Name: pupal and adult injections for rnai and crispr gene editing in nasonia vitripennis
Uploaded: 2020-12-04T14:00:00
Description: Watch this Scientific Journal Video about Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis at JoVE.com

This work was supported in part by UCSD startup funds directed to O.S.A. and NSF/BIO grant 1645331 to J.L.R.

1. Nasonia rearing
<ol>
	<li>Set up ~20 mated females singularly in small glass test tubes plugged with cotton. 
	NOTE: To reduce rearing space, 10 x 85 cm, 4 mL tubes are optimal. Females are easily distinguished from males because of the larger wings and the presence of the ovipositor (Figure 1A). Detailed protocols for rearing N. vitripennis can also be found in other literature13,14,15...

<ol>
	<li>Verhulst, E. C., Beukeboom, L. W., van de Zande, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maternal+control+of+haplodiploid+sex+determination+in+the+wasp+Nasonia.">Maternal control of haplodiploid sex determination in the wasp Nasonia.</a> Science. 328 (5978), 620-623 (2010).</li><li>Verhulst, E. C., Lynch, J. A., Bopp, D., Beukeboom, L. W., van de Zande, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+component+of+the+Nasonia+sex+determining+cascade+is+maternally+silenced+and+regulates+transformer+expression.">A new component of the Nasonia sex determining cascade is maternally silenced and regulates transformer expression.</a> PloS One. 8 (5), 63618 (2013).</li><li>Dalla Benetta, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genome+elimination+mediated+by+gene+expression+from+a+selfish+chromosome.">Genome elimination mediated by gene expression from a selfish chromosome.</a> Science Advances. 6 (14), (2020).</li><li>Aldrich, J. C., Leibholz, A., Cheema, M. S., Ausi&#972;, J., Ferree, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+"selfish"+B+chromosome+induces+genome+elimination+by+disrupting+the+histone+code+in+the+jewel+wasp+Nasonia+vitripennis.">A "selfish" B chromosome induces genome elimination by disrupting the histone code in the jewel wasp Nasonia vitripennis.</a> Scientific Reports. 7, 42551 (2017).</li><li>Ferree, P. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+genes+uniquely+expressed+in+the+germ-line+tissues+of+the+jewel+wasp+Nasonnia+vitripennis.">Identification of genes uniquely expressed in the germ-line tissues of the jewel wasp Nasonnia vitripennis.</a> G3. 3 (12), 2647-2653 (2015).</li><li>Akbari, O. S., Antoshechkin, I., Hay, B. A., Ferree, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transcriptome+profiling+of+Nasonia+vitripennis+testis+reveals+novel+transcripts+expressed+from+the+selfish+B+chromosome,+paternal+sex+ratio.">Transcriptome profiling of Nasonia vitripennis testis reveals novel transcripts expressed from the selfish B chromosome, paternal sex ratio.</a> G3. 3 (9), 1597-1605 (2013).</li><li>Dalla Benetta, E., Beukeboom, L. W., van de Zande, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adaptive+differences+in+circadian+clock+gene+expression+patterns+and+photoperiodic+diapause+induction+in+Nasonia+vitripennis.">Adaptive differences in circadian clock gene expression patterns and photoperiodic diapause induction in Nasonia vitripennis.</a> The American Naturalist. 193 (6), 881-896 (2019).</li><li>Paolucci, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Latitudinal+variation+in+circadian+rhythmicity+in+Nasonia+vitripennis.">Latitudinal variation in circadian rhythmicity in Nasonia vitripennis.