Variation in RNAi efficacy among insect species is attributable to dsRNA degradation in vivo

Insect Biochemistry and Molecular Biology 77 (2016) 1e9

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Insect Biochemistry and Molecular Biology journal homepage: www.elsevier.com/locate/ibmb

Variation in RNAi efficacy among insect species is attributable to dsRNA degradation in vivo Kangxu Wang, Yingchuan Peng, Jian Pu, Wenxi Fu, Jiale Wang, Zhaojun Han* Department of Entomology, College of Plant Protection, Nanjing Agricultural University, Jiangsu/The Key Laboratory of Monitoring and Management of Plant Diseases and Insects, Ministry of Agriculture, Nanjing, 210095 Jiangsu, China

a r t i c l e i n f o

a b s t r a c t

Article history: Received 28 December 2015 Received in revised form 12 July 2016 Accepted 16 July 2016 Available online 20 July 2016

RNA interference (RNAi) has become an essential technique in entomology research. However, RNAi efficiency appears to vary significantly among insect species. Here, the sensitivity of four insect species from different orders to RNAi was compared to understand the reason for this variation. A previously reported method was modified to monitor trace amounts of double-stranded RNA (dsRNA). After the administration of dsRNA, the dynamics of its content was determined in the hemolymph, in addition to the capability of its degradation in both the hemolymph and the midgut juice. The results showed that injection of dsRNA targeting the homologous chitinase gene in Periplaneta americana, Zophobas atratus, Locusta migratoria, and Spodoptera litura, with doses (1.0, 2.3, 11.5, and 33.0 mg, respectively) resulting in the same initial hemolymph concentration, caused 82%, 78%, 76%, and 20% depletion, respectively, whereas feeding doses based on body weight (24, 24, 36, and 30 mg) accounted for 47%, 28%, 5%, and 1% depletion. The sensitivity of insects to RNAi was observed to be as follows: P. americana > Z. atratus >> L. migratoria >> S. litura. In vivo monitoring revealed that RNAi effects among these insect species were highly correlated with the hemolymph dsRNA contents. Furthermore, in vitro experiments demonstrated that the hemolymph contents after dsRNA injection were dependent on hemolymph degradation capacities, and on the degradation capabilities in the midgut juice, when dsRNA was fed. In conclusion, the RNAi efficacy in different insect species was observed to depend on the enzymatic degradation of dsRNA, which functions as the key factor determining the inner target exposure dosages. Thus, enzymatic degradation in vivo should be taken into consideration for efficient use of RNAi in insects. © 2016 Elsevier Ltd. All rights reserved.

Keywords: RNA interference RNAi efficacy dsRNA degradation Insect species dsRNA delivery

1. Introduction RNA interference (RNAi) has become an important technique for manipulating cellular phenotypes and mapping genetic pathways and could potentially be used for pest control (Li et al., 2015; San Miguel and Scott, 2015; Scott et al., 2013). To trigger the RNAi pathway, double-stranded RNA (dsRNA) should be delivered to organisms, first. After cellular uptake and recognition, dsRNA molecules are cleaved by a protein, Dicer, into short interfering RNAs (siRNAs) comprising 20e25 bp, which guide a multi-protein RNA-induced silencing complex (RISC) to the complementary mRNA for degradation (Filipowicz, 2005; Mello and Conte, 2004). Injection and feeding of in vitro synthesized dsRNA are the most wildly used approaches for administering RNAi in insects. However,

* Corresponding author. E-mail address: [email protected] (Z. Han). http://dx.doi.org/10.1016/j.ibmb.2016.07.007 0965-1748/© 2016 Elsevier Ltd. All rights reserved.

the efficiency of RNAi varies considerably with the insect species and the mode of dsRNA delivery (Scott et al., 2013; Terenius et al., 2011). Many caterpillars are not very amenable to RNAi using both the injection and feeding approach (Terenius et al., 2011). However, RNAi in the German cockroach, Blattella germanica, is effective for many genes and at different life stages, both by injection and feeding methods (Huang et al., 2013; Irles et al., 2013). In Aedes aegypti, dsRNA injection is the most successful way for RNAi (Cirimotich et al., 2009; Gillen et al., 2015), whereas dsRNA feeding has been successful in only a few cases (Coy et al., 2012; Mysore et al., 2013). In the desert locust, Schistocerca gregaria, significant knockdown of target genes was observed with dsRNA injection, but the RNAi response was much less sensitive to dsRNA feeding (Wynant et al., 2014b). Effective RNAi has been achieved by dsRNA injection in the red flour beetle, Tribolium castaneum (Miller et al., 2012; Wang et al., 2013; Xiao et al., 2015); successful gene suppression by dsRNA feeding has also been evidenced in this beetle (Whyard et al., 2009).

