On Methods of Germ-line Transformation and RNA Interference in African Malaria Mosquito, Anopheles gambiae

J. Asia-Pacific Entomol. 6(2): 111-117 (2003) www.entomology.or.kr

REVIEW

On Methods of Germ-line Transformation and RNA Interference in African Malaria Mosquito, Anopheles gambiae Young S. Hong, Lucas Q. Ton and Frank H. Collins* Center for Tropical Disease Research and Training, Department of Biological Sciences, University of Notre Dame, Notre Dame, IN 46556, U.S.A.

Abstract Germ-linetransformationand RNA interference (RNAi) are useful molecular tools that make functional genetic studies feasible because through these techniques target genes can be altered in vivo and their phenotypes can be investigated. Germ-line transformation allows establishment of a stable transformed line while injected double-stranded RNAi (dsRNAi) transiently knocks out an endogenous gene of interest by post-transcriptionally silencing its transcripts. Recently, both these techniques have been demonstrated workable in Anopheles gambiae, a most deadly African malaria mosquito. Availability of these techniques in A. gambiae is highly expected to help researchers gain more insight into gene functions in A. gambiae. As a result, attempts to control malaria transmission by genetically manipulating A. gambiae can be made in the future. In particular, dsRNAi should be useful to elucidate biological functions of those conceptually predicted genes in the A. gambiae genome as suggested in other organisms such as Caenorhabditis elegans and Drosophila melanogaster. Therefore, we provide the technical details of these two experimental procedures so that they will become accessible to more vector biologists. Key words Germ-line transformation, RNA interference (RNAi), Anopheles gambiae, malaria

Introduction Anopheles gambiae is the major African mosquito vector for the malaria parasite, Plasmodiumfalciparum. Malaria claims 1-2 million human lives and imposes a huge socioeconomic burden on endemic countries in the sub-Saharan region in Africa (WHO, 1998). However, malaria control efforts have been hampered by resistance problems because both malaria parasites and mosquitoes have become resistant to anti-malarial ·Corresponding author. E-mail: [email protected] Tel: +1-574-631-8045; Fax: +1-574-631-3996 (Received September 15, 2003; Accepted September 26, 2003)

drugs and insecticides, respectively. Therefore, alternative control means for malaria transmission are urgently needed. One of the possibilities to abate malaria transmission is to genetically manipulate the mosquito vectors so that they will become incompetent for malaria transmission. For the genetic manipulation of A. gambiae, molecular tools allowing the germ-line transformation have been rigorously sought. However, it was not until recently that A. gambiae was stably transformed using a piggyBac transformation vector (Grossman et al., 2001). Moreover, transient gene silencing was also achieved in the adult A. gambiae by employing double-stranded RNA interference (dsRNAi) (Blandin et al., 2002). For the adult dsRNAi, dsRNA molecules of a defensin gene synthesized by in vitro transcription were injected into the adult female A. gambiae, degraded the corresponding endogenous mRNA and consequently generated a phenotypic knockout of the target molecule (Blandin et al., 2002). The RNAi technique is particularly attractive in that it can mimic functional knockouts of affected genes in vivo in a relatively short time period compared to that of the germ-line transformation. This can lead to more functional studies of the mosquito genes (Barstead, 2001). For example, among the 14,653 predicted genes as of August 2003, many of them still remain to be annotated for their functions (Barstead, 2001). Therefore, having both the germ-line transformation and RNAi techniques available in A. gambiae is very useful for not only the genetic manipulation but also reverse genetic studies of A. gambiae. In several mosquito species, various versions of the technical procedures for germ-line transformation have been successfully used (Coates et al., 1998; Jasinskiene et al., 1998; Catteruccia et al., 2000; Allen et al., 2001; Grossman et al., 2001). In a practical way, germ-line transformation has become a routine molecular procedure in Aedes aegypti and A. stephensi, both having been transformed with different transposonbased vectors. For instance, Ae. aegypti has been transformed by mariner, Hermes and piggyBac vectors (Coates et al., 1998; Jasinskiene et al., 1998; Kokoza et al., 2001); A. stephensi by Minos and

112 J. Asia-Pacific Entomol. Vol. 6 (2003)

piggyBac (Catteruccia et al., 2000; Ito et al., 2002;

Needle preparation

Nolan et al., 2002). However, most of these reports

are lacking technical details, making it difficult for an inexperienced researcher to perform the experiment based on the published descriptions. Therefore, herein we provide the technical details of germline transformation and adult RNAi, focusing mainly on A. gambiae for which germ-line transformation is not yet routine. It is our intention to make these techniques more widely available for mosquito vector biology laboratories so that further improvement of these genetic manipulation or knockout techniques can be made in A. gambiae. Also, some of the procedures provided here can be easily modified to be used for other insect species.

