Research reported in this project was supported by an Institutional Development Award (IDeA) from the National Institute of General Medical Sciences of the National Institutes of Health under grant number P20GM10342 to HH3, and by NSF award number IOS-1257217 to CGE.

1. Hardware Setup and Preparation of Materials
NOTE: Please see Table 1 and Table of Materials for preparation of solutions, reagent, and equipment details.
<ol>
	<li>Set up a dissecting microscope in order to see eggs and guide the injection needle. (Figure 1A shows a dissecting microscope equipped with fluorescence.) Fluorescence capability is advantageous but not necessary.</li>
	<li>Position a 3-axis micromanipulator to maneuver the injection needle into place (Figure 1A).</li>
	<li>Set up a microinjector with tubing and a needle holder to inject small amounts of material (Figure 1A). Optionally use a foot pedal which is not shown in the figure.</li>
	<li>Design and create a stamp to create wells for eggs (Figure 1C) by printing the inverse of the desired pattern, either with a 3D printer or a laser engraver. 
	NOTE: The 3D printed sample stamp shown is 4.5 cm x 5 cm with 128, 3 cm x 1 cm x 1 mm protrusions for creating individual wells. Details on how to make a variety of customizable molds have been published elsewhere16.</li>
	<li>Prepare 10x Injection Buffer, HEPES buffered saline (HBS), and a stock solution of Tetramethylrhodamine Dextran dye and store at 4 &#176;C.</li>
	<li>Prepare experimental solutions to be injected, such as dsRNA, CRISPR/Cas912, transposable elements, in vitro capped mRNA7,8, pharmacological agents17, ZFNs, or TALENS9.</li>
	<li>Place double-stick tape on several layers of standard lab tape on a glass microscope slide in order to create a platform 1 mm or so above the glass slide, which will be used for holding needles for the assessment.</li>
	<li>Prepare a cricket container approximately 40 cm x 60 cm in size with a lid possessing a mesh-covered opening for air circulation.</li>
	<li>Add food, water, and sheltering materials.</li>
	<li>Add several dozen healthy male and female adult crickets. Identify adult cricket by the presence of wings. 
	NOTE: A range of post-imaginal ages may increase egg yields. The density of crickets should be relatively high in order to be able to collect enough eggs for an experiment, but not so crowded that cannibalism occurs. For example, approximately two dozen healthy crickets, with a roughly equal male to female ratio, in a container of the size noted above should be sufficient to collect about 200 eggs in a one to two h period. Crickets can be kept at room temperature, but development and behavior will be optimal if crickets are maintained in a warm (23.5&#8211;26 &#176;C), humid (40 - 60% relative humidity) incubator.</li>
</ol>
2. Making Needles for Injection
<ol>
	<li>Use Borosilicate Glass Capillaries with filament (see the Table of Materials for size details). Prepare the pull needles on the same day, or up to a few days before injections are completed.</li>
	<li>Place a capillary glass into the puller, center the glass on the filament, and tighten the knobs to secure the glass in place.</li>
	<li>Set the puller temperature near the glass ramp temp (-5 &#176;C to +10 &#176;C).</li>
	<li>Use a single-step, high-heat protocol to pull a long taper with a small opening but avoid melting the ends closed. 
	NOTE: For example, when using the micropipette puller detailed in the Table of Materials equipped with a box filament, use the following parameters for glass with a ramp temperature of 478: Heat 478; Pull 45; Vel 75; Del 120. Optimal conditions for producing a needle with the shape shown in Figure 2B should be determined empirically by each user.</li>
	<li>Bevel needle tips in preparation for the injections.</li>
	<li>Wet the spinning grinder plane of the beveler (Figure 2A) with dripping water. Use either reverse osmosis (RO) or diethylpyrocarbonate (DEPC)-treated distilled water.</li>
	<li>Place the pulled needle into the beveler and turn to a 20&#176; angle.</li>
	<li>Lower the needle until it just touches the grinding plane and bevel for 2 to 3 min.</li>
	<li>Assess the bevel by placing the beveled needle on double-stick tape that has been adhered to a glass microscope slide.</li>
	<li>Place the slide holding the needle on the stage of a compound microscope equipped with a camera and image acquisition software.</li>
	<li>Acquire an image of the needle tip using a 20x objective.</li>
	<li>Use the imaging software to measure the interior lumen diameter of the needle just proximal to the beveled opening (Figure 2C). The optimal size is 10 &#181;m + 2 &#181;m.</li>
	<li>Discard needles that have an opening below 8 &#181;m and above 12 &#181;m.</li>
	<li>Store the needles in a container with a lid to avoid getting dusty. Use a strip of wax or similar material to elevate the needles off the bottom of the container to prevent breaking the tips (Figure 2D).</li>
</ol>
3. Egg Collection and Preparation
<ol>
	<li>Maximize the number of eggs laid by depriving crickets of any moist laying material (water vials, egg dishes) overnight or for at least 8&#8211;10 h before attempting to collect eggs. 
