Juvenile hormone-induced histone deacetylase 3 suppresses apoptosis to maintain larval midgut in the yellow fever mosquito.

SignificanceJuvenile hormone (JH), a sesquiterpenoid, regulates many aspects of insect development, including maintenance of the larval stage by preventing metamorphosis. In contrast, ecdysteroids promote metamorphosis by inducing the E93 transcription factor, which triggers apoptosis of larval cells and remodeling of the larval midgut. We discovered that JH suppresses precocious larval midgut-remodeling by inducing an epigenetic modifier, histone deacetylase 3 (HDAC3). JH-induced HDAC3 deacetylates the histone H4 localized at the promoters of proapoptotic genes, resulting in the suppression of these genes. This eventually prevents programmed cell death of midgut cells and midgut-remodeling during larval stages. These studies identified a previously unknown mechanism of JH action in blocking premature remodeling of the midgut during larval feeding stages.

Arthropod-borne arboviruses are a major threat to human health. The yellow fever mosquito, Aedes aegypti, is a major vector of human arboviruses that cause dengue, Zika, yellow fever, chikungunya, and other diseases. A widespread epidemic of zika across Central and South America and the Caribbean has been linked to fetal brain abnormalities (1). Dengue is the most-common human arboviral disease infecting millions of people each year. Over the last decade, chikungunya emerged as one of the major epidemics in Asia, Europe, and America. Adult female mosquitoes transmit the above arboviruses during blood-feeding on hosts. Therefore, blocking the transition from the immature to an adult stage could be a useful strategy to prevent the spread of mosquito-borne diseases.

Epigenetic modifications regulate genome expression by turning on or off a network of genes resulting in different phenotypes from a common genotype (2). Aberrations in epigenetic regulation can affect growth and development and cause diseases such as cancer. The main classes of epigenetic modifiers are chromatin remodelers (affect DNA–histone interactions) and histone modifiers (affect histone posttranslational modifications) that alter chromatin organization. Acetylation is one of the major histone modifications carried by histone acetyltransferases (HATs) that neutralize the positive charge of histones, thereby relaxing chromatin structure. This provides target gene promoter access to the transcription factors and RNA polymerase complexes to activate gene expression. In contrast, histone deacetylases (HDACs) remove acetyl groups from the lysine residues in the histone tails, resulting in chromatin compaction and repression of the target gene expression. Histone acetylation regulates many developmental processes in insects. Histone acetylation by Creb-binding protein (CBP) initiates dendrite pruning in Drosophila melanogaster (3). The inhibition of HDAC activity alters caste specification in honeybees (4). HDACs also regulate mandibles and wing sizes in the broad-horned flour beetle, Gnatocerus cornutus (5). Equilibrium between CBP-mediated histone acetylation and HDAC-mediated histone deacetylation in the brain regulates caste-specific foraging and scouting behaviors in the ant, Camponotus floridanus (6). HATs and HDACs antagonistically regulate nuclear receptor expression and activity during insect metamorphosis (7, 8).

Previously, we showed that CBP acetylates histones and regulates the transcription of genes involved in JH and ecdysone signaling in A. aegypti and Tribolium castaneum (9–11). In contrast, the inhibition of HDAC activity using Trichostatin A inhibitor or knockdown of genes coding for HDACs affected the metamorphosis of T. castaneum (11–14). Specifically, HDAC1 knockdown increased expression of JH response genes, which in turn affected the larval–pupal metamorphosis of T. castaneum (13). HDAC3 knockdown affected the wing development in T. castaneum (14). In G. cornutus, knockdown of HDAC1 caused mandible size reduction, whereas HDAC3 knockdown induced mandible hypertrophy (5). In the brown planthopper, Nilaparvata lugens, knockdown of HDAC1 affected the female and male fertility and ovary development (15). Overexpression of HDAC4 impaired long-term courtship memory in D. melanogaster (16). These studies suggest that each HDAC may perform distinct functions within insect species. However, the role of HDACs in A. aegypti growth, development, and metamorphosis are not known. Hence, we investigated the function of HDACs in A. aegypti by knocking them down using RNA interference (RNAi). We found that the knockdown of each HDAC has a distinct effect on the growth, development, and metamorphosis of A. aegypti. The knockdown of HDAC3 had a severe impact on larval survival, and it is required for maintaining the larval stage in A. aegypti.

Results

HDACs Play Distinct Roles in Larval and Pupal Development in A. aegypti.