Germ-line Transformation of A. gambiae Basically, the principle of germ-line transformation in A. gambiae is the same as those of other mosquitoes including A. aegypti. Unlike A. aegypti, however, the embryo of A. gambiae is extremely susceptible to dehydration and also appears to be more sensitive to physical stresses like penetration of the injection needle. Therefore, careful handling of the embryo is highly recommended for a successful transformation experiment.

DNA preparation Each plasmid containing helper (or transposase) or target transgene in E. coli host cells can be prepared using a standard alkaline lysis method (Sambrook et al., 1989) with CsCl centrifugation or Qiagen plasmid preparation columns (Valencia, CA). Both the plasmid preparation methods yield high quality DNAs. If the CsCl method is chosen, however, it is recommended to perform the centrifugation step twice to enrich the supercoiled DNA. The plasmid DNAs are resuspended in sterile molecular biology grade (MBG) nucleasefree water. Both helper and transgene plasmids are adjusted to a working concentration (0.3 and 0.5 pglpl, respectively) in IX injection buffer (5mM KCl, 0.1 mM sodium phosphate, pH 6.8). In addition, blue food dye (Durkee Foods, Iowa) can be added to the injection solution, which helps to visualize the DNA solution while it is being injected into embryos. helper DNA

0.3 /lgl/ll

transgene DNA

0.5 /lgl/ll

lOX injection buffer

5 /ll

blue food dye

1 )11

MBG water

Bring to the final volume of 50 ill

Preparing a good needle is a very important step toward a successful microinjection experiment. Therefore, it is worthwhile investing time and effort to optimize the condition to make desirable needles. In general, there are two steps to produce microinjection needles: pulling glass capillaries and beveling needles. Pulling the needle: Aluminosilicated capillaries (World Precision Instruments, Sarasota, FL, cat# ASIOOF-4) are pulled to make two microinjection needles per capillary using a puller. We have used a Sutter puller (Model P-87, Sutter Instrument, Novato, CA) equipped with a tungsten filament for heating the glass capillary. The pulling parameters can vary depending on the preferred shapes of needles. We found a two-cycle method produces suitable needles (see below for details). Since the set-up conditions for each needle puller can differ, it is recommended to configure an adequate condition for pulling desired needles following the manufacturers guide. Generally, if the tip is tapered too thin and long, it will be hard to penetrate the chorion membrane because the tip is too flexible. In contrast, if the tip is tapered too short, the needle is most likely to have a larger opening, causing the cytoplasm of the embryo to leak through the hole after injection. This is detrimental to the embryo. In Figure lA, an example of a proper needle is shown. Once needles are pulled, they can be stored in a commercial (World Precision Instruments, Sarasota, FL, cat# E21O) or a makeshift container that is made using a large petri dish and playdough (Fig. lB). Beveling the needle: Pulled needles need beveling, which produces needles with pointed and consistent openings. Alternatively, one can break the tips of needles by gently touching embryos, but it is difficult to control the sizes of the tip openings. We have used a beveler from Sutter Instrument (Model BV-lO). Needles are beveled by gently touching the revolving grinder supplied with 1% Photo-Flo (Kodak, Rochester, NY, cat# 146 4510). Beveling a needle at an angle between 35-45° has been determined to make suitable openings. In order to help see the needle tips through the microscope mounted on the beveler, the beveler is illuminated with an incandescent lamp. Consequently with illumination, two images can be seen, one the needle itself and the other the shadow of the needle. To bevel needles, the needle should be lowered until the tip is about to touch the grinder, which can be judged by observing the tip of needle and its shadow barely contacting each other. Then the needle is slowly further lowered by turning the fine tuner 180-360°. As the tip of a needle touching the grinder and being