</a> Behavioral Sciences. 9 (11), (2019).</li><li>Werren, J. H., Loehlin, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+parasitoid+wasp+Nasonia:+an+emerging+model+system+with+haploid+male+genetics.">The parasitoid wasp Nasonia: an emerging model system with haploid male genetics.</a> Cold Spring Harbor Protocols. 2009 (10), (2009).</li><li>Lynch, J. A., Desplan, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+for+parental+RNA+interference+in+the+wasp+Nasonia+vitripennis.">A method for parental RNA interference in the wasp Nasonia vitripennis.</a> Nature Protocols. 1 (1), 486-494 (2006).</li><li>Li, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Generation+of+heritable+germline+mutations+in+the+jewel+wasp+Nasonia+vitripennis+using+CRISPR/Cas9.">Generation of heritable germline mutations in the jewel wasp Nasonia vitripennis using CRISPR/Cas9.</a> Scientific Reports. 7 (1), 901 (2017).</li><li>Chaverra-Rodriguez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Germline+mutagenesis+of+Nasonia+vitripennis+through+ovarian+delivery+of+CRISPR-Cas9+ribonucleoprotein.">Germline mutagenesis of Nasonia vitripennis through ovarian delivery of CRISPR-Cas9 ribonucleoprotein.</a> Insect Molecular Biology. , (2020).</li><li>Li, M., Bui, M., Akbari, O. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryo+microinjection+and+transplantation+technique+for+Nasonia+vitripennis+genome+manipulation.">Embryo microinjection and transplantation technique for Nasonia vitripennis genome manipulation.</a> Journal of Visualized Experiments. (130), (2017).</li><li>Werren, J. H., Loehlin, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+Sarcophaga+bullata+fly+hosts+for+Nasonia+(Parasitoid+Wasp).">Rearing Sarcophaga bullata fly hosts for Nasonia (Parasitoid Wasp).</a> Cold Spring Harbor Protocols. 2009 (10), (2009).</li><li>Werren, J. H., Loehlin, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Strain+maintenance+of+Nasonia+vitripennis+(Parasitoid+Wasp).">Strain maintenance of Nasonia vitripennis (Parasitoid Wasp).</a> Cold Spring Harbor Protocols. 2009 (10), (2009).</li><li>Beukeboom, L. W., van de Zande, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetics+of+sex+determination+in+the+haplodiploid+wasp+Nasonia+vitripennis+(Hymenoptera:+Chalcidoidea).">Genetics of sex determination in the haplodiploid wasp Nasonia vitripennis (Hymenoptera: Chalcidoidea).</a> Journal of Genetics. 89 (3), 333-339 (2010).</li><li>Leung, K., van de Zande, L., Beukeboom, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Life-history+traits+of+the+Whiting+polyploid+line+of+the+parasitoid.">Life-history traits of the Whiting polyploid line of the parasitoid.</a> Entomologia experimentalis et applicata. 167 (7), 655-669 (2019).</li><li>Kennerdell, J. R., Carthew, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+dsRNA-mediated+genetic+interference+to+demonstrate+that+frizzled+and+frizzled+2+act+in+the+wingless+pathway.">Use of dsRNA-mediated genetic interference to demonstrate that frizzled and frizzled 2 act in the wingless pathway.</a> Cell. 95 (7), 1017-1026 (1998).</li><li>Yang, J., Zhao-jun, H. A. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Efficiency+of+different+methods+for+dsrna+delivery+in+cotton+bollworm+(Helicoverpa+armigera).">Efficiency of different methods for dsrna delivery in cotton bollworm (Helicoverpa armigera).