The authors would like to thank all the members of the "Insect Genetics and Genomics" Laboratory, in particular to Lorenzo Ghiringhelli who has worked at developing, adapting and maintaining the rearing of the medfly over the past thirty years. Part of the representative results of this paper have been reprinted from N. Biotechnology, 25(1) by Scolari F. et al., Fluorescent sperm marking to improve the fight against the pest insect Ceratitis capitata (Wiedemann; Diptera: Tephritidae), 76-84, 2008, with permission from Elsevier (License number 3796240759880). This work received support from Cariplo-Regione Lombardia "IMPROVE" (FS).

1. Experimental Set-up
<ol>
	<li>Insectary requirements
	<ol>
		<li>Maintain all C. capitata life stages at 25 &#176;C, 65% humidity and 12/12 hr light/dark photoperiod.</li>
		<li>Place about 1,500-2,000 medfly pupae in a 6 L cage. Use a cage with a brass mesh on one side with holes small enough to stimulate oviposition42. Insert a sponge strip through a small opening in the cage&#39;s base to provide flies with water by means of capillary action. Use a mixture of yeast and sugar (1:10) as food source for the adults (Figure 1A). Increase the amount of yeast (up to 1:3, yeast:sugar) should the females fail to lay eggs.</li>
		<li>Place a plate filled with water below the cage to collect the eggs deposited through the mesh. As virgin females can oviposit, wait until the flies are at least 6-7 days old before collecting eggs for the microinjection experiments. This will ensure that the females will have mated and that the eggs will have been fertilized.</li>
		<li>Collect the eggs by filtering the water with a fine net strainer. Transfer the eggs from the strainer in a plastic box containing standard larval food (1.5 L H2O, 100 ml HCl 1%, 5 g broad-spectrum antimicrobial agent dissolved in 50 ml ethanol, 400 g sugar, 175 g demineralized brewer&#39;s yeast, 1 kg soft wheat bran) using a Pasteur pipette.</li>
		<li>Place each box in a transparent container closed with a net lid to allow air circulation and containing a layer of bran to favor pupation (Figure 1B).</li>
		<li>After about 10 days, check for 3rd instar larvae that emerge from the larval food and jump down into the bran layer for pupation.</li>
		<li>24 hr later, collect the pupae in a small cup using a soft brush and transfer them to a new adult cage. Adults normally emerge 10 days after pupation. The emerging fly uses the ptilinum, an inflatable pocket located on its head above the base of the antennae, to force off the end of the puparium. After emergence, the ptilinum permanently collapses back inside the head.</li>
	</ol>
	</li>
	<li>Preparation of 1 L food for microinjected larvae
	<ol>
		<li>Dissolve 2.5 g agar in 300 ml H2O.</li>
		<li>Warm 500 ml H2O on a hot stirring plate and dissolve 4 g Sodium Benzoate, 4.5 ml 37% HCl, 42 g yeast extract and 115 g dehydrated carrot powder.</li>
		<li>Add to the mixture a solution of 2.86 g broad-spectrum antimicrobial agent dissolved in 10 ml absolute ethanol.</li>
		<li>Distribute the medium in 15 cm round Petri dishes and allow to cool. If necessary, store at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Slide preparation
	<ol>
		<li>Place a 2 cm strip of double-sided tape on a microscope slide (75 x 26 mm2) and mark the edges with a china marker, to prevent oil spread and the consequent desiccation of the injected embryos.</li>
	</ol>
	</li>
	<li>Nucleic Acid Preparation
	<ol>
		<li>Isolate plasmid DNA using commercially available kits34. Precipitate the plasmid using ethanol and re-suspend in injection buffer (0.1 mM phosphate buffer pH 7.4, 5 mM KCl) at the desired concentration.</li>
		<li>Purify in vitro synthetized long dsRNA using phenol-chloroform, precipitate using isopropanol and re-suspend in injection buffer43.</li>
	</ol>
	</li>
</ol>
2. Embryo Preparation
<ol>
	<li>Place a tray filled with water under a cage containing 6-7 day old adults.</li>
	<li>Allow the females to oviposit for 30 min and then collect the embryos by filtering the water using a fine strainer. Always keep the strainer in water to avoid desiccation of the embryos.</li>
	<li>Dechorionate the embryos by immerging the strainer for 5 sec in a commercial 50% bleach solution.</li>
	<li>Wash the embryos carefully by repeatedly immerging the strainer in clean ultrapure water (at least 4-5 times). Finally, place the strainer in clean ultrapure water. Use the embryos within maximum 2 hr. 
	NOTE: The timing of microinjection is related to the necessity to inject into pre-blastoderm embryos, during a phase of nuclear divisions prior to cellularization. This allows the injected DNA to be taken-up into the nuclei, specifically into the primordial germ cell nuclei that will originate the gametes. In Drosophila, cellularization at the pole cells begins at about 90 min after fertilization (at 25 &#176;C), with blastoderm formation occurring about 30 min later44, whereas in the medfly cellularization takes place between 9 and 12 hr45,46.</li>
