pPV540 and pPV402 were kind gifts from Dr. James Lok at the University of Pennsylvania. We thank Astra Bryant for helpful comments on the manuscript. This work was funded by a Burroughs-Wellcome Fund Investigators in the Pathogenesis of Disease Award, a Howard Hughes Medical Institute Faculty Scholar Award, and National Institutes of Health R01 DC017959 (E.A.H.).

<ol>
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The authors declare no conflicts of interest.

This microinjection protocol details the methods for introducing constructs for transgenesis and CRISPR/Cas9-mediated mutagenesis into S.&#160;stercoralis and S.&#160;ratti. For both S.&#160;stercoralis and S.&#160;ratti, post-injection survival and the rate of transgenesis or mutagenesis are subject to several variables that can be fine-tuned.
The first critical consideration for successful transgenesis is how plasmid transgenes are constructed. Previous st...

Strongyloides stercoralis has long been overlooked as an important human pathogen compared to the more widely recognized hookworms and the roundworm Ascaris lumbricoides1. Previous studies of worm burden often severely underestimated the prevalence of S.&#160;stercoralis due to the low sensitivity of common diagnostic methods for S.&#160;stercoralis2. In recent years, epidemiological studies based on improved diagnostic tools have estimated that the true prevalence of S.&#160;stercoralis infections is much higher than previously reported, approximately 610 million people worldwide2.
Both S.&#160;stercoralis and other Strongyloides species, including the closely related rat parasite and common lab model S.&#160;ratti, have an unusual life cycle that is advantageous for experimental genomic studies because it consists of both parasitic and free-living (environmental) generations3 (Figure 1). Specifically, both S.&#160;stercoralis and S.&#160;ratti can cycle through a single free-living generation. The free-living generation consists of post-parasitic larvae that develop into free-living adult males and females; all progeny of the free-living adults develop into infective larvae, which must infect a host to continue the life cycle. Furthermore, this environmental or free-living generation can be experimentally manipulated in the laboratory. Because free-living Strongyloides adults and C. elegans adults share similar morphology, techniques such as intragonadal microinjection that were originally developed for C. elegans can be adapted for use with free-living adult Strongyloides4,5. While DNA is generally introduced into free-living adult females, both males and females of Strongyloides can be microinjected6. Thus, functional genomic tools are available to interrogate many aspects of the biology of Strongyloides. Other parasitic nematodes lack a free-living generation, and as a result, are not as readily amenable to functional genomic techniques3.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63023/63023fig01.jpg" /> 
Figure 1: The Strongyloides stercoralis life cycle. The S.&#160;stercoralis parasitic females inhabit the small intestine of their mammalian hosts (humans, non-human primates, dogs). The parasitic females reproduce by parthenogenesis and lay eggs within the small intestine. The eggs hatch while still inside the host into post-parasitic larvae, which are then passed into the environment with feces. If the post-parasitic larvae are male, they develop into free-living adult males. If the post-parasitic larvae are female, they can either develop into free-living adult females (indirect development) or third-stage infective larvae (iL3s; direct development). The free-living males and females reproduce sexually to create progeny that are constrained to become iL3s. Under certain conditions, S.&#160;stercoralis can also undergo autoinfection, in which some of the post-parasitic larvae remain inside the host intestine rather than passing into the environment in feces. These larvae can develop into autoinfective larvae (L3a) inside the host, penetrate through the intestinal wall, migrate through the body, and eventually return to the intestine to become reproductive adults. The life cycle of S.&#160;ratti is similar, except that S.&#160;ratti infects rats and does not have an autoinfective cycle. The environmental generation is key to using Strongyloides species for genetic studies. The free-living adult females (P0) can be microinjected; their progeny, which will all become iL3s, are the potential F1 transgenics. This figure has been modified from Castelletto et al.3. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/63023/63023fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
S.&#160;stercoralis shares many aspects of its biology with other gastrointestinal human-parasitic nematodes, including host invasion and host immune modulation. For example, human-parasitic hookworms in the genera Necator and Ancylostoma also infect by skin penetration, navigate similarly through the body, and ultimately reside as parasitic adults in the small intestine7. Thus, many gastrointestinal nematodes likely use common sensory behaviors and immune evasion techniques. As a result, the knowledge gleaned from Strongyloides will complement findings in other less genetically tractable nematodes and lead to a more complete understanding of these complex and important parasites.
This microinjection protocol outlines the method for introducing DNA into Strongyloides free-living adult females to make transgenic and mutant progeny. The strain maintenance requirements, including the developmental timing of adult worms for microinjections and the collection of transgenic progeny, are described. Protocols and a demonstration of the complete microinjection technique, along with protocols for culturing and screening transgenic progeny, are included, along with a list of all necessary equipment and consumables.