</a> Journal of Integrative Agriculture. 13 (1), 115-123 (2014).</li><li>Brown, S., Holtzman, S., Kaufman, T., Denell, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+Tribolium+Deformed+ortholog+and+its+ability+to+directly+regulate+Deformed+target+genes+in+the+rescue+of+a+Drosophila+Deformed+null+mutant.">Characterization of the Tribolium Deformed ortholog and its ability to directly regulate Deformed target genes in the rescue of a Drosophila Deformed null mutant.</a> Development Genes and Evolution. 209 (7), 389-398 (1999).</li><li>Hughes, C. L., Kaufman, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=RNAi+analysis+of+Deformed,+proboscipedia+and+Sex+combs+reduced+in+the+milkweed+bug+Oncopeltus+fasciatus:+novel+roles+for+Hox+genes+in+the+hemipteran+head.">RNAi analysis of Deformed, proboscipedia and Sex combs reduced in the milkweed bug Oncopeltus fasciatus: novel roles for Hox genes in the hemipteran head.</a> Development. 127 (17), 3683-3694 (2000).</li><li>Dzitoyeva, S., Dimitrijevic, N., Manev, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intra-abdominal+injection+of+double-stranded+RNA+into+anesthetized+adult+Drosophila+triggers+RNA+interference+in+the+central+nervous+system.">Intra-abdominal injection of double-stranded RNA into anesthetized adult Drosophila triggers RNA interference in the central nervous system.</a> Molecular Psychiatry. 6 (6), 665-670 (2001).</li><li>Chaverra-Rodriguez, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+delivery+of+CRISPR-Cas9+ribonucleoprotein+into+arthropod+ovaries+for+heritable+germline+gene+editing.">Targeted delivery of CRISPR-Cas9 ribonucleoprotein into arthropod ovaries for heritable germline gene editing.</a> Nature Communications. 9 (1), 3008 (2018).</li><li>Macias, V. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cas9-mediated+gene-editing+in+the+malaria+mosquito+anopheles+stephensi+by+ReMOT+control.">Cas9-mediated gene-editing in the malaria mosquito anopheles stephensi by ReMOT control.</a> G3. 10 (4), 1353-1360 (2020).</li><li>Heu, C. C., McCullough, F. M., Luan, J., Rasgon, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CRISPR/Cas9-based+genome+editing+in+the+silverleaf+whitefly+(Bemisia+tabaci).">CRISPR/Cas9-based genome editing in the silverleaf whitefly (Bemisia tabaci).</a> CRISPR Journal. 3 (2), 89-96 (2020).</li><li>Shirai, Y., Daimon, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mutations+in+cardinal+are+responsible+for+the+red-1+and+peach+eye+color+mutants+of+the+red+flour+beetle+Tribolium+castaneum.">Mutations in cardinal are responsible for the red-1 and peach eye color mutants of the red flour beetle Tribolium castaneum.</a> Biochemical and Biophysical Research Communications. 529 (2), 372-378 (2020).</li><li>Hunter, W. B., Gonzalez, M. T., Tomich, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BAPC-assisted+CRISPR/Cas9+system:+targeted+delivery+into+adult+ovaries+for+heritable+germline+gene+editing+(Arthropoda:+Hemiptera).">BAPC-assisted CRISPR/Cas9 system: targeted delivery into adult ovaries for heritable germline gene editing (Arthropoda: Hemiptera).</a> bioRxiv. , 478743 (2018).</li><li>Chaverra Rodriguez, D., Bui, M., Li, M., Raban, R., Akbari, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developing+genetic+tools+to+control+ACP.">Developing genetic tools to control ACP.</a> Citrus Research Board Citrograph Magazine. 11 (1), 64-68 (2020).</li></ol>

