The authors acknowledge Channa Aluvihare and Yonus Gebermicale, ITF, UMD, for insight and support during the initial phase of protocol development. Tungsten needles were a generous gift from David O&#39;Brochta, ITF, UMD. We are thankful to Dr. Ladislav Simo for testing this protocol in I. ricinus and for insightful discussions. This project was funded by NIH-NIAID R21AI128393 and Plymouth Hill Foundation, NY to MG-N, startup funds from the University of Nevada to AN, the National Science Foundation Grant No. 2019609 to MG-N and AN, and a Peer-to-Peer Grant from IGTRCN to AS.

Ixodes scapularis adults were either purchased from Oklahoma State University (OSU) or reared at the University of Nevada, Reno (UNR) (IACUC protocol #21-001-1118).
1. Preparation of female ticks for embryo collection
NOTE: To collect eggs of appropriate age, it is important to synchronize egg-laying. Although egg-laying cues in ticks remain unclear, under the standard insectary conditions (27 &#176;C temperature and &#62;90% relative humidity (RH)), I. scapularis females begin laying eggs approximately 8 days post-host detachment. This timeline can be lengthened by storing replete females at 4 &#176;C. We have stored blood-fed females at 4 &#176;C up to 8 weeks without any negative effect on egg-laying. These conditions may need to be modified for each insectary.
<ol>
	<li>Store all replete females at 4 &#176;C with &#62;90% RH in a 6-quart plastic box lined with moist filter paper until used for microinjections.</li>
	<li>A week before injections, transfer females from 4 &#176;C to a 27 &#176;C incubator for the initiation of egg-laying.</li>
	<li>When the females start laying eggs (3-4 days after moving them to 27 &#176;C), remove any eggs using a fine-tipped paintbrush and prepare females for Gen&#233;&#39;s organ (wax gland) ablation. 
	NOTE: The Gen&#233;&#39;s organ extends from the area between the scutum and capitulum (dorsal base of the mouthparts), mouthparts bend over the ventral body surface (parallel to the surface), and the extended Gene&#39;s organ coats eggs as soon as the eggs are laid (Figure 1). Eggs laid before the Gen&#233;&#39;s organ removal or emptying of wax will have a wax coating and are not suitable for injections and must be discarded.</li>
	<li>To either remove or empty the Gen&#233;&#39;s organ, use clay to position the engorged female on a glass slide under a microscope oriented so&#160;that mouthparts are visible on both ventral and dorsal sides (Figure 1B,C).
	<ol>
		<li>Carefully poke the area exactly between the scutum and mouthparts using fine forceps and tungsten needles made in the lab using tungsten wire following a previously published protocol16 (needles can also be purchased commercially, and attached to a micro dissecting probe, see Table of Materials).</li>
		<li>Working the way inside with the tungsten needle, pull the wax gland out (Figure 1D,E). It may take several minutes to remove the gland. Wipe away any liquid wax secretion using lab wipes. 
		NOTE: The gland can also be removed from the ventral side right below the mouthparts; by reaching in the back towards scutum with a needle and forceps. Application of silicon glue around mouthparts also blocks Gene&#39;s organ eversion and can be used instead of removing or emptying the wax (personal communication with Dr. Ladislav Simo, INRAE, France). Engorged ticks will not stay on double-sided tape. Using clay to &#34;swaddle&#34; the tick helps keep them in place during manipulation. Use forceps to hold the tick and tungsten needle for dissection. The ventral part is easier to pierce with a needle and preferred by the authors.</li>
	</ol>
	</li>
	<li>To empty the gland, arrange the gravid tick as mentioned in step 1.4. Carefully puncture the area behind the mouthparts on the dorsal side and apply pressure on the ventral side using forceps. 
	NOTE: Alternatively, it&#160;is not necessary to remove the wax gland; emptying the gland of wax will last for several days. Egg-laying females have a white area around the mouth parts that indicates the location of the wax gland and can be used as a reference.
	<ol>
		<li>Place the edge of a lint-free lab wipe and remove the liquid wax coming out of the puncture site. Ensure to avoid hemolymph secretion, which is a clear liquid compared to slightly yellowish viscous wax. It may take 5-10 min until most of the wax is removed.</li>
	</ol>
	</li>
	<li>After manipulation, place females in an incubator at 27 &#176;C (&#62;90% RH) and allow them to recover for 1-2 days. 
	NOTE: Treated females usually take 1-2 days to recover and start laying eggs again.</li>
	<li>When the females begin laying eggs again, use a fine paintbrush to collect freshly deposited eggs (0-18 h post-egg-laying) and place them in a 1.5 mL microcentrifuge tube. 
	&#8203;NOTE: If using eggs from the same female over time, this procedure must be repeated after 4-5 days of egg-laying if glands were not removed.</li>
</ol>
2. Embryo treatment for microinjections
<ol>
	<li>Add ~200 &#181;L (or sufficient volume to cover the eggs, dependent on the number of eggs) of 5% (weight/volume) benzalkonium chloride/water (see Table of Materials) in the tube containing 0-18 h old eggs. Gently swirl the eggs with the paintbrush for 5 min to avoid the eggs settling into the bottom of the tube (for details, see Sharma et al.4). Remove the supernatant using a micropipette, leaving the eggs behind in tube. 
	CAUTION: Benzalkonium chloride is corrosive to skin and eyes, wear gloves and eye protection.</li>