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In initial studies, the RNAi phenomenon was observed to be different among insects and nematodes (Kennerdell and Carthew, 1998). However, substantial difference was not noticed in the core elements and mode of action in these organisms (Prentice et al., 2015; Tomoyasu et al., 2008; Wynant et al., 2015). In the present study, we investigated the RNAi action in insects, with the aim of enhancing its efficiency and usage. Insects from four different orders were selected and experiments were carried out in parallel to compare their response to RNAi. A sensitive method was established by improving on the one reported by Garbutt et al. (2013) to monitor the in vivo dynamics of dsRNA content after injection and feeding. The RNAi efficacy among the insect species was found to be highly correlated with the dsRNA content in the hemolymph; in vitro incubation assays demonstrated that the major reason for this was the varied dsRNA degradation capability of different insect species. 2. Materials and methods 2.1. Insects Four species of insects from different orders, generally known to vary in their sensitivity to RNAi (Huang et al., 2013; Irles et al., 2013; Miller et al., 2012; Terenius et al., 2011; Wang et al., 2013; Wynant et al., 2014b; Whyard et al., 2009; Xiao et al., 2015), were selected for comparison; these included the migratory locust (Locusta migratoria, Orthoptera), the American cockroach (Periplaneta americana, Blattaria), the tobacco caterpillar (Spodoptera litura, Lepidoptera), and the tenebrionid beetle (Zophobas atratus, Coleoptera). Adults of the migratory locusts were reared in the gregarious phase under a photoperiod with 14 h light: 10 h dark at 30
C, with continuous supply of reed and corn leaves diet. Adults of the American cockroaches were fed daily with oats and dog food and kept in complete darkness at 30
C; larvae of the tobacco caterpillars were reared on semi-artificial food (Huang and Han, 2007) at 25
C and a photoperiod of 17 h light: 7 h dark; larvae of the tenebrionid beetles were reared under 24 h darkness at 28
C and fed food (lettuce, carrot, powder milk) and water ad libitum. 2.2. dsRNA, cDNA synthesis, and RNA extraction The dsRNA was generated by in vitro transcription using a T7 RiboMAX system (Promega, Madison, WI, USA), according to the manufacturer's manual. A DNA template flanked by two T7 promoter sequences was synthesized for the production of dsRNA. Primers used in the dsRNA synthesis are listed in Table 1. The concentration of each transcript was determined using a NanoDrop ND-1000 spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). In view of the lack of Z. atratus genome sequence, the homologous chitinase gene of Z. atratus was partially cloned by using degenerate primers, designed on the basis of the sequences of chitinase genes from related species whereas the sequences of target genes of S. litura, L. migratoria, and P. americana used in the dsRNA synthesis were retrieved from the transcriptome databases with reciprocal tBLASTn (National Center for Biotechnology

Information). The specificity of these genes was confirmed by BLAST analysis (NCBI). For each kind of insect, cDNA was prepared from their whole-body. Total RNA was extracted with an RNeasy Mini Kit (Qiagen, Valencia, CA, USA), according to the manufacturer's protocol. The quality and quantity of RNA was measured using 1% gel electrophoresis and a NanoDrop ND-1000 spectrophotometer. The cDNA was reverse-transcribed from 1 mg of total RNA with MMLV reverse transcriptase (TaKaRa, Dalian, Liaoning, China). The dsRNA was stored at 70
C, until further use. 2.3. Treatment with dsRNA For comparing RNAi effects induced by dsRNA injection, different doses of dsRNA were used for different insect species to achieve similar initial hemolymph dsRNA concentration, thereby, reducing the influence of body size and hemolymph volume. First, 4 mg dsRNA was intra-abdominally injected into test insects of the four species (S. litura: middle 6th instar larvae; L. migratoria: 2days-old adults; P. americana: 3-days-old adults, and Z. atratus: middle 5th instar larvae), which were selected for convenient operation and had similar body weight. Five minutes later, after even mixing of dsRNA, the hemolymph was sampled and dsRNA concentration was determined with the method described below. The ratio calculated from the obtained dsRNA concentration in S. litura, L. migratoria, Z. atratus, and P. americana (29, 82, 405, and 943 pg/ml, respectively) was used to adjust the dsRNA doses for different insect species to achieve similar initial hemolymph dsRNA concentration. Thus, the injection doses employed for S. litura, L. migratoria, Z. atratus, and P. americana were 33, 11.5, 2.3, and 1 mg, respectively. For RNAi induction by feeding, dsRNA doses were calculated based on the body weight of the test insects (1.0, 1.2, 0.8, and 0.8 g, respectively). The doses used were 30, 36, 24, and 24 mg for S. litura, L. migratoria, Z. atratus, and P. americana, respectively. The insects were first subjected to starvation for 24 h to stimulate their appetite and to avoid vomiting. Thereafter, single insects were held between two fingers with their mouth pointed upwards and dsRNA solution was dropped into the mouth with a pipette. The insects administered dsRNA through injection or feeding were reared routinely and samples were collected for checking the hemolymph dsRNA concentration and RNAi effect. 2.4. Quantitative Real-Time PCR For checking the RNAi effect, the expression level of target gene was determined by quantitative Real-Time PCR (qPCR). Insect samples were collected 72 h after dsRNA treatment and kept in liquid nitrogen. For each treatment, total RNA was isolated from five individuals. RNA extraction and reverse-transcription was performed with the same methods as described above. The qPCR assays were conducted according to MIQE (Minimum Information for Publication of Quantitative Real-Time PCR Experiments) guidelines. To estimate the amplification efficiency and correlation coefficient of each primer pairs, a range of serial dilution of cDNA (10n-fold) was used to create the five-point standard curve. The equation