	NOTE: Since females have the ability to internally store sperm, a differently timed procedure may be necessary if carefully timed fertilization is experimentally essential18.</li>
	<li>Make 40 mL of 1% agarose in water and pour into a 10 cm petri dish.</li>
	<li>Place an egg-well stamp on the surface of the agarose before it solidifies.</li>
	<li>Remove the stamp, once the agarose is solidified, to reveal the wells (Figure 1C).</li>
	<li>Fill the agarose well plate with HBS containing 1% Penicillin/Streptomycin.</li>
	<li>Place the lid on the dish, wrap in parafilm and store at 4 &#176;C. 
	NOTE: Dishes can be stored for a few days at 4 &#176;C but should be warmed up to room temperature before use to avoid condensation.</li>
	<li>Make an egg collection dish for crickets to lay eggs in by filling a 35 mm petri dish with white playground sand (Figure 1B). 
	NOTE: The sand specified in the Table of Materials is of the appropriate grain size. If the sand grains are too large, the sand will damage the eggs.</li>
	<li>Cover the egg dish with a square of paper towel cut to be approximately 18 cm x 18&#160;cm, and place on the top of dish with sand. Through this paper towel cover the filled dish with tap water.</li>
	<li>Tilt the petri dish and gently squeeze the top to remove excess water.</li>
	<li>Tuck the corners of the paper towel square under the dish and place the egg dish in the inverted lid, which will help keep the paper towel cover in place.</li>
	<li>Place the egg dish into the cricket bin and allow the adult females to oviposit eggs for one to two h. 
	NOTE: The relatively short time here ensures that all the eggs will be similar in the stage of development at the time of injection. If injection before cellularization is important, injections should occur within 14 h of egg laying19.</li>
	<li>Meanwhile, allow the agarose dish (from step 3.6) to warm to the room temperature in preparation for holding eggs for injection.</li>
	<li>Remove egg collection dish, now containing freshly laid eggs, from the cricket bin, and remove the paper towel cover. Place a strainer with a pore size of 0.5&#8211;1 mm over a beaker of at least 500 mL capacity (Figure 1B).</li>
	<li>Rinse the contents of the egg-laying dish (sand and eggs) into the strainer under gently running tap water letting the sand grains fall through the strainer mesh into the water in the beaker below but leaving the cricket eggs in the strainer basket.</li>
	<li>Fill a container with RO water and place it in a tray. Invert the strainer over the container and tap it against the dish to dislodge the eggs into the water. The eggs will sink to the bottom of the container.</li>
	<li>Cut the tip off a P1000 pipette tip with scissors to make an opening approximately 3 mm in diameter. Place this tip on a P1000 pipettor and use it to transfer the eggs from the container to the agarose egg mold dish, transferring as little water as possible.</li>
	<li>Use plastic tweezers to line up eggs in agarose wells filled with HBS plus 1% Penicillin/Streptomycin. Each egg will sink to the bottom of an individual well. Cover with the petri dish lid until ready to inject.</li>
</ol>
4. Prepare the Microinjector
<ol>
	<li>Attach the microinjector to a compressed air or gas source (this protocol was optimized using either compressed air or N2). 
	NOTE: If using a portable air tank, fill to at least 75 psi. The tank will lose air pressure throughout the injections and may need to be refilled; injections become difficult below 40psi.</li>
	<li>Turn on the microinjector. Set the injection time to 0.17 s and the pressure to 10&#8211;15 psi. 
	NOTE: The exact pressure needed will depend on the opening of the needle and must be determined empirically for every round of injection and/or new needle used.</li>
	<li>Ensure that the BALANCE knob of the microinjector is turned to 0 (typically fully counter-clockwise) and that the microinjector is pressurized and in the injection mode.</li>
</ol>
5. Injections
<ol>
	<li>Filter about 500 &#181;L of 10x Injection Buffer using a 1 mL syringe and a 0.45 &#181;m syringe filter.</li>
	<li>Mix injection reagents (the details that follow are for the injection of dsRNA prepared as described in12). 
	NOTE: It may be advisable to design and perform several controls, especially when one is first learning this technique. Poking eggs without injecting, injecting buffer + dye alone, or injecting buffer + dye + a control reagent (such as eGFP dsRNA) are all good tests of how disruptive various elements of the injection protocol might be. See Figure 3 for the survivability of a number of different controls.</li>
	<li>Start with a 4 &#956;L dsRNA aliquot.</li>
	<li>Add 0.5 &#956;L of the filtered 10x Injection Buffer to the dsRNA aliquot.</li>
	<li>Add 0.5 &#956;L of 50 mg/mL Tetramethyl rhodamine Dextran dye stock solution. If working on a dissecting scope without fluorescence, use Phenol Red instead.</li>
	<li>Keep all materials on ice for the duration of the injection period.</li>
	<li>Load the injection solution into an injection needle using 20 &#956;L loading tips and a p10 pipet.</li>
	<li>Draw up a 1.5 &#956;L injection solution and insert the loading tip into the wide end of the injection needle.</li>