	<li>Add ~200 &#181;L of distilled (DI) water to the tube containing eggs, swirl with a paintbrush, and remove the water from the microcentrifuge tube using a micropipette. Repeat the washing with DI water one more time.</li>
	<li>After washing with DI water, add ~200 &#181;L of 5% (w/v) sodium chloride (NaCl) and gently swirl with a paintbrush for 5 min. Remove the solution from the tube after 5 min and wash the eggs twice with DI water.</li>
	<li>Add ~100 &#181;L of 1% (w/v) NaCl solution in the microcentrifuge tube containing eggs. Keep the eggs in 1% (w/v) NaCl solution until they are used for injections.</li>
	<li>Start microinjections after about an hour. This timeline helps in proper desiccation of the eggs and prevents them from bursting. 
	&#8203;NOTE: Eggs can be kept in 1% (w/v) NaCl solution for 7-8 h without significantly affecting&#160;survival. If the eggs are difficult to inject or burst, then the eggs are not desiccated enough and can be further treated with 5% (w/v) NaCl for an additional 5 min.</li>
</ol>
3. Preparation of injection needles
<ol>
	<li>Insert an aluminosilicate capillary glass (with filament, see Table of Materials) into the needle puller following the manufacturer&#39;s instructions.</li>
	<li>Adjust the needle puller to the following settings: Heat: 575, Pull: 20, Velocity: 50, time: 200, and Pressure: 700.</li>
	<li>Activate the pull function of the needle puller and repeat the process for additional needles.</li>
	<li>Store the pulled needles by sticking them into lines of modeler&#39;s clay in a clean Petri dish.</li>
	<li>Load the needle with the injection mixture using a micro loader tip (see Table of Materials). 
	&#8203;NOTE: The comparison of needles used for tick and mosquito embryo injection is shown in Figure 2.</li>
</ol>
4. Slide setup for the microinjections
<ol>
	<li>Using double-sided tape, adhere two glass microscope slides together, leaving a gap of about 0.5 cm to support aligning eggs and preventing them from rolling over during injections (Figure 3A).</li>
	<li>Apply a piece of transparent film dressing (see Table of Materials) on the double-sided tape in the gap between the slides. Ensure to align the film dressing perfectly with the gap.</li>
</ol>
5. Embryo microinjections
<ol>
	<li>Align 8-10 eggs at a time on the slide setup (above) using a single-haired paintbrush. 
	NOTE: Eggs aligned with the longer axis perpendicular (Figure 3B) to the slide edge are easier to inject than eggs whose longitudinal axis is aligned parallel to the edge (Figure 3C). Eggs in the perpendicular orientation (Figure 3B) can better withstand injection resulting in higher survival.</li>
	<li>Place the slide with aligned eggs on the stage of a compound microscope. Use a piece of lint-free wipe to remove the 1% NaCl solution in which they are stored.</li>
	<li>Attach the filled injection needle to a microinjector connected to a micromanipulator (see Table of Materials).</li>
	<li>Open the needle by gently rubbing it against the egg surface using a high-pressure setting on the microinjector (&#62;5,000 hPa).</li>
	<li>After the needle is opened, reduce the pressure (1,000-2,500 hPa) of the microinjector depending on the opening of the needle. 
	NOTE: A small opening will require comparatively higher pressure, while a larger opening will require a lower pressure. This is to control the volume of the construct injected. Depending on the size of the needle opening, the injection volume will vary from 1-5 pL.</li>
	<li>Inject all the eggs on the slide at a 10&#176;-15&#176; angle as quickly as possible and immediately move the slide to high humidity conditions in a Petri dish lined with moist paper. As soon as the eggs are injected, they start to desiccate. 
	&#8203;NOTE: Injecting a small number of eggs at a time increases the injection success and the survival probability. Needles do get clogged by embryo reflux. If the tip is still sharp, to clear clogged needles, move the needle into a small droplet of 1% NaCl added to the edge of the slide. The capillary action will dislodge the small clog. If the needle is already blunt, change it.</li>
</ol>
6. Post-injection care of embryos
<ol>
	<li>Place the slide containing injected embryos in a large Petri dish lined with moist filter paper. 
	NOTE: It is critical not to disturb the eggs for at least 5-6 h. This allows the injection wound to seal properly. The injection wound may open up when agitated, and cytoplasm may begin to ooze out, resulting in egg mortality.</li>
	<li>After 5-6 h, add a small drop of DI water to the transparent film dressing with injected embryos attached. Using a paintbrush, gently displace the eggs. Transfer the eggs to a small Petri dish (10 cm) and submerge them in DI water. 
	NOTE: The film dressing allows easy removal of eggs after injections. The moment a water drop is added to the film dressing, the eggs will start to move along with the water.</li>
	<li>Keep the eggs submerged in water and place the Petri dish in a 6-quart plastic box in an incubator at 27 &#176;C and &#62;90% RH.</li>
	<li>Check the eggs regularly, daily for the first few days, and then every 3-4 days until larvae start hatching. 
	NOTE: If a large number of eggs are damaged during microinjections, gently draining the DI water after a week will prevent fungal growth on the dead eggs.</li>
	<li>When the larvae begin to hatch from the injected embryos (21-25 days after injection for the present laboratory setup), check them daily and transfer any hatched larvae to glass vials with a screen on top. Larvae can hatch underwater and survive for ~1-2 weeks.</li>
	<li>Screen the larvae4 if the eggs were injected with a construct that may result in a visible phenotype.</li>
</ol>

<ol>
	<li>Hinckley, A. F. et al. Lyme disease testing by large commercial laboratories in the United States. Clinical Infectious Diseases: An Official Publication of the Infectious Diseases Society of America. 59 (5), 676-681 (2014).</li>
	<li>Jongejan, F., Uilenberg, G. The global importance of ticks. Parasitology. 129 Suppl, S3-14 (2004).</li>
	<li>Eisen, R. J., Eisen, L. The blacklegged tick, Ixodes scapularis: An increasing public health concern. Trends in Parasitology. 34 (4), 295-309 (2018).</li>
	<li>Sharma, A. et al. Cas9-mediated gene editing in the black-legged tick, Ixodes scapularis, by embryo injection and ReMOT Control. iScience. 25 (3), 103781 (2022).</li>
	<li>Nuss, A., Sharma, A., Gulia-Nuss, M. Genetic manipulation of ticks: A paradigm shift in tick and tick-borne diseases research. Frontiers in Cellular and Infection Microbiology. 11, 678037 (2021).</li>
	<li>Heinze, S. D. et al. CRISPR-Cas9 targeted disruption of the yellow ortholog in the housefly identifies the brown body locus. Scientific Reports. 7 (1), 4582 (2017).</li>
	<li>Kistler, K. E., Vosshall, L. B., Matthews, B. J. Genome engineering with CRISPR-Cas9 in the mosquito Aedes aegypti. Cell Reports. 11 (1), 51-60 (2015).</li>
	<li>Criscione, F., O&#39;Brochta, D. A., Reid, W. Genetic technologies for disease vectors. Current Opinion in Insect Science. 10, 90-97 (2015).</li>
	<li>Jasinskiene, N. et al. Stable transformation of the yellow fever mosquito, Aedes aegypti, with the Hermes element from the housefly. Proceedings of the National Academy of Sciences of the United States of America. 95 (7), 3743-3747 (1998).</li>
	<li>Dermauw, W. et al. Targeted mutagenesis using CRISPR-Cas9 in the chelicerate herbivore Tetranychus urticae. Insect Biochemistry and Molecular Biology. 120, 103347 (2020).</li>
	<li>Santos, V. T. et al. The embryogenesis of the tick Rhipicephalus (Boophilus) microplus: the establishment of a new chelicerate model system. Genesis (New York, N.Y. 2000). 51 (12), 803-818 (2013).</li>
	<li>Ginsberg, H. S., Lee, C., Volson, B., Dyer, M. C., Lebrun, R. A. Relationships between maternal engorgement weight and the number, size, and fat content of larval Ixodes scapularis (Acari: Ixodidae). Journal of Medical Entomology. 54 (2), 275-280 (2017).</li>
	<li>Feldman-Muhsam, B., Havivi, Y. Accessory glands of Gene&#39;s organ in ticks. Nature. 187 (4741), 964 (1960).</li>
	<li>Kakuda, H., Koga, T., Mori, T., Shiraishi, S. Ultrastructure of the tubular accessory gland in Haemaphysalis longicornis (Acari: Ixodidae). Journal of Morphology. 221 (1), 65-74 (1994).</li>
	<li>Booth, T. F. Wax lipid secretion and ultrastructural development in the egg-waxing (Gen&#233;&#39;s) organ in ixodid ticks. Tissue &#38; Cell. 21 (1), 113-122 (1989).</li>
	<li>Arrieta, M. C., Leskiw, B. K., Kaufman, W. R. Antimicrobial activity in the egg wax of the African cattle tick Amblyomma hebraeum (Acari: Ixodidae). Experimental &#38; Applied Acarology. 39 (3-4), 297-313 (2006).</li>
	<li>Brady, J. A simple technique for making very fine, durable dissecting needles by sharpening tungsten wire electrolytically. Bulletin of the World Health Organization. 32 (1), 143-144 (1965).</li>
</ol>