Table 1 Primers used for dsRNA synthesis. Fragment

Forward

Reversed

dsRNA length

dsSlChi dsLmChi dsZaChi dsPaChi dsEGFP

TCTACGGTCAGTCCTATAG TCAGCCAGTGAGAATGAC CTCTTTACGGTCAGTCCT TTCTTGAGCCATCCGAAAG AAGTTCAGCGTGTCCG

AGGTTCTGGTGGTCTTT GGATTTGAAGGACTAGACC CTTCCTCAATGGAAGGAATC CTTCTTCGAAATGATGGCG CACCTTGATGCCGTTC

419 422 417 423 414

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(E ¼ [10(1/slope)1]  100%) was used to calculate the qPCR efficiency for each primer. The qPCR was performed with SYBR Premix Ex Taq™ (TaKaRa) on an ABI 7500 Real Time PCR system (Applied Biosystems, Foster City, CA, USA). The PCR conditions were as follows: 95
C for 30 s, followed by 40 cycles at 95
C for 5 s and 60
C for 34 s. For each gene, at least five biological and three technical replicates were tested. Beta-actin for S. litura (GenBank: DQ494753.1), L. migratoria (GenBank: KC118986.1), and P. americana (GenBank: AY116670.1) and ribosomal protein, RpS18, for Z. atratus (Marciniak et al., 2013), were used as the internal controls, as described in previous reports (Jung et al., 2013; Lu et al., 2013; Marciniak et al., 2013; Yang and Han, 2014). The 2△△Ct method was used for calculating the relative expression level with ABI 7500 analysis software. The primers used for qPCR are listed in Table 2. 2.5. In vivo monitoring of dsRNA dynamics The dsRNA solution was injected or fed through the drink, as described above in the dsRNA treatment. The solution for hemolymph collection was prepared by addition of 10 ml b-mercaptoethanol (Promega) into 1 ml RLT-buffer present in the RNeasy Micro Kit (Qiagen). A single insect was picked up and held with its abdomen facing outwards. A sterilized tiny needle was used to prick the insect at the coxal cavity of its hind leg or at the base of the 1st proleg of caterpillars. Ten microliters of hemolymph was drained from the wound and mixed with 350 ml of the collection solution immediately by spinning at 16,000  g for 5 min. The resultant lysate was stored at 70
C prior to the RNA extraction. The insect was put back and reared normally for continuous sampling over the experimental time course. The RNeasy Micro Kit was used to extract the trace dsRNA in the samples, according to the protocol and a 5 min denaturation step at 65
C was added directly preceding the reverse transcription reaction in order to denature the secondary structure of the dsRNA. The cDNA was synthesized from this RNA according to the instructions provided in the PrimeScript™ Reverse Transcriptase (TaKaRa) manual. The dsRNA residues in the hemolymph were tested by qPCR as described above and quantified using the formula derived from the calibration experiments with serially diluted dsRNA. The primers for cDNA synthesis and qPCR are listed in Tables 1 and 2 The experiment was performed at least five times independently. 2.6. Incubation of dsRNA in the collected hemolymph or midgut juice Hemolymph was collected from individual insects and kept in different microcentrifuge tubes containing phenylthiourea and cooled on ice in order to avoid melanization. Hemocytes were removed by centrifugation at 16,000  g for 10 min at 4
C. The procedure for collecting the midgut juice was adapted from that

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described by Yang and Han (2014). The juice was stored at 20
C until use. For in vitro incubation assays, 1 mg dsEGFP dissolved in 1 ml water was added to 5 ml hemolymph or midgut juice diluted with nuclease-free water. The control used 5 ml of nuclease-free water instead of hemolymph or midgut juice. After incubation for the indicated time, the samples were mixed with loading dye and run on a 1% agarose EtBr gel. The dsRNA was visualized with a VersaDoc MP 4000 (Bio-Rad, Hercules, CA, USA) under UV light. The results were analyzed using the QuantityOne v4.62 software (BioRad). The relative band intensity was calculated according to the method of Henriques et al. (2012). 2.7. Statistical analysis Significance analyses were performed with the Student's t-test. Differences were considered significant at P < 0.05. Values are reported as mean ± SE. Both linear regression models and curve models were constructed with Curve Estimation process. Bivariate correlations were adopted for correlation analyses. All statistical analyses were carried out using SPSS 19.0 software (SPSS Inc., Chicago, IL, USA). 3. Results 3.1. Efficacy of RNAi in S. litura, L. migratoria, P. americana, and Z. atratus The sensitivity of S. litura, L. migratoria, P. americana, and Z. atratus to RNAi was compared by administration of dsRNA through feeding and injection. With the homologous chitinase genes (SlChi, LmChi, PaChi, and ZaChi) of these four insects as the templates, four dsRNAs, about 417e423 bp in length, were synthesized targeting the same fragment (Fig. 1). These were then fed to or injected into different insect species as described in section 2.3. The qPCR was used to check the expression levels of chitinase genes in the treated insects. The results in Fig. 2 demonstrate that the depletion caused by dsRNA injection was dramatic in P. americana (82%), Z. atratus (78%), and L. migratoria (76%), whereas it was not significant in S. litura (20%). For dsRNA feeding, significant depletion was found only in P. americana and Z. atratus (by 47% and 29%, respectively), but not in S. litura and L. migratoria (by 1% and 5%, respectively). This implied that among the four tested insect species, S. litura was highly insensitive to RNAi, L. migratoria was sensitive only to dsRNA injection, P. americana and Z. atratus were sensitive to both dsRNA injection and feeding, and P. americana was the most sensitive species. 3.2. Method for monitoring hemolymph content dynamics of dsRNA in vivo For monitoring of the hemolymph content dynamics of dsRNA