	<li>Eject solution as far down into the needle as possible and eliminate air bubbles by flicking gently; be careful not to break the needle.</li>
	<li>Place the dish of eggs under the dissecting microscope and select a low magnification of around 10x (1x magnification viewed through 10x eye objectives).</li>
	<li>Insert the needle into the injection housing and tighten. 
	NOTE: Take care that the needle is properly and firmly inserted into the housing. To do this, pull the wide end of the needle back out of the metal housing, bringing the silicon tubing in the housing with it. Adjust the silicon tubing so that the glass end of the needle protrudes just beyond the silicon. If this is not done, the silicon tubing may get pinched between the glass and the metal housing, blocking the end of the needle.</li>
	<li>Carefully insert the injection housing with an attached needle into the micromanipulator. Be aware of the sharp end of the needle and be careful that it is not bumped into surfaces and broken.</li>
	<li>While looking at both the eggs and the needle through the microscope, move the needle near the eggs in the top left corner of the dish grid.</li>
	<li>Lower the needle until the tip enters the HBS plus 1% Penicillin/Streptomycin buffered solution in the dish.</li>
	<li>Center the needle in the field of vision and move the egg dish so that the needle is a few mm closer to the edge of the dish than the eggs.</li>
	<li>Do not obstruct the view of the grid of eggs with the needle.</li>
	<li>Set the microscope to the filter appropriate for Rhodamine so it is possible to observe the fluorescence in the needle and focus on the tip of the needle.</li>
	<li>On the microinjector, slowly turn the BALANCE knob clockwise until the injection solution starts to leak out of the needle into the suspension solution. The balance number on the screen of the microinjector will begin to increase, though the numerical value is not important. Next, turn the knob back counterclockwise slightly just until the dye stops leaking out of the needle. 
	NOTE: It is critical to set this balance each time a new needle is used. Without the balance set appropriately, the microinjector will continue to inject for some time after the injection is triggered. It is even possible that a single push of the injection button with an unbalanced needle will result in the expulsion of the entire contents of the loaded needle.</li>
	<li>With the needle centered in the field of view, move the egg dish so that the needle is aimed at the egg to be injected first. It is recommended that injections start in one corner of the grid of arranged eggs, for example, the upper left corner.</li>
	<li>Adjust the magnification to about 50x; at this magnification, a single egg will fill most of the field of view.</li>
	<li>Use both the micromanipulator and one&#8217;s hand on the egg dish to move the needle into position for the injection.</li>
	<li>Advance the needle using the micromanipulator and insert the tip of the needle into the first egg to be injected.</li>
	<li>Insert the needle at 20&#8211;30% egg length from the posterior (blunter) end of the egg (Figure 1E), perpendicular to the long axis of the egg.</li>
	<li>Inject solution either with the injection foot pedal or the injection button on the microinjector. A small bolus of fluorescent material inside the egg will indicate a successful injection (Figure 1D). 
	NOTE: It is not uncommon for some yolk to be ejected from the egg during injection (Figure 1F), and this does not necessarily indicate that the injected egg will be inviable.</li>
	<li>Retract the needle from the egg. If the egg is unintentionally lifted out of its well upon retraction of the needle, use small forceps to push the egg back into its well while retracting the needle.</li>
	<li>If the needle clogs during injection, remove the needle from the egg and, keeping the needle submerged in the HBS plus 1% Penicillin/Streptomycin solution covering the eggs, clear the clogged needle using either the Clear function or by pushing the inject button.</li>
	<li>Continuing to use the same needle, repeat the injection procedure for as many eggs as possible. At first, this may be only about 20 or 30 eggs but will increase to over 100 with practice.</li>
	<li>If the needle becomes clogged and cannot be cleared, discard the needle, fill a new needle and begin again (i.e., return to step 5.7).</li>
	<li>Store the dish of injected eggs in a warm incubator (23.5&#8211;26 &#176;C) that is slightly humid (to minimize the evaporation) until the embryos have reached the desired stage of development for the analysis.</li>
	<li>At least every 24 h, remove eggs that are no longer viable (Figure 3A,B), and replace the suspension solution with fresh HBS plus 1% Penicillin/Streptomycin.</li>
	<li>Two to three days after injection, transfer the eggs by gentle pipetting or with plastic forceps to paper towels moistened with HBS plus 1% Penicillin/Streptomycin in a lidded petri dish to continue development in the warm, humid incubator.</li>
	<li>Monitor the eggs, and remove eggs that are no longer viable each day (Figure 3A,B).</li>
</ol>

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The authors have nothing to disclose.

The two main challenges with this technique are the related issues of optimal needle size and survivability. Though smaller needles improve survivability, needles with narrower lumens have a greater degree of capillary forces at work, which makes it more likely that yolk will move into the needle causing it to clog. In the best case, blockages can be cleared simply by injecting another egg or by clearing the needle as described above. One can also attempt to increase the balance pressure on the microinjector until the yo...

The ability to modify the genome or influence gene expression in organisms is the basis for the design of many types of experiments testing functional causality. It is also critical for the comparative and evolutionarily-relevant work that genomic and non-genomic modification techniques be available in organisms outside traditional genetic laboratory animal model systems (e.g., Mus musculus, Danio rerio, Drosophila melanogaster, and Caenorhabditis elegans). Whether it is the desire to understand organismal diversity1 or one&#39;s adherence to Krogh&#39;s principle, that for every biological question there is an organism best suited to its solution2,3, the ability to modify genomes or influence the gene expression is essential for modern experimental designs.
The cricket Gryllus bimaculatus is an emerging model system. Used for the last century in neuroethology experiments4, the last two decades have witnessed an increased experimental interest in the cricket, particularly focused on the evolution and development of this organism5. The cricket is a hemimetabolous insect that branches basally to well-studied holometabolous insects, such as D. melanogaster and Tribolium castaneum6. Due to its useful position on the evolutionary tree, scientists are interested in asking modern, sophisticated experimental questions in this insect, which has led to a growing interest in adapting molecular tools for the use in G. bimaculatus.
Injections of molecular reagents into cricket eggs can be used for genomic modification experiments as well as non-genomic manipulations of gene expression in embryos. For example, transgenic G. bimaculatus carrying eGFP insertions have been created using the transposase piggyBac7,8. Investigators have successfully created knockout G. bimaculatus using Zinc-finger nucleases (ZFNs) and transcription activator-like (TAL) effector nucleases (TALENs) to introduce double-stranded breaks in specific genomic regions9. Though ZFNs and TALENs allow site-specific targeting in animals beyond the big four model systems, these reagents have quickly been surpassed by the CRISPR/Cas9 system, which is simpler to use, more efficient, and highly flexible10. CRISPR has been used in G. bimaculatus to produce knock-out11 as well as knock-in lines12,13 In addition to genomic modification, dsRNA can be injected into eggs to knock down mRNA expression in developing embryos, allowing investigators to understand the role of specific transcripts throughout development14,15. Some limited details on how to inject cricket eggs have been published previously12.
Here, we describe a detailed protocol for injecting early G. bimaculatus eggs. This protocol is effective and easily adaptable to various laboratory settings, injection materials, and possibly to other insects. While additional details for designing and implementing genomic modification and knockdown experiments have been published elsewhere12,13, these approaches will ultimately rely on the injection protocol detailed here.

<table><tbody><tr><td>Fluorescent dissecting microscope</td><td>Leica</td><td>M165 FC</td><td>Stereomicroscope with fluorescence</td></tr><tr><td>External light source for fluorescence</td><td>Leica</td><td>EL 6000</td><td></td></tr><tr><td>Microinjector</td><td>Narishige</td><td>IM-300</td><td>-Accessories may include Injection Needles Holder, Input Hose (with a hose connector), AC Power Cord, Foot Switch, Silicone Rubber Gasket-</td></tr><tr><td>mCherry filter cube</td><td>Leica</td><td>M205FA/M165FC</td><td>Filter cube for mCherry or similar red dye will work</td></tr><tr><td>Micromanipulator</td><td>World Precision Instruments, Inc.</td><td>M3301R</td><td>Used with Magnetic Stand (Narishige, Type GJ-8)</td></tr><tr><td>Magnetic stand</td><td>Narishige</td><td>MMO-202ND</td><td></td></tr><tr><td>Pipette Holder (Needle holder)</td><td>Narishige</td><td>HD-21</td><td></td></tr><tr><td>Tubing to connect air source to microinjector</td><td></td><td></td><td></td></tr><tr><td>Egg well stamp</td><td>3D printed</td><td>custom</td><td>3D printed on a Lulzbot Taz 5 using Poly Lactic Acid thermoplastic</td></tr><tr><td>Microwave</td><td>various</td><td></td><td></td></tr><tr><td>Incubator or temperature controlled room</td><td>various</td><td></td><td>Temperatures of 23.5-26 &amp;deg;C are needed.</td></tr><tr><td>Cricket food</td><td>various</td><td></td><td>cat food or fish flakes are appropriate food.&amp;nbsp;</td></tr><tr><td>Cricket water</td><td>vairous</td><td></td><td>Water can be held in vials and presented to crickets through cotton balls</td></tr><tr><td>Cricket shelter</td><td>arious</td><td></td><td>Shelter materials can include crumpled paper towels or egg cartons</td></tr><tr><td>Glass capillary tubes</td><td>World Precision Instruments, Inc.</td><td>Item no. 1B100F-4</td><td>Kwik-Fil&amp;trade; Borosilicate Glass Capillaries, 100 mm length, 0.58 mm ID, 1.0 mm OD, with filament</td></tr><tr><td>Micropipette puller</td><td>Flaming/Brown</td><td>Model P-97</td><td>Distributed by Sutter Instrument Co.</td></tr><tr><td>Beveller/Micro grinder</td><td>Narishige</td><td>Model EG-45/EG-400</td><td>EG-400 includes a microscope head</td></tr><tr><td>Petri dishes</td><td>CellTreat</td><td>Product code 229693</td><td>90 mm diameter</td></tr><tr><td>Play Sand</td><td>Sandtastik Products Ltd.</td><td>B003U6QLVS</td><td>White play sand</td></tr><tr><td>Agarose</td><td>American Bioanalytical</td><td>AB000972</td><td>Agarose GPG/LE ultrapure</td></tr><tr><td>Egg Strainer: Extra Fine Twill Mesh Stainless Steel Conical Strainers</td><td>US Kitchen Supply</td><td>Model SS-C123</td><td>Pore size should be between 0.5 - 1.0 mm</td></tr><tr><td>Penicillin Streptomycin</td><td>Gibco by Life Technologies</td><td>Ref 15070-063</td><td>Pen Strep</td></tr><tr><td>Plastic tweezers</td><td>Sipel Electronic SA</td><td>P3C-STD</td><td>Black Static Dissipative, 118 mm</td></tr><tr><td>Syringe filters, 25 mm diameter, 0.45 &amp;micro;m</td><td>Nalgene</td><td>725-2545</td><td>Use with 1 mL syringe</td></tr><tr><td>1 mL syringe, with Tuberculin Slip Tip</td><td>Becton Dickinson</td><td>309602</td><td>Use with syring filter to filter Injection Buffer , Luer-Lok tip syringes would also work</td></tr><tr><td>Air tank (optional)</td><td>Midwest Products</td><td>Air Works&amp;reg;</td><td>Portable air tank</td></tr><tr><td>Rhodamine dye</td><td>Thermofisher</td><td>D-1817</td><td>dextran, tetramethylrhodamine 10,000MW,</td></tr><tr><td>20 mL loading tips</td><td>Eppendorf</td><td>Order no. 5242 956.003</td><td>epT.I.P.S. 20 &amp;mu;L Microloader</td></tr><tr><td>Compound microscope</td><td>Zeiss</td><td>Axioskope 2 plus</td><td></td></tr><tr><td>20x objective</td><td>Ziess</td><td>Plan-Apochromat 20x/0.75 M27</td><td></td></tr><tr><td>Camera</td><td>Leica</td><td>DMC 5400</td><td></td></tr><tr><td>Leica Application Suite&amp;nbsp; software</td><td>Leica</td><td>LAS</td><td>Version 4.6.2 used here</td></tr></tbody></table>

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.

Crickets readily lay eggs in the moist material, and providing adequate material, such as moist sand or dirt, induces them to lay a large number of eggs. This is especially effective if crickets are first deprived of egg-laying material for 8&#8211;10 h. Eggs laid in clean sand can be easily separated, collected (Figure 1B) and placed into custom-designed egg wells for injection (Figure 1C). A dissecting microscope, a microinject...

Watch this Scientific Journal Video about Injecting Gryllus bimaculatus Eggs at JoVE.com

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.

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 ...

Research

JoVE Journal

Developmental Biology

16.4K Views.  Colby College. 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.

Video: Injecting Gryllus bimaculatus Eggs

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.

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.

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

Genetics

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Schistosoma parasites are blood flukes that infect an estimated 200 million people worldwide 1. In chronic infection with Schistosoma, the severe pathology, including liver fibrosis and splenomegaly, is caused by the immune response to the parasite eggs rather than the parasite itself 2. Parasite eggs induce a Th2 response characterized by the production of IL-4, IL-5 and IL-13, the alternative activation of macrophages and the recruitment of eosinophils. Here, we describe injection of Schistosoma mansoni eggs as a model to examine parasite-specific Th2 cytokine responses in the lung and draining lymph nodes, the formation of pulmonary granulomas surrounding the egg, and airway inflammation. 
Following intraperitoneal sensitization and intravenous challenge, S. mansoni eggs are transported to the lung via the pulmonary arteries where they are trapped within the lung parenchyma by granulomas composed of lymphocytes, eosinophils and alternatively activated macrophages 3-6. Associated with granuloma formation, inflammation in the broncho-alveolar spaces, expansion of the draining lymph nodes and CD4 T cell activation can be observed. Here we detail the protocol for isolating Schistosoma mansoni eggs from infected livers (modified from 7), sensitizing and challenging mice, and recovering the organs (broncho-alveolar lavage (BAL), lung and draining lymph nodes) for analysis. We also include representative histologic and immunologic data and suggestions for additional immunologic analysis. 
Overall, this method provides an in vivo model to investigate helminth-induced immunologic responses in the lung, which is broadly applicable to the study of Th2 inflammatory diseases including helminth infection, fibrotic diseases, allergic inflammation and asthma. Advantages of this model for the study of type 2 inflammation in the lung include the reproducibility of a potent Th2 inflammatory response in the lung and draining lymph nodes, the ease of assessment of inflammation by histologic examination of the granulomas surrounding the egg, and the potential for long-term storage of the parasite eggs.

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Schistosoma mansoni eggs are potent stimulators of the T helper type 2 (Th2) immune response, characteristic of parasite infection, asthma and allergic inflammation. This protocol utilizes S. mansoni egg injection to generate a CD4 Th2 cytokine-induced inflammatory response in the lung, characterized by lung granuloma formation around the egg, eosinophilia and macrophage alternative activation.

Schistosoma parasites are blood flukes that infect an estimated 200 million people worldwide 1. In chronic infection with Schistosoma, the severe ...

Immunology and Infection

Manipulation of Gene Function in Mexican Cavefish

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, including eye degeneration, albinism, sleep loss, hyperphagia, and sensory processing. Species of cavefish from around the world display a convergent evolution of morphological and behavioral traits due to shared environmental pressures between different cave systems. Diverse cave species have been studied in the laboratory setting. The Mexican tetra, Astyanax mexicanus, with sighted and blind forms, has provided unique insights into biological and molecular processes underlying the evolution of complex traits and is well-poised as an emerging model system. While candidate genes regulating the evolution of diverse biological processes have been identified in A. mexicanus, the ability to validate a role for individual genes has been limited. The application of transgenesis and gene-editing technology has the potential to overcome this significant impediment and to investigate the mechanisms underlying the evolution of complex traits. Here, we describe a different methodology for manipulating gene expression in A. mexicanus. Approaches include the use of morpholinos, Tol2 transgenesis, and gene-editing systems, commonly used in zebrafish and other fish models, to manipulate gene function in A. mexicanus. These protocols include detailed descriptions of timed breeding procedures, the collection of fertilized eggs, injections, and the selection of genetically modified animals. These methodological approaches will allow for the investigation of the genetic and neural mechanisms underlying the evolution of diverse traits in A. mexicanus.&#160;

We describe approaches for the manipulation of genes in the evolutionary model system Astyanax mexicanus. Three different techniques are described: Tol2-mediated transgenesis, targeted manipulation of the genome using CRISPR/Cas9, and knockdown of expression using morpholinos. These tools should facilitate the direct investigation of genes underlying the variation between surface- and cave-dwelling forms.

