at 48 h after WSSV infection. We infer that the lack of significant difference in survival rates between groups could be attributable to the overlying outcome of LvCapsase-1 proapoptosis-induced death and its antiviral function in favor of survival, which needs to be confirmed by further studies. Taken together, these results suggest that LvCaspase-1 could play a key role in antiviral apoptosis against WSSV infection.

In recent years, growing evidence has indicated that AhR is an immunomodulatory factor that has great potential in areas of antiviral immunity, bacterial infection, autoimmune diseases, and cancer immunotherapy (9, 5557). Research has mainly been conducted into the effects of AhR on immunity in vertebrates, and works focusing on invertebrate AhRs are scarce and fragmentary. In this study, to our knowledge, we identified an AhR homolog from shrimp L. vannamei (LvAhR) and uncovered an AhR–caspase axis in an arthropod (shrimp) for the first time that plays a crucial role in inducing antiviral apoptosis against WSSV infection (Fig. 7). Our findings extend the knowledge of invertebrate AhR signaling that can connect to apoptosis signaling by directly targeting the regulation of caspase.

FIGURE 7.

Model for LvAhR/LvARNT-LvCaspase-1 axis–mediated anti-WSSV mechanism. In shrimp, LvAhR was activated by WSSV invasion, and then LvAhR and LvARNT formed a heterodimer in the nucleus that aroused caspase expression via binding to the XRE motif on caspase promoter, such as LvCaspase-1. The high expression of caspases promoted the activation of caspase-mediated apoptosis signaling that restricted WSSV replication.

FIGURE 7.

Model for LvAhR/LvARNT-LvCaspase-1 axis–mediated anti-WSSV mechanism. In shrimp, LvAhR was activated by WSSV invasion, and then LvAhR and LvARNT formed a heterodimer in the nucleus that aroused caspase expression via binding to the XRE motif on caspase promoter, such as LvCaspase-1. The high expression of caspases promoted the activation of caspase-mediated apoptosis signaling that restricted WSSV replication.

Close modal

AhR was identified originally for its role in toxicology, but in recent decades, it has been increasingly recognized as an important modulator for immune and inflammatory responses. A large number of studies have shown that AhRs from vertebrates exert antiviral effects via multiple mechanisms, such as modulating intracellular lipid state (58), reducing cellular dNTP levels (59), and changing the balance between effector and regulatory T cells (60). Although the antiviral ability of vertebrate AhR has been gradually revealed, the function of invertebrate AhR against viruses is still unclear. Our data showed that the expression of LvAhR was substantially induced in response to WSSV infection, and silencing of LvAhR resulted in elevated viral loads and rendered shrimp more susceptible to virus infection, which strongly suggest that LvAhR is an essential component of the shrimp immune defense system. Subsequently, we observed that apoptosis was reduced in shrimp with LvAhR knockdown at the early stage of WSSV infection. The elimination of infected cells via programmed cell death (apoptosis) is one of the most ancestral defense mechanisms against viral infection (61). Thus, we reason that LvAhR exerts antiviral effects via inducing apoptosis. Many studies have shown that AhR signaling could alter apoptosis progress, whereas the effects are commonly indirect. For example, in classical AhR signaling, the activated AhR regulated the transcription of CYP1A1 and CYP1A2, which oxidize toxic and carcinogenic chemicals that can cause oxidative stress to induce apoptosis (14, 62). The AhR was activated after influenza virus infection and induced the oxidation of DNA, protein, and lipid components within cells to produce reactive oxygen species at a higher level, thus leading to apoptosis (63, 64).