Table 2 The primers used for qPCR. Fragment

Forward

Reversed

PCR efficiency (%)

Size (bp)

qSlChi qSlAct qLmChi qLmAct qZaChi qZaRps18 qPaChi qPaAct qEGFP

CGCAACGGCAAGTAGTGAAGC ATCCTCCGTCTGGACTTGG CAACCACAGCGATTGCGGAAG CGAAGCACAGTCAAAGAGAGGTA ATGGACAATGGGACAAACAAACCG CGAAGAGGTCGAGAAAATCG AGGCAGTGGCAAATTGAGAAACC GCTATCCAGGCTGTGCTTTC GACGACGGCAACTACAAGAC

GTCGTAGTGGTGGTCGTAGTGG CGCACGATTTCCCTCTCA ACAAAGGATGGACTACGGTAAGGG GCTTCAGTCAAGAGAACAGGATG GGACTGACCGTAAAGAGGCATACC CGTGGTCTTGGTGTGTTGAC GGCAGACGCAATGAGTTGTAGG ACCGGAATCCAGCACAATAC GTCCTCCTTGAAGTCGATGC

96.7 96.1 94.3 93.8 92.7 97.4 93.1 94.2 95.9

105 214 190 156 161 235 104 66 96

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Fig. 1. Multiple alignment of the homologous chitinase gene targets for dsRNAs from four different insect species. SL: Spodoptera litura; LM: Locusta migratoria; ZA: Zophobas atratus and PA: Periplaneta americana.

Fig. 2. RNAi effects in four insect species after ingestion or injection of dsCHI specific to each species. (A) Relative expression of chitinase mRNA after injection of dsCHI; (B) Relative expression of chitinase mRNA after ingestion of dsCHI. The negative control used was dsEGFP and the blank control used was water. Each treatment contained at least 5 replicates, with 5 individuals included in each replicate. The relative expression was calculated based on the blank control. Statistical analysis was performed with Student's t-test (mean ± SE; *, p < 0.05; **, p < 0.01).

in vivo, the method reported by Garbutt et al. (2013) was adopted and modified. An RNeasy micro kit was employed to extract the dsRNAs from 10 ml hemolymph samples (see section 2.5 for more details). It was further checked if an endogenous gene interfered with the detection of the corresponding dsRNA; as a representative, dsSlChi (419 bp) was employed for S. litura and its content was determined in the hemolymph of S. litura before and after feeding, over the time course (Fig. 3). It was observed that the ingested dsSlChi was detected in the hemolymph and the peak level (15 pg/ ml) appeared after about 8 min of feeding. However, the endogenous gene provided at a base level of 3 pg/ml appeared to have no interference on the monitoring of the foreign dsRNA. However, different insects might have different base levels because of the endogenous level of gene expression or degradation. For comparison in the present study, a 414-bp dsEGFP was synthesized and used for treatment. As this dsRNA was unrelated to any endogenous gene in the different insects, a uniform method could be used to monitor the same dsRNA among the various insect species. The calibration line for dsEGFP quantification developed by calibration experiments with serial dilutions of known dsEGFP amounts proved to be perfect (Fig. 4). The linear range detected in the sample was approximately from 1 pg to 100 ng dsEGFP. Another

serial dsEGFP dilution with 10 ml of S. litura hemolymph was also tested for each sample. It was observed that the hemolymph medium did depress the sensitivity of the test, especially at lower concentrations. However, the calibration lines in the two cases were nearly parallel, indicating a minor difference. Thus, we selected the line obtained with water dilutions as the work line instead of setting different work lines for each insect species because of their different hemolymph. 3.3. In vivo content dynamics of injected dsEGFP in insect hemolymph In the present study, a uniform 414-bp dsEGFP was employed for injection and as the detection target to avoid the possible interference of endogenous genes in the insects; 33, 11.5, 2.3, and 1 mg dsEGFP was used for S. litura, L. migratoria, Z. atratus, and P. americana, respectively, to achieve similar initial hemolymph dsRNA concentration (see section 2.3). The hemolymph dsEGFP contents in vivo were monitored after the injection. The results in Fig. 5 show that the maximum hemolymph dsEGFP concentrations monitored first time (at 5 min) after the injection were around 310 pg/ml in the four species, which then decreased at different

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Fig. 3. Content dynamics of ingested dsSlChi in S. litura. (A) Calibration curve for dsSlChi quantification. Ct values were plotted against the log10 transformed quantity (pg) of dsEGFP in 10 ml samples. (B) Content dynamics curve. The SlChi background was detected 5 min before feeding 30 mg dsSlChi. After the treatment, a sample of 10 ml hemolymph was collected from a single insect at different time points. RNA was isolated from the 10-ml hemolymph and the sample was incubated at 65
C for 5 min, just before the reverse transcription, to denature the dsRNA. The remaining dsSlChi was then transformed to cDNA and analyzed using qPCR. The data were described by constructing curve model (y ¼ 0.04592x2 þ 0.70706x - 1.9135, R ¼ 0.8586). Values are mean ± SE; n  5.