To identify the genes coding for HDACs in A. aegypti, HDAC protein-coding sequences from T. castaneum were used to search the A. aegypti genome database. A total 10 HDAC homologs belonging to four classes were identified in A. aegypti (). To determine functions of identified HDACs in A. aegypti, these genes were knocked down by feeding second instar larvae on food pellets containing nanoformulated double-stranded RNA (dsRNA targeting each HDAC gene. The control larvae were fed on nanoformulated dsRNA targeting the maltose-binding protein gene (malE) from Escherichia coli. Knockdown of class I HDACs (HDAC1 and HDAC3 except for HDAC8) affected the larval growth and survival. Specifically, the HDAC1 knockdown larvae exhibited growth retardation and had smaller bodies than those in control and they eventually died. Although some of the treated larvae developed into pupae, they could not shed their old head capsule and eventually died (Fig. 1). Knockdown of HDAC3 caused the highest larval mortality; the treated insects remained in the larval stage for an extended period and eventually died (Fig. 1 and ). Knockdown of HDAC8 affected pupal survival, but no effect on larvae was detected (). Knockdown of Class II HDACs (HDAC4 and HDAC6) had a moderate impact on larval and pupal survival. HDAC4 knockdown caused 40% larval mortality, and the remaining larvae pupated but experienced incomplete ecdysis of the head, similar to the phenotype detected in HDAC1 knockdown (). HDAC6 knockdown caused late larval and early pupal mortality. The knockdown of Class III HDACs (Sirtuins 2, 4, 6, and 7) had a major impact on pupal growth and survival. Notably, Sirtuin-2 knockdown increased melanization of pupal cuticle before their death. Knockdown of Sirtuin-4 affected the compound eye development, as the pupae which developed from dsSirtuin-4-treated larvae showed no compound eyes (Fig. 1). Sirtuin-6 and -7 knockdown had low to moderate effects on pupal survival. The knockdown of Class IV HDAC, HDAC11, caused moderate larval and high pupal mortality (). To confirm that the observed phenotype changes are due to the knockdown of target HDAC genes, we extracted the total RNA from the treated larvae on the fifth day after dsRNA feeding and determined their messenger RNA (mRNA) levels. We observed more than 55% knockdown of the target HDAC genes in respective dsRNA treatments compared to their levels in control larvae fed on a diet containing nanoformulated dsmalE (). We also checked whether the dsRNAs used in these experiments have any unintended off-target effects on the other HDAC genes. In this regard, the dsRNA target sequence was aligned with the sequences of the other HDAC genes following the method described recently (17). This alignment did not find any sequences with a continuous stretch of identity between the dsRNA target sequence with the other HDAC gene sequences. We also reconfirmed no off-target effects of the selected HDACs dsRNAs by determining relative mRNA levels of other HDAC genes in HDAC3 knockdown larvae (). Similarly, no significant changes in the mRNA levels of HDAC3 gene were detected in other HDACs dsRNAs treated larval samples (). Together, these data suggest that each HDAC has a distinct role in the growth and development of A. aegypti; Class I and II HDACs are required for larval growth and development, while Class III and IV HDACs function in pupal and adult development. Among the HDACs tested, HDAC3 knockdown caused 100% mortality and severe phenotypes; thus, we focused our further studies on HDAC3.

Fig. 1.

Knockdown of HDACs affects growth, development, and metamorphosis of A. aegypti larvae and pupae. Phenotypes of larvae or pupae fed on food pellets containing nanoformulated dsRNA targeting HDAC genes of A. aegypti. (A) Phenotype of control larvae fed on a dsmalE-nanoformulated diet. (B) Knockdown of HDAC1 resulted in larval growth retardation, a decrease in body size and death. (C) Some of the dsHDAC1-treated larvae pupated, but they were unable to completely shed their old integument and died. (D) Knockdown of HDAC3 caused the highest larval mortality. These larvae stayed in the larval stage for a long period and eventually died. (E) Knockdown of HDAC4 prevented shedding of old integument after metamorphosis into the pupal stage. (F and G) HDAC6 knockdown caused late (fourth instar) larval and early pupal mortality. (H) HDAC8 knockdown caused pupal mortality. (I) Sirtuin-2 knockdown manifested an increased melanization of pupae before they died. (J) Sirtuin-4 knockdown inhibited the compound eye development in the pupae. (K) Sirtuin-6 knockdown caused pupal mortality but did not affect larval survival. (L) Sirtuin-7 knockdown caused pupal mortality. (M) The control larvae fed on a dsmalE-nanoformulated diet developed as normal pupae. Scale bar is 1 millimeter in all images. (N) Knockdown of HDACs induced larval and pupal mortality in A. aegypti. n = 90 (30 × 3 independent experiments). Pupation, adult eclosion rates, and corrected mortality data are included in .

JH Induces the HDAC3 Expression during the Larval Stage of A. aegypti.

To understand why HDAC3 knockdown insects died during the larval stage, HDAC3 mRNA levels were quantified during larval, pupal, and adult stages of A. aegypti. We found that HDAC3 mRNA levels were higher during larval stages than in other developmental stages. Notably, high HDAC3 mRNA levels were recorded in the fourth instar larvae up to 30 h after ecdysis to this stage, which then gradually decreased by the end of the larval period (). Moreover, the HDAC3 mRNA levels showed a positive correlation (R-value: +0.76) with the JH titers during the A. aegypti larval stage (18–20), suggesting that JH may induce HDAC3 expression. To test this hypothesis, we exposed Aag-2 cells to 10 µM of Juvenile hormone III (JH III) for 3 h and quantified the HDAC3 mRNA levels. Aag-2 cells exposed to JH III showed an increase in HDAC3 mRNA levels compared to its levels in Aag-2 cells exposed to the control solvent, dimethyl sulfoxide (DMSO) (Fig. 2). Also, newly molted fourth instar larvae grown in water containing 100 ng of JH analog, methoprene, per ml showed significantly higher mRNA levels of HDAC3 and Kr-h1 (which is a JH-induced gene—used as a positive control) at 36, 42, and 48 h after molting into fourth instar larvae compared to their expression levels in the control larvae exposed to DMSO (Fig. 2 ). The methoprene-treated larvae died during the pupal stage (). We also knocked down genes coding for JH acid methyltransferase (JHAMT), which is involved in the biosynthesis of JH, and assessed its impact on the expression of HDAC3. A significant reduction in the HDAC3 mRNA levels was detected in the JHAMT knockdown larvae (). To confirm that the JH levels were reduced in the JHAMT knockdown larvae, the expression of Kr-h1 (primary JH response gene) was determined. Kr-h1 is sensitive to subnanomolar levels of JH and its expression levels are directly correlated with the JH titers (21, 22). The Kr-h1 mRNA levels decreased in the JHAMT knockdown larvae compared to those in control larvae. These results together confirm that JH induces HDAC3 expression.

Fig. 2.