Interestingly, we found that the expression level of LvCaspase-1 was decreased in LvAhR-knockdown shrimp, which indicates a potential link between AhR signaling and caspase-mediated apoptosis. AhR functions as a transcription factor via forming heterodimers with another bHLH protein, ARNT, that binds to XREs in target gene promoters. We found that a putative XRE motif exists in the promoter region of LvCaspase-1, an ortholog of caspase-3 in mammals. The complete N-terminal region of LvAhR—consisting of the bHLH domain, PAS A domain, and PAS B domain—is required for both protein dimerization and DNA binding (promoter activity), which is consistent with AhR homologs in other reports (65). Although four other caspases (LvCaspase-2–LvCaspase5) are not confirmed to be directly regulated by the LvAhR/LvARNT heterodimer, the expressions of LvCaspase-2–LvCaspase5 are markedly restrained in the LvAhR silencing shrimp. Additionally, XRE motifs and others are also found in the promoter region of LvCaspase-2, LvCaspase-3, and LvCaspase-5 (data not shown), and AhR is reported to regulate the expression of genes that do not bear XREs in their regulatory regions (1). We infer that these caspases could be regulated by LvAhR directly or cooperate with other transcription factors, which needs to be further confirmed. We also demonstrated that LvCaspase-1 can induce antiviral apoptosis that markedly inhibits virus replication, despite the protective effect not having a significant impact on survival. Considering that multiple caspase proteins could be targeted by LvAhR, a cooperative effect was therefore constructed to efficiently activate an apoptotic cascade response.

Collectively, we identified an AhR homolog from the arthropod (shrimp) L. vannamei and explored its function during WSSV infection. The results demonstrated that LvAhR restricted WSSV replication by promoting antiviral apoptosis through the AhR–caspase-1 axis (Fig. 7). However, the method by which LvAhR is activated by WSSV infection has not yet been determined. Nevertheless, our current data may provide clues for therapeutic options for WSSV, such as the exploration of ligands that can activate AhR signaling.

This work was supported by National Natural Science Foundation of China Grant 32022085/31930113, National Key Research and Development Program of China Grant 2018YFD0900600/2018YFD0900500, Independent Research and Development Projects of Maoming Laboratory Grant 2021ZZ007/2021TDQD004, Southern Marine Science and Engineering Guangdong Laboratory (Zhuhai) Grant SML2021SP301, Key-Area Research and Development Program of Guangdong Province Grant 2018B020204001, and by Fundamental Research Funds for the Central Universities, Sun Yat-sen University Grant 22lglj05. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

The online version of this article contains supplemental material.

C.L., J.H., and Q.F. conceived and designed the experiments; Q.F., H.L., S.W., S.C., X.L., B.X., and C.L. performed the experiments and analyzed data; Q.F., J.H., and C.L. wrote the draft manuscript; C.L. and J.H. acquired funding; X.J. and R.C. provided experimental animals (shrimp); and C.L. was responsible for forming the hypothesis, project development, data coordination, and writing, finalizing, and submitting the manuscript. All authors discussed the results and approved the final version of the manuscript.

The sequences presented in this article have been submitted to GenBank under accession number OM046573.

Abbreviations used in this article:

     
  • AhR

    aryl hydrocarbon receptor

  •  
  • ARNT

    AhR nuclear translocator

  •  
  • bHLH

    basic helix-loop-helix

  •  
  • DmDcp-1

    D. melanogaster Dcp-1

  •  
  • DmGrim

    D. melanogaster Grim

  •  
  • DmReaper

    D. melanogaster Reaper

  •  
  • FSC

    forward scatter

  •  
  • HA

    hemagglutinin

  •  
  • LvAhR

    L. vannamei AhR

  •  
  • LvARNT

    L. vannamei ARNT

  •  
  • LvCaspase

    L. vannamei caspase

  •  
  • ORF

    open reading frame

  •  
  • PAS

    Per–Arnt–Sim

  •  
  • PI

    propidium iodide

  •  
  • qRT-PCR

    quantitative RT-PCR

  •  
  • RACE

    rapid amplification of cDNA ends

  •  
  • RNAi

    RNA interference

  •  
  • SSC

    side scatter

  •  
  • TAD

    transcriptional activation domain

  •  
  • WSS

    white spot syndrome

  •  
  • WSSV

    WSS virus

  •  
  • XRE

    xenobiotic response element

1.
Rothhammer
V.
,
F. J.
Quintana
.
2019
.
The aryl hydrocarbon receptor: an environmental sensor integrating immune responses in health and disease.
Nat. Rev. Immunol.
19
:
184
197
.
2.
McIntosh
B. E.
,
J. B.
Hogenesch
,
C. A.
Bradfield
.
2010
.
Mammalian Per-Arnt-Sim proteins in environmental adaptation.
Annu. Rev. Physiol.
72
:
625
645
.
3.
Cella
M.
,
M.
Colonna
.
2015
.
Aryl hydrocarbon receptor: linking environment to immunity.
Semin. Immunol.
27
:
310
314
.
4.
Ohtake
F.
,
K.
Takeyama
,
T.
Matsumoto
,
H.
Kitagawa
,
Y.
Yamamoto
,
K.
Nohara
,
C.
Tohyama
,
A.
Krust
,
J.
Mimura
,
P.
Chambon
, et al
2003
.
Modulation of oestrogen receptor signalling by association with the activated dioxin receptor.
Nature
423
:
545
550
.
5.
McBerry
C.
,
R. M. S.
Gonzalez
,
N.
Shryock
,
A.
Dias
,
J.
Aliberti
.
2012
.
SOCS2-induced proteasome-dependent TRAF6 degradation: a common anti-inflammatory pathway for control of innate immune responses.
PLoS One
7
:
e38384
.
6.
Yeste
A.
,
M. C.
Takenaka
,
I. D.
Mascanfroni
,
M.
Nadeau
,
J. E.
Kenison
,
B.
Patel
,
A.-M.
Tukpah
,
J. A. B.
Babon
,
M.
DeNicola
,
S. C.
Kent
, et al
2016
.
Tolerogenic nanoparticles inhibit T cell-mediated autoimmunity through SOCS2.
Sci. Signal.
9
:
ra61
.
7.
Wilson
S. R.
,
A. D.
Joshi
,
C. J.
Elferink
.
2013
.
The tumor suppressor Kruppel-like factor 6 is a novel aryl hydrocarbon receptor DNA binding partner.
J. Pharmacol. Exp. Ther.
345
:
419
429
.
8.
Jaronen
M.
,
F. J.
Quintana
.
2014
.
Immunological relevance of the coevolution of IDO1 and AHR.
Front. Immunol.
5
:
521
.
9.
Moura-Alves
P.
,
K.
Faé
,
E.
Houthuys
,
A.
Dorhoi
,
A.
Kreuchwig
,
J.
Furkert
,
N.
Barison
,
A.
Diehl
,
A.
Munder
,
P.
Constant
, et al
2014
.
AhR sensing of bacterial pigments regulates antibacterial defence.
Nature
512
:
387
392
.
10.
Gao
J.
,
K.
Xu
,
H.
Liu
,
G.
Liu
,
M.
Bai
,
C.
Peng
,
T.
Li
,
Y.
Yin
.
2018
.
Impact of the gut microbiota on intestinal immunity mediated by tryptophan metabolism.
Front. Cell. Infect. Microbiol.
8
:
13
.
11.
Feng
S.
,
Z.
Cao
,
X.
Wang
.
