Peptides 53 (2014) 307–314

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DsRNA degradation in the pea aphid (Acyrthosiphon pisum) associated with lack of response in RNAi feeding and injection assay Olivier Christiaens a , Luc Swevers b , Guy Smagghe a,∗ a b

Department of Crop Protection, Faculty of Bioscience Engineering, Ghent University, 9000 Ghent, Belgium Insect Molecular Genetics and Biotechnology, Institute of Biosciences & Applications, NCSR “Demokritos”, Aghia Paraskevi, Athens, Greece

a r t i c l e

i n f o

Article history: Received 28 October 2013 Received in revised form 18 December 2013 Accepted 18 December 2013 Available online 3 January 2014 Keywords: Insects Hemiptera Acyrthosiphon pisum Aphids RNAi dsRNA degradation

a b s t r a c t Over the past decade, RNA interference (RNAi), the sequence-specific suppression of gene expression, has proven very promising for molecular research in many species, including model insects as Tribolium castaneum and Apis mellifera. It showed its usefulness to analyze gene function and its potential to manage pest populations and reduce disease pathogens. However, in several insects, the efficiency of RNAi is low or very variable at best. One of the factors that could influence RNAi efficiency in insects is degradation of dsRNA after administration to the insect. In this paper, we report on the importance of dsRNA breakdown in the pea aphid (Acyrthosiphon pisum) associated with the absence of an RNAi response upon oral feeding and injection with dsRNA targeting different genes such as the ecdysone hormone receptor and ultraspiracle. In essence, we discovered that both the salivary secretions of aphids and the hemolymph were able to degrade the dsRNA. In parallel, introduction of dsRNA in the aphid body was not able to provoke a response in the expression of the siRNA core machinery genes. © 2013 Elsevier Inc. All rights reserved.

1. Introduction Over the last decade, the post-transcriptional silencing technique RNA interference (RNAi) has proven to be an intriguing prospect for molecular research as well as possible applications in pest management and reduction of disease pathogens. It was first discovered in the nematode Caenorhabditis elegans at the end of the last century and allows a very specific as well as a strong and systemic silencing effect of the targeted gene in that animal [14]. The technique has since been used successfully as a functional genomics tool in a vast array of other species, including the red flour beetle Tribolium castaneum, the honeybee Apis mellifera, the cockroach Blattella germanica and the fruitfly Drosophila melanogaster [4,9,21,29,30,33,36,41,44]. However, a systemic and long term effect including an amplification of the RNAi signal similar to the one that is observed in nematodes, does not seem to occur in insects [44]. Additionally, the success rate of RNAi in insects seems to be quite variable between different species, target genes and delivery methods [34,42,48]. Over the last few years, more and more research is being conducted toward some of the factors that determine RNAi efficiency in these arthropods, such as dsRNA uptake [18], dsRNA degradation [2,16,26], expression of

∗ Corresponding author. Tel.: +32 92646150. E-mail address: [email protected] (G. Smagghe). 0196-9781/$ – see front matter © 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.peptides.2013.12.014

RNAi machinery genes [17,26,44], virus interaction [15,25] and the effect of the target gene and dsRNA fragment itself [3,6]. Aphids are sap-sucking insects of great economic importance, and recently the pea aphid Acyrthosiphon pisum had its genome sequenced and annotated [43]. Therefore, RNAi would be a very interesting tool to add more biologically relevant information to all these sequence data. However, our experience as well as those from other laboratories (Douglas and Carolan, personal communications) indicated that RNAi in the pea aphid is difficult to achieve, despite a few publications reporting successful RNAi in the pea aphid [19,31,35,47]. In this research paper, we first report on a number of RNAi experiments targeting the insect molting hormone receptor (i.e. ecdysone receptor; EcR) and ultraspiracle (USP) which are two essential nuclear receptors involved in growth and development (including molting), to evaluate the responsiveness of aphids for RNAi [8]. We also tested dsRNAs targeting C002 and vATPase E subunit, as silencing of these two target genes has been reported before [31,47]. In a next step, we investigated the importance of degradation of dsRNA. This was done with three bioassays where we tested breakdown of dsRNA: first by aphid salivary secretions in the artificial diet containing dsRNA, and second by incubation of dsRNA ex vivo with hemolymph which was collected from the aphid body hemocoel. Third, we tested the degradation of dsRNA in vivo under the natural conditions present inside the insect body. To do this, we used a qPCR approach to follow the degradation of injected GFP dsRNA upon injection in the aphid hemocoel. The expression of a

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Table 1 Primers used for dsRNA synthesis and for quantitative RT-PCR. Fragment

Forward

Reversed

Fragment size

dsEcR dsUSP dsvATPase dsC002 dsGFP qTubulin qActin qRPL7 qEcR qDcR2 qAgo2 qR2D2 qEri1 qSid1 qGFP

a

a

390 412 185 620 455 100 122 76 184 137 199 140 187 197 100

a

GACCTCCCGAAGAGCTTTGT a CAATGGGTCCTCAGTCACCT a TTAGCCAACACTGGAATAAACGTC a GGGAAGTTACAAATTATACG a TACGGCGTGCAGTGCT TGGACAATCAGGTGCTGGA CGTGAAAAGATGACCCAAATC TTGAAGAGCGTAAGGGAACTG CAACTGTCATTCAGTCGGTTT AACCCATCCAACCTACCAAT AGAGATGGTGTAAGCGAAGG CAAAAATCGCCTTGTTCCTC ATGGCTCGGTTTCTTTATGG TTATGCAATGGGATCAGCAC CCGACCACTACCAGCAGAAC

TAGGCGTGCCCATTATCATT a TGCACGGCTTCTCTTTTCAT a CCAAACAGTCCATGCATATTATT a CTCCCATAGCCATCTTG a TGATCGCGCTTCTCG CTCGGCTTCTTTCCTCACAA CCAGAGTCCAAAACGATACCA TATTGGTGATTGGAATGCGTTG TTTTCTCCACTTTCCAACCA CAGTTATTTCACCAGGAGTTTTGTG TGCCAGAAGGGACATTAGAAA CCACATGCTTGGCTTCTTTT GAGGGTTGCCTTCAAATTCC CAAATGCCAAAACACCAAAC TTGGGGTCTTTGCTCAGG

Only gene-specific parts of the primer are listed. These are preceded by the T7 adaptor TAATACGACTCACTATAGGG for dsRNA synthesis.

number of RNAi-related genes was also investigated, namely dicer2, argonaute-2, r2d2 and sid-1 [4,7,17,37–40,42]. Here we speculate that introduction of dsRNA causes an up-regulation of these genes in RNAi sensitive insects. In the case of no up-regulation, this would imply there is no activation of the siRNA pathway. Finally, we searched for the presence of an endocellular nuclease, Eri-1, in the aphid genome. Such Eri-1 enzyme is present in nematodes where it acts as a dsRNase [20]. Subsequent phylogenetic analysis was done to evaluate whether this aphid Eri-1-like nuclease belongs to the same subclass as the Eri-1 in nematodes, or to a different group or Eri-1-like nucleases to which the homologs in holometabolic insects such as Tribolium and Drosophila belong. The data contribute to a better understanding of the importance of breakdown and cellular uptake of dsRNA in the success of RNAi. 2. Materials and methods

In a second experiment, dsEcR, dsUSP and dsGFP (control) were delivered to second-instar aphid nymphs through microinjection. Twenty nanoliters of 4 ␮g/␮L dsRNA solutions were injected into each aphid and then aphids were placed on V. faba bean leaves. The following five days, potential effects on mortality and molting were scored. Finally, in a third experiment dsC002 (microinjection) and dsvATPase E subunit (feeding) were administered to pea aphids according to the same procedures which were used in the publications by Mutti et al. [31] and Whyard et al. [47]. DsvATPase was administered to neonates through feeding while the dsC002 was injected into adults. In both cases, the same primers were used for dsRNA synthesis and the same amounts/concentrations were administered to the aphids. In the COO2 experiment, the longer dsRNA was used instead of the pre-diced siRNA as previously [31,47].

2.1. Aphid colony

2.4. RNA isolation and real-time qPCR

Aphids were taken from a continuous culture at Ghent University which is maintained on Vicia faba bean plants at standardized conditions of 23 ◦ C, 45% RH and a 16:8 photoperiod [7]. Neonates were collected by placing adults on fresh faba bean leaves for a few hours and collecting the newborn aphid nymphs.

For the expression analysis studies, total RNA was extracted using the Rneasy Mini kit (Qiagen, Venlo, Netherlands), following the manufacturer’s instructions. The RNA concentration was quantified and verified using a Nanodrop ND-1000 (Thermo Scientific, Zellik, Belgium) and 1.5% agarose gel electrophoresis. Subsequently, cDNA was synthesized using the Superscript II kit (Invitrogen, Merelbeke, Belgium). Real-time qPCR was performed in a CFX96TM Real-Time System and the CFX Manager software (both from Bio-Rad, Nazareth, Belgium). Primers used for RT-qPCR are listed in Table 1; these were all validated with a standard curve based on a serial dilution of cDNA to determine efficiency and a melt curve analysis with temperature range from 60 ◦ C to 95 ◦ C to ensure specificity. Reactions of 20 ␮L were performed in duplicate or triplicate and contained 10 ␮L SsoFastTM EvaGreen Supermix (Bio-Rad), 0.4 ␮L of 10 ␮M forward primer (Invitrogen), 0.4 ␮L of 10 ␮M reverse primer (Invitrogen), 8.2 ␮L MilliQ water and 1 ␮L cDNA. Three reference genes, rpl7, actin and ˛-tubulin were chosen based on previous reports [19,35] and our optimization. A “no template control” and “no RT control” reactions were included to exclude interference by foreign DNA or genomic DNA. The relative transcript levels were normalized to the amount of the three reference genes. Amplification conditions were 3 min at 95 ◦ C followed by 39 cycles of 10 s at 95 ◦ C and 30 s at 58 ◦ C.

2.2. Synthesis of dsRNA DsRNA was synthesized using the MEGAScript dsRNA synthesis kit (Ambion, Huntingdon, UK) according to the manufacturer’s instructions. After purification, elution from the column was done with nuclease-free water instead of the elution solution which is supplied in the kit. The transcription reaction was allowed to proceed overnight. The primers used for the synthesis are listed in Table 1. 2.3. RNAi experiments with A. pisum For the RNAi experiments, both feeding as well as a microinjection were used to administer the dsRNA to aphids of different stages. In a first experiment, dsEcR was administered to neonate aphid nymphs via a feeding assay adapted from Sadeghi et al. [32]. The final concentration of dsRNA in the diet was 200 ng/␮L. Water and dsGFP were used as controls. Nymphal survival and development were followed daily for 3 days while they were supplied with diet ad libitum.

2.5. Degradation of dsRNA in feeding artificial diet To test the stability of dsRNA in the artificial diet used for the RNAi bioassays, feeding cages were setup containing 130 ␮L of