JH induces HDAC3 expression through its receptor, Met. (A) HDAC3 expression levels were elevated in Aag-2 cells exposed to 10 µM of JH III compared to its expression levels in control cells exposed to DMSO. (B) The expression levels of HDAC3 in A. aegypti larvae treated with 100 ng/mL of JH analog, methoprene, or DMSO. (C) Kr-h1 (used as a positive control) expression levels in methoprene or DMSO-treated larvae. Newly molted fourth instar A. aegypti larvae were treated with 100 ng/mL methoprene or DMSO. The HDAC3 and Kr-h1 mRNA levels were determined at 36, 42, and 48 h after treatment. (D) Knockdown of Met in Aag-2 cells prevented JH induction of HDAC3 expression. (E) Overexpression of Met receptor in Aag-2 cells induced the HDAC3 expression. VC: pIEx-4 empty vector control; Met: pIEx-4 vector containing AaMet complete ORF. The target gene-expression levels were normalized using the reference gene, RPS7, expression levels. Mean + SE (n = 9) are shown. Ns, Not significant. * denotes the significant differences in the expression levels of target genes between treatment and control at P < 0.05 analyzed using one-way ANOVA.

To determine whether JH induces HDAC3 expression through its receptor, Met, the Met gene was knocked down in Aag-2 cells by exposing them to dsMet followed by treatment with JH III. The HDAC3 mRNA levels increased in Aag-2 exposed to dsmalE and JH III but not in cells exposed to dsMet and JH III (Fig. 2). Moreover, the lowest HDAC3 mRNA levels were recorded in Met knockdown and DMSO-exposed cells compared to its levels in control cells treated with dsmalE and DMSO. To reconfirm these results, we transfected Aag-2 cells with AaMet expression construct and exposed them to JH III or DMSO. An increase of HDAC3 mRNA levels was detected in cells transfected with Met expression construct and exposed to DMSO. Notably, the HDAC3 mRNA levels were further increased in Aag-2 cells transfected with Met expression construct and exposed to JH III (Fig. 2). These results together confirm that JH induces the HDAC3 expression through its receptor, Met.

HDAC3 Knockdown Induces the Expression of Genes Involved in A poptosis.

Identification of HDAC3 regulated genes and pathways could provide information on its functions and mechanisms involved in the death of HDAC3 knockdown larvae. Hence, we knocked down HDAC3 in Aag-2 cells and performed RNA-sequencing (RNA-seq) to identify its targets. The differential gene-expression analysis of RNA-seq data identified 103 up-regulated and 66 down-regulated genes in HDAC3 knockdown cells compared to their expression levels in control cells treated with dsmalE ( and Dataset S1). Gene enrichment analysis revealed that genes involved in apoptosis, postembryonic development, DNA-binding transcription factors, etc. are significantly enriched in the up-regulated gene group (), while Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis showed that genes involved in Hippo, MAPK, and Notch signaling pathways are significantly enriched in the HDAC3 knockdown samples ().

To verify the RNA-seq data, we selected 15 up-regulated genes (predicted to be involved in different developmental processes) and determined their expression levels in HDAC3 knockdown and control larvae using the qRT-PCR. Out of the 15 genes tested, 14 showed similar expression patterns, with the only exception being the gene-coding for mitogen-activated kinase kinase (Fig. 3). Notably, RNA-seq and qRT-PCR data identified one important regulator of apoptosis (ubiquitin regulator of apoptosis, URA), which is overexpressed in HDAC3 knockdown larvae and Aag-2 cells (Fig. 3). URA regulates the inhibitor of apoptosis proteins (IAP), which blocks the activation of caspases and prevents apoptosis (23). Hence, we hypothesized that HDAC3 knockdown might have altered the expression of genes involved in apoptosis and that might have caused larval death. To test this hypothesis, we determined the mRNA levels of apoptotic genes: URA, IAP, Anti-IAP protein-Hid, Michelob (reaper), and initiator and executor caspases in HDAC3 knockdown larvae. The IAP mRNA levels were decreased, while URA, Hid, and the executor caspase 3 mRNA levels were increased in HDAC3 knockdown larvae (Fig. 3). However, the expression of the Michelob and initiator caspases 8 and 9 were not affected by HDAC3 knockdown ().

Fig. 3.

HDAC3 suppresses apoptosis-triggering gene expression. (A) To identify HDAC3 targets and validate their expression, we selected 15 up-regulated genes from the HDAC3 knockdown RNA-seq data and determined their mRNA levels in the A. aegypti larvae fed on a dsHDAC3- or dsmalE-nanoformulated diet using the RT-qPCR. AAEL002390: Kruppel homolog 1, AAEL007329: Held out wings protein, AAEL014622: Zinc finger transcription factor, AAEL026297: Toll, AAEL000328: Zinc-binding oxidoreductase, AAEL008138: ATP-binding cassette G1, AAEL019982: Ras GTPase-activating protein, AAEL014749: Ras-related protein, AAEL012899: Meiotic nuclear division protein 1, AAEL008379: Mitogen-activated kinase 14B, AAEL001094: Heat shock protein beta-1, AAEL014439: Juvenile hormone-inducible protein, AAEL001126: REST corepressor, AAEL008306: Mitogen-activated kinase kinase 15 (MAP3K15), and AAEL020878: Ubiquitin regulator of apoptosis (URA). (B) Knockdown of HDAC3 in A. aegypti larvae induced the expression of genes involved in apoptosis. The mRNA levels of URA, Yorkie, Hid, IAP, and cell death enzyme, caspase 3, were determined in the A. aegypti larvae fed on dsHDAC3-nanoformulated diet and the control larvae fed on dsmalE-nanoformulated diet. (C) HDAC3 deacetylates histone H4—acetylation levels were determined by Western blot using the proteins from Aag-2 cells treated with dsHDAC3 or dsmalE (control). (a) β-actin (used as a loading control) levels in dsHDAC3- or dsmalE-treated cells. (b) Histone H2, H3, and H4 acetylation levels in dsHDAC3- or dsmalE-treated cells detected using the Acetylated-Lysine (Ac-K2-100) MultiMab Rabbit mAb mix. (c) Histone H4 acetylation levels were detected using the Ac-Histone H4 specific mAb antibody (E-5). (D) ChIP assay revealed that knockdown of HDAC3 enriched the histone H4 acetylation levels at the promoter regions of URA, Hid, and caspase 3. Aag-2 cells were treated with dsmalE or dsHDAC3 for 72 h. The chromatin was cross linked and enriched using the Ac-Histone H4 specific mAb antibody (E-5). Purified DNA was used in qPCR to quantify the enrichment levels of promoters of target genes and control genes, β-actin and HSP70. Normalization was performed using the Rabbit IgG antibody–enriched chromatin DNA (negative control). Data are shown as fold enrichment mean ± SE (n = 8). * denotes significant differences in the promoter enrichment levels between treatment and control analyzed using one-way ANOVA at P < 0.005. (E) Both HDAC3 and SMRTER regulate the expression of genes involved in apoptosis. The mRNA levels of URA, Yorkie, Hid, IAP, and caspase 3 were determined in the Aag-2 cells exposed individually to dsHDAC3, dsSMRTER, or both dsHDAC3 and dsSMRTER. The target gene-expression levels were normalized using the reference gene, RPS7, expression levels. Mean + SE (n = 9) are shown. * denotes the significant differences in the mRNA levels of target genes between treatment and control at P < 0.05 analyzed using one-way ANOVA. dsHD3, dsHDAC3; dsSMT, dsSMRTER; ACT, β-Actin; and YKI, Yorkie. HDAC3 and SMRTER expression levels in the cognate dsRNA treatments are provided in .