The authors have nothing to disclose.

This is the first protocol developed to inject early tick embryos successfully. A survival rate of ~4%-8% has been achieved, which is comparable to embryo injection in other well-established insect models5.
As this is the initial protocol, it is anticipated that this protocol will be further refined and specialized to individual tick species. In particular, injection timing will vary from species to species, dependent upon embryogenesis, especially the timing of cellula...

Ticks are vectors of medical and veterinary importance, capable of transmitting a variety of viral, bacterial, protozoan pathogens and nematodes1,2. In the eastern United States, the black-legged tick, Ixodes scapularis, is an important vector of the Lyme disease (LD) pathogen, the spirochete Borrelia burgdorferi. Over 400,000 cases of LD are reported each year in the United States, making it the top vector-borne infectious disease in the US1. In addition to B. burgdorferi, six other microorganisms are transmitted by I. scapularis- including four bacteria (Anaplasma phagocytophilum, B. mayonii, B. miyamotoi, and Ehrlichia muris eauclarensis), one protozoan parasite (Babesia microti), and one virus (Powassan virus), making this tick species a major public health concern3. While tick-borne diseases have become more prevalent in recent years, research on ticks has fallen behind other arthropod vectors, such as mosquitoes, due to the unique biology of ticks&#160;and challenges associated with applying genetic and functional genomic tools4,5.
Gene-editing techniques, particularly CRISPR/Cas9, have now made functional genomics studies feasible in non-model organisms. For creating heritable mutations in an organism, embryo injection remains the preferred method for delivering constructs for altering the germline6,7,8,9. However, until recently4, tick eggs were considered too difficult or even impossible to inject without killing the embryo10,11. A thick wax layer on eggs, hard chorion, and high intra-oval pressure were some of the main obstacles that prevented embryo injection in ticks. Adult, blood-fed I. scapularis deposit a single clutch of up to 2,000 eggs12 over 3-4 weeks (approximately 100 eggs/day). Eggs are laid singly, and each egg is coated with wax that is secreted by protrusions or &#34;horns&#34; of the glandular Gen&#233;&#39;s organ13,14,15&#160;of the mother. This wax protects the eggs from desiccation and contains antimicrobial compounds15. To successfully inject tick eggs, it is important to remove the wax layer, soften the chorion, and desiccate the eggs to decrease the intraoval pressure so that the injection does not irreversibly damage the egg. Understanding the critical importance of embryo injections for successful germline transformation, a protocol for I. scapularis is developed, which can be used to deliver a CRISPR/Cas9 construct and generate stable germline mutations4. In addition to its contribution to I. scapularis research, this protocol could also be optimized for other tick species.

<table><tbody><tr><td>Aluminum silicate capillaries, with filament</td><td>Sutter instruments</td><td>AF100-64-10</td><td>Embryo injection</td></tr><tr><td>Benzalkonium chloride 50% in water, 25 g</td><td>TCI-America</td><td>B0414</td><td>Embryo treatment, 25 g is approximately 25 mL</td></tr><tr><td>Filter paper</td><td>Whatman</td><td>1001-090</td><td>Post-injection care</td></tr><tr><td>Forceps</td><td>Thomas Scientific</td><td>300-101</td><td>Gene`s organ manipulation</td></tr><tr><td>Lab Wipes</td><td>Genesee Scientific</td><td>88-115</td><td></td></tr><tr><td>Microloader tips</td><td>Eppendorf</td><td>930001007</td><td>Loading the pulled needles</td></tr><tr><td>Micromanipulator</td><td>Sutter instruments</td><td>ROE-200</td><td>Embryo injection</td></tr><tr><td>Microscopic slides- plain, ground edges</td><td>Genesee Scientific</td><td>29-100</td><td>Embryo alignment, ground edges are preferred, beveled edges could obscure the eggs from view</td></tr><tr><td>NaCl</td><td>Research Products International</td><td>S23020-500.0</td><td>Embryo treatment</td></tr><tr><td>Needle Puller</td><td>Sutter Instruments</td><td>P-1000</td><td></td></tr><tr><td>Permanent Double sided tape</td><td>Scotch</td><td>34-8716-3417-5</td><td>Embryo alignment</td></tr><tr><td>Petri plates</td><td>Genesee Scientific</td><td>32-107G</td><td>Post-injection care</td></tr><tr><td>Tegaderm/ Transparent film dressing</td><td>3M Healthcare</td><td>1628</td><td>Embryo alignment</td></tr><tr><td>Tungsten needles</td><td>Fine Science Tools</td><td>10130-10</td><td>Gene`s organ manipulation</td></tr><tr><td>Tungsten Wire</td><td>Amazon</td><td>B08DNT7ZK3</td><td>Gene`s organ manipulation</td></tr><tr><td>XenoWorks Digital Microinjector</td><td>Sutter instruments</td><td>MPC-200</td><td>Embryo injection</td></tr></tbody></table>

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.

A successful embryo injection protocol for I. scapularis is described in this article. Egg-laying females were kept at high humidity to avoid desiccation of partially-waxed eggs. The wax layer was removed to inject tick embryos by ablating the Gene&#39;s organ (wax gland) of the gravid female (Figure 1A-E). We used aluminosilicate glass needles with a shorter neck (Figure 2). This shape was ideal for tick egg injection ...

Watch this Scientific Journal Video about Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis at JoVE.com

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.

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 ...

embryo-injection-technique-for-gene-editing-black-legged-tick-ixodes

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

2.4K Views.  University of Nevada, Reno. The present protocol describes a method for injecting tick embryos. Embryo injection is the preferred technique for genetic manipulation to generate transgenic lines.

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

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

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever.  The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue fever. The techniques for correctly preparing microinjection needles, dessicating embryos, and performing microinjection are demonstrated.

In this video, Nijole Jasinskiene demonstrates the methodology employed to generate transgenic Aedes aegypti mosquitoes, which are vectors for dengue ...

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 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, ...