<table><tbody><tr><td>(&amp;minus;)-Nicotine, &amp;ge;99% (GC), liquid</td><td>Sigma-Aldrich</td><td>N3876-5ML</td><td>nicotine for paralyzing worms</td></tr><tr><td>3&quot; iron C-clamp, 3&quot; x 2&quot; (capacity x depth)</td><td>VWR</td><td>470121-790</td><td>C-clamp to secure setup to bench top</td></tr><tr><td>Agarose LE</td><td>Phenix</td><td>RBA-500</td><td>agarose for slides</td></tr><tr><td>Bone char, 4 lb pail, 10 x 28 mesh</td><td>Ebonex</td><td>n/a</td><td>charcoal for fecal-charcoal cultures</td></tr><tr><td>Bone char, granules, 10 x 28 mesh</td><td>Reade</td><td>bonechar10x28</td><td>charcoal for fecal-cultures (alternative to the above)</td></tr><tr><td>Coarse micromanipulator</td><td>Narishige</td><td>MMN-1</td><td>coarse micromanipulator</td></tr><tr><td>Corning Costar Spin-X centrifuge tube filters</td><td>Fisher</td><td>07-200-385</td><td>microfilter column</td></tr><tr><td>Cover glass, 48 x 60 mm, No. 1 thickness</td><td>Brain Research Lab</td><td>4860-1</td><td>coverslips (48 x 60 mm)</td></tr><tr><td>Deep Petri dishes, heavy version with 6 vents, 100 mm diameter</td><td>VWR</td><td>82050-918</td><td>10 cm Petri dishes (for fecal-charcoal cultures)</td></tr><tr><td>Eisco retort base w/ rod</td><td>Fisher</td><td>12-000-101</td><td>stand for Baermann apparatus</td></tr><tr><td>Eppendorf FemtoJet microinjector microloader tips</td><td>VWR</td><td>89009-310</td><td>for filling microinjection needles</td></tr><tr><td>Fisherbrand absorbent underpads</td><td>Fisher</td><td>14-206-62</td><td>bench paper (for prepping)</td></tr><tr><td>Fisherbrand Cast-Iron Rings</td><td>Fisher</td><td>14-050CQ</td><td>Baermann o-ring</td></tr><tr><td>Fisherbrand tri-cornered polypropylene beakers</td><td>Fisher</td><td>14-955-111F</td><td>Plastic beaker (for mixing)</td></tr><tr><td>Fisherbrand tri-cornered polypropylene beakers</td><td>Fisher</td><td>14-955-111F</td><td>Plastic beaker (for catch bucket/water bucket)</td></tr><tr><td>Fisherbrand tri-cornered polypropylene beakers</td><td>Fisher</td><td>14-955-111F</td><td>Plastic beaker (x2) (to make holder)</td></tr><tr><td>Gorilla epoxie in syringe</td><td>McMaster-Carr</td><td>7541A51</td><td>glue (to attach tubing)</td></tr><tr><td>Halocarbon oil 700</td><td>Sigma-Aldrich</td><td>H8898-50ML</td><td>halocarbon oil</td></tr><tr><td>High-temperature silicone rubber tubing for food and beverage, 1/2&quot; ID, 5/8&quot; OD</td><td>McMaster-Carr</td><td>3038K24</td><td>tubing (for funnel)</td></tr><tr><td>KIMAX funnels, long stem, 60&amp;deg; Angle, Kimble Chase</td><td>VWR</td><td>89001-414</td><td>Baermann funnel</td></tr><tr><td>Kimberly-Clark Professional Kimtech Science benchtop protectors</td><td>Fisher</td><td>15-235-101</td><td>bench paper (for prepping)</td></tr><tr><td>Leica stereomicroscope with fluorescence</td><td>Leica</td><td>M165 FC</td><td>GFP stereomicroscope for identifying and sorting transgenic worms</td></tr><tr><td>microINJECTOR brass straight arm needle-holder</td><td>Tritech</td><td>MINJ-4</td><td>microinjection needle holder</td></tr><tr><td>microINJECTOR system</td><td>Tritech</td><td>MINJ-1</td><td>microinjection system</td></tr><tr><td>Mongolian Gerbils</td><td>Charles River Laboratories</td><td>213-Mongolian Gerbil</td><td>gerbils for maintenance of S. stercoralis, male 4-6 weeks</td></tr><tr><td>Nasco Whirl-Pak easy-to-close bags, 18 oz</td><td>VWR</td><td>11216-776</td><td>Whirl-Pak sample bags</td></tr><tr><td>Nylon tulle (mesh)</td><td>Jo-Ann Fabrics</td><td>zprd_14061949a</td><td>nylon mesh for Baermann holder</td></tr><tr><td>Platinum wire, 36 Gauge, per inch</td><td>Thomas Scientific</td><td>1233S72</td><td>platinum/iridium wire for worm picks</td></tr><tr><td>Puritan tongue depressors, 152 mm (L) x 17.5 mm (W)</td><td>VWR</td><td>62505-007</td><td>wood sticks (for mixing samples)</td></tr><tr><td>QIAprep Spin Miniprep Kit (250)</td><td>QIAGEN</td><td>27106</td><td>QIAGEN miniprep kit</td></tr><tr><td>Rats-Long Evans</td><td>Envigo</td><td>140 HsdBlu:LE Long Evans</td><td>rats for maintenance of S. ratti, female 4-6 weeks</td></tr><tr><td>Rats-Sprague Dawley</td><td>Envigo</td><td>002 Hsd:Sprague Dawley SD</td><td>rats for maintenance of S. ratti, female 4-6 weeks</td></tr><tr><td>Really Useful Boxes translucent storage boxes with lids, 1.6 L capacity, 7-5/8&quot; x 5-5/16&quot; x 4-5/16&quot;</td><td>Office Depot</td><td>452369</td><td>plastic boxes for humidified chamber</td></tr><tr><td>Shepherd techboard, 8 x 16.5 inches</td><td>Newco</td><td>999589</td><td>techboard</td></tr><tr><td>Stainless steel raised wire floor</td><td>Ancare</td><td>R20SSRWF</td><td>wire cage bottoms</td></tr><tr><td>StalkMarket compostable cutlery spoons, 6&quot;, white, pack of 1,000</td><td>Office Depot</td><td>9587303</td><td>spoons</td></tr><tr><td>Stender dish, stacking type, 37 x 25 mm</td><td>Carolina (Science)</td><td>741012</td><td>watch glasses (small, round)</td></tr><tr><td>Stereomicroscope</td><td>Motic</td><td>K-400 LED</td><td>dissecting prep scope</td></tr><tr><td>Storage tote, color clear/white, outside height 4-7/8 in, outside length 13-5/8 in, Sterilite</td><td>Grainger</td><td>53GN16</td><td>plastic boxes for humidified chamber</td></tr><tr><td>Sutter P-30 micropipette puller</td><td>Sutter</td><td>P-30/P</td><td>needle puller with platinum/iridium filament</td></tr><tr><td>Syracuse watch glasses</td><td>Fisher</td><td>S34826</td><td>watch glasses (large, round)</td></tr><tr><td>Thermo Scientific Castaloy fixed-angle clamps</td><td>Fisher</td><td>05-769-2Q</td><td>funnel clamps (2x)</td></tr><tr><td>Three-axis hanging joystick oil hydrolic micromanipulator</td><td>Narishige</td><td>MM0-4</td><td>fine micromanipulator</td></tr><tr><td>United Mohr pinchcock clamps</td><td>Fisher</td><td>S99422</td><td>Pinch clamps (2x)</td></tr><tr><td>Vented, sharp-edge Petri dishes (60 mm diameter)</td><td>Tritech Research</td><td>T3308P</td><td>6 cm Petri dishes (for small-scale fecal-charcoal cultures)</td></tr><tr><td>VWR light-duty tissue wipers</td><td>VWR</td><td>82003-820</td><td>lining for Baermann holder</td></tr><tr><td>watch glass, square, 1-5/8 in</td><td>Carolina (Science)</td><td>742300</td><td>watch glasses (small, square)</td></tr><tr><td>Whatman qualitative grade plain circles, grade 1, 5.5 cm diameter</td><td>Fisher</td><td>09-805B</td><td>filter paper (for 6 cm Petri dishes)</td></tr><tr><td>Whatman qualitative grade plain circles, grade 1, 9 cm diameter</td><td>Fisher</td><td>09-805D</td><td>filter paper (for 10 cm Petri dishes)</td></tr><tr><td>World Precision Instrument borosilicate glass capillary, 1.2 mm x 4 in</td><td>Fisher</td><td>50-821-813</td><td>glass capillaries for microinjection needles</td></tr><tr><td>X-Acto Knives, No. 1 Knife With No. 11 Blade</td><td>Office Depot</td><td>238816</td><td>X-Acto knives without blades to hold worm picks</td></tr><tr><td>Zeiss AxioObserver A1</td><td>Zeiss</td><td>n/a</td><td>inverted microscope</td></tr></tbody></table>

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.

If the experiment was successful, the F1 larvae will express the transgene and/or mutant phenotype of interest (Figure 4). However, transformation rates are highly variable and depend on the constructs, the health of the worms, the post-injection culturing conditions, and the skill of the experimenter. In general, a successful experiment will yield &#62;15 F1 larvae per injected female and a transformation rate of &#62;3% for fluorescent markers. If the total number of ...

Watch this Scientific Journal Video about Generating Transgenics and Knockouts in Strongyloides Species by Microinjection at JoVE.com

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.

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 ...

generating-transgenics-knockouts-strongyloides-species

Research

JoVE Journal

Immunology and Infection

2.6K Views.  University of California, Los Angeles. 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. stercoralis and S. ratti.

Video: Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

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

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 Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

Biology

Generation of Genetically Modified Mice through the Microinjection of Oocytes

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear injection of DNA expression constructs into fertilized oocytes remains the most commonly used technique to generate transgenic mice for overexpression. With the introduction of CRISPR technology for gene targeting, pronuclear injection into fertilized oocytes has been extended to the generation of both knockout and knockin mice. This work describes the preparation of DNA for injection and the generation of CRISPR guides for gene targeting, with a particular emphasis on quality control. The genotyping procedures required for the identification of potential founders are critical. Innovative genotyping strategies that take advantage of the "multiplexing" capabilities of CRISPR are presented herein. Surgical procedures are also outlined. Together, the steps of the protocol will allow for the generation of genetically modified mice and for the subsequent establishment of mouse colonies for a plethora of research fields, including immunology, neuroscience, cancer, physiology, development, and others.

The microinjection of mouse oocytes is commonly used for both classic transgenesis (i.e., the random integration of transgenes) and CRISPR-mediated gene targeting. This protocol reviews the latest developments in microinjection, with a particular emphasis on quality control and genotyping strategies.

The use of genetically modified mice has significantly contributed to studies on both physiological and pathological in vivo processes. The pronuclear ...

Developmental Biology

phiC31 Integrase-Mediated Embryo Transgenesis in Drosophila

Embryo Microinjection for Transgenesis in Drosophila

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the construction of transgenic flies. Microinjection requires some types of equipment, including a microinjector, a micromanipulator, an inverted microscope, and a stereo microscope.&#160;Plasmids isolated with a plasmid miniprep kit are qualified for microinjection. Embryos at the pre-blastoderm or syncytial blastoderm stage, where nuclei share a common cytoplasm, are subjected to microinjection. A cell strainer eases the process of dechorionating embryos. The optimal time for dechorionation and desiccation of embryos needs to be determined experimentally. To increase the efficiency of embryo microinjection, needles prepared by a puller need to be beveled by a needle grinder. In the process of grinding needles, we utilize a foot air pump with a pressure gauge to avoid the capillary effect of the needle tip. We routinely inject 120-140 embryos for each plasmid and obtain at least one transgenic line for around 85% of plasmids. This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and&#160;presents a detailed protocol for embryo microinjection for transgenesis in Drosophila.

Author Spotlight: Optimizing Embryo Microinjection for Transgenesis in Drosophila

This article takes the phiC31 integrase-mediated transgenesis in Drosophila as an example and presents an optimized protocol for embryo microinjection,&#160;a crucial step for creating transgenic flies.

Transgenesis in Drosophila is an essential approach to studying gene function at the organism level. Embryo microinjection is a crucial step for the ...