Germ-line Transformation and RNA Interference

113

A

I

11

Fig. 1. (A) Comparison of the needle tips for embryo (top) and adult (bottom) microinjections. The embryo injection needle has a longer taper with a finer tip than the adult needle. (B) Commercial (left) and makeshift (right) needle storage containers. The makeshift container is made with a disposable petri dish and playdough.

beveled, one can see Photo-Flo solution coming into the needle by the capillary force. The speed of influx can be an indicator of how the needle is being beveled. It is, therefore, advised to watch the tip carefully and to bevel the needles as little as possible. After beveling, needles can be back-filled with 95% ethanol. The tips can then be examined to check how ethanol comes out of the tip with application of a positive pressure. A good needle should allow ethanol to smoothly trickle out at the tip with an application of a working injection pressure (15-50 psi). Beveled needles can be stored for several weeks or months prior to actual injection.

(150 mM NaCl, 5 mM KCl; 10 mM HEPES; 2.5 mM CaCb; pH 7.2) containing 0.1 mM p-nitrophenyl p' -guanidinobenzoate (pNPGB) (Sigma-Adrich, St. Louis, MO, Cat# N-8010). pNPGB delays hardening and darkening of the chorionic membrane of the embryo by inhibiting phenoloxidase (Catteruccia et al., 2000). As a result, the chorionic membrane remains soft for an extended period of time. However, one caveat to using pNPGB is that female mosquitoes tend to lay fewer eggs than on plain water. pNPGB can be prepared as lOX solution and diluted to the working concentration with the injection buffer. To maintain freshness, pNPGB also needs to be kept on ice during the injection. Otherwise, it turns yellow from oxidation and should be discarded.

Embryo collection Anopheline mosquitoembryosare particularlyvulnerable to dehydration. Therefore, it is critical to maintain embryos well hydrated during the entire microinjection procedure. In order to collect embryos for injection, about 100 mosquitoes are kept in a cage (~20 x 20 em, diameter x height). It is not necessary to separate males and females. Female mosquitoes about 5 days old post-emergence are fed on anesthetized mammals (i.e., rats, mice or guinea pig) of choice three days before the experiment. If females have a blood meal for the first time, it is usually better for her to have a second blood meal about 1 or two days after the first one. This helps the female mosquitoes lay more eggs. After a blood meal, mosquitoes are kept at 25°C or room temperature with 75-80% relative humidity. On the third day post-blood meal, mosquitoes are dark-adapted for 30-60 min. prior to egg collection. Embryos can be collected for about 30 minutes by putting a petri dish layered with a wet filter paper into the cage kept in the dark. The embryos can be collected repeatedly from a single cage during injection until no embryos are deposited. As an option, the filter paper can be moistened with isotonic buffer

Injecting embryos The preblastoderm stage of the embryo lasts up to 60-90 minutes post-oviposition at room temperature and this stage can be used for germline transformation. Newly laid embryos have a pale milky color, gradually turn gray and eventually become very dark. In general, embryos with light gray or gray color are ideal for injection. On the embryo, the anterior end is wider and rounder than the opposite posterior end that is more pointed (Fig. 2A). It is the posterior end where germ cells form and DNA is to be injected. For a single round of injection, a batch of 20-30 embryos are lined up along the edge of a piece of cut filter paper (Fig. 2A) with the posterior ends placed along the edges. This lining-up procedure should be done on the same petri dish used to collect embryos. The filter paper on the petri dish needs to be well moistened to keep embryos from dehydration. However, excess water can cause embryos to float, making lining-up more difficult. Either a fine soft paintbrush or forceps can be used to handle embryos. Whatever tool is chosen, however, embryo handling