	<li>Embryo orientation in rows
	<ol>
		<li>Using a fine paintbrush, collect about 50 embryos and place them on a disk of black filter paper soaked with water.</li>
		<li>Under a dissecting stereomicroscope, arrange the embryos in a row on the upper surface of the double-sided tape on the microscope slide using a fine paintbrush (000) (Figure 1C). 
		NOTE: Align the embryos all in the same orientation, as injection is performed in the posterior pole, where germ-line will originate; the posterior pole is opposite to the micropyle region.</li>
		<li>Cover the embryos with a layer of chlorotrifluoroethylene oil making sure that the oil does not spill over china marker borders (see 1.3). 
		NOTE: Before covering the embryos with oil, a further step can be performed: the embryos can be desiccated for a few minutes to reduce their liquid content and thus facilitate the entry of the injected DNA solution. This may help avoid embryo bursting during injection or excessive leakage of the injected solution, as a consequence of the high liquid pressure.</li>
	</ol>
	</li>
</ol>
3. Embryo Microinjection
<ol>
	<li>Fill a needle (1-2 &#181;l) with plasmid DNA or dsRNA solution (1-2 &#181;g/&#181;l) using a micropipette.</li>
	<li>Connect the needle to the microinjection apparatus, which controls the injection time (0.5 sec), the pressure (800 hPa) and the backpressure (150 hPa).</li>
	<li>Perform microinjection under a stereomicroscope, using a micromanipulator to control the movements of the needle (Figure 1D). Use the microscope stage to move the slide.
	<ol>
		<li>Place the slide on the stage of the scope.</li>
		<li>Insert the tip of the needle into the posterior pole of the embryo ( Figure 1E ).</li>
		<li>Inject the solution by activating the microinjection system with the pedal. The injected liquid induces an increase in internal pressure, resulting in a slight movement of the embryo.</li>
		<li>Release the pedal and immediately, but gently, remove the needle from the embryo.</li>
		<li>Observe a little cytoplasmic leakage at the injection site. 
		NOTE: To discriminate between potential lethal effects of DNA/dsRNA/siRNA and mortality due to the microinjection procedure itself, control experiments must be performed. These comprise the injection of buffer only, or, in the case of reverse genetics experiments, of a green fluorescent protein (GFP)-derived double-stranded RNA (dsRNA-GFP), or siRNA.</li>
	</ol>
	</li>
	<li>Following injection incubate the embryos at 25 &#176;C.</li>
</ol>
4. Post-injection Procedures
<ol>
	<li>Two days after injection, check the slides for hatched larvae (Figure 1F) under a stereomicroscope. Larvae can move in the oil but need to be transferred to larval food as soon as possible. For this reason, check the slides several times a day.</li>
	<li>Using a fine paintbrush (000), gently transfer the hatched larvae to the carrot-based larval food. Transfer up to a maximum of 200 larvae per each Petri dish.</li>
	<li>Incubate the larvae at 65% humidity (25 &#176;C). Ensure that the lid does not stick to the dish, impairing air exchange.</li>
	<li>Keep the Petri dishes with the lid mostly closed so that there is still air flow to the larvae. To avoid excessive drying, check the dishes daily and, if needed, add water to compensate for evaporation.</li>
	<li>Place the Petri dishes in a clear container closed by a net-lid and containing a layer of bran to favor pupation. As for routine insectary rearing, 3rd instar larvae jump out of the larval food and pupate in the bran.</li>
	<li>Collect the pupae and place them in a small cage for adult rearing and screening (this is Generation 0, G0). Rear the progeny on standard larval food.</li>
	<li>In the case of germ-line transformation experiments, check for possible transformed individuals in the next generation (G1), as injected individuals usually are chimeras. Transposition events, indeed, might have occurred in any of the nuclei of the embryonic syncytium, which will form either the germ-line or the soma.
	<ol>
		<li>Soon after emergence, anesthetize G1 individuals using CO2 and check for transformation events under an epifluorescence stereomicroscope. To screen for the presence of red fluorescence, use a DsRedwide filter (Ext. 546/12; Emm. 605/75), while to screen for green fluorescence use the EYFP filter (Ext. 500/20; Emm. 535/30).</li>
		<li>Place transformed individuals in a small cage with wild-type adults of the opposite sex and originate a new, initially heterozygous, strain.</li>
	</ol>
	</li>
</ol>