2013
.
Role of aryl hydrocarbon receptor in cancer.
Biochim. Biophys. Acta
1836
:
197
210
.
12.
Dong
W.
,
H.
Teraoka
,
S.
Kondo
,
T.
Hiraga
.
2001
.
2,3,7,8-Tetrachlorodibenzo-p-dioxin induces apoptosis in the dorsal midbrain of zebrafish embryos by activation of arylhydrocarbon receptor.
Neurosci. Lett.
303
:
169
172
.
13.
Wójtowicz
A. K.
,
K. A.
Szychowski
,
A.
Wnuk
,
M.
Kajta
.
2017
.
Dibutyl phthalate (DBP)-induced apoptosis and neurotoxicity are mediated via the aryl hydrocarbon receptor (AhR) but not by estrogen receptor alpha (ERα), estrogen receptor beta (ERβ), or peroxisome proliferator-activated receptor gamma (PPARγ) in mouse cortical neurons.
Neurotox. Res.
31
:
77
89
.
14.
Du
F.
,
T.
Zhao
,
H. C.
Ji
,
Y. B.
Luo
,
F.
Wang
,
G. H.
Mao
,
W. W.
Feng
,
Y.
Chen
,
X. Y.
Wu
,
L. Q.
Yang
.
2019
.
Dioxin-like (DL-) polychlorinated biphenyls induced immunotoxicity through apoptosis in mice splenocytes via the AhR mediated mitochondria dependent signaling pathways.
Food Chem. Toxicol.
134
:
110803
.
15.
Turski
W. A.
,
A.
Wnorowski
,
G. N.
Turski
,
C. A.
Turski
,
L.
Turski
.
2020
.
AhR and IDO1 in pathogenesis of Covid-19 and the “systemic AhR activation syndrome:” a translational review and therapeutic perspectives.
Restor. Neurol. Neurosci.
38
:
343
354
.
16.
Chattopadhyay
S.
,
G. C.
Sen
.
2017
.
RIG-I-like receptor-induced IRF3 mediated pathway of apoptosis (RIPA): a new antiviral pathway.
Protein Cell
8
:
165
168
.
17.
Chattopadhyay
S.
,
M.
Yamashita
,
Y.
Zhang
,
G. C.
Sen
.
2011
.
The IRF-3/Bax-mediated apoptotic pathway, activated by viral cytoplasmic RNA and DNA, inhibits virus replication.
J. Virol.
85
:
3708
3716
.
18.
Riedl
S. J.
,
Y.
Shi
.
2004
.
Molecular mechanisms of caspase regulation during apoptosis.
Nat. Rev. Mol. Cell Biol.
5
:
897
907
.
19.
Verbrugge
I.
,
R. W.
Johnstone
,
M. J.
Smyth
.
2010
.
SnapShot: extrinsic apoptosis pathways.
Cell
143
:
1192.e1–2
.
20.
Zhao
H.
2012
.
Extrinsic and intrinsic apoptosis signal pathway review.
In
Apoptosis and Medicine.
T.
Ntuli
.
IntechOpen
,
London
, p.
3
21
.
21.
Salvamoser
R.
,
K.
Brinkmann
,
L. A.
O’Reilly
,
L.
Whitehead
,
A.
Strasser
,
M. J.
Herold
.
2019
.
Characterisation of mice lacking the inflammatory caspases-1/11/12 reveals no contribution of caspase-12 to cell death and sepsis.
Cell Death Differ.
26
:
1124
1137
.
22.
Zhirnov
O. P.
,
T. E.
Konakova
,
W.
Garten
,
H.
Klenk
.
1999
.
Caspase-dependent N-terminal cleavage of influenza virus nucleocapsid protein in infected cells.
J. Virol.
73
:
10158
10163
.
23.
Volkmann
X.
,
M.
Cornberg
,
H.
Wedemeyer
,
F.
Lehner
,
M. P.
Manns
,
K.
Schulze-Osthoff
,
H.
Bantel
.
2006
.
Caspase activation is required for antiviral treatment response in chronic hepatitis C virus infection.
Hepatology
43
:
1311
1316
.
24.
Guillermo Bardera
G.
,
N.
Usman
,
M.
Owen
,
D.
Pountney
,
K. A.
Sloman
,
M. E.
Alexander
.
2018
.
The importance of behaviour in improving the production of shrimp in aquaculture.
Rev. Aquacult.
1
:
29
.
25.
Naylor
R. L.
,
R. W.
Hardy
,
A. H.
Buschmann
,
S. R.
Bush
,
L.
Cao
,
D. H.
Klinger
,
D. C.
Little
,
J.
Lubchenco
,
S. E.
Shumway
,
M.
Troell
.
2021
.
A 20-year retrospective review of global aquaculture. [Published errata appear in 2021 Nature 593: E12 and E36.]
Nature
591
:
551
563
.
26.
Li
C.
,
S.
Weng
,
J.
He
.
2019
.
WSSV-host interaction: host response and immune evasion.
Fish Shellfish Immunol.
84
:
558
571
.
27.
Arulmoorthy
M. P.
,
E.
Anandajothi
,
S.
Vasudevan
,
E.
Suresh
.
2020
.
Major viral diseases in culturable penaeid shrimps: a review.
Aquacult. Int.
28
:
1939
1967
.
28.
Durand
S.
,
D. V.
Lightner
,
R. M.
Redman
,
J. R.
Bonami
.
1997
.
Ultrastructure and morphogenesis of white spot syndrome baculovirus (WSSV).
Dis. Aquat. Organ.
29
:
205
211
.
29.
Li
C.-Y.
,
Y.-J.
Wang
,
S.-W.
Huang
,
C.-S.
Cheng
,
H.-C.
Wang
.
2016
.
Replication of the shrimp virus WSSV depends on glutamate-driven anaplerosis.
PLoS One
11
:
e0146902
.
30.
Leu
J. H.
,
F.
Yang
,
X.
Zhang
,
X.
Xu
,
G. H.
Kou
,
C. F.
Lo
.
2009
.
Whispovirus.
In
Lesser known large dsDNA viruses.
W. H.
Wilson
,
J. L.
Van Etten
,
M. J.
Allen
.
Springer-Verlag
,
Berlin, Heidelberg
, p.
197
227
.
31.
Sánchez-Paz
A.
2010
.
White spot syndrome virus: an overview on an emergent concern.
Vet. Res.
41
:
43
.
32.
Sahul Hameed
A.
,
M.
Sarathi
,
R.
Sudhakaran
,
G.
Balasubramanian
,
S. S.
Musthaq
.
2006
.
Quantitative assessment of apoptotic hemocytes in white spot syndrome virus (WSSV)-infected penaeid shrimp, Penaeus monodon and Penaeus indicus, by flow cytometric analysis.
Aquaculture
256
:
111
120
.
33.
Hu
W. Y.
,
C. L.
Yao
.
2016
.
Molecular and immune response characterizations of a novel AIF and cytochrome c in Litopenaeus vannamei defending against WSSV infection.
Fish Shellfish Immunol.
56
:
84
95
.
34.
Rijiravanich
A.
,
C. L.
Browdy
,
B.
Withyachumnarnkul
.
2008
.
Knocking down caspase-3 by RNAi reduces mortality in Pacific white shrimp Penaeus (Litopenaeus) vannamei challenged with a low dose of white-spot syndrome virus.
Fish Shellfish Immunol.
24
:
308
313
.
35.
Li
C.
,
S.
Weng
,
Y.
Chen
,
X.
Yu
,
L.
,
H.
Zhang
,
J.
He
,
X.
Xu
.
2012
.
Analysis of Litopenaeus vannamei transcriptome using the next-generation DNA sequencing technique.
PLoS One
7
:
e47442
.
36.
Li
C.
,
Y.
Chen
,
S.
Weng
,
S.
Li
,
H.
Zuo
,
X.
Yu
,
H.
Li
,
J.
He
,
X.
Xu
.
2014
.
Presence of Tube isoforms in Litopenaeus vannamei suggests various regulatory patterns of signal transduction in invertebrate NF-κB pathway.
Dev. Comp. Immunol.
42
:
174
185
.
37.
Letunic
I.
,
T.
Doerks
,
P.
Bork
.
2015
.
SMART: recent updates, new developments and status in 2015.
Nucleic Acids Res.
43
(
D1
):
D257
D260
.
38.
Larkin
M. A.
,
G.
Blackshields
,
N. P.
Brown
,
R.
Chenna
,
P. A.
McGettigan
,
H.
McWilliam
,
F.
Valentin
,
I. M.
Wallace
,
A.
Wilm
,
R.
Lopez
, et al
2007
.
Clustal W and Clustal X version 2.0.
Bioinformatics
23
:
2947
2948
.
39.
Nicholas
K.
,
H.