To verify the inverse relationship between the expression of HDAC3 and its target genes, the developmental expression profiles of HDAC3, URA, Hid, and caspase 3 were determined. A strong negative correlation was observed between HDAC3 and URA (correlation coefficient, R-value: −0.906) and HDAC3 and Hid expression levels (R-value: −0.874) during the larval stage (). Similarly, the expression of HDAC3 and caspase 3 were negatively correlated (R-value: −0.62059) during the larval stage. These results suggest that HDAC3 is required for suppression of URA, Hid, and caspase 3 and induction of IAP, thereby preventing apoptosis in actively feeding and growing Ae. aegypti larvae.

HDAC3 Deacetylates Histone H4 Localized near the Promoters of the Proapoptotic Genes.

HDACs suppress target gene expression by deacetylating histones near the promoters of target genes, preventing their access to the transcription machinery. To determine whether deacetylation of histones is the reason for the altered expression of genes involved in apoptosis, we knocked down HDAC3 in Aag-2 cells and performed Western blot hybridization using the antibody that detects total lysine acetylation levels. Histone H4 acetylation levels increased in HDAC3 knockdown cells compared to its acetylation levels in control cells treated with dsmalE (Fig. 3 and ). This result was reconfirmed using the Acetyl-Histone H4 specific antibodies that detect acetylated Ser-1, Lys-5, Lys-8, and Lys-12 amino acids in the N-terminal tail of histone H4 (Fig. 3). To determine whether the HDAC3 deacetylated histone H4 is associated with the promoter regions of genes involved in apoptosis, Aag-2 cells were treated with dsHDAC3 or dsmalE and performed chromatin immunoprecipitation (ChIP) assay using the Acetyl-Histone H4 specific antibody. ChIP assay revealed that key proapoptotic genes, URA, Hid, and Caspase 3 promoters were enriched in HDAC3 knockdown cells compared to their promoter enrichment levels in the control cells treated with dsmalE (Fig. 3). In contrast, IAP and control genes, β-actin, and HSP70 promoters were not enriched in HDAC3 knockdown cells. These results demonstrate that HDAC3 deacetylates histone H4 localized near the promoters of proapoptotic genes, URA, Hid, and caspase 3.

HDAC3 and SMRTER Together Suppress the Expression of Genes Involved in Apoptosis.

HDAC3 was found to interact with the silencing mediator of retinoid and thyroid hormone receptor (SMRT) to form a corepressor complex that represses the genes regulated by the thyroid hormone receptor (24). The D. melanogaster ecdysone receptor–interacting protein, SMRTER, is functionally similar to the vertebrate nuclear corepressors, SMRT and N-CoR (nuclear receptor corepressor) (25). To test whether SMRTER is required for HDAC3 to regulate the expression of genes involved in apoptosis, we knocked down SMRTER alone or along with HDAC3 in Aag-2 cells and quantified the expression of target proapoptotic genes. Interestingly, SMRTER knockdown had a similar effect as HDAC3 knockdown on the expression of the proapoptotic genes (Fig. 3 and ). The URA, Hid, and caspase 3 genes were up-regulated in cells exposed to dsSMRTER or both dsSMRTER and dsHDAC3 compared to their expression levels in control cells treated with dsmalE. Contrastingly, IAP mRNA levels were decreased in dsSMRTER or both dsSMRTER- and dsHDAC3-treated cells. Further, to verify whether SMRTER is indeed required for HDAC3 to mediate the suppression of apoptosis genes, we transfected Aag-2 cells with AaHDAC3 or AaSMRTER expression constructs individually or together and determined URA, IAP, Hid, and caspase 3 mRNA levels. The overexpression of HDAC3 alone did not alter URA, IAP, Hid, and caspase 3 mRNA levels (Fig. 4). However, simultaneous overexpression of HDAC3 and SMRTER in Aag-2 cells resulted in the suppression of URA, Hid, and caspase 3 genes (Fig. 4). These results suggest that HDAC3 and SMRTER may act cooperatively to suppress apoptosis.

Fig. 4.