Fig. 4. Calibration curve for dsEGFP quantification. The assay was generated by spiking the hemolymph with serially diluted dsEGFP (depicted with circles) or by only using the serially diluted dsEGFP solution (diamonds). Ct values were plotted against the log10 transformed quantity of dsEGFP (pg) in 10 ml samples. Values are mean ± SE; n  5.

speeds within 2 h of injection. The decreasing speed index was determined by using the linear regression model (the log concentration with the post injection time) and the slope of the decrease line was used as the parameter for comparison. The result suggests that the dsRNA decrease speed for S. litura was fastest, and that for P. americana was slowest. The relative dsRNA decrease speed indexes for S. litura, L. migratoria, Z. atratus, and P. americana were 2.06, 1.44, 1.20, and 1, respectively, which were negatively correlated (0.9501) with the RNAi depletion (20%, 76%, 78%, and 82%) among these insect species. 3.4. In vivo content dynamics of ingested dsEGFP in the insect hemolymph

Fig. 5. Content decreasing dynamics of injected dsEGFP in S. litura, L. migratoria, Z. atratus, and P. americana. DsEGFP was injected into individual insects and then 10 ml hemolymph was collected from a single insect over the time course. RNA was isolated from the 10 ml hemolymph and the sample was incubated at 65
C for 5 min, just before reverse transcription, to denature the dsRNA. The remaining dsEGFP was then transformed to cDNA and its content was analyzed with qPCR. The linear regression models were established to analyze the hemolymph dsRNA decrease speed index in the four insect species (Sl, y ¼ 0.0268x þ 2.4317, R ¼ 0.9642; Lm, y ¼ 0.0187x þ 2.4610, R ¼ 0.9616; Za, y ¼ 0.0156x þ 2.3771, R ¼ 0.9725; Pa, y ¼ 0.0130x þ 2.3456, R ¼ 0.9698). Values are mean ± SE; n  5.

The test insects of S. litura, L. migratoria, Z. atratus, and P. americana were fed 30, 36, 24, and 24 mg dsEGFP, respectively according to their body weight, and then the hemolymph dsEGFP content dynamics were monitored (Fig. 6). The dsEGFP could be detected in the hemolymph of the tested insects at around 3e5 min after feeding. Thereafter, the dsEGFP content increased for a while. However, the length of the increase phase, the peak level reached, and the persistence time of the detectable dsEGFP concentration in the hemolymph of different insect species was quite different. The peak level appeared at about 7 min in S. litura (10 pg/ml), at 13 min in L. migratoria (10 pg/ml), at 45 min in Z. atratus (25 pg/ml), and at 65 min in P. americana (600 pg/ml). Correspondingly, the detectable period extended by about 10 min in S. litura, 15 min in L. migratoria, 95 min in Z. atratus, and 120 min in P. americana. The dsRNA

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Fig. 6. Hemolymph content dynamics of ingested dsEGFP in S. litura, L. migratoria, Z. atratus, and P. americana. DsEGFP was fed to insects of the four species and then 10 ml hemolymph was collected from each individual insect at different time points. RNA was isolated from the 10 ml hemolymph and the sample was incubated at 65
C for 5 min, just before the reverse transcription, to denature the dsRNA. The remaining dsEGFP was then transformed to cDNA and its content was analyzed with qPCR. The curve models were constructed to analyze the dsRNA hemolymph concentration-time integrals in the four insect species (Sl, y ¼ 0.0538x2 þ 0.7908x  2.1092, R ¼ 0.9284; Lm, y ¼ 0.0144x2 þ 0.3598x  1.3946, R ¼ 0.9954; Za, y ¼ 0.0007x2 þ 0.0684x  0.2676, R ¼ 0.9885; Pa, y ¼ 0.0009x2 þ 0.1104x  0.6533, R ¼ 0.9894). Values are mean ± SE; n  5.

hemolymph concentration-time integral were calculated as hemolymph content for comparison. It was observed that the relative hemolymph dsEGFP contents in the feeding treatment were 1, 1.17, 2.01, and 2.90 for S. litura, L. migratoria, Z. atratus, and P. americana, respectively. This showed a good correlation (0.9983) between the hemolymph contents of dsEGFP and the RNAi depletions by feeding dsRNA among the insect species (1, 5, 28, and 47%). 3.5. Degradation of dsRNA in the insect hemolymph For determining the hemolymph degradation of dsRNA, in vitro experiments were performed by incubation of 1 mg dsEGFP (in 1 ml water) with 5 ml of a serial dilution of cell-free hemolymph collected from S. litura, L. migratoria, Z. atratus, and P. americana, respectively. After one hour, the remaining dsEGFP in the samples was visualized by electrophoresis on a 1% agarose gel. In the control, 1 mg dsEGFP was incubated with 5 ml nuclease-free water. As shown in Fig. 7, among the four insect species tested, S. litura displayed the strongest ability to degrade dsRNA, followed by L. migratoria, Z. atratus, and P. americana. Bands observed in the incubation with 4 dilution of hemolymph were then quantified using the QuantityOne v4.62 software. The hemolymph degradation was 95, 72.6, 24.7, and 9%, respectively for S. litura, L. migratoria, Z. atratus, and P. americana, respectively, as compared to the control, showing strong correlation (0.9350) with the in vivo hemolymph dsRNA decreasing speed index after the injection (relatively, 2.06, 1.44, 1.20, and 1). 3.6. Degradation of dsRNA in insect midgut juice In vitro experiments were performed to evaluate the dsRNA degradation in insect midgut juice. One microgram dsEGFP (in 1 ml