Nicholas
.
1997
.
GeneDoc: a tool for editing and annotating multiple sequence alignments.
Available at: http://www.nrbsc.org/gfx/genedoc/. Accessed: December 26, 2020
.
40.
Tamura
K.
,
D.
Peterson
,
N.
Peterson
,
G.
Stecher
,
M.
Nei
,
S.
Kumar
.
2011
.
MEGA5: molecular evolutionary genetics analysis using maximum likelihood, evolutionary distance, and maximum parsimony methods.
Mol. Biol. Evol.
28
:
2731
2739
.
41.
Zhang
X.
,
J.
Yuan
,
Y.
Sun
,
S.
Li
,
Y.
Gao
,
Y.
Yu
,
C.
Liu
,
Q.
Wang
,
X.
Lv
,
X.
Zhang
, et al
2019
.
Penaeid shrimp genome provides insights into benthic adaptation and frequent molting.
Nat. Commun.
10
:
356
.
42.
Qiu
W.
,
S.
Zhang
,
Y. G.
Chen
,
P. H.
Wang
,
X. P.
Xu
,
C. Z.
Li
,
Y. H.
Chen
,
W. Z.
Fan
,
H.
Yan
,
S. P.
Weng
, et al
2014
.
Litopenaeus vannamei NF-κB is required for WSSV replication.
Dev. Comp. Immunol.
45
:
156
162
.
43.
Tassanakajon
A.
,
P.
Amparyup
,
K.
Somboonwiwat
,
P.
Supungul
.
2011
.
Cationic antimicrobial peptides in penaeid shrimp.
Mar. Biotechnol. (NY)
13
:
639
657
.
44.
Lin
F.
,
H.
Huang
,
L.
Xu
,
F.
Li
,
F.
Yang
.
2011
.
Identification of three immediate-early genes of white spot syndrome virus.
Arch. Virol.
156
:
1611
1614
.
45.
Li
F.
,
M.
Li
,
W.
Ke
,
Y.
Ji
,
X.
Bian
,
X.
Yan
.
2009
.
Identification of the immediate-early genes of white spot syndrome virus.
Virology
385
:
267
274
.
46.
Marks
H.
,
M.
Mennens
,
J. M.
Vlak
,
M. C. W.
van Hulten
.
2003
.
Transcriptional analysis of the white spot syndrome virus major virion protein genes.
J. Gen. Virol.
84
:
1517
1523
.
47.
Nainu
F.
,
A.
Shiratsuchi
,
Y.
Nakanishi
.
2017
.
Induction of apoptosis and subsequent phagocytosis of virus-infected cells as an antiviral mechanism.
Front. Immunol.
8
:
1220
.
48.
Du
Z. J.
,
G. Q.
Cui
,
J.
Zhang
,
X. M.
Liu
,
Z. H.
Zhang
,
Q.
Jia
,
J. C.
Ng
,
C.
Peng
,
C. X.
Bo
,
H.
Shao
.
2017
.
Inhibition of gap junction intercellular communication is involved in silica nanoparticles-induced H9c2 cardiomyocytes apoptosis via the mitochondrial pathway.
Int. J. Nanomedicine
12
:
2179
2188
.
49.
Guo
S.
,
Z.
Wen
,
X.
Zhang
,
F.
Li
,
X.
You
.
2021
.
Characterization of five caspase genes and their transcriptional changes in response to exogenous iridescent virus challenge in the whiteleg shrimp (Litopenaeus vannamei).
Aquaculture
534
:
736192
.
50.
Wang
P. H.
,
D. H.
Wan
,
Y. G.
Chen
,
S. P.
Weng
,
X. Q.
Yu
,
J. G.
He
.
2013
.
Characterization of four novel caspases from Litopenaeus vannamei (Lvcaspase2–5) and their role in WSSV infection through dsRNA-mediated gene silencing.
PLoS One
8
:
e80418
.
51.
Phongdara
A.
,
W.
Wanna
,
W.
Chotigeat
.
2006
.
Molecular cloning and expression of caspase from white shrimp Penaeus merguiensis.
Aquaculture
252
:
114
120
.
52.
Larigot
L.
,
L.
Juricek
,
J.
Dairou
,
X.
Coumoul
.
2018
.
AhR signaling pathways and regulatory functions.
Biochim. Open
7
:
1
9
.
53.
Hernández-Palomares
M. L. E.
,
J. A.
Godoy-Lugo
,
S.
Gómez-Jiménez
,
L. A.
Gámez-Alejo
,