With the recent increased use of Nasonia vitripennis as a model organism for various biological questions2,3,7,17, there is a need to develop and optimize injection methods to enable a simplified and efficient protocol for the functional analysis of N. vitripennis genes. The current methods involving embryonic microinjection of gene editing reagents are challenging<sup class="...

CRISPR/Cas9 gene editing is a powerful technology for functional genetic studies, especially in many rising model organisms such as the jewel wasp, Nasonia vitripennis. The ease of rearing and the availability of a complete genome makes the jewel wasp an important experimental system for elucidating the molecular mechanisms of different biological processes. For example, N. vitripennis has recently been used to unravel the epigenetic basis of the haplodiploid sex determination system1,2, the biology of B-chromosomes3,4,5,6, and the genetic basis for circadian and seasonal regulation7,8. Some of the features that make N. vitripennis amenable to work with include short generation time (~2 weeks at 25 &#176;C), high reproduction rates, easy sex separation at the pupal stage, and the ability to diapause and store strains at 4 &#176;C. The life cycle begins with female wasps parasitizing the pupae of the blowfly, Sarcophaga bullata. Through their ovipositor, females lay up to 50 eggs in the pupal case of the blowfly. Eggs develop into larvae that feed on the S. bullata pupa, continue to develop over the next several days, and then pupate, followed by adult eclosion and emergence from the host puparium9.
Molecular tools to perform functional genetic studies in N. vitripennis, such as RNA interference (RNAi)10 and CRISPR/Cas911,12, are available, but are limited, primarily due to difficulties in performing embryonic microinjections13. As N. vitripennis eggs require a pupal host for development, egg manipulation is very challenging. Pre-blastoderm stage embryos must be collected from the host blowfly pupae, quickly microinjected, and immediately transferred back to the host for development13. These steps require precision and specialized training to avoid damaging the microinjected embryos or the pupal hosts13. Moreover, the eggs are very small and fragile, especially after microinjections, with a very viscous cytoplasm causing a continuous clogging of the injecting needle13. These features make embryonic microinjections exceptionally challenging, requiring highly trained operators and specialized equipment that is absent in most N. vitripennis laboratories.
The optimization of alternative injection methods for the delivery of CRISPR reagents would contribute to the consolidation of N. vitripennis as a model organism. The manipulation of pupae and adults is less challenging than manipulating embryos and can be accomplished with a basic injection setup. Here, two protocols are described for injection of pupae and adults: one involving specialized equipment for injections, and the other involving the use of an aspirator tube assembly fitted with a glass capillary needle. The use of an aspirator tube is particularly suited for laboratories that do not have access to specialized equipment for embryo microinjections. Efficient injections of different developmental stages of N. vitripennis, including white or black pupae and adult wasps, are demonstrated. Wasps at the white pupal stage are particularly suited for RNAi-mediated knockdown experiments. Although RNAi in Nasonia was first described by Lynch and Desplan in 200610, there is no visual procedure available for how RNAi injections are performed. RNAi was recently used to discover the haploidizer gene of the B-chromosome PSR (Paternal Sex Ratio)3 and to study the involvement of the clock gene, period, in N. vitripennis biological rhythms7.
Black pupae and adult wasps can be used to induce CRISPR/Cas9 germline gene editing using ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) and BAPC (Branched Amphiphilic Peptide Capsules) protocols. These two ovary delivery methods have been recently described to be effective in Nasonia for generating germline mutations in the target gene, cinnabar7,12. Here, a simplified protocol is provided for injections including a visual procedure of a step-by-step methodology for both pupal and adult injections that can be utilized to generate functional genetics studies in Nasonia and likely in other parasitoid wasps, without requiring specialized equipment and bypassing embryonic microinjections.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 x 15 mm Stackable Petri Dishes, Polystyrene, Mono, Sterile</td>
			<td>Sigma</td>
			<td>960-97693-083</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminosilicate glass capillary tubing 1 mm(outside diameter) x 0.58 mm (inner diameter)</td>
			<td>Sutter Instruments</td>
			<td>BF100-58-10</td>
			<td>Can also use Borosilicate or Quartz</td>
		</tr>
		<tr>
			<td>Aspirator tube assemblies for calibrated microcapillary pipettes</td>
			<td>Sigma</td>
			<td>A5177-5EA</td>
			<td></td>
		</tr>
		<tr>
			<td>BAPC</td>
			<td>phoreus biotech</td>
			<td>BAPtofect-25 0.5 mg Kit</td>
			<td></td>
		</tr>
		<tr>
			<td>Cas9 Protein with NLS</td>
			<td>PNABio</td>
			<td>CP01</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting needle</td>
			<td>VWR</td>
			<td>10806-330</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase/RNase-Free distilled Water</td>
			<td>Invitrogen</td>
			<td>10977-015</td>
			<td></td>
		</tr>
		<tr>
			<td>Femtojet Express programmable microinjector</td>
			<td>Eppendorf</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Femtotips Microloader tips</td>
			<td>Fisher Scientific</td>
			<td>E5242956003</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine-tip paintbrush</td>
			<td>ZEM</td>
			<td>2595</td>
			<td></td>
		</tr>
		<tr>
			<td>Flesh fly pupae,&#160;Sarcophaga bullata</td>
			<td>Ward&#39;s Science</td>
			<td>470180-392</td>
			<td></td>
		</tr>
		<tr>
			<td>Food colorant dye</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Test Tubes</td>
			<td>Fisher Scientific</td>
			<td>982010</td>
			<td></td>
		</tr>
		<tr>
			<td>Glue</td>
			<td>Elmer&#39;s</td>
			<td></td>
			<td>washable, no toxic school glue</td>
		</tr>
		<tr>
			<td>Micropipette Puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000 or P-2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisherbrand</td>
			<td>12-550-A3</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Olympus</td>
			<td>SZ51</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

This paper presents two easy methods for pupal and adult microinjection, either using a femtojet or an aspirator tube. The first method allows a more precise injection of liquid, which is important for RNAi consistency, whereas the second one allows the injection of larger amounts of liquid into Nasonia pupae or adults.
Representative results presented in Table 1 show good survival rates (from 20% to 89...

Watch this Scientific Journal Video about Pupal and Adult Injections for RNAi and CRISPR Gene Editing in Nasonia vitripennis at JoVE.com

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.

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, ...

Research on non-canonical insect model, like parasitoid wasp, is limited because embryonic microinjection is very challenging. Our method bypasses this problem, enabling site-specific and heritable germline mutations. Injecting pupae or adult females is fast and removes the requirement for expensive equipment or specialized training required to achieve successful embryonic microinjections.This technique will allow no specialized labs that do not have access to expensive equipment to do functional studies in any insect species they are using as a model. After the optimization, this method can be used to study functional genetics of many other genes in Nasonia. We also think this method may be used to deliver nucleic acids for genetic transformation of Nasonia or other insects.Begin by setting up approximately 20 mated females singularly in small glass test tubes plugged with cotton. Add two S.bullata hosts per tube and maintain the wasps at 25 degrees Celsius and 30%relative humidity with a 12:12 light dark cycle for two to three days. After two to three days, gently remove the females using a fine tipped brush to avoid continuous over position and a synchrony in offspring development.Removed females can optionally be re-hosted or stored at five degrees Celsius for up to a month. Maintain the parasitized host at 25 degrees Celsius and 30%relative humidity with a 12:12 light dark cycle for seven days if the desired injection stage requires white pupae, for 13 days, if the required developmental stage is black pupae, or 14 days for young, newly emerged adults. When the desired stage is reached, crack open the host puparium with a dissecting needle to recover N.vitripennis pupae and adults.Prepare a glass slide by applying a line of school glue at the center. Spread the glue with the dissecting needle to obtain a thick layer, which will be used to align the pupae. Let the glue dry for approximately two minutes before transferring the pupae.Under a dissecting microscope, use a dissecting needle to apply glue on the back of the head of the pupa. Attach the pupa to the glue layer on the slide with its abdomen facing up. Align 20 to 30 pupae side-by-side on the slide.Place the slide with the pupae and a Petri dish to let the glue drive for approximately 10 minutes. Before starting the injections, test the adherence of the pupae to the glue by pushing them gently with the dissecting needle. If most of the pupae are loosened, prepare a new slide.Alternatively, remove all loose pupae and proceed to inject those that are properly attached to the slide. Load one capillary glass tube into a needle puller and pull needles following the manufacturer's instructions. Load the needle with five microliters of the prepared injection mixture using microloader pipette tips.Open the needle, sliding the tip on a surface made from two overlapping slides. Alternatively, open the tip of the needle with a pair of fine forceps to create a sharp edge. Place the slide with the aligned pupae under a dissecting microscope, then insert the needle into the femtojet's injection tube and tighten the grip head.Looking at the needle under the microscope, turn on the femtojet and set the compensation pressure and injection pressure parameters by rotating the rotary knobs. Carefully insert the needle between the second and third visible abdominal segments with a vertical angle of about 30 degrees. Inject with a continuous flow until the whole abdomen turns green in the case of white stage pupae or until the abdomen increases in size in the case of black pupae.Stop injecting when it is clear that the abdomen cannot take any more liquid or when the liquid starts flowing out of the body. Move carefully to the next pupa and repeat these steps. Transfer the slide with the injected pupae to a Petri dish.Cover the dish with its lid and place it at 25 degrees Celsius until wasp emergence. For adult preparation, separate groups of 20 virgin females in a clean small test tube with a fine paint brush. Place a glass slide on top of ice and align the female side by side on the cold slide with abdomens facing up under a dissecting scope.Load one needle with the ribonucleoprotein solution using a microloading tip and insert the needle into the connector pack of the aspirator tube assembly. Support the females from the opposite side with blunted dissecting needles, while injecting slowly into the abdomen. Avoid touching the ovipositor.This will severely damage this critical reproductive structure. Carefully insert the needle between the second and third visible abdominal segments with a vertical angle of about 30 degrees. Inject the solution into the female abdomen, stopping when no more liquid can enter or when leaking of surplus solution is observed.Move carefully to the next female and repeat these steps. When finished, gently transfer a single injected females to a new tube with one host using a paintbrush. Leave them to recover at room temperature for approximately one hour, confirm survival, and return the tubes to the rearing incubator.After adult eclosion from the injected pupae, place single wasps into a glass to plugged with cotton and insert one S.bullata host. For CRISPR/Cas9 knockout experiments, allow females to parasitize the host for one day and replace the host each day for three consecutive days. Place the parasitized host, set 25 degrees Celsius until emergence of the G0 male offspring, approximately 13 to 14 days.Under a dissecting microscope, screen G0 males for the mutated phenotypes. The cinnabar gene is responsible for eye pigmentation. Wasps with brown eyes are wild-type and wasps with bright red eyes or variations between red and brown eyes are mutants.This protocol can be used for either pupal or adult microinjection. Survival rates of this procedure range from 15 to 100%The efficiency of achieving the desired phenotype via RNAi or CRISPR without requiring laborious egg injections is shown here. When injecting pupae or adult females, the needle should be sharp and be introduced in a way that liquid should not leak immediately after injecting.If this happens, remove the needle, ensure that the tip is fine and sharp, change it if necessary or inject in a different location. Function analysis can be performed on the gene of interest targeted from this procedure. This will answer a question about the gene function and if the gene is necessary for the specific phenotype studied.

pupal-adult-injections-for-rnai-crispr-gene-editing-nasonia

Research

JoVE Journal

Biology

Protocol for RNAi Assays in Adult Mosquitoes (A. gambiae)

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite.  The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Protocol for RNAi Assays in Adult Mosquitoes A. gambiae

Reverse genetic approaches have proven extremely useful for determining which genes underly resistance to vector pathogens in mosquitoes. This video protocol illustrates a method used by the Dimopoulos lab to inject dsRNA into Anopheles gambiae mosquitoes, which harbor the malaria parasite. The technique manipulating the injection setup and injecting dsRNA into the thorax is illustrated.

Reverse genetic approaches have proven extremely useful for determining  which genes underly resistance to vector pathogens in mosquitoes.  This video ...

Localized RNAi and Ectopic Gene Expression in the Medicinal Leech

In this video, we show the use of a pneumatic capillary gun for the accurate biolistic delivery of reagents into live tissue. We use the procedure to perturb gene expression patterns in selected segments of leech embryos, leaving the untreated segments as internal controls.
The pneumatic capillary gun can be used to reach internal layers of cells at early stages of development without opening the specimen. As a method for localized introduction of substances into living tissues, the biolistic delivery with the gun has several advantages: it is fast, contact-free and non-destructive. In addition, a single capillary gun can be used for independent delivery of different substances. The delivery region can have lateral dimensions of ~50-150 &#181;m and extends over ~15 &#181;m around the mean penetration depth, which is adjustable between 0 and 50 &#181;m. This delivery has the advantage of being able to target a limited number of cells in a selected location intermediate between single cell knock down by microinjection and systemic knockdown through extracellular injections or by means of genetic approaches.
For knocking down or knocking in the expression of the axon guidance molecule Netrin, which is naturally expressed by some central neurons and in the ventral body wall, but not the dorsal domain, we deliver molecules of dsRNA or plasmid-DNA into the body wall and central ganglia. This procedure includes the following steps: (i) preparation of the experimental setup for a specific assay (adjusting the accelerating pressure), (ii) coating the particles with molecules of dsRNA or DNA, (iii) loading the coated particles into the gun, up to two reagents in one assay, (iv) preparing the animals for the particle delivery, (v) delivery of coated particles into the target tissue (body wall or ganglia), and (vi) processing the embryos (immunostaining, immunohistochemistry and neuronal labeling) to visualize the results, usually 2 to 3 days after the delivery.
When the particles were coated with netrin dsRNA, they caused clearly visible knock-down of netrin expression that only occurred in cells containing particles (usually, 1-2 particles per cell). Particles coated with a plasmid encoding EGFP induced fluorescence in neuronal cells when they stopped in their nuclei.

In this video, we show a procedure for an accurate biolistic delivery of reagents into live tissue with a novel miniature gene gun. We are knocking down the expression of the axon guidance molecule Netrin in leech embryos by delivering molecules of dsRNA into the ventral body wall and ganglia of single segments.

In this video, we show the use of a pneumatic capillary gun for the accurate biolistic delivery of reagents into live tissue. We use the procedure to ...

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

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 ...

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion rate. This method can be used to obtain a better understanding of the process of diuresis in insects and is especially useful in the study of diuresis in blood-feeding arthropods that are able to take up huge amounts of liquid in a single blood meal. 
This RNAi-mediated gene knockdown combined with an in vivo diuresis assay was developed by the Hansen lab to study the effects of RNAi-mediated knockdown of aquaporin genes on Aedes aegypti mosquito diuresis1. 
The protocol is setup in two parts: the first demonstration illustrates how to construct a simple mosquito injection device and how to prepare and inject dsRNA into the thorax of mosquitoes for RNAi-mediated gene knockdown. The second demonstration illustrates how to determine excretion rates in mosquitoes using an in vivo bioassay.

RNAi-mediated Gene Knockdown and In Vivo Diuresis Assay in Adult Female Aedes aegypti Mosquitoes

In this protocol we combine RNAi-mediated gene silencing with an in vivo diuresis assay to study the effects knockdown of genes of interest has on mosquito fluid excretion.

This video protocol demonstrates an effective technique to knockdown a particular gene in an insect and conduct a novel bioassay to measure excretion ...

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter implantation. The technical limitation of this surgical technique lies in the finesse of the hands. The injection is rapid, especially for a trained experimenter, and since tissue disruption with this technique is minimal, repeated injections are possible; moreover immune reaction to foreign tools (e.g. catheter) does not occur, thereby giving a better and more specific read out of spinal cord modulation. Since the application of the substance is largely limited to the target region of the spinal cord, drugs do not need to be applied in large dosages, and more importantly unwanted effects on other tissue, as observed with a systemic delivery, could be circumvented1,2. Moreover, we combine this technique with in vivo transfection of nucleic acid with the help of polyethylenimine (PEI) reagent3, which provides tremendous versatility for studying spinal functions via delivery of pharmacological agents as well as gene, RNA, and protein modulators.

In Vivo SiRNA Transfection and Gene Knockdown in Spinal Cord via Rapid Noninvasive Lumbar Intrathecal Injections in Mice

This report describes a simple and rapid technique of intrathecal needle puncture for a localized transfection of siRNA in the lumbar spinal cord in mouse under short lasting light anesthesia.

This report describes a step-by-step guide to the technique of acute intrathecal needle injections in a noninvasive manner, i.e. independent of catheter ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

Silencing the Spark: CRISPR/Cas9 Genome Editing in Weakly Electric Fish

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these independently derived phenotypes, which share a common genetic architecture. This is perhaps best exemplified by the numerous convergent features of gymnotiforms and mormyrids, two species-rich teleost clades that produce and detect weak electric fields and are called weakly electric fish. In the 50 years since the discovery that weakly electric fish use electricity to sense their surroundings and communicate, a growing community of scientists has gained tremendous insights into evolution of development, systems and circuits neuroscience, cellular physiology, ecology, evolutionary biology, and behavior. More recently, there has been a proliferation of genomic resources for electric fish. Use of these resources has already facilitated important insights with regards to the connection between genotype and phenotype in these species. A major obstacle to integrating genomics data with phenotypic data of weakly electric fish is a present lack of functional genomics tools. We report here a full protocol for performing CRISPR/Cas9 mutagenesis that utilizes endogenous DNA repair mechanisms in weakly electric fish. We demonstrate that this protocol is equally effective in both the mormyrid species Brienomyrus brachyistius and the gymnotiform Brachyhypopomus gauderio by using CRISPR/Cas9 to target indels and point mutations in the first exon of the sodium channel gene scn4aa. Using this protocol, embryos from both species were obtained and genotyped to confirm that the predicted mutations in the first exon of the sodium channel scn4aa were present. The knock-out success phenotype was confirmed with recordings showing reduced electric organ discharge amplitudes when compared to uninjected size-matched controls.

Here, a protocol is presented to produce and rear CRISPR/Cas9 genome knockout electric fish. Outlined in detail are the required molecular biology, breeding, and husbandry requirements for both a gymnotiform and a mormyrid, and injection techniques to produce Cas9-induced indel F0 larvae.

Electroreception and electrogenesis have changed in the evolutionary history of vertebrates. There is a striking degree of convergence in these ...

Nuclei Isolation from Adult Mouse Kidney for Single-Nucleus RNA-Sequencing

The kidneys regulate diverse biological processes such as water, electrolyte, and acid-base homeostasis. Physiological functions of the kidney are executed by multiple cell types arranged in a complex architecture across the corticomedullary axis of the organ. Recent advances in single-cell transcriptomics have accelerated the understanding of cell type-specific gene expression in renal physiology and disease. However, enzyme-based tissue dissociation protocols, which are frequently utilized for single-cell RNA-sequencing (scRNA-seq), require mostly fresh (non-archived) tissue, introduce transcriptional stress responses, and favor the selection of abundant cell types of the kidney cortex resulting in an underrepresentation of cells of the medulla.
Here, we present a protocol that avoids these problems. The protocol is based on nuclei isolation at 4 &#176;C from frozen kidney tissue. Nuclei are isolated from a central piece of the mouse kidney comprised of the cortex, outer medulla, and inner medulla. This reduces the overrepresentation of cortical cells typical for whole-kidney samples for the benefit of medullary cells such that data will represent the entire corticomedullary axis at sufficient abundance. The protocol is simple, rapid, and adaptable and provides a step towards the standardization of single-nuclei transcriptomics in kidney research.

Here, we present a protocol to isolate high-quality nuclei from frozen mouse kidneys that improve the representation of medullary kidney cell types and avoids the gene expression artifacts from enzymatic tissue dissociation.

The kidneys regulate diverse biological processes such as water, electrolyte, and acid-base homeostasis. Physiological functions of the kidney are ...

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent adult integument. Their immobility and transparency make them ideal for in vivo live imaging experiments. Many studies have focused on the dorsal epithelial monolayer of the pupal&#160;notum because of its accessibility and relatively large size. In addition to the studies of epithelial mechanics and development, the notum has been an ideal tissue to study wound healing. After an injury, the entire epithelial repair process can be captured by live imaging over 6-12 h. Despite the popularity of the notum for live imaging, very few published studies have utilized fixed notum samples. Fixation and staining are common approaches for nearly all other Drosophila tissues, taking advantage of the large repertoire of simple cellular stains and antibodies. However, the pupal notum is fragile and prone to curling and distortion after removal from the body, making it challenging to complement live imaging. This protocol offers a straightforward method for fixing and staining the pupal notum, both intact and after laser-wounding. With this technique, the ventral side of the pupa is glued down to a coverslip to immobilize the pupa, and the notum is carefully removed, fixed, and stained. The notum epithelium is mounted on a slide or between two coverslips to facilitate imaging from the tissue&#39;s dorsal or ventral side.

Dissecting, Fixing, and Visualizing the Drosophila Pupal Notum

The present protocol details the preparation and visualization of fixed tissue of the Drosophila pupal notum. It can be used for either intact or wounded tissue, and the original architecture of the tissue is preserved. The procedures for dissecting, fixing, and staining are all described in this article.

The pupae of Drosophila melanogaster are immobile for several days during metamorphosis, during which they develop a new body with a thin transparent ...