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to visceral pathology. Deciphering the nature of the vector/parasite interactions is of primary importance for better understanding of Leishmania transmission to their hosts. Among the parameters controlling the sand fly vector competence (i.e. their ability to carry and transmit pathogens), parameters intrinsic to these insects were shown to play a key role. Insect immune response, for example, impacts sand fly vector competence to Leishmania. The study of such parameters has been limited by the lack of methods of gene expression modification adapted for use in these non-model organisms. Gene downregulation by small interfering RNA (siRNA) is possible, but in addition to being technically challenging, the silencing leads to only a partial loss of function, which cannot be transmitted from generation to generation. Targeted mutagenesis by CRISPR/Cas9 technology was recently adapted to the Phlebotomus papatasi sand fly. This technique leads to the generation of transmissible mutations in a specifically chosen locus, allowing to study the genes of interest. The CRISPR/Cas9 system relies on the induction of targeted double-strand DNA breaks, later repaired by either Non-Homologous End Joining (NHEJ) or by Homology Driven Repair (HDR). NHEJ consists of a simple closure of the break and frequently leads to small insertion/deletion events. In contrast, HDR uses the presence of a donor DNA molecule sharing homology with the target DNA as a template for repair. Here, we present a sand fly embryo microinjection method for targeted mutagenesis by CRISPR/Cas9 using NHEJ, which is the only genome modification technique adapted to sand fly vectors to date.

Sand Fly Phlebotomus papatasi Embryo Microinjection for CRISPR/Cas9 Mutagenesis

This protocol details the steps of CRISPR/Cas9 targeted mutagenesis in sand flies: embryo collection, injection, insect rearing, and identification as well as selection of mutations of interest.

Sand flies are the natural vectors for Leishmania species, protozoan parasites producing a broad spectrum of symptoms ranging from cutaneous lesions to ...

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how genes regulate social traits, we have developed mutant ants via delivery of CRISPR complex into young embryos during their syncytial stage. Here, we provide a protocol of CRISPR-mediated mutagenesis in Harpegnathos saltator, a ponerine ant species that displays striking phenotypic plasticity. H. saltator ants are readily reared in a laboratory setting. Embryos are collected for microinjection with Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs) using home-made quartz needles. Post-injection embryos are reared outside the colony. Following emergence of the first larva, all embryos and larvae are transported to a nest box with a few nursing workers for further development. This protocol is suitable for inducing mutagenesis for analysis of caste-specific physiology and social behavior in ants, but may also be applied to a broader spectrum of hymenopterans and other insects.

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Many characteristics of insect eusociality rely on within-colony communication and division of labor. Genetic manipulation of key regulatory genes in ant embryos via microinjection and CRISPR-mediated mutagenesis provides insights into the nature of altruistic behavior in eusocial insects.

The unique traits of eusocial insects, such as social behavior and reproductive division of labor, are controlled by their genetic system. To address how ...

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 ...

Developmental Biology

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (H&#252;bner)

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional genomics, and behavioral studies has made H. armigera a model species in Lepidoptera Noctuidae. To study the in vivo functions of and interactions between different genes, clustered regularly interspaced short palindromic repeats (CRISPR)/ associated protein 9 (Cas9) genome editing technology is a convenient and effective method used for performing functional genomic studies. In this study, we provide a step-by-step systematic method to complete gene knockout in H. armigera using the CRISPR/Cas9 system. The design and synthesis of guide RNA (gRNA) are described in detail. Then, the subsequent steps consisting of gene-specific primer design for guide RNA (gRNA) creation, embryo collection, microinjection, insect rearing, and mutant detection are summarized. Finally, troubleshooting advice and notes are provided to improve the efficiency of gene editing. Our method will serve as a reference for the application of CRISPR/Cas9 genome editing in H. armigera as well as other Lepidopteran moths.

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera H&#252;bner

Presented here is a protocol of Helicoverpa armigera (H&#252;bner) embryo microinjection and knockout mutant identification created by CRISPR/Cas9 genome editing. Mutant insects enable further research of gene function and interaction among different genes in vivo.

The cotton bollworm, Helicoverpa armigera, is one of the most destructive pests in the world. A combination of molecular genetics, physiology, functional ...

Behavior

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 ...

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, ...

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

Summary

Explore More Videos

Chapters in this video

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

Microinjection of A. aegypti Embryos to Obtain Transgenic Mosquitoes

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

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

Sand Fly (Phlebotomus papatasi) Embryo Microinjection for CRISPR/Cas9 Mutagenesis

Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator

Injecting Gryllus bimaculatus Eggs

Embryo Microinjection and Knockout Mutant Identification of CRISPR/Cas9 Genome-Edited Helicoverpa Armigera (Hübner)

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

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