R. M.
Ortiz
,
J. F.
Muñoz-Valle
,
A. B.
Peregrino-Uriarte
,
G.
Yepiz-Plascencia
,
J. A.
Rosas-Rodríguez
,
J. G.
Soñanez-Organis
.
2018
.
Regulation of lactate dehydrogenase in response to WSSV infection in the shrimp Litopenaeus vannamei.
Fish Shellfish Immunol.
74
:
401
409
.
54.
Soñanez-Organis
J. G.
,
A. B.
Peregrino-Uriarte
,
S.
Gómez-Jiménez
,
A.
López-Zavala
,
H. J.
Forman
,
G.
Yepiz-Plascencia
.
2009
.
Molecular characterization of hypoxia inducible factor-1 (HIF-1) from the white shrimp Litopenaeus vannamei and tissue-specific expression under hypoxia.
Comp. Biochem. Physiol. C Toxicol. Pharmacol.
150
:
395
405
.
55.
Hong
W.
,
W.
Cheng
,
T.
Zheng
,
N.
Jiang
,
R.
Xu
.
2020
.
AHR is a tunable knob that controls HTLV-1 latency-reactivation switching.
PLoS Pathog.
16
:
e1008664
.
56.
Pernomian
L.
,
M.
Duarte-Silva
,
C. R.
de Barros Cardoso
.
2020
.
The aryl hydrocarbon receptor (AHR) as a potential target for the control of intestinal inflammation: insights from an immune and bacteria sensor receptor.
Clin. Rev. Allergy Immunol.
59
:
382
390
.
57.
Cheong
J. E.
,
L.
Sun
.
2018
.
Targeting the IDO1/TDO2–KYN–AhR pathway for cancer immunotherapy—challenges and opportunities.
Trends Pharmacol. Sci.
39
:
307
325
.
58.
Fusco
D. N.
,
H.
Pratt
,
S.
Kandilas
,
S. S.
Cheon
,
W.
Lin
,
D. A.
Cronkite
,
M.
Basavappa
,
K. L.
Jeffrey
,
A.
Anselmo
,
R.
Sadreyev
, et al
2017
.
HELZ2 is an IFN effector mediating suppression of dengue virus.
Front. Microbiol.
8
:
240
.
59.
Kueck
T.
,
E.
Cassella
,
J.
Holler
,
B.
Kim
,
P. D.
Bieniasz
.
2018
.
The aryl hydrocarbon receptor and interferon gamma generate antiviral states via transcriptional repression.
eLife
7
:
e38867
.
60.
Veiga-Parga
T.
,
A.
Suryawanshi
,
B. T.
Rouse
.
2011
.
Controlling viral immuno-inflammatory lesions by modulating aryl hydrocarbon receptor signaling.
PLoS Pathog.
7
:
e1002427
.
61.
Galluzzi
L.
,
C.
Brenner
,
E.
Morselli
,
Z.
Touat
,
G.
Kroemer
.
2008
.
Viral control of mitochondrial apoptosis.
PLoS Pathog.
4
:
e1000018
.
62.
Nebert
D. W.
,
A. L.
Roe
,
M. Z.
Dieter
,
W. A.
Solis
,
Y.
Yang
,
T. P.
Dalton
.
2000
.
Role of the aromatic hydrocarbon receptor and [Ah] gene battery in the oxidative stress response, cell cycle control, and apoptosis.
Biochem. Pharmacol.
59
:
65
85
.
63.
Wheeler
J. L. H.
,
K. C.
Martin
,
E.
Resseguie
,
B. P.
Lawrence
.
2014
.
Differential consequences of two distinct AhR ligands on innate and adaptive immune responses to influenza A virus.
Toxicol. Sci.
137
:
324
334
.
64.
Chen
K.-K.
,
M.
Minakuchi
,
K.
Wuputra
,
C.-C.
Ku
,
J.-B.
Pan
,
K.-K.
Kuo
,
Y.-C.
Lin
,
S.
Saito
,
C.-S.
Lin
,
K. K.
Yokoyama
.
2020
.
Redox control in the pathophysiology of influenza virus infection.
BMC Microbiol.
20
:
214
.
65.
Abel
J.
,
T.
Haarmann-Stemmann
.
2010
.
An introduction to the molecular basics of aryl hydrocarbon receptor biology.
Biol. Chem.
391
:
1235
1248
.

The authors have no financial conflicts of interest.

Supplementary data