HDAC3- and SMRTER-mediated suppression of apoptosis genes is prevented by 20E. (A) Overexpression of both AaHDAC3 and AaSMRTER in Aag-2 cells suppresses URA, Hid, and caspase 3 and induces IAP and Yorkie expression. The HDAC3- and SMRTER-mediated suppression of URA, Hid, and caspase 3 genes in Aag-2 cells is blocked by 20E. Error bars show the SE mean of the four biological replicates. Bars with the same letter are not significantly different at 95% CI. (B) Presence of 20E induces the expression of URA, Hid, and caspase 3 genes but does not affect the expression of HDAC3 and Yorkie in Aag-2 cells. JH III induced the expression of HDAC3 and Yorkie in Aag-2 cells. However, Aag-2 cells simultaneously exposed to both JH III and 20E hormones revealed that JH was unable to suppress the proapoptotic genes, URA, Hid, and caspase 3 expressions in the presence of ecdysone. (C) SEA induces the expression of URA, Hid, and caspase 3 genes but no effect on the expression of HDAC3 and Yorkie. JH analog, Methoprene, induces the expression of HDAC3 and Yorkie. Fourth instar larvae 12 h old were treated individually with 100 ng/mL of methoprene, SEA, or a mix of both hormone analogs. The mRNA levels of target genes were determined at 6 and 12 h after treatment. The target gene-expression levels were normalized using the reference gene, RPS7, expression levels. Mean + SE (n = 9) are shown. * denotes the significant differences in the mRNA levels of target genes between treatment and control at P < 0.05 analyzed using one-way ANOVA. JH III, Juvenile hormone III; 20E, 20-hydroxyecdysone; DMSO, Dimethyl sulfoxide; SEA, Stable ecdysone agonist (RH-102240); HD3, HDAC3 overexpression; and SMT, SMRTER overexpression.

Ecdysone Prevents the HDAC3-Mediated Suppression of URA, Hid, and Caspase 3 Genes.

During larval–pupal metamorphosis, JH levels decrease and ecdysone levels increase, which in turn triggers apoptosis of midgut cells (20, 26). To test the influence of ecdysone on HDAC3 and its target gene expression, Aag-2 cells were exposed to 10 µM of 20-hydroxyecdysone (20E) or DMSO for 3 h, and mRNA levels of HDAC3 and proapoptotic genes were quantified. The URA, Hid, and caspase 3 mRNA levels were increased in Aag-2 cells exposed to 20E but not in cells exposed to DMSO (Fig. 4). To confirm this result in in vivo, the 12-h-old fourth instar larvae were treated with 100 ng/mL of stable ecdysone agonist (SEA), and mRNA levels of proapoptotic genes were quantified. The larvae were treated with SEA-induced URA, Hid, and caspase 3 mRNA levels (Fig. 4), while HDAC3 mRNA levels were not affected by 20E or SEA treatments in Aag-2 cells or A. aegypti larvae, respectively. Next, to test whether the presence of JH III has any influence on the 20E induction of URA, Hid, and caspase 3 genes, Aag-2 cells were simultaneously exposed to both 20E and JH III, and mRNA levels of proapoptotic genes were determined. Notably, 20E induced URA, Hid, and caspase 3 mRNA levels even in the presence of JH III (Fig. 4). Similarly, an increase in URA, Hid, and caspase 3 mRNA levels were observed in A. aegypti larvae simultaneously exposed to both SEA and methoprene (Fig. 4). Further, to determine the effect of 20E on HDAC3- and SMRTER-mediated suppression of URA, Hid, and caspase 3 genes, Aag-2 cells were transfected with both AaHDAC3 and AaSMRTER expression constructs and exposed to 20E or DMSO. In DMSO-exposed Aag-2 cells overexpressing AaHDAC3 and AaSMRTER, suppression of URA, Hid, and caspase 3 genes were observed (Fig. 4). However, 20E blocked AaHDAC3- and AaSMRTER-mediated suppression of URA, Hid, and caspase 3 genes. These results suggest that HDAC3 and SMRTER suppress URA, Hid, and caspase 3 genes in the presence of JH, but 20E prevents HDAC3- and SMRTER-mediated suppression of proapoptotic genes.

HDAC3 Regulates Midgut Cell Apoptosis Independent of Ecdysone Response Genes, E93 (Adult Specifier), and Broad Complex (Pupal Specifier).

Previous studies showed that the overexpression of the primary ecdysone response gene, E93, induces apoptosis in midgut cells during metamorphosis (27). To determine whether HDAC3 knockdown–induced midgut cell apoptosis is mediated through E93, the expression levels of E93 and genes coding for broad complex (Br-C) isoforms were analyzed in HDAC3 knockdown larvae. The E93 and Br-C mRNA levels were lower in HDAC3 knockdown larvae when compared to their levels in control larvae treated with dsmalE (). These data suggest that HDAC3 independently regulates the genes involved in apoptosis to control midgut cell apoptosis.

Midgut Size Is Reduced in HDAC3 Knockdown Larvae.

To determine whether apoptosis is the cause of larval death in HDAC3 knockdown larvae, the larval alimentary canals were dissected from 42-h-old fourth instar larvae that fed on dsHDAC3 or dsmalE and observed under a microscope. HDAC3 knockdown and control larvae exhibited a similar body size (Fig. 5 and ). However, a shorter alimentary canal was detected in HDAC3 knockdown larvae compared to its size in the control larvae (Fig. 5). To identify which part of the alimentary canal size was reduced in HDAC3 knockdown larvae, the experiment was repeated with A. aegypti larvae expressing the enhanced green fluorescent protein (EGFP) under IE1-hr5 promoter-enhancer and DsRed (Discosoma red fluorescent) protein driven by the IE2 promoter. The second instar larvae of this strain were fed on food pellets containing nanoformulated dsHDAC3 or dsmalE. The alimentary canals were dissected at 42 h after molting into fourth instar larvae and observed under a fluorescence microscope. As shown in Fig. 5, the midgut of dsHDAC3-treated larvae is significantly shorter than in the control larvae treated with dsmalE. Further, TUNEL (Terminal deoxynucleotidyl transferase dUTP nick end labeling) assay was used to identify apoptotic cells in the larval midguts dissected from the fourth instar larvae that fed on dsHDAC3 or dsmalE. Midgut cells undergoing apoptosis (as indicated by the green fluorescence of fragmented nuclear DNA labeled with fluorescent dUTP) were detected in 12-h-old fourth instar larvae, which fed on the dsHDAC3-containing diet (Fig. 5). Moreover, the number of fluorescent nuclei increased in 24-h-old larvae, and this number continued to increase until 48 h after molting into the fourth instar larvae that fed on the dsHDAC3-containing diet. Meanwhile, the control larvae which fed on dsmalE-containing diet showed fluorescently labeled nuclei only at 48 h after molting into fourth instar larvae. Moreover, the number of fluorescently labeled nuclei detected is lower than those in HDAC3 knockdown larvae (Fig. 5 and ). These results demonstrate that HDAC3 knockdown induces apoptosis of midgut cells in feeding larvae and causes a decrease in midgut size.