The authors declare that they have no competing financial interests.

Microinjection of nucleic acids in insect embryos is a universal technique that facilitates both the analysis of gene function and biotechnological applications.
The recent publication of the genome sequences from an increasing number of insect species leads to an urgent need for tools for the functional characterization of genes of yet unknown function. RNA-interference has proven to be one of the most valuable methods to infer molecular functions49 and embryo microinjection facili...

The Mediterranean fruit fly (medfly) Ceratitis capitata is a cosmopolitan species that extensively damages fruits and cultivated crops. It belongs to the Tephritidae family, which includes several pest species, such as those belonging to the genera Bactrocera and Anastrepha. The medfly is the most studied species of this family, and it has become a model not only for the study of insect invasions1, but also for optimizing pest management strategies2.
The medfly is a multivoltine species that can attack more than 300 species of wild and cultivated plants3,4. The damage is caused by both the adults and the larval stages: mated females pierce the surface of the fruit for oviposition, allowing microorganisms to affect their commercial quality, whereas the larvae feed on the fruit pulp. After three larval stages, larvae emerge from the host and pupate into the soil. Ceratitis capitata displays an almost worldwide distribution, including Africa, the Middle-East, Western Australia, Central and South America, Europe, and areas of the United States5.
The most common strategies to limit medfly infestations involve the use of insecticides (e.g., Malathion, Spinosad) and the environmentally-friendly Sterile Insect Technique (SIT)6. The latter approach involves the release into the wild of hundreds of thousands of males rendered sterile by exposure to ionizing irradiation. The mating of such sterilized males to wild females results in no progeny, causing a reduction in population size, eventually leading to eradication. Although SIT has proven effective in multiple campaigns worldwide, its major drawbacks include the high costs of rearing and sterilizing millions of insects to be released. Marking of released individuals is necessary to distinguish sterile from wild insects captured in the field during monitoring activities and it is currently achieved using fluorescent powders. These procedures are costly and have undesirable side-effects7.
In order to optimize and/or to develop more effective approaches for the control of this pest, medfly biology and genetics have been widely explored by numerous researchers worldwide. The availability of the medfly genome sequence8,9, will facilitate novel investigations on gene functions. RNA interference is a powerful tool for such studies and it can be achieved through the microinjection of dsRNA (double-stranded RNA) or siRNA (small interfering RNA). This technique has been used, for example, to demonstrate that the sex determination molecular cascade in C. capitata is only partially conserved with respect to that of Drosophila10.
The development of protocols to microinject medfly embryos allowed C. capitata to be the first non-Drosophilid fly species to be genetically modified. As its eggs are similar to those of Drosophila, both in terms of morphology and resistance to desiccation11, the protocol to deliver plasmid DNA into pre-blastoderm embryos first developed for D. melanogaster12,13 was initially adapted for use in C. capitata. These first experiments allowed medfly germ-line transformation based on the transposable element Minos11. Subsequently, the original system was modified14 using other transposon-based approaches. This is the case of&#160;piggyBac from the Lepidoptera Trichoplusia ni15. The protocol has since been further optimized and this has permitted the transformation of other tephritid species16-21 and also of many other Diptera22-31. All these systems rely on the use of a typical binary vector/helper plasmid transformation system: artificial, defective transposons containing desired genes are assembled into plasmid DNA and integrated into the genome of the insect by supplying the transposase enzyme32. A number of transgenic medfly lines have been generated, with multiple features including strains carrying a conditional dominant lethal gene that induces lethality, strains producing male-only progeny and thus not requiring additional sexing strategies, and strains with fluorescent sperm, which may enhance the accuracy of the SIT monitoring phase33-37. Although the release in the wild of transgenic organisms has occurred in pilot tests against mosquitoes only38,39, at least one company is evaluating a number of transgenic medfly strains for their use in the field40.
Embryo microinjection can also favor the development of new genome-editing tools, such as transcription activator-like effector nucleases (TALENs), clustered regularly interspaced short palindromic repeats (CRISPR) / CRISPR associated protein 9 nuclease (Cas9) and homing endonucleases genes (HEGs), which will enable novel evolutionary and developmental studies, as well as expanding the available biotechnological toolbox. Genome-editing approaches already allowed the generation of gene-drive systems in mosquitoes41, and their transfer to the medfly is imminent. Here we describe a universal protocol for microinjecting nucleic acids in medfly embryos that can be useful for all the above mentioned applications.

