The authors thank&#160;Danny Reinberg&#39;s and Claude Desplan&#39;s labs at New York University and J&#252;rgen Liebig&#39;s lab at Arizona State University for their support on ant genetics. Hua Yan acknowledges support from the National Science Foundation I/UCRC, the Center for Arthropod Management Technologies under Grant No. IIP-1821914 and by industry partners. Maya Saar was supported by the United States - Israel Binational Agricultural Research and Development Fund, Vaadia-BARD Postdoctoral Fellowship No. FI-595-19.

1. Regular maintenance of Harpegnathos saltator colonies
<ol>
	<li>Maintain wild-type colonies of H. saltator in transparent plastic boxes in an ant rearing room at 22-25 &#176;C and a photoperiod of 12 hours light: 12 hours dark (12L:12D) lighting schedule.
	<ol>
		<li>Use small boxes (9.5 x 9.5 cm2) to rear individual workers or small colonies. Use medium boxes (19 x 13.5 cm2) or large boxes (27 x 19 cm2) to rear larger colonies (Figure 1).</li>
		<li>To create nest boxes, use plaster to make floors for the boxes. As the wet plaster is drying in the medium and the large boxes, press a foam block into the plaster a few centimeters deep and a few centimeters from the back of the box to designate a lower nest region. Once the plaster has dried, cover the designated nest region with a square piece of glass. 
		NOTE: In small boxes, there is no need to designate a lower nest region. If ants are neglecting to use the designated lower nest region, it may help to cover the glass with a square piece of red cellophane. This gives the impression of a dark underground space resembling nests that H. saltator use in the wild and may encourage them to move their brood to the designated region.</li>
	</ol>
	</li>
	<li>Feed colonies with live crickets twice per week. 
	NOTE: Colonies should be fed enough that they consume all crickets prior to their next feeding. Feed any isolated ants and mutant ant colonies with crickets pre-stung by workers of a regular colony 2-3 times every week.</li>
	<li>Apply water regularly to the plaster nest box flooring using a wash bottle. 
	NOTE: The plaster should be moist enough that it does not feel dusty to the touch, but it should be dry enough that all added water is absorbed by the plaster. It is important that the nests are not watered excessively. On average, nest boxes will need a small amount of water added once per week.</li>
	<li>Whenever feeding occurs, remove trash and dead individuals. Freeze all waste and dead ants overnight at -30 &#176;C before disposing of these materials as regular garbage.</li>
	<li>Add a pinch of dried sawdust to colonies periodically; this helps larvae as they undergo pupation and helps workers keep the nest box clean.</li>
</ol>
2. Preparation of quartz glass microinjection needles
<ol>
	<li>Use a micropipette puller to pull glass microinjection needles.</li>
	<li>Select the glass to be pulled. Ensure that the glass being used has been stored in a dust-free and clean environment. 
	NOTE: Here, thin-walled filamentous quartz glass with an outer diameter of 1.0 mm, inner diameter of 0.5 mm, and length of 7.5 cm has been used to produce microinjection needles. If injecting a soft-bodied embryo, borosilicate needles may also be applicable, but borosilicate needles are not able to penetrate hard chorion.</li>
	<li>Set the parameter settings of the puller. Use a two-step process to pull microinjection needles for H. saltator embryos: parameters for the first step include heat of 575, filament of 3, velocity of 35, delay of 145, and a pull of 75; parameters of the second step include heat of 425, filament of 0, velocity of 15, delay of 128, and a pull of 200. Following the second step, ensure that the resulting needle has a 2 mm taper and a tip of 0.5 &#181;m (Figure 2). 
	NOTE: This set of parameters, along with parameters for other needle types, can be found in the Operation Manual17. Manuals for pipette pullers often provide recommended parameters for a variety of techniques. Some trial and error may be required to determine which parameters generate the best needles for specific needs. A short taper is ideal for H. saltator injections, as it can penetrate the hard chorion of H. saltator embryos. If injecting a soft embryo, such as one that has been dechorionated (e.g., Drosophila), a longer taper around 10 mm may yield better results. Operation manuals for pipette pullers usually provide specific parameters for pulling needles suitable for different needs. It is important that gloves are worn while handling glass filaments and pulling needles. Oils from bare hands may transfer to the glass if gloves are not worn.</li>
	<li>Once parameters are set, use a micropipette puller to pull needles for microinjection. Ensure that needles are kept in a dust-free and clean environment until used. 
	&#8203;NOTE: It is recommended to use freshly pulled microinjection needles. If pre-pulled needles are used, they should be stored properly in a box to prevent damage of needle tips and potential contamination. Needles that have undergone long-term storage are not recommended for embryo injections.</li>
</ol>
3. Preparation of microinjector
<ol>
	<li>Use a microinjector to inject desired materials into ant embryos.</li>
	<li>Prepare the microinjection mixture of Cas9 proteins and in vitro synthesized small guide RNAs (sgRNAs)10. Keep the mixture on ice until it is time to load a microinjection needle. When not in use, store the microinjection mixture at -80 &#176;C. 
	NOTE: The concentrations vary in different species. High concentration may cause high mortality, whereas low concentration may reduce efficiency. We use 0.2 &#181;g/&#181;L Cas9 proteins and 0.2 &#181;g/&#181;L sgRNAs for H. saltator embryo injection. Design of our sgRNAs followed a previously established protocol18. Gene sequences were obtained from DNA databases. The genome sequence of H. saltator was also previously reported19,20.</li>
	<li>Adjust the injection parameters for H. saltator embryos: an injection pressure of 140 hectopascal (hPa), a constant pressure of 70 hPa, and a time of 0.4 seconds. Adjust constant pressure such that material only flows in one direction. Adjust the injection pressure and time only if no material is flowing from the needle into the embryo. 
	NOTE: If injecting a different type of embryo, the primary parameter that may change is constant pressure, which is responsible for ensuring that fluid from the embryo does not flow back into the needle. Volume is not controlled for in this protocol. Setting the parameters of the microinjector is sufficient for obtaining consistent injections.</li>
	<li>Load a microinjection needle with 2 &#181;L of the mixture using microloader pipette tips. Do this slowly to ensure that no bubbles are formed in the mixture. 
	NOTE: If bubbles form, it may be difficult to maintain consistent microinjections.</li>
	<li>Break just the tip of the needle by breaking along the edge of the tape such that a narrow taper is still maintained. Ensure that the needle is broken just enough that the tip is opened, but not so much that the taper is broken off. 
	NOTE: Importantly, if the opening of the needle is too wide after breaking, the user will see liquid run out of the needle when mounted to the pressurized microinjector prior to application of injection pressure. Some microinjection protocols advise breaking needles by cutting the tip with scissors. This method is not advised, as scissors may cause the tip of the needle to shatter. Injecting embryos with a shattered needle will be highly damaging.</li>
	<li>Mount the needle to the micromanipulator</li>
</ol>
4. Injection of embryos
<ol>
	<li>Select embryos for microinjection from the syncytial stage: the time during development in which nuclei divide without cytokinesis. 
	NOTE: This is the ideal time during development for genome editing by microinjection, as discovered previously in Drosophila21. H. saltator embryos pass the syncytial stage and reach cellularization around 36 h after egg deposition10. Higher efficiency is achieved if younger embryos are used for injections.</li>
	<li>Line embryos on a piece of double-sided tape stuck to a glass microscope slide. Ensure that embryos are secured well to the tape to prevent movement during injection. Lay the embryos in a vertical orientation, such that the lateral side of the embryo is at the edge of the tape (Figure 3). Place the slide and lined embryos onto the stage of the microscope at a designated microinjection workstation. 
	NOTE: It is recommended that the embryos are arranged such that successive injections can be performed by moving the slide on the stage instead of adjusting the needle position with every injection. This allows for successive injections to be performed more efficiently.</li>
	<li>Align the needle with the first embryo to be injected using the micromanipulator (Figure 4a).</li>
	<li>Laterally puncture the needle into the first embryo along its dorsal/ventral axis under a microscope.</li>
	<li>Inject the microinjection mixture. Look for slight movement of the embryo, indicating an increase in internal pressure due to the injected liquid. Additionally, watch for the formation of a small droplet containing visible trace of tissue and/or lipid on the outer membrane of the embryo (Figure 4b). 
	&#8203;NOTE: Presence of trace of tissue and/or lipid in the droplet indicates that the needle has successfully punctured both chorion and vitelline membrane of the embryo. If traces of these materials are not present, the injection was not performed successfully and should be repeated. After a few seconds, the droplet will be reabsorbed by the embryo and will no longer be visible.</li>
	<li>Gently remove the needle from the embryo immediately, and proceed to the next by adjusting the position of the microscope slide. Repeat until all the embryos have been injected.</li>
	<li>Once all embryos on the slide have been successfully injected, transfer the slide to a humid box for 1 hour to give the embryos time to recover from the injection process prior to being removed from the slide.</li>
</ol>
5. Rearing of injected embryos
<ol>
	<li>After 1 hour&#160;of incubation in a humid box, gently remove the injected embryos from the tape using featherweight forceps, and transfer them to a tube filled with a small amount of 70% ethanol. Invert the tube several times to transfer the embryos to the bottom of the tube. Repeat the ethanol wash once, followed by three washes with autoclaved water.</li>
	<li>Using a small and soft paintbrush, transfer all injected embryos to 1% agar plates with 2% Antibiotic-Antimycotic. Apply the Antibiotic-Antimycotic after poured agar plates have cooled by spreading over the surface of the plate using a cell spreader. Seal the agar plate with parafilm to prevent agar desiccation. 
	NOTE: Do not attempt to return injected embryos to a colony after injections as workers may destroy most injected embryos. Survival is therefore optimized by allowing embryos to develop on agar plates outside of a normal colony environment.</li>
	<li>Incubate the agar plates at 25 &#176;C for approximately 4 weeks. Check regularly for hatching.</li>
	<li>Once the first embryo has hatched into a larva, return all embryos and larvae to a nest box with a few young nurse workers to care for the hatchlings. Feed using crickets pre-stung by a larger wild-type colony, remove waste products, and add water following the same protocol discussed in Section 1. 
	NOTE: Small boxes (9.5 x 9.5 cm2) are ideal for such colonies.&#160;H. saltator reproduces well in captivity. Therefore, mutant embryos can be reared to adulthood. Isolation of mutant adult(s) induces transition to the reproductive gamergate stage. Controlled crosses are used to establish mutant colonies with heterozygous or homozygous individuals (Figure 5).</li>
</ol>

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The authors have nothing to disclose.

The evolution of eusociality amongst insects, including ants, bees, wasps, and termites, has resulted in the appearance of novel behavioral and morphological traits, many of which are understood to be influenced by a combination of environmental and genetic factors1,2,3,4. Unfortunately, the attractiveness and usefulness of eusocial insects as research models in the field of genetics has been h...

The evolution of eusociality in insects, namely those of the orders Hymenoptera and Blattodea (formerly Isoptera), has resulted in unique and often sophisticated behavioral traits that manifest on both the individual and the colony levels. Reproductive division of labor, a trait characterizing the most advanced groups of social insects, often involves caste systems composed of several behaviorally and often morphologically distinctive groups. Such behavioral and morphological diversity between castes is controlled not only by their genetic system, but also often by the environment1,2,3,4, making eusocial insects attractive subjects for genetic and epigenetic research.
The ability to manipulate the genetic system of eusocial insects has proven to be challenging as many species do not mate and reproduce in laboratory settings. Most eusocial insects also have very few reproductive individuals in a colony, limiting the number of offspring that can be produced and consequently, limiting the sample size for genetic manipulation5. Additionally, many eusocial insects have long generation times compared to insects commonly used for genetic studies (such as Drosophila), adding to the difficulty of establishing genetic lines5. Some eusocial species, however, can generate a large proportion of reproductively active individuals in a colony, which alleviates the challenges and provides opportunities to establish mutant or transgenic lines.
In the case of the ponerine ant species, Harpegnathos saltator, all female workers can become reproductively active upon the death of a queen or social isolation. These workers are referred to as &#34;gamergates&#34; and can be used to generate new colonies6. Furthermore, there may be more than one gamergate present in a colony, thus increasing offspring production5,7,8. Thus far, mutant and/or transgenic lines have been developed in the European honeybee, Apis mellifera, and in the ant species, H. saltator, Ooceraea biroi, and Solenopsis invicta9,10,11,12,13,14,15. Genetic analyses in social bees and ants have paved the way toward a better understanding of eusociality, providing an array of opportunities to study genes and their impacts on eusocial insect behavior and caste-specific physiology.
Here, we provide a protocol for genetic modification via the CRISPR/Cas9 system in H. saltator. Specifically, this technique was used to generate a germline mutation in orco, the gene encoding the obligate co-receptor of all odorant receptors (ORs)10. OR genes have been remarkably expanded in hymenopteran eusocial insects16, and orco plays an essential role in insect olfaction; in its absence, ORs do not assemble or function normally. Mutations of the orco gene therefore disrupt olfactory sensation, neural development, and associated social behaviors9,10.
In this protocol, Cas9 proteins and small guide RNAs (sgRNAs) are introduced into ant embryos using microinjection for the purpose of inducing mutagenesis of a target gene. Here, we will describe the microinjection procedure in detail along with directions regarding the care of colonies and injected embryos. These methods are appropriate for inducing mutagenesis in a variety of different genes in H. saltator ants and may be applied to a broader spectrum of hymenopteran insects.

<table><tbody><tr><td>Antibiotic-Antimycotic (100X)</td><td>ThermoFisher</td><td>15240-062</td><td></td></tr><tr><td>Cas9 protein with NLS, high concentration</td><td>PNA Bio</td><td>CP02</td><td></td></tr><tr><td>Cellophane Roll 20 inch X 5 feet</td><td>Hypogloss Products</td><td>B00254CNJA</td><td>The product has many color variations. Purchase it in red for use in making ant nests.</td></tr><tr><td>Eclipse Ci-S upright microscope&amp;nbsp;</td><td>Nikon</td><td>Ci-S</td><td></td></tr><tr><td>Featherweight forceps, narrow tip</td><td>BioQuip</td><td>4748</td><td></td></tr><tr><td>FemtoJet ll microinjector</td><td>Eppendorf</td><td>920010504</td><td>This product is no longer sold or supported by Eppendorf. A comparable microinjector may be used instead.</td></tr><tr><td>Microloader pipette tips</td><td>Eppendorf</td><td>930001007</td><td></td></tr><tr><td>NCBI database</td><td>National Center for Biotechnology Information</td><td>Gene ID: 105183395&amp;nbsp;</td><td></td></tr><tr><td>P-2000 Micropipette Puller</td><td>Sutter Instruments</td><td>P-2000/G</td><td></td></tr><tr><td>Plastic boxes (19 X 13.5 cm&lt;sup&gt;2&lt;/sup&gt;)</td><td>Pioneer Plastics</td><td>079C&amp;nbsp;</td><td></td></tr><tr><td>Plastic boxes (27 X 19 cm&lt;sup&gt;2&lt;/sup&gt;)</td><td>Pioneer Plastics</td><td>195C</td><td></td></tr><tr><td>Plastic boxes (9.5 X 9.5 cm&lt;sup&gt;2&lt;/sup&gt;)</td><td>Pioneer Plastics</td><td>028C&amp;nbsp;</td><td></td></tr><tr><td>Quartz glass without filament</td><td>Sutter Instruments</td><td>Q100-50-7.5</td><td></td></tr><tr><td>Vannas scissors, 8.5 cm</td><td>World Precision Instruments</td><td>500086</td><td></td></tr><tr><td>Winsor &amp;amp; Newton Cotman Water Colour Series 111 Short Handle Synthetic Brush - Round #000</td><td>Winsor and Newton</td><td>5301030</td><td></td></tr></tbody></table>

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.

Using the protocol provided here, genome editing in Harpegnathos saltator embryos was performed successfully. These results were validated via polymerase chain reaction and pGEM cloning of DNA extracted from injected embryos followed by DNA sequencing. Efficiency of somatic mutagenesis using this protocol reached approximately 40%. F1 mutant males were mated to wild type females to produce heterozygous F2 females which, if not mated, produced F3 males. Mutant F3 males were mated ...

Watch this Scientific Journal Video about Embryo Injections for CRISPR-Mediated Mutagenesis in the Ant Harpegnathos saltator at JoVE.com

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.

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 ...

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2.5K Views.  University of Florida. Dit protocol stelt ons in staat om genetisch gemodificeerde mieren te genereren om de functie van genen in communicatie en gedrag in een sociale omgeving te begrijpen. Het manipuleren van het genetische systeem van eusociale insecten is een uitdaging, omdat veel soorten niet paren en zich voortplanten in laboratoriumomgevingen. De opvallende fenotypeplasticiteit in de hypernasale celvloer van de mier stelt ons in staat om CRISPR-gemedieerde mutantlijnen vast te stellen.Kayli Sieber, ee...

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

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration of transgenes. However, due to the low efficiency of homologous recombination (HR) and various indel mutations of non-homologous end joining (NHEJ)-based strategies in non-dividing cells, in vivo genome editing remains a great challenge. Here, we describe a homology-mediated end joining (HMEJ)-based CRISPR/Cas9 system for efficient in vivo precise targeted integration. In this system, the targeted genome and the donor vector containing homology arms (~800 bp) flanked by single guide RNA (sgRNA) target sequences are cleaved by CRISPR/Cas9. This HMEJ-based strategy achieves efficient transgene integration in mouse zygotes, as well as in hepatocytes in vivo. Moreover, a HMEJ-based strategy offers an efficient approach for correction of fumarylacetoacetate hydrolase (Fah) mutation in the hepatocytes and rescues Fah-deficiency induced liver failure mice. Taken together, focusing on targeted integration, this HMEJ-based strategy provides a promising tool for a variety of applications, including generation of genetically modified animal models and targeted gene therapies.

CRISPR/Cas9-mediated Targeted Integration In Vivo Using a Homology-mediated End Joining-based Strategy

The clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 (CRISPR/Cas9) system provides a promising tool for genetic engineering, and opens up the possibility of targeted integration of transgenes. We describe a homology-mediated end joining (HMEJ)-based strategy for efficient DNA targeted integration in vivo and targeted gene therapies using CRISPR/Cas9.

As a promising genome editing platform, the CRISPR/Cas9 system has great potential for efficient genetic manipulation, especially for targeted integration ...

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 ...

Generation of Chimeric Axolotls with Mutant Haploid Limbs Through Embryonic Grafting

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as axolotls, to regenerate entire limbs as adults. Here, we outline techniques used to generate chimeric axolotls with Cas9-mutagenized haploid forelimbs that can be used for exploring gene function and the fidelity of limb regeneration. We combine several embryological and genetic techniques, including haploid generation via in vitro activation, CRISPR/Cas9 mutagenesis, and tissue grafting into one protocol to produce a unique system for haploid genetic screening in a model organism of regeneration. This strategy reduces the number of animals, space, and time required for the functional analysis of genes in limb regeneration. This also permits the investigation of regeneration-specific functions of genes that may be required for other essential processes, such as organogenesis, tissue morphogenesis, and other essential embryonic processes. The method described here is a unique platform for conducting haploid genetic screening in a vertebrate model system.

This goal of this protocol is to produce chimeric axolotls with haploid forelimbs derived from Cas9-mutagenized donor tissue using embryonic tissue grafting techniques.

A growing set of genetic techniques and resources enable researchers to probe the molecular origins of the ability of some species of salamanders, such as ...

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 ...

Determining the Role of Maternally-Expressed Genes in Early Development with Maternal Crispants

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary for development until zygotic genome activation. Typically, genetic targeting of these maternal factors requires an additional generation to identify maternal-effect phenotypes, hindering the ability to determine the role of maternally-expressed genes during development. The discovery of the biallelic editing capabilities of CRISPR-Cas9 has allowed screening of embryonic phenotypes in somatic tissues of injected embryos or "crispants," augmenting the understanding of the role zygotically-expressed genes play in developmental programs. This article describes a protocol that is an extension of the crispant method. In this method, the biallelic editing of germ cells allows for the isolation of a maternal-effect phenotype in a single generation, or "maternal crispants." Multiplexing guide RNAs to a single target promotes the efficient production of maternal crispants, while sequence analysis of maternal crispant haploids provides a simple method to corroborate genetic lesions that produce a maternal-effect phenotype. The use of maternal crispants supports the rapid identification of essential maternally-expressed genes, thus facilitating the understanding of early development.

Early development is dependent on maternally-inherited products, and the role of many of these products is currently unknown. Herein, we described a protocol that uses CRISPR-Cas9 to identify maternal-effect phenotypes in a single generation.

Early development depends on a pool of maternal factors incorporated into the mature oocyte during oogenesis that perform all cellular functions necessary ...

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 ...

Developmental 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

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by filarial nematodes such as canine heartworm and elephantasis. Recently, CRISPR/Cas9 genome editing has been used to induce site-directed mutations by injecting a Cas9 protein that has been complexed with a guide RNA (gRNA) into freshly laid embryos of several insect species, including mosquitoes that belong to the genera Anopheles and Aedes. Manipulating and injecting Culex mosquitoes is slightly more difficult as these mosquitoes lay their eggs upright in rafts rather than individually like other species of mosquitoes. Here we describe how to design gRNAs, complex them with Cas9 protein, induce female mosquitoes of Culex pipiens to lay eggs, and how to prepare and inject newly laid embryos for microinjection with Cas9/gRNA. We also describe how to rear and screen injected mosquitoes for the desired mutation. The representative results demonstrate that this technique can be used to induce site-directed mutations in the genome of Culex mosquitoes and, with slight modifications, can be used to generate null-mutants in other mosquito species as well.&#160;

Preparing and Injecting Embryos of Culex Mosquitoes to Generate Null Mutations using CRISPR/Cas9

CRISPR/Cas9 is increasingly used to characterize gene function in non-model organisms. This protocol describes how to generate knock-out lines of Culex pipiens, from preparing injection mixes, to obtaining and injecting mosquito embryos, as well as how to rear, cross, and screen injected mosquitoes and their progeny for desired mutations.

Culex mosquitoes are the major vectors of several diseases that negatively impact human and animal health including West Nile virus and diseases caused by ...

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

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