114

J. Asia-Pacific Entomol. Vol. 6 (2003)

requires extreme caution to avoid any physical damages to embryos. After the line-up, embryos are transferred onto a slide glass using a double stick tape (Fig. 2B). Placing the embryos on the tape immobilizes them so that a needle can penetrate the chorion membrane during the injection. This transfer procedure can be performed by first placing a piece of wet filter paper with embryos on an 8-layered Kimwipe'" or similar paper tissues. Embryos are then transferred to a double stick tape on the slide glass by gently pressing down at the long ends of the slide glass (Fig. 2B). The proper amount of pressure to pick up embryos onto the tape without damaging embryos should be learned empirically. The slide is then submerged under water in a petri dish (Fig. 2e). As seen in Figure 2C, one side of the petri dish is cut to make an access for the needle to the embryos. Alternatively, embryos can be covered with Halocarbon oil 27 (Sigma-Aldrich, H-8773) and the slide glass can be placed on the injection microscope. However, hydrocarbon oil should be removed after injection. Otherwise, embryos may suffocate, lowering the survival rate. The injection

needle is maneuvered to be on the same focal plane as embryos using a manual (Narishige, MN-151) or motorized micromanipulator (Eppendorf, Model 5171) (Fig. 2D). Embryos are injected by gently moving them toward the needle connected to a microinjector (Narishige, Model 1M 300). After penetration, it is very critical to release DNA solution into embryos with a pressure as minimal as possible without clogging the needle. The injection pressure may be adjusted for each needle every time a new needle is used to achieve an optimum pressure. When a needle penetrates an embryo, the cytoplasm may influx into the needle due to the positive internal pressure of the embryo. This could result in clogging the needle. To prevent this, a balance pressure is applied to the needle that can offset the internal pressure of embryos. The balance pressure of 5-10 psi can be used to prevent the cytoplasm influx. During the injection, some embryos may not be possible to inject mainly due to their loose attachment to the tape. Those uninjected embryos should be discarded. Otherwise, they will hatch and cause unnecessary screening for transformants.

B posterior

D

Fig. 2. (A) A. gambiae embryos lined up for microinjection. It is the posterior end (bottom) where a transgene is injected. (B) Lined embryos are being transferred onto a double-stick tape on a slide glass. The slide needs to be gently pushed down to avoid rupturing the embryos. (C) The slide with embryos is placed under water in the injection container (yellow). (D) The embryos can be injected by moving them toward the needle.

Germ-line Transformation and RNA Interference

Post-injection care and screening for transformants After injection, embryos on the tape can be kept underwater in a petri dish until the first instars to hatch, which usually takes about 2-3 days at 2YC. The first instars are transferred to a new container and reared until they become adults. It is advised not to overfeed the first instars. Overfeeding the first instars tends to promote bacterial growth in the water, reducing the survival of the larvae. However, the later larval stages (i.e., 2nd through 4th instars) can be fed more freely. In fact, the fourth instars require to consume a large amount of food (see Appendix for the composition). In about 10-12 days post-hatching, the pupae are picked and kept in a cup inside an adult cage. The emerging adults (GO) are sexed and each sex is out-crossed en mass to the opposite sex of the background stock that is used for the injection. Out-crossing is necessary because of the swarm mating behavior of A. gambiae. The resulting progeny (G 1) can be screened for transformants according to the marker used.

dsRNAi in the adult mosquitoes Our dsRNAi method for the adult A. gambiae is based on that of Blandin et al. (2002) with some modifications. Major alterations include the use of ice and food dye. Ice is used to keep mosquitoes anesthetized during injection after a brief initial exposure to C02, and food dye in the injection solution helps to visualize the DNA solution. In our own experience, these changes have not affected the survivorship of injected mosquitoes. However, experimenters may test this procedure so that it can better serve their experimental purposes.

115

polymerase bind to its priming sites on the amplified fragments, resulting in a higher yield of in vitro transcription of DNA templates. In addition, the template DNA in LITMUS 28i can be linearized using two different restriction enzymes, one with BgIII and the other with Stu 1. This will generate two DNA templates, each with a T7 promoter-binding site either in anti- or sense orientation. Using the prepared DNA fragments as template, dsRNA can synthesized in a single reaction by in vitro transcription using a MEGAscript® T7 kit according to the manufacturer s recommendation (Ambion, cat# 1626). The resulting dsRNA is cleaned up by column purification provided with the kit. The dsRNA is resuspended in MBG water in the final concentration of 0.5-1.0 ug/ul. A food color can be added to the dsRNA about 1/30 of the dsRNA solution. This is now ready for injection.

Injection of dsRNA into adult mosquitoes Mosquitoes are anesthetized with a brief exposure to CO2 and kept on the ice-cold stage to extend immobilization during the injection (Fig. 3A). The injection stage can be made using a petri dish with a wet filter paper and placed on an ice container. With shorttapered needles (Fig. lA), dsRNA is injected into the ventral thorax area near the neck (Fig. 3B) using a standard dissecting microscope with illumination. An

Preparation of dsRNA A DNA fragment of interest is cloned into a plasmid vector with a common priming site flanking both ends of the cloning site (i.e., T7 or SP6) such that the insert can be amplified using a single primer of choice. We have used LITMUS 28i (NEB, N3528S) vector with the T7 promoter sequence flanking the multicloning site. The insert can be PCR-amplified using the T7 primer (TAATACGACTCACTATAG) following a standard PCR method. Alternatively, the insert can be prepared using LITMUS sequencing primers (forward, CTGCAGGATATCTGGATCCAC and reverse, GTGGATCCAGATATCCTGCAG), which provide additional bases at both upstream ends of the T7 RNA polymerase priming sites. This can help T7 RNA

B

Fig. 3. (A) dsRNAi microinjection into the adult female A. gambiae. The mosquitoes are anesthetized with brief exposure to C02 and kept on an ice-chilled petri dish during the injection. (B) A female mosquito is being injected in the ventral forethorax area. The dsRNAi solution contains blue food dye to help visualize the solution in the mosquito.

116 J. Asia-Pacific Entomol. Vol. 6 (2003)

injection pressure can be set between 15-25 psi with 5 psi back-pressure. After injection, mosquitoes are transferred to a cage supplied with sugar-cotton balls on top of the cage screen. After a recovery time of about 48 hours post-injection, mosquitoes can be checked for the phenotype(s) of interest. In addition, reduction of the target transcript can be checked using real-time quantitative PCR such as TaqMan® technology (Applied Biosystems Inc., CA).

Perspectives For the functional genomics studies in A. gambiae, germ-line transformation and RNAi are imperative techniques. As shown in the A. gambiae genome sequencing, quite a large number of genes still remain to be annotated. Particularly to this end, RNAi is an invaluable asset to investigate gene functions because it does not require two generation times before examining the phenotypes of target genes. For example, in D. melanogaster functions of CG5652 transcript were previously unknown but its functions have recently been demonstrated in both embryo and adult flies using dsRNAi (Dzitoyeva et al., 2003). When silenced in the embryo, CG 5652 resulted in partial or complete loss of denticle belts of the abdominal segments A2-A4. Accordingly, CG5652 was named beltless. In the adult, silencing of beltless caused the shrinkage of ovaries. Nonetheless, RNAi via dsRNA injection into the adult mosquito or any other developmental stages is a transient knockout method lacking a stably transformed line. In contrast, germ-line transformation will allow us to obtain a stable transformant line. The feasibility of stable germ-line transformation of A. gambiae has been demonstrated by using the piggyBac element (Grossman et al., 2001). However, transformation efficiency is less than 0.1% of injected embryos (per. comm. W. Kim). To be used more practically, transformation efficiency needs to be improved, which may require development of alternative transformation vectors and further optimization of the transformation procedures. Acknowledgment The authors wish to thank Dr. A. Sarkar and C. Sim for their critical reading of the paper. This work was supported by a grant, P01-AI45123-02 from NIHINIAID to F.H.e.

Literature Cited Allen, M.L., D.A. O'Brochta, P.W. Atinson and C.S. Levesque. 2001. Stable, germ-line transformation of Culex quinquefasciatus (Diptera: Culicidae). J. Med. Entomol. 38: 701-710. Barstead, R. 2001. Genome-wide RNAi. Curro Opin. Chern. Bio!. 5: 63-66. Blandin, S., L.F. Moita, T. Kocher, M. Wilm, F.C. Kafatos and E.A Levashina. 2002. Reverse genetics in the mosquito Anopheles gambiae: targeted disruption of the Defensin gene. EMBO Rep. 3: 852-856. Catteruccia, F., T. Nolan, T.G. Loukeris, C. Blass, C. Savakis, F.e. Kafatos and A Crisanti. 2000. Stable germline transformation of the malaria mosquito Anopheles stephensi. Nature 405: 959-962. Coates, C. J., N. Jasinskiene, L. Miyashiro and AA. James. 1998. Mariner transposition and transformation of the yellow fever mosquito, Aedes aegypti. Proc. Nat!. Acad. Sci. USA 95: 3748-3751. Dzitoyeva, S., N. Dimitrijevicand H. Manev. 2003. Identification of a novel Drosophila gene, beltless, using injectable embryonic and adult RNA interference (RNA i). BMC Genomics 4: 33. Grossman, G.L., C.S. Rafferty, lR. Clayton, T.K. Stevens, O. Mukabayire and M.Q. Benedict. 2001. Germline transformation of the malaria vector, Anopheles gambiae, with the piggyBac transposable element. Insect Mol. Bio!. 10: 597-604. Hoffman, S.L., G.M. Subramanian, F.H. Collins and J.C. Venter. 2002.Plasmodium, humanand Anopheles genomics and malaria. Nature 415: 702-709. Ito, J., A Ghosh, L.A Moreira, E.A Wimmer and M. Jacobs-Lorena. 2002. Transgenic anopheline mosquitoes impaired in transmission of a malaria parasite. Nature 417: 452-455. Jasinskiene, N., C.J. Coates, M.Q. Benedict, AJ. Cornel, C.S. Rafferty, A.A James and F.H. Collins. 1998. Stable transformation of the yellow fever mosquito, Aedes aegypti, with the Hermes element from the housefly. Proc. Nat!. Acad. Sci. USA 95: 3743-3747. Kokoza, V., A. Ahmed, E.A Wimmer and AS. Raikhel. 2001. Efficient transformation of the yellow fever mosquito Aedes aegypti using the piggyBac transposable element vector pBac[3xP3-EGFP afm]. Insect Biochem. Mol. BioI. 31: 1137-1143. Nolan, T., T.M. Bower, A.E. Brown, A Crisanti and F. Catteruccia 2002. piggyBac-mediated germline transformation of the malaria mosquito Anopheles stephensi using the red fluorescent protein dsRED as a selectable marker. J. BioI. Chern. 277: 8759-8762. Sambrook, J., E.F. Fritsch and T. Maniatis. 1989. Molecular Cloning: A laboratory manual, Ed. Cold Spring Harbor. Cold Spring Harbor Laboratory Press. New York. WHO. 1998. Fact Sheet No. 94, Ed. Geneva: World Health Organization.

Germ-line Transformation and RNA Interference

Appendix A. Reagents - Injection Buffer(5 mM KCI, 0.1 mM sodium phosphate, pH 6.8) - Isotonic Buffer(l50 mM NaCI, 5 mM KCI; 10 mM HEPES; 2.5 mM CaCh; pH 7.2) - lOX p-nitrophenyl p'-guanidinobenzoate (pNPGB, 1.0 mM) • Dissove 33.7 mg pNPGB in I ml Dimethyl Sulphoxide (DMSO). • Add pNPGB/DMSO solution into 99 ml isotonic buffer. • Aliquot lOX pNPGB and store at-20'C until use. • When use, thaw lOX pNPGB on ice and dilute it to the working concentration. - Mosquito food (preparation for 150 g) • Whole wheat flour 90.0 g 37.5 g • Baker's yeast

117

• Beef Blood (defibrinated) 15.0 g (lCN, cat# 105081) • Non-fat dried milk 7.5 g • Mix well the above ingredients and keep it refrigerated.

B. Suppliers' Web Pages Arnbion: http://www.ambion.com/ Applied Biosystems lnc.: http://www.appliedbiosystems.com! ICN Biochemicals: httpi//www.icnbiomed.com/ Kodak Company: http://www.kodak.com! Narishige Instrument: http://www.narishige.co.jp/ New England Biolabs: http://www.neb.com/ Qiagen: httpt//www.sigmaaldrich.com/ Sigma-Aldrich Chemicals: Sutter Instrument: http://www.sutter.com! World Precision Instruments: http://www.wpiinc.com/