Fig. 5.

Knockdown of HDAC3 induces apoptosis, resulting in midgut size reduction in A. aegypti larvae. (A) HDAC3 knockdown and control larvae are similar in size. (B) The alimentary canals in HDAC3 knockdown larvae are shorter than those in the control larvae. The alimentary canals were dissected at 42 h after molting into fourth instar larvae fed on dsHDAC3- or dsmalE-nanoformulated diet. The alimentary canals were visualized under a fluorescence microscope using a white filter. (C) The alimentary canals in HDAC3 knockdown larvae expressing EGFP and DsRed are shorter than those in the control larvae fed on dsmalE. The alimentary canals were dissected 42 h after molting into fourth instar larvae. These alimentary canals were observed under a fluorescence microscope using the GFP filter. (Scale bar, 1,000 μm). (D) HDAC3 knockdown triggered apoptosis of midgut cells. Longitudinal sections 4 μM thick were cut from the midgut dissected from dsHDAC3- or dsmalE-treated larvae at 12, 24, and 48 h after ecdysis to the fourth instar larval stage. The sections were stained with the TUNEL assay reagent and DAPI nuclear stain and photographed under a confocal microscope. (E) No background green fluorescence was detected in the control larval section stained only with DAPI without TUNEL reaction mixture. (Scale bar, 100 μm.) The larval body and alimentary canal sizes and the mean fluorescence intensity values of apoptotic cells are provided in .

Discussion

One of the major contributions of this paper is the discovery that HDACs are important players in the regulation of growth, development, and metamorphosis of A. aegypti. Interestingly, the knockdown of each HDAC gene showed a distinct effect on the growth, development, and metamorphosis of A. aegypti. Out of the 10 HDAC genes tested, knockdown of HDAC3 showed the most-severe phenotypes during the larval stages, suggesting that it is required for maintaining the larval stage in A. aegypti. Our previous studies in T. castaneum also showed that HDACs have distinct functions (13). The knockdown of HDAC3 affected the wing development, and HDAC1 knockdown showed the most-severe phenotypes in T. castaneum (14). The developmental expression pattern of AaHDAC3 suggests its major role during the larval stage of A. aegypti. Moreover, HDAC3 mRNA levels correlate with JH titers during the larval stage, and JH induces HDAC3 expression through its receptor, Met. In our previous studies, JH suppressed the expression of HDAC1, HDAC3, and HDAC11 in T. castaneum, suggesting that the JH regulation of HDACs is different between A. aegypti and T. castaneum (13, 14, 28). In the current study, we also identified conserved JH response elements in the promoter region (−815 to −803 bp) of the HDAC3 gene, suggesting that the JH receptor complex may bind to the upstream region of HDAC3 and induce its expression in the presence of JH, which is similar to JH induction of Kr-h1 and other JH response genes (21, 22, 29–31). However, further studies are required to test binding of the Met complex to the upstream region of HDAC3.

Studies in mammalian systems suggest that HDAC3 activity is influenced by its association with the corepressors, SMRT and N-CoR (32). Here, we report that transcriptional repression of apoptosis genes by HDAC3 requires SMRTER, which is functionally similar to the vertebrate nuclear corepressors; SMRT and N-CoR. SMRTER may be recruited to proapoptotic gene promoters and serve as an anchoring molecule for recruiting HDAC3 (). The recruited HDAC3 could deacetylate histones resulting in the suppression of proapoptotic gene expression. Previous studies showed that the interaction of SMRT/N-CoR with HDAC3 potentiates HDAC3 activity (32). Similarly, we found that overexpression of HDAC3 alone is not sufficient to suppress the expression of the URA, Hid, and caspase 3. However, simultaneous overexpression of both HDAC3 and SMRTER suppressed these genes, suggesting that SMRTER is required for HDAC3 action.

We observed a decreased midgut size in the HDAC3 knockdown larvae, perhaps due to the premature death of midgut cells. HDAC3 was shown to be involved in suppressing genes involved in apoptosis (32). Apoptosis is initiated after decreased expression of antiapoptotic genes such as IAP and increased expression of proapoptotic genes such as Hid and caspases. In our study, knockdown of HDAC3 induced the overexpression of URA, Hid, and caspase 3 genes and suppressed expression of IAP. We showed that HDAC3 knockdown increased acetylation of histone H4 near the promoters of URA, Hid, and caspase 3 genes, suggesting that HDAC3 regulates the expression of these genes through chromatin modifications. In contrast, the acetylation levels of H4 near the IAP promoter were not affected by HDAC3 knockdown, suggesting that HDAC3 may not directly affect IAP gene expression. RNA-seq analysis revealed that the Hippo-signaling pathway is enriched in HDAC3 knockdown cells (). Hippo-signaling controls growth and development in D. melanogaster and mammals by regulating the tumor suppressor genes (33). Hippo-signaling dysfunction induces a growth-promoting oncogene, Yorkie, which is known to induce the IAP expression. Notably, HDAC3 is required for Yorkie expression (Fig. 4). Therefore, HDAC3 knockdown might have increased Hippo-signaling that resulted in the suppression of Yorkie and its downstream regulator, IAP (Fig. 3).