Generation of Transgenic C. elegans by Biolistic Transformation

The number of laboratories using the free living nematode C. elegans is rapidly growing. The popularity of this biological model is attributed to a rapid generation time and short life span, easy and inexpensive maintenance, fully sequenced genome, and array of RNAi resources and mutant animals. Additionally, analysis of the C. elegans genome revealed a great similarity between worms and higher vertebrates, which suggests that research in worms could be an important adjunct to studies performed in whole mice or cultured cells. A powerful and important part of worm research is the ability to use transgenic animals to study gene localization and function. Transgenic animals can be created either via microinjection of the worm germline or through the use of biolistic bombardment. Bombardment is a newer technique and is less familiar to a number of labs. Here we describe a simple protocol to generate transgenic worms by biolistic bombardment with gold particles using the Bio-Rad PDS-1000 system. Compared with DNA microinjection into hermaphrodite germline, this protocol has the advantage of not requiring special skills from the operator with regards to identifying worm anatomy or performing microinjection. Further multiple transgenic lines are usually obtained from a single bombardment. Also in contrast to microinjection, biolistic bombardment produces transgenic animals with both extrachromosomal arrays and integrated transgenes. The ability to obtain integrated transgenic lines can avoid the use of mutagenic protocols to integrate foreign DNA. In conclusion, biolistic bombardment can be an attractive method for the generation of transgenic animals, especially for investigators not interested in investing the time and effort needed to become skilled at microinjection.

Generation of Transgenic C. elegans by Biolistic Transformation

Transgenic worms are commonly used in C. elegans research. Described is a simple, yet effective, protocol to introduce transgenes into worms using biolistic bombardment with DNA-coated gold particles. The effort involved and results of bombardment compare favorably with microinjection for the generation of transgenic animals.

The number of laboratories using the free living nematode C. elegans is rapidly growing. The popularity of this biological model is attributed to a rapid ...

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily produced by electroporation of CRISPR factors (and single-stranded oligo DNA donors) into the zygote. In contrast, gene cassette (&#62;1 kb) knock-in and floxed mice are mainly generated by microinjection of CRISPR factors and double-stranded DNA donors into zygotes. Genome editing technologies have also increased the flexibility of genetically modified mice production. It is now possible to introduce the intended mutations in the target genomic regions in a number of beneficial inbred mouse strains. Our team has produced over 200 gene cassette knock-in mouse lines, and over 110 floxed mouse lines by zygote microinjection of CRISPR-Cas9 following requests from several countries, including Japan. Some of these genome editing used BALB/c, C3H/HeJ, and C57BL/6N inbred strains, however most used C57BL/6J. Unlike the electroporation method, genome editing by zygote microinjection in various inbred strains of mice is not that easy. However, gene cassette knock-in and floxed mice on single inbred genetic backgrounds are as critical as genetic humanized, fluorescent reporter, and conditional knockout mouse models. Therefore, this article presents the protocol for the zygote microinjection of CRISPR factors and double-stranded DNA donors in C57BL/6J mice for generating gene cassette knock-in and floxed mice. This article exclusively focuses on nuclear injection rather than cytoplasmic injection. In addition to zygote microinjection, we outline the timeline for the production process and peripheral techniques such as induction of superovulation and embryo transfer.

The present protocol describes zygote microinjection of CRISPR-Cas9 and donor DNA to efficiently produce gene cassette knock-in and floxed mice.

CRISPR-Cas technology has enabled the rapid and effortless generation of genetically modified mice. Specifically, mice and point mutant mice are readily ...

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 ...

A Method for Microinjection of Patiria minata Zygotes

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

A Method for Microinjection of Patiria minata Zygotes

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

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

Gonad Microinjection: A Method of Compound Delivery Directly into the Germline of C. elegans

Source: Farboud, B. et al, <a href="https://www-jove-com.remotexs.ntu.edu.sg/video/57350/enhanced-genome-editing-with-cas9-ribonucleoprotein-diverse-cells">Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms.</a>&#160;J. Vis. Exp. (2018).
This video describes microinjection, a common method of creating transgenic C. elegans strains.&#160; In the example, we will see microinjection used to introduce a ribonucleoprotein (RNP) complex for CRISPR-bases gene editing.

Source: Farboud, B. et al, Enhanced Genome Editing with Cas9 Ribonucleoprotein in Diverse Cells and Organisms.&#160;J. Vis. Exp. (2018).
This video ...

Generating Transgenics and Knockouts in Strongyloides Species by Microinjection

Summary

Explore More Videos

Chapters in this video

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

Embryo Microinjection and Transplantation Technique for Nasonia vitripennis Genome Manipulation

Generation of Stable Transgenic C. elegans Using Microinjection

Generation of Genetically Modified Mice through the Microinjection of Oocytes

Embryo Microinjection for Transgenesis in Drosophila

Generation of Transgenic C. elegans by Biolistic Transformation

Zygote Microinjection for Creating Gene Cassette Knock-in and Flox Alleles in Mice

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

A Method for Microinjection of Patiria minata Zygotes

Gonad Microinjection: A Method of Compound Delivery Directly into the Germline of C. elegans