<table><tbody><tr><td>1 x injection Buffer</td><td></td><td></td><td>0.1 mM phosphate buffer pH 7.4, 5&amp;nbsp;mM KCl</td></tr><tr><td>Construct Plasmid</td><td></td><td></td><td></td></tr><tr><td>Helper Plasmid</td><td></td><td></td><td></td></tr><tr><td>dsRNA</td><td></td><td></td><td>Phenol-Chloroform purified</td></tr><tr><td>Standard Larval food</td><td></td><td></td><td>1.5 L H&lt;sub&gt;2&lt;/sub&gt;O, 100 ml HCl 1%, 5 g broad-spectrum antimicrobial agent used in pharmaceutical products dissolved in 50&amp;nbsp;ml of ethanol, 400 g sugar, 175 g demineralized brewer&#039;s yeast, 1 kg soft wheat bran</td></tr><tr><td>Carrot Larval Food</td><td></td><td></td><td>2.5 g agar, 4 g sodium benzoate, 4.5 ml 37% HCl, 42 g yeast extract, 115 g carrot powder, 2.86&amp;nbsp;g broad-spectrum antimicrobial agent , water to 1 L</td></tr><tr><td>Adult Food</td><td></td><td></td><td>yeast extract and sugar (1:10)</td></tr><tr><td>Microscope slides</td><td>Sigma-Aldrich</td><td>Z692247</td><td></td></tr><tr><td>Injection needles&amp;nbsp;</td><td>Eppendorf</td><td>5242956000</td><td></td></tr><tr><td>Microloaders</td><td>Eppendorf</td><td>5242956003</td><td></td></tr><tr><td>Double slided tape</td><td></td><td></td><td></td></tr><tr><td>Whatman Black circle paper</td><td>Sigma-Aldrich</td><td>Z695505</td><td></td></tr><tr><td>Bleach</td><td></td><td></td><td>Diluite 1:2 before use</td></tr><tr><td>Paintbrush (000)</td><td></td><td></td><td></td></tr><tr><td>Micromanipulator</td><td>Narishige</td><td>MN-153</td><td></td></tr><tr><td>Microinjector</td><td>Eppendorf</td><td></td><td>Femtojet</td></tr><tr><td>Adult cages</td><td></td><td></td><td></td></tr><tr><td>Chlorotrifluoroethylene oil</td><td>Sigma-Aldrich</td><td>H8898</td><td></td></tr><tr><td>&lt;em&gt;Ceratitis capitata&lt;/em&gt;</td><td></td><td></td><td>The strain used is ISPRA</td></tr></tbody></table>

delivery of nucleic acids through embryo microinjection in the worldwide agricultural pest insect, ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Here we report two applications of embryo microinjection directed at the functional characterization of a gene of interest (Case 1), and at the generation of transgenic strains (Case 2), respectively.
Delivery of dsRNA into embryos to unravel gene function.
The innexin-5 gene encodes a gap-junction that, in insects, is expressed specifically in the male and fe...