The other apoptosis protein, Hid, is also induced by the HDAC3 knockdown. Hid binds to IAP and blocks its action. It is also possible that URA induced by HDAC3 knockdown may regulate IAP protein activity by cleaving it to prevent IAP binding to caspases (23). In the absence of IAP, the initiator caspase 9 undergoes autocatalytic processing and becomes active. HDAC3 knockdown also induced hyperacetylation of the caspase 3 promoter that results in increased caspase 3 levels. The activated initiator caspase 9 cleaves the interchain of the inactive caspase 3 to make it catalytically active. Activated caspase 3 degrades many cellular proteins and causes apoptosis. Previous studies showed that inhibition of HDACs activity decreased cholangiocarcinoma cell growth and increased caspase-dependent apoptosis (34, 35).

Interestingly, caspase 3 was shown to cleave the C-terminal region of HDAC3 (which results in loss of its activity) and nuclear localization signal peptide (that leads to accumulation of HDAC3 protein in the cytoplasm) (36). The truncated HDAC3 was unable to deacetylate histones associated with proapoptotic gene promoters. Then URA-mediated ubiquitin-dependent degradation of HDAC3 results in the induction of caspase 3-mediated apoptosis (36). A previous study demonstrated that blocking of DPP-signaling induces autophagy in midgut cells and eventually removes midgut cells (37), while application of JH analog, methoprene, prevents apoptosis of larval midgut cells during larval–pupal metamorphosis and retains larval midgut in A. aegypti pupae (38). In this study, we showed that JH-induced, HDAC3-mediated deacetylation suppresses proapoptotic genes during the larval stage to prevent premature induction of apoptosis in larval midgut cells.

During larval–pupal metamorphosis, a decrease in JH titers and an increase in ecdysteroid titers may cause an exchange of the corepressor complex with a coactivator complex near the ecdysone response gene promoters to activate their transcription (39). We observed that HDAC3 was unable to suppress genes involved in apoptosis in the presence of 20E. This suggests that 20E acts as a switch to induce proapoptotic genes by exchanging corepressor complex (SMRTER/HDAC3) with a coactivator complex and recruiting ecdysone receptors to the promoters of these genes (40–42). This leads to transcription of proapoptotic genes and apoptosis of larval midgut (). Similarly, in the absence of ligand, thyroid hormone, retinoic acid receptors recruit SMRT/N-CoR corepressor complexes that bind to the target gene promoters and repress gene transcription. However, the presence of thyroid hormone dissociates corepressor complexes and recruits coactivator complexes to the promoters to induce target gene expression (43).

In conclusion, we showed that antagonistic actions of JH and ecdysone on proapoptotic gene expression are mediated by the HDAC3 complex. JH induces an epigenetic modifier, HDAC3, to prevent premature death of larval cells during the feeding stage. During metamorphosis, the decline in JH titers and increase in ecdysone titers result in the suppression of HDAC3 and promotion of apoptosis of larval midgut cells. This study could serve as a foundation for understanding the role of epigenetic modifications in the antagonistic action of JH and ecdysone involved in the regulation of the expression of apoptosis genes during larval development and metamorphosis.

Materials and Methods

Larval Bioassays.

A. aegypti Liverpool (LVP) strain mosquitoes were reared as described previously (9). To knockdown target genes, dsRNA was synthesized using the MEGAscript RNAi kit (Life Technologies) as described previously (44). Poly-L-lysine (PLL):Epigallocatechin gallate (EGCG):dsRNA nanoparticle complexes were prepared following the methods described previously (9, 45). Briefly, food pellets were prepared by mixing PLL:EGCG:dsRNA complexes with mosquito larval food and bovine liver powder. These pellets were fed to second instar larvae in 12-well plates (five larvae/well; n = 30) containing 2 mL nuclease-free water. Each food pellet contained 40 μg dsRNA conjugated with PLL:EGCG nanoparticles. Each food pellet was divided into three equal pieces and distributed to three wells. Fresh food pellets containing PLL:EGCG:dsRNA complexes were added to each well every day until pupation or death. Mortality and phenotypes were recorded every day until adult eclosion. Corrected mortality was calculated using Abbott's formulae (46).

Cell Culture, Target Gene Knockdown, and Hormone Treatment.

A. aegypti embryo-derived Aag-2 cells were cultured at 28 °C in Schneider's Drosophila medium (Life Technologies) supplemented with 10% fetal bovine serum (Life Technologies). Aag-2 cells were seeded in 6-well culture plates at a density of 200,000 cells/well. Cells were treated with 10 µg dsRNA of target or dsmalE (control) for 3 consecutive d. On the fourth day, cells were exposed to 10 µM of JH III (Sigma-Aldrich) or 20E (Sigma-Aldrich) or DMSO for 3 h. Then the total RNA was isolated, and target genes mRNA levels were quantified.

JH Analog, Methoprene, and Ecdysone Agonist Treatments.

Technical-grade methoprene (isopropyl (E,E)-(RS)-11-methoxy-3,7,11-trimethyldodeca-2,4-dienoate) was dissolved in DMSO. Technical-grade SEA, RH-102240 (N-(1,1-dimethylethyl)-N¢-(2-ethyl-3-methoxybenzoyl)-3,5-dimethylbenzohydrazide) was dissolved in ethanol. In total, 100 ng/mL final concentration of methoprene or SEA was added to nuclease-free water and dispensed into 12-well plates and released newly molted five fourth instar larvae/well (n = 20: four replicates). Control larvae were treated with 1 μL/mL DMSO or ethanol. RNA was extracted from the treated and control larvae at 36, 42, and 48 h after treatment.

RNA Isolation, cDNA Preparation, and qRT-PCR.

Total RNA was extracted from A. aegypti larvae or pupae or Aag-2 cells using the TRI reagent by following the manufacturer's protocol (Molecular Research Center, Inc.). Developmental expression of target genes was determined by extracting the total RNA from the whole insects collected at different time points during development. First-strand complementary DNA (cDNA) synthesis using 2 µg total RNA and qRT-PCR assays were performed as described previously (9). RPS7 and β-tubulin housekeeping genes were used as internal reference genes to normalize the target gene mRNA levels (). The target gene-expression levels were normalized with the reference gene-expression levels by subtracting the target gene CT value from the reference gene CT value followed by determination of −ΔΔCT value (47). All experiments were performed using at least three biological replicates and repeated three times.

RNA-Seq and Annotation.

Three RNA-seq libraries per treatment were prepared using RNA isolated from three independent biological samples of Aag-2 cells treated with dsHDAC3 or dsmalE. KAPA Stranded mRNA-Seq Kit (Illumina) was used to prepare RNA-seq libraries by following manufacturer protocol. RNA-seq was performed using the Illumina HiSeq 4000 sequencer platform (Sequencing and Genomics Technologies Center of Duke University). RNA-seq data were analyzed using the CLC Genomics Workbench (version 10.1.1, Qiagen Bioinformatics) as described previously (44). Briefly, the sequence data were subjected to quality control to remove low-quality reads and adapters. Clean reads were mapped to the LVP strain transcriptome data retrieved from VectorBase (https://vectorbase.org/vectorbase/app/record/dataset/TMPTX_aaegLVP_AGWG). Mapping was performed using the CLC Genomics Workbench employing default mapping parameters. To find the differentially expressed genes between dsHDAC3 treatment and dsmalE-treated control, the empirical analysis of the differential gene-expression tool in the CLC Genomics Workbench was used by employing standard parameters along with greater- or equal-than-twofold expression variation and a P value < 0.05 cutoff values. For annotation, the cloud blast feature in the Blast2GO software was used to check against the arthropod nonredundant protein database with a Blast expectation value (e-value) 1.0 × 10−6. Gene Ontology (GO) enrichment analysis was performed using the Web Gene Ontology Annotation Plot (WEGO) online tool (48). KEGG pathway enrichment analysis was performed using the KEGG Orthology-Based Annotation System (KOBAS) 3.0 online tool (49).

Western Blot Hybridization and ChIP Assay.

To identify HDAC3 deacetylated histones, Aag-2 cells were treated with dsHDAC3 or dsmalE for 3 consecutive d, and chromatin-bound proteins were isolated by following the protocol described previously (50). Western blotting was performed using 10 μg total protein resolved on 12% SDS-PAGE (sodium dodecyl sulfate–polyacrylamide gel electrophoresis) gels. Blotting, hybridization, and detection were carried out as described previously (9). Band intensities of three replicate Western blots were measured using the ImageJ software, and statistical analysis was performed to determine the significance of difference (18).

ChIP assays were performed using the Pierce Magnetic ChIP Kit (Thermo Fisher Scientific) as described previously (9). Briefly, 2 × 106 cells were treated with 10 µg dsHDAC3 or dsmalE for 3 consecutive d. On the fourth day, cells were cross linked using formaldehyde. Cross-linked DNA was digested with micrococcal nuclease and immunoprecipitated using antibodies that recognize histone H4 acetylation (Ac-Histone H4 Antibody [E-5]: sc-377520; Santa Cruz Biotechnology, Inc., Dallas) or the rabbit IgG antibody (used as a negative control). Immunoprecipitated DNA was purified, and the target genes promoter enrichment levels were quantified using the qPCR.

Transfection and Overexpression of Target Genes.

To overexpress AaMet in Aag-2 cells, the AaMet construct reported previously was used (21). Full-length AaHDAC3 and AaSMRTER cDNAs were cloned into the pIEx-4 vector and used for transfection. Aag-2 cells were transfected with 500 ng of pIEx-4 vector containing the gene of interest or empty pIEx-4 vector (control) using the Cellfectin II transfection reagent. At 48 h after transfection, the cells were exposed to DMSO or 10 µM of JH III or 20E for an additional 3 h. Total RNA was isolated from these cells used to quantify mRNA levels of target genes. Three independent experiments were performed, and each experiment included four biological replicates.

Preparation of Whole-Insect Longitudinal Sections and TUNEL Assay.

TUNEL assays were performed on whole-insect longitudinal sections as described previously (38). Briefly, alimentary canals were dissected from larvae that fed on dsHDAC3- or dsmalE-nanoformulated diet, then washed with 1× phosphate-buffered saline (PBS) and fixed in 4% paraformaldehyde overnight at 4° C. After fixation, the tissues were washed with 1× PBS twice, then dehydrated using a series of grades of ethanol (25%, 50%, 75%, 95%, and 100% in 1× PBS) and infiltrated through a series of grades of xylene (25%, 50%, 75%, 95%, and 100% in ethanol) and embedded in Paraplast Plus at 56° C. Longitudinal sections 4 μM thick were cut using a microtome (Leica RM 2135, Germany) and mounted on a glass slide. The sections were deparaffinized by incubating glass slides at 60° C for 1 h, followed by rehydration using a series of grades of xylene and ethanol. Then the sections were incubated in 0.3% Triton X-100 in 0.1% sodium citrate buffer for 10 min and washed with 1× PBS twice and used to perform TUNEL assay using the In Situ Cell Death Detection Kit, Fluorescein (Roche) by following manufacturer’s instructions. Sections were exposed to the TUNEL reaction mixture (30 µl/slide) for 60 mins at 37° C, then washed with 1× PBS and mounted using DAPI nuclear-staining reagent. The sections were examined under the Leica TCS SP8 DLS (Digital LightSheet) confocal microscope. The mean fluorescence intensities for control and treated samples were determined by following the method described by Shihan et al. (51).

Statistical Analysis of Data.

One-way ANOVA was used for all statistical analysis unless otherwise stated. P < 0.05 was considered significant.