water) was incubated for 1 h with 5 ml of 20 dilutions of the midgut juice collected from S. litura, L. migratoria, Z. atratus, and P. americana, respectively. The remaining dsEGFP in the samples was visualized by electrophoresing on 1% agarose gel. The control consisted of nuclease-free water instead of the midgut juice. Band quantification was done using the QuantityOne v4.62 software. The 20 dilution of the midgut juice collected from S. litura and L. migratoria could degrade the dsRNA efficiently (88 and 80%, respectively), whereas the midgut juice of Z. atratus and P. americana could only degrade it to a little (17 and 16%, respectively) extent (Fig. 8). The degradation activity in the midgut juice of S. litura, L. migratoria, Z. atratus, and P. americana negatively correlated (0.9143) with their hemolymph contents (relatively 1, 1.17, 2.01, and 2.90) after the feeding treatments.

4. Discussion Numerous studies have shown that RNAi efficacy varies with the insect species (Scott et al., 2013; Terenius et al., 2011). This phenomenon was confirmed in our study, conducted in parallel on four insect species from different orders. On administration of 33, 11.5, 2.3, and 1 mg dsRNA injections to S. litura, L. migratoria, Z. atratus, and P. americana, respectively, the lower treatment doses induced significant reduction in gene expression in L. migratoria, Z. atratus, and P. americana (76%, 78%, and 82%, respectively), but the highest treatment dose did not cause much reduction the expression in S. litura, the decrease being only by 20%. When 30 and 36 mg dsRNA were fed to S. litura and L. migratoria, respectively, no significant RNAi effect was observed (1% and 5% depletion in expression). However, feeding 24 mg dsRNA to Z. atratus and P. americana induced significant (29% and 47%) depletion. It was clear that although the treatment dose was higher, the caterpillar S. litura

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Fig. 8. Incubation of 1 mg dsEGFP (in 1 ml water) with 5 ml of £20 dilution of the midgut juice collected from the indicated insects for 10 min. The control used nuclease-free water instead of the midgut juice. The remaining dsEGFP in the samples was visualized by electrophoresis on a 1% agarose gel. Band quantification was performed using the QuantityOne v4.62 software. Values are relative to the control arbitrarily fixed at 100%. Results are means of at least three independent experiments. Statistical analysis was performed with Student's t-test (mean ± SE; **, p < 0.01).

Fig. 7. dsEGFP degradation upon incubation with insect hemolymph. (A) Incubation of 1 mg dsEGFP (in 1 ml water) with either 5 ml of a serial dilution of the hemolymph from 1. S. litura, 2. L. migratoria, 3. Z. atratus, 4. P. americana, or 5.5 ml nucleasefree water for 1 h. The samples were visualized by electrophoresis on a 1% agarose gel. The collected hemolymph was diluted with nuclease-free water, as indicated; (B) Band quantification of the dsEGFP in 4 hemolymph dilutions was analyzed by using the QuantityOne v4.62 software, the relative band intensity was calculated according to Henriques et al. (2012). Values are relative to the control, arbitrarily fixed at 100%. Results are means of at least three independent experiments. Statistical analysis was performed with Student's t-test (mean ± SE; *, p < 0.05; **, p < 0.01).

showed no obvious response to both the dsRNA injection and feeding, whereas significant RNAi effect could be induced by a lower treatment dose in the cockroach, P. americana, in both the treatment cases. Similar RNAi result was also obtained in the beetle, Z. atratus; however, the interference was a little weaker than in the cockroach. In the locust, L. migratoria, although significant RNAi effect could be induced by the dsRNA injection, no such effect was observed by dsRNA feeding. Thus, the RNAi sensitivity of these four insect species was clearly in the order: P. americana > Z. atratus >> L. migratoria >> S. litura. This suggests that it is easy to induce RNAi in the cockroach, P. americana, and the beetle, Z. atratus, whereas it is much difficult in the caterpillar, S. litura. This is consistent with previous results demonstrating successful RNAi induction in cockroaches and beetles (Miller et al., 2012; Huang et al., 2013; Irles et al., 2013; Wang et al., 2013; Xiao et al., 2015). However, in caterpillars the induction was possible only when much higher doses of dsRNA were injected or fed