Watch this Scientific Journal Video about Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata at JoVE.com

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

delivery-nucleic-acids-through-embryo-microinjection-worldwide

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8.9K Views.  University of Pavia. The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

Video: Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

Laser-assisted Cytoplasmic Microinjection in Livestock Zygotes

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can create genetic modifications that will be present in every cell of an adult organism. It can also be used to deliver siRNA, mRNAs or blocking antibodies to study gene function in preimplantation embryos. The conventional technique for microinjecting embryos used in rodents consists of a very thin micropipette that directly penetrates the plasma membrane when advanced into the embryo. When this technique is applied to livestock animals it usually results in low efficiency. This is mainly because in contrast to mice and rats, bovine, ovine, and porcine zygotes have a very dark cytoplasm and a highly elastic plasma membrane that makes visualization during injection and penetration of the plasma membrane hard to achieve. In this protocol, we describe a suitable microinjection method for the delivery of solutions into the cytoplasm of cattle zygotes that has proved to be successful for sheep and pig embryos as well. First, a laser is used to create a hole in the zona pellucida. Then a blunt-end glass micropipette is introduced through the hole and advanced until the tip of the needle reaches about 3/4 into the embryo. Then, the plasma membrane is broken by aspiration of cytoplasmic content inside the needle. Finally, the aspirated cytoplasmic content followed by the solution of interest is injected back into the embryonic cytoplasm. This protocol has been successfully used for the delivery of different solutions into bovine and ovine zygotes with 100% efficiency, minimal lysis, and normal blastocysts development rates.

This protocol shows how to perform cytoplasmic microinjection in farm animal zygotes. This technique can be used to deliver any solution into the one-cell embryo such as genome editing tools to generate knockout animals.

Cytoplasmic microinjection into one-cell embryos is a very powerful technique. As an example, it enables the delivery of genome editing tools that can ...

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex determination, venom synthesis, and host-symbiont interactions, among others. A major limitation of working with this organism is the lack of effective protocols to perform directed genome modifications. An important part of genome modification is delivery of editing reagents, including CRISPR/Cas9 molecules, into embryos through microinjection. While microinjection is well established in many model organisms, this technique is particularly challenging to perform in N. vitripennis primarily due to its small embryo size, and the fact that embryonic development occurs entirely within a parasitized blowfly pupa. The following procedure overcomes these significant challenges while demonstrating a streamlined, visual procedure for effectively removing wasp embryos from parasitized host pupae, microinjecting them, and carefully transplanting them back into the host for continuation and completion of development. This protocol will strongly enhance the capability of research groups to perform advanced genome modifications in this organism.

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Microinjection of Nasonia vitripennis embryos is an essential method for generating heritable genome modifications. Described here is a detailed procedure for microinjection and transplantation of Nasonia vitripennis embryos, which will greatly facilitate future genome manipulation in this organism.

The jewel wasp Nasonia vitripennis has emerged as an effective model system for the study of processes including sex determination, haplo-diploid sex ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Developmental Biology

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, Eastern equine encephalitis, lymphatic filariasis, and Saint Louis encephalitis. Notably, avian malaria has played a major role in the extinction of numerous endemic island bird species, while WNV has become an important vector-borne disease in the United States. To gain further insight into C. quinquefasciatus biology and expand their genetic control toolbox, we need to develop more efficient and affordable methods for genome engineering in this species. However, some biological traits unique to Culex mosquitoes, particularly their egg rafts, have made it difficult to perform microinjection procedures required for genome engineering. To address these challenges, we have developed an optimized embryo microinjection protocol that focuses on mitigating the technical obstacles associated with the unique characteristics of Culex mosquitoes. These procedures demonstrate optimized methods for egg collection, egg raft separation and other handling procedures essential for successful microinjection in C. quinquefasciatus. When coupled with the CRISPR/Cas9 genome editing technology, these procedures allow us to achieve site-specific, efficient and heritable germline mutations, which are required to perform advanced genome engineering and develop genetic control technologies in this important, but currently understudied, disease vector.

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

This protocol describes microinjection procedures for Culex quinquefasciatus embryos that are optimized to work with CRISPR/Cas9 gene editing tools. This technique can efficiently generate site-specific, heritable, germline mutations that can be used for building genetic technologies in this understudied disease vector.

Culex quinquefasciatus is a vector of a diverse range of vector-borne diseases such as avian malaria, West Nile virus (WNV), Japanese encephalitis, ...

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is applicable to a wide variety of organisms and is highly efficient in the species-rich order Coleoptera (beetles). Here, we summarize the necessary steps for developing this technique in a novel organism and illustrate its application to the different developmental stages of the aquatic diving beetle Thermonectus marmoratus. Target gene sequences can be obtained cost-effectively through the assembly of transcriptomes against a close relative with known genomics or de novo. Candidate gene cloning utilizes a specific cloning vector (the pCR4-TOPO plasmid), which allows the synthesis of double-stranded RNA (dsRNA) for any gene with the use of a single common primer. The synthesized dsRNA can be injected into either embryos for early developmental processes or larvae for later developmental processes. We then illustrate how RNAi can be injected into aquatic larvae using immobilization in agarose. To demonstrate the technique, we provide several examples of RNAi experiments, generating specific knockdowns with predicted phenotypes. Specifically, RNAi for the tanning gene laccase2 leads to cuticle lightening in both larvae and adults, and RNAi for the eye pigmentation gene white produces a lightening/lack of pigmentation in eye tubes. In addition, the knockdown of a key lens protein leads to larvae with optical deficiencies and a reduced ability to hunt prey. Combined, these results exemplify the power of RNAi as a tool for investigating both morphological patterning and behavioral traits in organisms with only transcriptomic databases.

RNA interference is a widely applicable, powerful technique for manipulating gene expression at specific developmental stages. Here, we describe the necessary steps for implementing this technique in the aquatic diving beetle Thermonectus marmoratus, from the acquisition of gene sequences to the knockdown of genes that affect structure or behavior.

RNA interference (RNAi) remains a powerful technique that allows for the targeted reduction of gene expression through mRNA degradation. This technique is ...

Environment

Injecting Gryllus bimaculatus Eggs

Altering gene function in a developing organism is central to different kinds of experiments. While tremendously powerful genetic tools have been developed in traditional model systems, it is difficult to manipulate genes or messenger RNA (mRNA) in most other organisms. At the same time, evolutionary and comparative approaches rely on an exploration of gene function in many different species, necessitating the development and adaptation of techniques for manipulating expression outside currently genetically tractable species. This protocol describes a method for injecting reagents into cricket eggs to assay the effects of a given manipulation on embryonic or larval development. Instructions for how to collect and inject eggs with beveled needles are described. This relatively straightforward technique is flexible and potentially adaptable to other insects. One can gather and inject dozens of eggs in a single experiment, and survival rates for buffer-only injections improve with practice and can be as high as 80%. This technique will support several types of experimental approaches including injection of pharmacological agents, in vitro capped mRNA to express genes of interest, double-stranded RNA (dsRNA) to achieve RNA interference, use of clustered regularly interspaced short palindromic repeats (CRISPR) in concert with CRISPR-associated protein 9 (Cas9) reagents for genomic modification, and transposable elements to generate transient or stable transgenic lines.

Injecting Gryllus bimaculatus Eggs

Here we present a protocol to inject cricket eggs, a technique which serves as a foundational method in many experiments in the cricket, including, but not limited to, RNA interference and genomic manipulation.

Altering gene function in a developing organism is central to different kinds of experiments. While tremendously powerful genetic tools have been ...

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although RNA interference (RNAi) has proved to be a powerful tool for studying WCR biology, it has its limitations. Specifically, RNAi itself is transient (i.e. does not result in long-term Mendelian inheritance of the associated phenotype), and it requires knowing the DNA sequence of the target gene. The latter can be limiting if the phenotype of interest is controlled by poorly conserved, or even novel genes, because identifying useful targets would be challenging, if not impossible. Therefore, the number of tools in WCR&#39;s genomic toolbox should be expanded by the development of methods that could be used to create stable mutant strains and enable sequence-independent surveys of the WCR genome. Herein, we detail the methods used to collect and microinject precellular WCR embryos with nucleic acids. While the protocols described herein are aimed at the creation of transgenic WCR, CRISPR/Cas9-genome editing could also be performed using the same protocols, with the only difference being the composition of the solution injected into the embryos.

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

Here we present protocols for collecting and microinjecting precellular western corn rootworm embryos for the purpose of performing functional-genomic assays such as germline transformation and CRISPR/Cas9-genome editing.

The western corn rootworm (WCR) is an important pest of corn and is well known for its ability to rapidly adapt to pest management strategies. Although ...

Bioengineering

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the triggering of RNA interference (RNAi) in P. maidis through microinjection of double-stranded RNAs (dsRNAs) into nymphs and adults. Despite the power of RNAi, phenotypes generated via this technique are transient and lack long-term Mendelian inheritance. Therefore, the P. maidis toolbox needs to be expanded to include functional genomic tools that would enable the production of stable mutant strains, opening the door for researchers to bring new control methods to bear on this economically important pest. However, unlike the dsRNAs used for RNAi, the components used in CRISPR/Cas9-based genome editing and germline transformation do not easily cross cell membranes. As a result, plasmid DNAs, RNAs, and/or proteins must be microinjected into embryos before the embryo cellularizes, making the timing of injection a critical factor for success. To that end, an agarose-based egg-lay method was developed to allow embryos to be harvested from P. maidis females at relatively short intervals. Herein are provided detailed protocols for collecting and microinjecting precellular P. maidis embryos with CRISPR components (Cas9 nuclease that has been complexed with guide RNAs), and results of Cas9-based gene knockout of a P. maidis eye-color gene, white, are presented. Although these protocols describe CRISPR/Cas9-genome editing in P. maidis, they can also be used for producing transgenic P. maidis via germline transformation by simply changing the composition of the injection solution.

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

Herein are protocols for collecting and microinjecting precellular corn planthopper embryos for the purpose of modifying their genome via CRISPR/Cas9-based genome editing or for the addition of marked transposable elements through germline transformation.

The corn planthopper, Peregrinus maidis, is&#160;a pest of maize and a vector of several maize viruses. Previously published methods describe the ...

A Method for Microinjection of Patiria minata Zygotes

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and evolution of developmental processes. The sea star, Patiria miniata, is gaining prevalence as a model system for these types of studies which were previously performed almost exclusively in the sea urchins, Strongylocentrotus purpuratus and Lytechinus variegatus. An advantage of these model systems is the ease of producing modified embryos in which a particular gene is up or downregulated, labeling a group of cells, or introducing a reporter gene. A single microinjection method is capable of creating a wide variety of such modified embryos. Here, we present a method for obtaining gametes from P. miniata, producing zygotes, and introducing perturbing reagents via microinjection. Healthy morphant embryos are subsequently isolated for quantitative and qualitative studies of gene function. The availability of genome and transcriptome data for this organism has increased the types of studies that are performed and the ease of executing them.

A Method for Microinjection of Patiria minata Zygotes

Methods that produce morphant embryos are essential to study developmental mechanisms and gene regulatory networks. The sea star Patiria miniata is an emerging model system for these studies. Here we present a protocol for obtaining gametes, producing cultures of embryos, and rapid microinjection of zygotes from this species.

Echinoderms have long been a favorite model system for studies of reproduction and development, and more recently for the study of gene regulation and ...

Biology

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, host-symbiont interactions, speciation, and venom synthesis. Despite the availability of several molecular tools, including CRISPR/Cas9, functional genetic studies are still limited in this organism. The major limitation of applying CRISPR/Cas9 technology in N. vitripennis stems from the challenges of embryonic microinjections. Injections of embryos are particularly difficult in this organism and in general in many parasitoid wasps, due to small embryo size and the requirement of a host pupa for embryonic development. To address these challenges, Cas9 ribonucleoprotein complex delivery into female&#160;ovaries by adult injection, rather than embryonic microinjection, was optimized, resulting in both somatic and heritable germline edits. The injection procedures were optimized in pupae and female wasps using either ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) or BAPC (Branched Amphiphilic Peptide Capsules). These methods are shown to be effective alternatives to embryo injection, enabling site-specific and heritable germline mutations.

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis

Here, we describe methods for efficient pupal and adult injections in Nasonia vitripennis as accessible alternatives to embryo microinjection, enabling functional analysis of genes of interest using either RNA-silencing via RNA interference (RNAi) or gene knockout via CRISPR/Cas9 genome editing.

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, ...

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

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Laser-assisted Cytoplasmic Microinjection in Livestock Zygotes

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Embryo Microinjection Techniques for Efficient Site-Specific Mutagenesis in Culex quinquefasciatus

RNA Interference in Aquatic Beetles as a Powerful Tool for Manipulating Gene Expression at Specific Developmental Time Points

Injecting Gryllus bimaculatus Eggs

Microinjection of Western Corn Rootworm, Diabrotica virgifera virgifera, Embryos for Germline Transformation, or CRISPR/Cas9 Genome Editing

Microinjection of Corn Planthopper, Peregrinus maidis, Embryos for CRISPR/Cas9 Genome Editing

A Method for Microinjection of Patiria minata Zygotes

Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis