Introduction

Current high-throughput DNA sequencing technologies and improved gene databases have facilitated the molecular analyses of many non-model insects, including pest species. However, gene functional analyses available for non-model insects are quite limited. RNA interference (RNAi) is an invaluable, widely used tool for gene silencing in various non-model insects (Futahashi et al. 2011; Miyazaki et al. 2014; Ohde et al. 2013). However, the efficiency of RNAi is low or very variable in some insects (Christiaens et al. 2014; Tomoyasu et al. 2008). Genome editing technologies such as TALENs and CRISPR/Cas9 have recently become popular for manipulating target genes in insect genomes (Ma et al. 2014; Watanabe et al. 2012); however, their use is restricted to early embryos. The phenotypic effects should then be checked at particular developmental stages. Therefore, in some species, it is difficult to microinject RNA into the eggs without significantly damaging their growth.

In vivo electroporation introduces plasmid DNA encoding a gene of interest or short-hairpin RNA into various tissues, enabling both gain-of-function and loss-of-function analyses. This technique is useful for the rapid functional analyses of non-model insects (Ando and Fujiwara 2013; Golden et al. 2007; Kunieda and Kubo 2004). In vivo electroporation is very powerful technology; however, a special and expensive device system (around US$10,000 in general) is required for the method. Ando and Fujiwara (2013) reported a low-cost (under $200) handmade electroporator. However, it is not easy for non-experts to construct it because it requires a program with C language to control voltage pulse duration, and an additional special power supply such as that used in electrophoresis.

To overcome these limitations, we have developed a simple, very inexpensive, easily constructed, all-in-one electroporation device for the gene functional analyses of insects. Here, we report the design and features of the device. Using the simple electroporator, we successfully introduced exogenous fluorescence protein genes into silkworm, Bombyx mori (Linnaeus), and two important pest species, adzuki bean borer moth, Ostrinia scapulalis (Walker) and corn borer moth, Ostrinia furnacalis (Guenée).

Materials and methods

Figure 1 is a circuit diagram of the electroporation device. All electronic parts (listed in Table 1) and their connecting wires are soldered onto the circuit board. A schematic of the in vivo electrode of the device is presented in Fig. 2a. When the toggle switch is on, a direct current flows through Circuit 1, turning on the blue LED, and showing a voltage on the voltmeter (Fig. 1). The electrode voltage is controlled by operating the potentiometer, while checking the voltage registered on the voltmeter. Depressing the footswitch reroutes electricity to the electrodes in Circuit 2, switching off the LED and the voltmeter in Circuit 1.

Fig. 1
figure 1

Circuit diagram of the simple electroporation device. Details of all designators (except GND) are shown in Table 1. The ground (GND) connects to the negative terminal of the PWR1

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Table 1 Parts list
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Fig. 2
figure 2

E lectroportation of O. scapulalis larva. a Schematic illustration of the in vivo electrode. b The ring-shaped electrode easily holds phosphate buffered saline (PBS) when dipped in the buffer. PBS colored with an orange dye was used in this image for easy understanding. c Platinum electrodes holding PBS droplets were placed near the injection site and on the opposite side of the body and were supplied with electric pulses

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The B. mori strain Kinsyu × Showa (Ueda-Sanshu), and field-collected O. scapulalis and O. furnacalis were maintained on an artificial diet (Silkmate 2 M, Nihon–Nosan) at 23 ± 1 °C in a long-day regime of 16 h light and 8 h dark.

A vector and helper plasmids pPIG-A3GR and pHA3PIG system allows stable expression of exogenous enhanced-green (EGFP) and red fluorescence protein (DsRed2) genes in targeted tissues (Ando and Fujiwara 2013; Tamura et al. 2000). Injection of both plasmid solution and electroporation were conducted as described by Ando and Fujiwara (2013) with some modifications. Injection was performed under a dissecting microscope using a glass capillary needle (GD-1, Narishige) made by a glass microcapillary puller (PN-31, Narishige) connected to an air pump (FP15 N, Tokyo Glass Kikai). The capillary was inserted into third instar nymphs of the three moth species that were anesthetized in a stream of carbon dioxide. Approximately 0.5 µl of each donor and helper plasmid solution (1 µg/µl) was co-injected into the body cavity by using air pressure. Immediately after injection, platinum electrodes holding phosphate-buffered saline (PBS) droplets (Fig. 2b) were placed on the larval body: one near the injection site, the other on the opposite side of the body (Fig. 2c). The larvae were loaded with five 15-V pulses (each lasting approximately 250 ms) by manually controlling the simple electroporation device with the footswitch.

All insect images were taken with a digital camera (DFC310 FX, Leica) connected to a fluorescence stereomicroscope (M205FA, Leica).

RNA was isolated from the electroporated sites of O. scapulalis and O. furnacalis 5 days after electroporation using RNeasy plus Mini Kit (QIAGEN). First strand cDNAs were synthesized using pd(N)6 primers and PrimeScript reverse transcriptase (Takara-Bio). Quantitative reverse transcription-PCR (RT-PCR) was performed as previously described (Kayukawa and Ishikawa 2009) using a Mx3005P QPCR system (Agilent) and THUNDERBIRD SYBR qPCR MIX (Toyobo) with primers EGFP-q-F1 (5′-GGGCATGGCGGACTTGAAGA-3′) and EGFP-q-R1 (5′-ACGGCAAGCTGACCCTGAAG-3′) for EGFP, with primers DsRed2-q-F1 (5′-TGCAGGACGGCTGCTTCATC-3′) and DsRed2-q-R1 (5′-TTCAGGGCCTTGTGGGTCTC-3′) for dsRed2, and with primers Osef1α-q-F1 (5′-GACTCCGGCAAGTCCACCAC-3′) and Osef1α-q-R1 (5′-CCTGGGCCTCCTTCTCGAAC-3′) for elongation factor 1α (ef1α). The PCR program consisted of 98 °C for 1 min, followed by 50 cycles of 98 °C for 10 s, 62 °C for 15 s, 68 °C for 30 s, with final dissociation curve analysis. The relative expression levels were normalized by the transcript levels of ef1α.

Results and discussion

The construction of the simple electroporation device is shown in Fig. S1. The device can be constructed in 3 h from materials costing under US$85 (Table 1), and it requires no additional power supply, unlike the previous device reported by Ando and Fujiwara (2013). Whereas commercially available in vivo electrodes are often inappropriately large for application to small insects, the size and shape of our electrodes can be adjusted to the target insects. Moreover, our ring-shaped electrode readily holds the PBS (Fig. 2b) that prevents epidermal burn injury. The potentiometer controls the electrode voltage output from 1.5 to 22.5 V. When the more than 32 V AC adaptor is used, a maximum voltage of 30.5 V can be outputted. Appropriate adaptors can realize a 100–240 V input, compliant with the power supplies of many countries.

Using this simple electroporation device, we first confirmed the successful expression of exogenous fluorescence proteins in the larval epidermis of B. mori by fluorescence microscopy (Fig. S2). We then applied the same method to two important pest and model species in chemical ecology, O. scapulalis and O. furnacalis (Ishikawa et al. 1999; Lassance 2010). The larval epidermis of both species electroporated with EGFP and dsRed2 exhibited green and red fluorescence (Figs. 3a and S3a). DsRed2 fluorescence was more strongly observed in both Ostrinia species. Expressions of the exogenous fluorescent genes were confirmed by quantitative RT-PCR analyses in the electroporated sites in both Ostrinia species (Fig. S4). As expected, no fluorescence was observed in the no-injection control or in the negative control with distilled water (data not shown). The insects injected with the donor and helper plasmids, but not treated with an electric pulse, showed no fluorescence (Figs. 3b and S3b) and very low mRNA expressions of EGFP and DsRed2 were detected (Fig. S4). Twenty-five days post-treatment, the survival rates were not significantly lower in electroporated insects [O. scapulalis, 69.2 % (18/26); O. furnacalis, 65.2 % (15/23)] than in non-electroporated controls [O. scapulalis, 86.7 % (13/15); O. furnacalis, 80.0 % (13/15)]. These results indicate the effectiveness of our electroporation device for the somatic transformation of the examined moth species.

Fig. 3
figure 3

Fluorescence microscopy analysis of overexpressed exogenous EGFP and dsRed2 in O. scapulalis epidermis. a 9 days after electroporation and b non-electroporated controls injected with the donor and helper plasmids

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The simple electroporation device presented in this study has several good properties: e.g. low cost, easily constructed and universally applicable. A previous study indicated that various insect tissues are successfully electroporated by 20–30 V pulses; namely, the larval epidermis and pupal wing of B. mori, and the larval epidermis of the Asian swallowtail butterfly, Papilio xuthus (Linnaeus) and the red flour beetle, Tribolium castaneum (Herbst) (Ando and Fujiwara 2013). This voltage range is within the usable range of our electroporator, suggesting that our device is applicable not only to the moths but also to diverse insect species. Thus far, electroporation methods that enable gain-of-function analyses have required expensive equipment. This simple low-cost electroporator will render gene function analyses accessible to many non-model insects, including pest species.