continuously (Terenius et al., 2011; Yang and Han, 2014). For locusts, S. gregaria has been reported to be sensitive to dsRNA injection, but not to dsRNA feeding (Wynant et al., 2014b). In the present work, similar results were observed for L. migratoria. Thus, the ingestion system might play an important protective role for such kind of insects. Chemical sensitivity of organisms depends on the inner dosage (target exposure dose and the persistent time integral) after treatment and the target affinity. Insects have an open circulatory system with all their organs immersed in the hemolymph. Thus, the dsRNA content in hemolymph might be an important pharmacological parameter that directly indicates not only the inner doses of dsRNA on the different organs and cells, but also the dynamics of its metabolism in the insects. In the present study, the results obtained in the RNAi experiments showed that treatment doses did not correlate with the depletions among the different insect species. However, monitoring of the hemolymph concentration revealed that the inner hemolymph dsRNA dosage (concentration and persistence) was the key for RNAi effect in different insect species. When the same dose (4 mg) was injected, the initial hemolymph concentration in S. litura, L. migratoria, Z. atratus, and P. americana was 29, 82, 405, and 943 pg/ml, respectively. When the treatment doses were adjusted accordingly to 33, 11.5, 2.3, and 1 mg for S. litura, L. migratoria, Z. atratus, and P. americana, respectively, similar initial hemolymph concentration (310 pg/ml) was achieved; thereafter the hemolymph concentration decreased at different speeds in these insects with the calculated relative speed index being 2.06, 1.44, 1.20, and 1, respectively. Because of long persistence time, the concentration and persistence time integral for different insect species were not calculated and compared. However, it was clear that faster decreasing speed shortened the persistence of hemolymph dsRNA and resulted in considerably lower inner dosages. Thus, the negative correlation (0.9501) between the hemolymph dsRNA decreasing speed index and the RNAi effects implies that the inner hemolymph dsRNA dosage was key for the RNAi effect among the different insect species. A similar phenomenon was observed with dsRNA feeding. The

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results demonstrated that the foreign hemolymph dsRNA in all the tested insects could be detected at around 3e5 min after feeding. However, the peak level and persistence time varied considerably in different insect species. The hemolymph dsRNA contents calculated using hemolymph concentration and persistence time integral were in the ratio 1, 1.17, 2.01, and 2.90 for S. litura, L. migratoria, Z. atratus, and P. americana, respectively. This showed a good correlation (0.9983) with the RNAi depletions induced by feeding dsRNA. Using different experiments, recent studies have shown that the injected dsRNA was rapidly degraded in the tobacco hornworm, Manduca sexta, and the pea aphid, Acyrthosiphon pisum, which are characterized by a low efficacy for RNAi, but persisted much longer in the German cockroach, Blattella germanica, and in the migratory locust, L. migratoria, which are thought to be sensitive to RNAi induced through dsRNA injection (Christiaens et al., 2014; Garbutt et al., 2013; Ren et al., 2014). All these results unanimously demonstrate that the hemolymph dsRNA content might be the key factor for RNAi efficacy. Since RNAi is a cellular process, adequate level of hemolymph dsRNA must persist for sufficiently long time to allow for the cellular uptake. Thus, the inner dosage exposure to targets could be the key for RNAi. Some in vivo factors could influence the inner dosage or the dsRNA content in the hemolymph, including degradation by some extracellular enzymes (Garbutt et al., 2013; Wynant et al., 2014b), adherence by lipophorins (Wynant et al., 2014a), and absorption by cells, in general. In the present study, the faster decrease in hemolymph dsRNA content could not have been caused by adherence by lipophorins, because the adhered dsRNA can be extracted and monitored. Previous reports showed that different insect cell lines varied in their dsRNA absorption. However, RNAi occurs in cells, and fast cell absorption could promote RNAi. If all the adsorbing cells could be the target cells, hemolymph dsRNA decreasing speed caused by absorption should not negatively correlate with the RNAi effect. Our in vitro cell-free hemolymph incubation assays suggested that the degradation activity of some extracellular enzymes was the key factor determining the hemolymph content of the treated dsRNA in the different insect species. Our results have demonstrated that dsRNA degradation capacity varied considerably among the insect species in both hemolymph and midgut juice. Correlation analysis found that in vitro hemolymph degradation capacities highly correlated (0.9350) with the hemolymph dsRNA decreasing speed index after injection. For feeding treatments, the ingested dsRNAs would be exposed to the enzyme in the midgut juice before entering the hemocoel (Whangbo and Hunter, 2008). A few studies have demonstrated that the efficiency of RNAi induced by dsRNA feeding depended on the activity of enzymes in the midgut (Luo et al., 2013; Wynant et al., 2014b). Here, our results demonstrated that the capacity of dsRNA degradation in the midgut juice had a high negative correlation with the hemolymph dsRNA contents (0.9143) after feeding. Thus, the enzymatic degradation of dsRNA should be taken into consideration for efficient use of RNAi in insects. 5. Conclusion In conclusion, the present study evaluated the efficacy of RNAi in four insect species from different orders with respect to the mode of administration of the dsRNA. We report that the RNAi efficacy in the different insect species depends on the enzymatic degradation of dsRNA in the medium around the target cells, which functions as the key factor in determining the hemolymph dsRNA contents after the exposure. In addition, our results suggest that injecting dsRNA is more effective in inducing RNAi than its administration through feeding. We believe that this contribution is theoretically and