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artificial diet. To this artificial diet, 40 ng of dsEcR was added. In each feeding cage, 15 neonate aphids were placed to feed on the diet. Three cages were used per dsRNA, and in the controls 3 feeding cages with dsRNA-diet but without aphids were used. After 84 h, dsRNA was extracted from the pooled diets with the RNeasy kit (Qiagen) and the dsRNA loaded on a 1.5% agarose gel. 2.6. Degradation of dsRNA ex vivo in collected hemolymph Hemolymph was collected from about 500 adult aphids as sampled ad random in colony. This was done by amputating one or more legs and then collecting the droplets of hemolymph coming out with microcapillaries. Hemolymph was collected in phenylthiourea (PTU) buffer in order to avoid melanization and kept on ice. Once enough hemolymph was collected, hemocytes were removed by centrifuging at 1000 × g for 8 min and collecting the cell-free supernatant. An amount of 500 ng dsEcR was incubated in either 3.5 ␮L of RNase-free water or 3.5 ␮L cell-free hemolymph at 23 ◦ C. Upon ex vivo incubation, samples were collected after 1, 3 and 12 h, and dsRNA was extracted using the RNeasy kit (Qiagen) and run on a 1.5% agarose gel. 2.7. Degradation of dsRNA in vivo in aphid body We used a qPCR-based method based on Garbutt et al. [16], to detect the persistence of intact dsGFP inside the aphid body. Thirty third–fourth-instar nymphs were injected with 180 ng dsGFP and whole body samples were taken at 0.5, 1, 2, 5 and 24 h after injection and stored in RLT-buffer supplemented with ␤-mercaptoethanol (Qiagen kit), at −80 ◦ C prior to RNA extraction. Subsequently, total RNA was extracted using the RNeasy Mini Kit (Qiagen) and a 10 min 75 ◦ C denaturing step was added directly preceding the reverse transcription reaction in order to denature the secondary structure of the dsRNA. Finally, cDNA was synthesized from this RNA according to the same methods described in Section 2.4, except that random hexamers were used instead of oligodT primers. Primers used for the qPCR are listed in Table 1 and qPCR methods and conditions were as described above. This experiment was performed 3 times independently. 2.8. Expression of RNAi-related genes upon injection of dsRNA To investigate whether or not the siRNA pathway is activated after dsRNA introduction in the aphid, a qPCR expression analysis was performed for the core machinery genes Dicer-2 (XP 003240109.1), Argonaute 2 (XP 001944852.2), R2D2 (NP 001155644.1) and Sid-1 (XP 001951907.1). These genes were identified in the pea aphid genome database and primers were designed for qPCR. Second-instar nymphs of A. pisum were injected with 50 ng dsEcR and then placed on V. faba leaves. Injection with water was used as a control. Samples of 5 pooled aphids were taken after 6 and 24 h. The RNA extraction, cDNA synthesis and qPCR were performed according to the methods described in Section 2.4 and the primers used are listed in Table 1. This experiment was 3 times performed independently. 2.9. Phylogenetic analysis of Eri-1 nucleases The amino acid sequences of Eri-1-like nucleases from Homo sapiens, Strongylocentrotus purpuratus, C. elegans, D. melanogaster, T. castaneum and A. pisum were retrieved from GenBank and aligned using Clustal W. The alignment of the exonuclease domain was used for the phylogenetic analysis using the maximum parsimony algorithm. These analyses were performed using Mega 5. The GenBank accession numbers are mentioned in the figure. The expression of

Fig. 1. DsRNA degradation by salivary secretions during feeding by the pea aphid (Acyrthosiphon pisum). Forty nanograms of dsEcR were added to 130 ␮L artificial diet and the stability of the dsRNA in the diet on which pea aphids were feeding was compared to that in the diet on which no aphids were feeding. Afterwards, dsRNA was purified from the diet using the RNeasy kit (Qiagen) and run on a 1.5% agarose gel.

Eri-1 was also analyzed after injection with dsRNA following the same methods as in Section 2.8. 3. Results 3.1. RNAi shows no response in the pea aphid The RNAi experiments conducted with dsEcR and dsUSP showed no effect on both the survival and development (molting) of treated aphids compared to the control group which was fed with diet containing the same concentration of dsGFP. For the dsEcR feeding experiment, no mortality was observed in both control and treatment groups and no difference was seen in the molting behavior as well. The control group went through 1.76 molts over three days, while the dsEcR fed aphids went through 1.70 molts over the same period. Also for the injection experiment, where dsEcR and dsUSP injection was compared to a control group (dsGFP injected), no mortality was observed after 5 days and also no molting defects were observed. The group injected with dsUSP, dsEcR and dsGFP had molted 2.33, 2.04 and 2.04 times on average, respectively. The RNAi experiments targeting the genes coding for C002 and vATPase E subunit lacked any phenotypic effect. DsvATPase-fed neonates showed only 10% mortality after 7 days which was the same as the control group. In contrast, Whyard et al. [47] had recorded 50% mortality after 7 days with the same concentration of dsRNA in the diet. DsC002-injected adults in our experiments also did not show any difference in mortality compared to the control group, which was injected with dsGFP, while Mutti et al. [31] showed that aphids injected with the C002 died much more rapidly than the control group. 3.2. Degradation of dsRNA in feeding artificial diet The dsRNA present in the diet on which aphids had been feeding for 84 h demonstrated a clear smear on agarose gel (Fig. 1), while in the controls (i.e. without aphid feeding) there were no signs of degradation. These results indicate that salivary secretions from the aphids while feeding, caused breakdown of dsGFP. 3.3. Degradation of dsRNA ex vivo in collected hemolymph In this experiment, dsGFP was incubated ex vivo in collected aphid hemolymph. As shown in Fig. 2, there was a rapid and strong

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used to follow the expression of a number of genes that code for proteins which are considered to be involved and even essential in this siRNA pathway. The sequences of four of these genes, namely the endoribonuclease Dicer-2, Argonaute-2 and R2D2 which are two partners of the RISC complex, and also Sid-1, which is involved in cellular uptake of dsRNA were retrieved from the database and their expression was analyzed after dsRNA injection into the aphid hemocoel. However, as shown in Fig. 4, none of these RNAi-related genes showed any up-regulation after dsRNA introduction compared to the control, for which aphids were injected with water. 3.6. Phylogenetic and transcriptional analysis of A. pisum Eri-1 Fig. 2. Ex vivo degradation of dsRNA by pea aphid (Acyrthosiphon pisum) hemolymph. An amount of 500 ng dsEcR was incubated in either 3.5 ␮L of RNase-free water during 12 h or in 3.5 ␮L cell-free hemolymph as collected from adult aphids, during 1, 3 and 12 h. Afterwards, dsRNA was extracted using RNeasy kit (Qiagen) and run on a 1.5% agarose gel.

degradation of dsRNA. Already after 1 h, there was clear dsRNA degradation, and after 3 h, the dsRNA band had totally disappeared and only a smear of smaller fragments was left. 3.4. Degradation of dsRNA in vivo in aphid body Given our discovery that an ex vivo incubation in collected hemolymph could rapidly degrade dsRNA, it was important to investigate whether a dsRNA degradation pattern could also be observed in vivo in the natural conditions inside the hemocoel of an aphid body and whether the speed of degradation was similar. This in vivo dsRNA persistence experiment confirmed that dsRNA is not staying intact in the aphid body for a long time. Fig. 3 presents the levels of intact dsGFP left in the whole body samples after injection into the hemocoel, and it is clear that these amounts already drop within hours after injection. At 5 h post-injection, these levels have fallen down to about 5% of those recorded at 0.5 h after injection, and at 24 h, these were almost undetectable, confirming that most – if not all – of the dsRNA was degraded. 3.5. Expression of RNAi-related genes upon injection of dsRNA In order to investigate if a response of the siRNA pathway could be observed after introduction of dsRNA into the aphid, qPCR was

Fig. 3. In vivo persistence of dsRNA in the pea aphid (Acyrthosiphon pisum). 180 ng dsGFP was injected into L3/L4 nymphs and the fate of dsRNA was followed by a qPCR approach. Samples were taken at the different time points, RNA was extracted and the sample was incubated at 75 ◦ C for 10 min just before reversed transcription to denature the dsRNA. The levels of remaining dsGFP in the cDNA were then analyzed with PCR. Data is normalized against a reference sample. Values are mean ± SEM based on three biological replicates.