Cave animals provide a compelling system for investigating the evolutionary mechanisms and genetic bases underlying changes in numerous complex traits, ...

Microinjection Method for Anopheles gambiae Embryos

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA fragments encoding genes of interest as well as favorable traits into the insect germline in a stable and heritable manner. The resulting transgenic strains can be studied for phenotypic changes resulting from the expression of the integrated DNA to answer basic questions or used in practical applications. Although the technology is straightforward, it requires of the investigator patience and practice to achieve a level of skill that maximizes efficiency. Shown here is a method for microinjection of embryos of the African malaria mosquito, Anopheles gambiae. The objective is to deliver by microinjection exogenous DNA to the embryo so that it can be taken up in the developing germline (pole) cells. Expression from the injected DNA of transposases, integrases, recombinases, or other nucleases (for example CRISPR-associated proteins, Cas) can trigger events that lead to its covalent insertion into chromosomes. Transgenic An. gambiae generated from these technologies have been used for basic studies of immune system components, genes involved in blood-feeding, and elements of the olfactory system. In addition, these techniques have been used to produce An. gambiae strains with traits that may help control the transmission of malaria parasites.

Microinjection Method for Anopheles gambiae Embryos

Microinjection techniques are essential to introduce exogenous genes into the genomes of mosquitoes. This protocol explains a method used by the James laboratory to microinject DNA constructs into Anopheles gambiae embryos to generate transformed mosquitoes.

Embryo microinjection techniques are essential for many molecular and genetic studies of insect species. They provide a means to introduce exogenous DNA ...

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to their successful evolutionary radiation on the earth. The application of genetic tools such as genome editing is the key to understanding genetic changes that are responsible for evolutionary transitions in an Evo-Devo approach. In this article, we describe our current protocol for generating and maintaining mutant strains of T. domestica. We report a dry injection method, as an alternative to the reported wet injection method, that allows us to obtain stably high survival rates in injected embryos. We also report an optimized environment setting to mate adults and obtain subsequent generations with high efficiency. Our method underlines the importance of taking each species&#8217; unique biology into account for the successful application of genome editing methods to non-traditional model organisms. We predict that these genome editing protocols will help in implementing T. domestica as a laboratory model and to further accelerate the development and application of useful genetic tools in this species.

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

We provide a detailed protocol for rearing, microinjection of eggs and for efficient mating of the firebrat Thermobia domestica to generate and maintain mutant strains after genome editing.

The firebrat Thermobia domestica is an ametabolous, wingless species that is suitable for studying the developmental mechanisms of insects that led to ...

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression analyses by RNA interference and stable transgenesis remain limited in many species due to the particular anatomy and molecular cellular biology of the organism. For example, outside of the crown group Caenorhabditis that includes Caenorhabditis elegans3, stably transmitted transgenic lines in non-Caenorhabditis species have not been reported in this specious phylum (Nematoda), with the exception of Strongyloides stercoralis4 and Pristionchus pacificus5. To facilitate the expanding role of P. pacificus in the study of development, evolution, and behavior6-7, we describe here the current methods to use microinjection for making transgenic animals and gene knock down by RNAi. Like the gonads of C. elegans and most other nematodes, the gonads of P. pacificus is syncitial and capable of incorporating DNA and RNA into the oocytes when delivered by direct microinjection. Unlike C. elegans however, stable transgene inheritance and somatic expression in P. pacificus requires the addition of self genomic DNA digested with endonucleases complementary to the ends of target transgenes and coinjection markers5. The addition of carrier genomic DNA is similar to the requirement for transgene expression in Strongyloides stercoralis4 and in the germ cells of C. elegans. However, it is not clear if the specific requirement for the animals' own genomic DNA is because P. pacificus soma is very efficient at silencing non-complex multi-copy genes or that extrachromosomal arrays in P. pacificus require genomic sequences for proper kinetochore assembly during mitosis. The ventral migration of the two-armed (didelphic) gonads in hermaphrodites further complicates the ability to inject both gonads in individual worms8. We also demonstrate the use of microinjection to knockdown a dominant mutant (roller,tu92) by injecting double-stranded RNA (dsRNA) into the gonads to obtain non-rolling F1 progeny. Unlike C. elegans, but like most other nematodes, P. pacificus PS312 is not receptive to systemic RNAi via feeding and soaking and therefore dsRNA must be administered by microinjection into the syncitial gonads. In this current study, we hope to describe the microinjection process needed to transform a Ppa-egl-4 promoter::GFP fusion reporter and knockdown a dominant roller prl-1 (tu92) mutant in a visually informative protocol.