practically relevant because it will help in more efficacious induction of RNAi for basic research and applied aspects. Acknowledgments We are grateful to Dr. Guanheng Zhu (NAU Department of Entomology) for his support in raising the insects. We thank Dr. Mohammed Esmail Abdalla Elzaki (NAU Department of Entomology) for his help in the detailed discussion on the manuscript, Prof. Baoping Li (NAU Department of Entomology) and Dr. Ting Sun (NAU National Engineering and Technology Center for Information Agriculture) for their assistance in the statistical analyses and model constructions. This work was supported by the National Natural Science Foundation of China (31130045) to ZJH, the Special Fund for Agro-scientific Research in the Public Interest of China (201303017) to ZJH and Postgraduate Research and Innovation Projects in Jiangsu Province (KYLX15_0621) to KXW. References Christiaens, O., Swevers, L., Smagghe, G., 2014. DsRNA degradation in the pea aphid (Acyrthosiphon pisum) associated with lack of response in RNAi feeding and injection assay. Peptides 53, 307e314. Cirimotich, C.M., Scott, J.C., Phillips, A.T., Geiss, B.J., Olson, K.E., 2009. Suppression of RNA interference increases alphavirus replication and virus-associated mortality in Aedes aegypti mosquitoes. BMC Microbiol. 9, 49. Coy, M., Sanscrainte, N., Chalaire, K., Inberg, A., Maayan, I., Glick, E., Paldi, N., Becnel, J., 2012. Gene silencing in adult Aedes aegypti mosquitoes through oral delivery of double-stranded RNA. J. Appl. Entomol. 136, 741e748. Filipowicz, W., 2005. RNAi: the nuts and bolts of the RISC machine. Cell 122, 17e20. s, X., Richards, E.H., Reynolds, S.E., 2013. Persistence of doubleGarbutt, J.S., Belle stranded RNA in insect hemolymph as a potential determiner of RNA interference success: evidence from Manduca sexta and Blattella germanica. J. Insect Physiol. 59, 171e178. Gillen, C., Akuma, D., Piermarini, P., 2015. Expression pattern and RNAi inhibition of putative sodium-coupled cation-chloride cotransporters in the mosquito Aedes aegypti. FASEB J. 29 (1Supplement), 7, 843. Henriques, A., Carvalho, F., Pombinho, R., Reis, O., Sousa, S., Cabanes, D., 2012. PCRbased screening of targeted mutants for the fast and simultaneous identification of bacterial virulence factors. Biotechniques. http://dx.doi.org/10.2144/ 000113906. Huang, J.H., Lozano, J., Belles, X., 2013. Broad-complex functions in postembryonic development of the cockroach Blattella germanica shed new light on the evolution of insect metamorphosis. BBA-Gen. Subjects 1830, 2178e2187. Huang, S., Han, Z., 2007. Mechanisms for multiple resistances in field populations of common cutworm, Spodoptera litura (Fabricius) in China. Pestic. Biochem. Physiol. 87, 14e22. Irles, P., Silva-Torres, F.A., Piulachs, M.D., 2013. RNAi reveals the key role of Nervana 1 in cockroach oogenesis and embryo development. Insect Biochem. Mol. Biol. 43, 178e188. Jung, J.W., Kim, J.H., Pfeiffer, R., Ahn, Y.J., Page, T.L., Kwon, H.W., 2013. Neuromodulation of olfactory sensitivity in the peripheral olfactory organs of the American cockroach, Periplaneta americana. PLoS One 7, e81361. Kennerdell, J.R., Carthew, R.W., 1998. Use of dsRNA-mediated genetic interference to demonstrate that frizzled and frizzled 2 act in the wingless pathway. Cell 95, 1017e1026. Li, H., Guan, R., Guo, H., Miao, X., 2015. New insights into an RNAi approach for plant defence against piercing-sucking and stem-borer insect pests. Plant Cell Environ. 38, 2277e2285. Lu, Y., Yuan, M., Gao, X., Kang, T., Zhan, S., Wu, X., Wan, W., Li, J., 2013. Identification and validation of reference genes for gene expression analysis using quantitative PCR in Spodoptera litura (Lepidoptera: Noctuidae). PLoS One 8, e68059. Luo, Y., Wang, X., Yu, D., Chen, B., Kang, L., 2013. Conserved repressive function of Krüppel homolog 1 on insect metamorphosis in hemimetabolous and holometabolous species. Differential responses of migratory locusts to systemic RNA interference via double-stranded RNA injection and feeding. Insect Mol. Biol. 22, 574e583. Marciniak, P., Szymczak, M., Pacholska-Bogalska, J., Audsley, N., Rosinski, G., 2013. Identification and localisation of selected myotropic neuropeptides in the ventral nerve cord of tenebrionid beetles. Comp. Biochem. Physiol. A 166, 44e51. Mello, C.C., Conte, D., 2004. Revealing the world of RNA interference. Nature 431, 338e342. Miller, S.C., Miyata, K., Brown, S.J., Tomoyasu, Y., 2012. Dissecting systemic RNA interference in the red flour beetle Tribolium castaneum: parameters affecting the efficiency of RNAi. PLoS One 7, e47431. Mysore, K., Flannery, E.M., Tomchaney, M., Severson, D.W., Duman-Scheel, M., 2013. Disruption of Aedes aegypti olfactory system development through chitosan/ siRNA nanoparticle targeting of semaphorin-1a. PLoS Negl. Trop. Dis. 7, e2215.

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