Besides extracellular breakdown in the insect hemocoel and gut, intracellular degradation of dsRNA can also play an important role. Phylogenetic analysis of the intracellular Eri-I-like nucleases, representing sequences from H. sapiens, the sea urchin S. purpuratus, the nematode C. elegans and the three insects Tribolium, Drosophila and A. pisum was performed and revealed three distinct subclasses: Eri-1/3 Exo, Snipper and Pint 1 (Fig. 5). The phylogenetic analysis clearly showed that the A. pisum Eri-1 homolog that was identified in the genome (NP 001155946.1) belongs to the group of the Eri-1/3 Exo nucleases, which is contrary to the homologs found in Drosophila and Tribolium, which group together in the Snipper group. Additionally, we investigated the expression of Eri-1 and found that no up-regulation was evident after introduction of dsRNA in the aphid body (Fig. 6). 4. Discussion The results of the pea aphid RNAi experiments reported here showed a clear lack of silencing. Attempts to silence EcR and USP, two nuclear receptors which form the functional receptor for the insect molting hormone ecdysone, did not result in any phenotypic effects on either survival or molting. In contrast, previous experiments with other insect species in which these hormone receptors were silenced, for example in T. castaneum and Helicoverpa armigera, resulted in clear molting defects and also in mortality of the insect juveniles [41,49]. Following these experiments, attempts were made to repeat previously published and successful knockdown experiments in the pea aphid, which targeted the genes coding for C002 and the vATPase E subunit [31,47]. In these publications, both experiments led to clear mortality effects upon silencing and these experiments could therefore act as a positive control in our RNAi experiments. However, administering dsC002 and dsvATPase subunit E according to the same methods to our aphids did not result in the same mortality effects as were reported in those publications [31,47], indicating that RNAi in our pea aphid strain is insensitive. RNAi insensitivity has been observed in many insect species, and several possible factors could contribute to this. Our investigations on dsRNA degradation in the aphid diet showed that aphid salivary products which are produced during feeding, are able to degrade dsRNA. While there still remains an amount of dsRNA intact in the feeding diet after 84 h, it is important to note that this degradation has only been caused by the salivary secretions brought into the artificial diet by the aphid and that the concentrations of these digestive products will most likely be much higher within the aphid’s digestive tract while the amount of dsRNA would be much lower. Previously, Allen and Walker [1] demonstrated a similar effect by salivary products on dsRNA in another hemipteran, the tarnished plant bug Lygus lineolaris. The authors collected saliva as well as extracts from the salivary glands, and both were shown to be able to degrade dsRNA rapidly, while hemolymph did not. Based on these data, it is difficult to speculate on the speed of intestinal

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Fig. 4. Relative expression of four RNAi-related genes (Dicer-1, Argonaute 1, R2D2 and Sid-1) in Acyrthosiphon pisum after injection with 50 ng dsEcR. Water was injected in the control population. After injection, the L2 nymphs were placed on Vicia faba leaves and samples were taken after 6 and 24 h for RNA extraction (5 individuals were pooled per sample). Afterwards, cDNA was synthesized and the expression of the four RNAi-related genes was analyzed by real-time qPCR, using tubulin, RPL7 and actin as reference genes. Values are mean ± SEM based on three biological replicates.

dsRNA degradation in aphids and further research investigating the stability of dsRNA in the gut itself should be carried out. Experiments in other species do suggest that some insects exhibit a very strong degradation of dsRNA in their digestive system [1,27]. An important consequence of this degradation by salivary secretions is that RNAi experiments using artificial diets where a constant supply of dsRNA is provided, should be designed carefully and the dsRNA-containing diet should be replaced regularly. Our results on the degradation of dsRNA in hemolymph and the in vivo degradation experiment are indicative that dsRNA can only stay intact in the hemocoel of the aphid body for a short period of time. This could have played an important role in the experienced difficulties achieving RNAi in the pea aphid, since successful RNAi requires enough dsRNA to persist in the hemolymph sufficiently long to allow uptake into the cells. Bolognesi et al. [6] recently reported that the dsRNA fragment length had to be longer than 60 bp for successful silencing of the Snf7 gene in Diabrotica virgifera virgifera corn rootworms. Interestingly, it turned out to be sufficient that 21 bp of those 60 or more base pairs were target gene-specific, suggesting that the importance of the fragment length is not linked to the RNAi machinery itself but might be linked for example to the uptake of the dsRNA by the cells. Therefore, a rapid degradation of

dsRNA to smaller fragments could lead to a lower RNAi efficiency. The speed of degradation in the hemocoel that was observed here in aphids is in agreement with an earlier report [16] where 200 ng dsRNA targeting moricin was degraded within 2–3 h in the tobacco hornworm Manduca sexta, which is another insect characterized by a low efficiency for RNAi. The most likely cause of this dsRNA degradation in the aphid body is the presence of ribonucleases targeting dsRNAs. However, so far not much is known about dsRNases in insects. In the silkmoth Bombyx mori, a DNA/RNA non-specific alkaline nuclease has recently been described [2,26] and this enzyme is expected to contribute to the rapid and strong degradation of dsRNA which was observed in the digestive system of this caterpillar, an insect which is known for its insensitivity for RNAi. While this protein was originally thought to be expressed in the insect midgut only, Liu et al. [26] demonstrated that this protein is in fact present in other tissues of B. mori larvae as well, including epidermis, fat body, thoracic muscles, Malpighian tubules, brain and silk glands. A search in the pea aphid genome using this lepidopteran nuclease as a template revealed the presence of two nuclease-coding genes that seem related to the one found in B. mori by Liu et al. [26]. We named the two nucleases ApDsNucl1 (XP 001945502.1) and ApDsNucl2

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Fig. 5. Phylogenetic analysis of the Eri1-like nucleases-containing sequences. The maximum parsimony tree is based on the alignment of the exonuclease domain from related sequences found in Homo sapiens, Strongylocentrotus purpuratus, Caenorhabditis elegans, Drosophila melanogaster, Tribolium castaneum and Acyrthosiphon pisum. The Eri-like nucleases cluster together into three distinct subclasses: Eri-1/3 hExo, Snipper and Pint 1. The A. pisum Eri-1-like nuclease is an Eri-1/3 hExo type nuclease, while Drosophila and Tribolium only have Snipper-type nucleases. Accession numbers for the sequences that were used are given on the tree.

(XP 001950681.1). Both exhibit a typical DNA/RNA non-specific endonuclease domain. Further research toward these two dsRNases will have to elucidate their potential role in dsRNA breakdown in the pea aphid. Another nuclease that is worth looking into is the Eri-1 homolog that we could identify in the pea aphid genome. In C. elegans, this nuclease appears to cause tissues to become refractory to RNAi [20]. Tomoyasu et al. [44] recently found a similar ortholog in Tribolium and compared this to the ones that are known to be present in C. elegans, D. melanogaster and a number of other species. These authors discovered through phylogenetic analysis that the Eri-1 homologs in Drosophila and Tribolium do not actually belong to the subgroup of the real Eri-1/3 hExo group to which the C. elegans and human Eri-1 nucleases belong, but rather to the related Snipper (SNP) subclass of nucleases. While these SNP-nucleases

Fig. 6. Relative expression of the Eri-1 gene in Acyrthosiphon pisum after injection with 50 ng dsEcR. Water was injected in the control population. After injection, the L2 nymphs were placed on Vicia faba leaves and samples were taken after 6 and 24 h for RNA extraction (5 individuals were pooled per sample). Afterwards, cDNA was synthesized and the expression Eri-1 was analyzed by real-time qPCR, using tubulin, RPL7 and actin as reference genes. Values are mean ± SEM based on three biological replicates.

are also capable of cutting both RNA and DNA, in Drosophila they seem to play no role in RNAi [23] and the presence of this nuclease does not seem to prevent Tribolium from exhibiting a very strong and systemic RNAi effect. Our own phylogenetic analysis demonstrated that A. pisum-Eri-1 does belong to the Eri-1/3 Exo subclass, in contrast to the homologs found in Tribolium and Drosophila. Therefore it is possible that this Eri-1 homolog in the pea aphid is capable of interfering with the RNAi process as it does in C. elegans. This enzyme is considered to be intracellular in C. elegans, and is therefore not linked to the observed hemolymph degradation in the pea aphid. However, we believe that this enzyme could be a part of a broader range of strategies this aphid has to deal with dsRNA. Expression studies performed in this research showed no up-regulation of this gene after introduction of dsRNA in the pea aphid. However, we want to remark here that it is possible that there is a constant high level expression of this gene in the pea aphid. Further research toward this enzyme should investigate whether it is involved in dsRNA breakdown or not. As far as we know, not much more is known about dsRNases in insects. Garbutt et al. [16] have performed a first analysis of the possible dsRNase present in M. sexta hemolymph. They found that preheating of the plasma to 100 ◦ C for 10 min strongly inhibited the ability of the hemolymph to degrade dsRNA. Furthermore, they demonstrated that EDTA has the ability to inhibit the degradation as well, indicating that a metal-dependent enzyme is responsible for the degradation of dsRNA in the hemolymph plasma of M. sexta [16]. Whether this is a homolog of one of the nucleases we have identified in the genome of the pea aphid has to be further investigated. Besides the presence of nucleases, we believe that other factors such as pH may influence the dsRNA stability inside the aphid’s body as well. The lepidopteran gut is known to be very alkaline with pH’s up to 12 [12,13] and dsRNA is unstable in these alkaline conditions. While not as extreme as in lepidopterans, the midgut of aphids is alkaline as well, having a pH ranging from 7.5 to 8.5 at several zones [10].

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Another possible cause of RNAi insensitivity was proposed by Belles [4] who suggested that a low response (up-regulation) of core RNAi genes after dsRNA introduction could also play a role. The expression of a selection of RNAi core machinery genes. In these experiments, pea aphids were injected with dsEcR and then the expression of Dicer-2, Argonaute 2 and R2D2, three crucial genes in the siRNA pathway, was investigated. Additionally, the expression of Sid-1 was investigated as well because this protein is known to play a role in cellular uptake of dsRNA in nematodes and in some insects as well [44,45]. No up- or down-regulation of these genes was observed at 6 and 24 h after injection of dsEcR, which is contrary to the results of similar experiments in some other insect species, including the rather insensitive Lepidopteran M. sexta [17,28], where dicer-2 and argonaute-2 were up-regulated 79–395-fold and 8–27-fold, respectively, depending on the specific tissue 6 h after injection. Interestingly, in the cockroach, which is characterized by an efficient RNAi response [4], injection of dsDcr2 induced two types of responses, a fast one that involved an increase of Dcr2 mRNA expression (as in Manduca), followed by a slower response that corresponded to the silencing of Dcr2 [28]. In the shrimp Litopenaeus vannamei, a 9-fold up-regulation of dicer-2 was detected 9 h after injecting Poly(C-G), a dsRNA analog [7]. One possible explanation for these results is that the RNAi machinery has not been activated, possibly because the dsRNA molecules did not reach the cells, either due to degradation and/or perhaps due to a lack of uptake into the target cells. Indeed, Terenius et al. [42] and Yu et al. [48] previously reported on the importance of cellular uptake. In the future, it will be of interest to investigate whether aphid cells are able to take up dsRNA molecules from their environment. This can be done in uptake experiments using labeled dsRNA and confocal microscopy similar to those previously reported [18]. Another possible explanation for the observed lack of up-regulation of these RNAi-related genes is that the expression of these genes is already at a very high level due to viral interactions for example. Viral infections have been shown to up-regulate these RNAi genes in a number of species [7,40]. The presence of viruses could then not only activate the RNAi machinery but also have saturated the machinery resulting in a lack of RNAi response. This interaction and the possible consequences for RNAi efficiency have been discussed recently [40]. Since some laboratories do seem to have some success with RNAi in the pea aphid, virus interaction could explain the variation in RNAi efficiency between laboratories. To investigate this, it would be interesting to look into these different aphid cultures, compare efficiency in different strains from different laboratories and investigate whether or not there are difference in expression of RNAi-related genes and in the virus population that is present in these cultures. Since RNAi is still relatively new as a research tool, not much is known about the many factors that could affect its efficiency in insects. In agreement with previous suggestions [34], more research toward these factors, such as dsRNA degradation, virus interaction, dsRNA uptake, and target genes and tissues is needed, especially given this technique’s potential as a possible tool in crop protection and in reducing disease pathogens as well. Therefore, more ring testing of protocols and use of positive (reference) controls could help to standardize this RNAi research more. More research toward novel ways to deliver the dsRNA or siRNA could also help in solving these problems. In mammals, serum nucleases are a major hindrance as well in the development of RNAi-based drugs. Recently, there have been some developments in this area, such as the development of nanoparticles and transfection agents used for intravenous delivery of siRNA in the bloodstream. Examples are the use of liposomes, micelles, solid lipid nanoparticles, chemically modified siRNAs (e.g. Stealth RNAiTM ) or the recently developed polyethyleniminebased in vivo-JetPIE® and STICKY siRNATM [5,11,22,24,46]. Perhaps

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