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

In model organisms, transgenesis can manipulate gene functions while RNAi can knockdown specific mRNA transcripts 1-2. This protocol aims to illustrate the techniques needed to introduce stably transmitted DNA and transient double stranded RNA into the necromenic nematode Pristionchus pacificus for studies in evolutionary, developmental, and behavioral biology.

Although it is increasingly affordable for emerging model organisms to obtain completely sequenced genomes, further in-depth gene function and expression ...

Biology

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 ...

Bioengineering

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and Strongyloides ratti. S.&#160;stercoralis is a human-parasitic, skin-penetrating nematode that infects approximately 610 million people, while the rat parasite S.&#160;ratti is closely related to S.&#160;stercoralis and is often used as a laboratory model for S.&#160;stercoralis. Both S.&#160;stercoralis and S.&#160;ratti are easily amenable to the generation of transgenics and knockouts through the exogenous nucleic acid delivery technique of intragonadal microinjection, and as such, have emerged as model systems for other parasitic helminths that are not yet amenable to this technique.
Parasitic Strongyloides adults inhabit the small intestine of their host and release progeny into the environment via the feces. Once in the environment, the larvae develop into free-living adults, which live in feces and produce progeny that must find and invade a new host. This environmental generation is unique to the Strongyloides species and similar enough in morphology to the model free-living nematode Caenorhabditis elegans that techniques developed for C. elegans can be adapted for use with these parasitic nematodes, including intragonadal microinjection. Using intragonadal microinjection, a wide variety of transgenes can be introduced into Strongyloides. CRISPR/Cas9 components can also be microinjected to create mutant Strongyloides larvae. Here, the technique of intragonadal microinjection into Strongyloides, including the preparation of free-living adults, the injection procedure, and the selection of transgenic progeny, is described. Images of transgenic Strongyloides larvae created using CRISPR/Cas9 mutagenesis are included. The aim of this paper is to enable other researchers to use microinjection to create transgenic and mutant Strongyloides.

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

The functional genomic toolkit for the parasitic nematodes Strongyloides stercoralis and Strongyloides ratti includes transgenesis, CRISPR/Cas9-mediated mutagenesis, and RNAi. This protocol will demonstrate how to use intragonadal microinjection to introduce transgenes and CRISPR components into S.&#160;stercoralis and S.&#160;ratti.

The genus Strongyloides consists of multiple species of skin-penetrating nematodes with different host ranges, including Strongyloides stercoralis and ...

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite the growing burden of tick-borne diseases, research on ticks has lagged behind insect disease vectors due to challenges in applying genetic transformation tools for functional studies to the unique biology of ticks. Genetic interventions have been gaining attention to reduce mosquito-borne diseases. However, the development of such interventions requires stable germline transformation by injecting embryos. Such an embryo injection technique is lacking for chelicerates, including ticks. Several factors, such as an external thick wax layer on tick embryos, hard chorion, and high intra-oval pressure, are some&#160;obstacles that previously prevented embryo injection protocol development in ticks. The present work has overcome these obstacles, and an embryo injection technique for the black-legged tick, Ixodes scapularis, is described here.&#160;This technique can be used to deliver components, such as CRISPR/Cas9, for stable germline transformations.

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

The present protocol describes a method for injecting tick embryos. Embryo injection is the preferred technique for genetic manipulation to generate transgenic lines.

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite ...

Injecting Gryllus bimaculatus Eggs

Summary

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Chapters in this video

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

Using Eggs from Schistosoma mansoni as an In vivo Model of Helminth-induced Lung Inflammation

Manipulation of Gene Function in Mexican Cavefish

Microinjection Method for Anopheles gambiae Embryos

Egg Microinjection and Efficient Mating for Genome Editing in the Firebrat Thermobia domestica

RNAi Mediated Gene Knockdown and Transgenesis by Microinjection in the Necromenic Nematode Pristionchus pacificus

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

High Throughput Microinjections of Sea Urchin Zygotes

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis