We would like to thank Michelle Moon (Vanderbilt University) for assistance with photography. Vanderbilt University provided support to PA and SSLB is supported by DGE-1445197.

1. Plant Cultures
<ol>
	<li>Purchase seeds from any commercial vendor or collect from mature plants in the field. 
	NOTE: This protocol is suitable for most commercially available milkweed species (e.g., Asclepias incarnata, A. syriaca, A. curassavica, Gomphocarpus physocarpus). Some seeds may need to be cold-stratified, and instructions from the seed supplier should be checked.</li>
	<li>Plant seeds in a fine germinating soil (60&#8211;70% fine peat moss, perlite, vermiculite, limestone).
	<ol>
		<li>Fill a standard seedling tray with germination mix soil; ensuring that the soil reaches the top of the wells. In each well, make an indentation to create a hole in the soil about a 3 cm deep.</li>
		<li>Place one seed in each hole and water very well with a watering can such that the soil covers the seeds and is saturated.</li>
		<li>Grow seeds in a greenhouse (see conditions below, 1.5). Water regularly, daily to every-other-day; enough to maintain the soil moisture to a moderate level.</li>
	</ol>
	</li>
	<li>When the seedlings have grown their first set of full leaves, re-pot seedlings in a general potting mix (50&#8211;60% peat moss, bark, and limestone) (Figure 1A).
	<ol>
		<li>Use 4-inch round pots that fit with a tight seal with the cup cages. Fill with general potting soil up to about 5 cm below the rim.</li>
		<li>Create a hole in the soil deep enough to reach the bottom of the pot.</li>
		<li>With the hand, gently scoop the mature seedling from its well and place it deep inside the hole in the 4 inch pot. Cover the seedling with the soil. Water very well.</li>
		<li>Grow plants in the greenhouse and water regularly, daily to every other day; enough to maintain moderate soil moisture.</li>
	</ol>
	</li>
	<li>Greenhouse conditions.
	<ol>
		<li>Set the greenhouse thermostats to maintain daytime temperatures between 18&#8211;28 &#176;C and nighttime temperatures between 16&#8211;22 &#176;C using the manufacturer&#39;s instructions.</li>
		<li>During winter months when the days are shorter, supplement the daylight with 600 W high-pressure sodium bulbs (12 h, 8 am &#8211; 8 pm).</li>
	</ol>
	</li>
	<li>Control unwanted pests (e.g., thrips, aphids) with a foliar organic soap solution, however, use these products with caution.
	<ol>
		<li>Make the soap solution according to the manufacturer&#39;s recommendation and apply using a spray bottle.</li>
		<li>Leave the soap on the plants for 4&#8211;24 h. Gently rinse the plants with water to remove the soap 4&#8211;24 h post-application and rinse them with water a second time prior to use with laboratory aphid cultures.</li>
	</ol>
	</li>
	<li>Culture the average aphid population on plants that have grown at least 3&#8211;4 sets of full leaves and are at least 10 cm tall (Figure 1B).</li>
</ol>
2. Aphid Cultures
<ol>
	<li>Start the laboratory aphid populations from an existing lab isoclonal population or start from the field-collected aphids following the directions below.
	<ol>
		<li>When starting a laboratory population from an existing lab isoclonal population, transfer aphids as described in 2.3.1&#8211;2.3.3.</li>
	</ol>
	</li>
	<li>When starting the new isoclonal, field-collected aphid populations and place a single, reproducing, adult aphid on a suitable host plant as mentioned in Step 1.6. 
	NOTE: Populations may be started from winged (alate) or unwinged (apterous) adults.
	<ol>
		<li>Manually inspect plants from the greenhouse for unwanted pests prior to the use with laboratory aphids. Freeze any plants with unwanted aphids. If desired, use an ethanol vacuum flask to remove thrips or other pests. 
		NOTE: Be sure to rinse plants that have been treated with soap prior to use as described in Step 1.5.2.</li>
		<li>Safely transfer a single adult aphid using a paintbrush or a mouth pipette created with 3/16&#34; ID x 1/4&#34; OD plastic tubing, a 1,000 &#181;L pipette tip, and a 2,00 &#181;L pipette tip (Figure 2A).</li>
		<li>Securely cover plants with aphids with a cup cage created with a plastic cup with the top cut off, covered with a fine mesh and secured with tape (Figure 2B).</li>
		<li>Place aphid-infested plants in a tray and keep in a controlled environmental chamber (16L:8D, 22 &#176;C, 70% humidity).</li>
	</ol>
	</li>
	<li>To maintain the stock populations, transfer aphids to fresh, new plants weekly (2.2.1&#8211;2.2.3).
	<ol>
		<li>Safely transfer 1&#8211;3 2nd or 3rd instar nymphs and 1 adult-aged aphids using a mouth pipette (Figures 2A, 3). 
		NOTE: Stocks are best maintained by transferring unwinged individuals.</li>
		<li>Securely cover aphid-infested plants with a cup cage created with a plastic cup with the top cut off, covered with a fine mesh and secured with tape.</li>
		<li>Place plants in a tray and keep aphids in an environmental chamber (16L:8D, 22 &#176;C, 70% humidity).</li>
		<li>Alternatively, if desired and if the host plant is of decent quality, use an ethanol vacuum flask to reduce populations leaving only one reproducing adult and two to three 2nd or 3rd instar nymphs.</li>
	</ol>
	</li>
	<li>To create same age populations for the use in experiments, place up to 5 adults (preferably unwinged) from the stock population onto a new host plant.
	<ol>
		<li>Remove the adults 24 h later.</li>
		<li>About 5&#8211;7 days later, once the F1 offspring have matured to adulthood, place up to 5 unwinged F1 adults on a new host plant. Remove the adults 24 h later.</li>
		<li>Once the F2 population has matured to adulthood, this population is ready to be used in experiments. 
		NOTE: This process ensures that the experimental population is roughly the same age and are born of roughly same age mothers.</li>
	</ol>
	</li>
	<li>Confirm the genotypic differences between field-caught isoclonal lines using microsatellite genotyping (described below, Sections 3 &#38; 4).</li>
</ol>
3. DNA Extraction
<ol>
	<li>Preparation
	<ol>
		<li>Use sterile techniques to prepare 1 L lysis buffer (0.1 M NaCl, 0.2 M sucrose, 0.1 M Tris (pH 9.1), 0.05 M EDTA, 0.05% SDS).</li>
		<li>Warm the heating block or water bath to 65 &#176;C.</li>
	</ol>
	</li>
	<li>Tissue homogenization and lysis
	<ol>
		<li>Place the aphid near the bottom of a sterile, 1.5 mL microcentrifuge tube.</li>
		<li>Place the sterile pestle in the tube with the aphid and immerse the bottom of the tube in liquid nitrogen. 
		NOTE: Optimal tissue disintegration is achieved when the aphid is positioned between the pestle and side of the tube.</li>
		<li>Grind the aphid with the pestle to initially lyse cells.</li>
		<li>For a single adult aphid, use 200 &#181;L (split into 2 x 100 &#181;L aliquots) of the lysis buffer. Add the first aliquot to grind and resuspend the crushed aphid until the sample is visibly disintegrated, then use the second aliquot to wash off the pestle.</li>
		<li>Incubate the crushed aphids in lysis buffer at 65 &#176;C in the water bath or heat block for 30 min.</li>
	</ol>
	</li>
	<li>DNA precipitation
	<ol>
		<li>While the tube is warm, add 14 &#181;L of 8 M KOAc. Invert the tube to mix.Store the sample on ice for 30 min. 
		Note: The protocol can be paused here, and samples can be stored at -20 &#176;C up to 24 h.</li>
		<li>Centrifuge at 13,000 x g for 15 min at room temperature. Transfer the supernatant to new 1.5 mL tube with a pipette. Be careful not to remove any of the pelleted debris.</li>
		<li>To improve DNA pellet visualization, add 2 &#181;L glycogen (20 &#181;g/mL) to the supernatant. If the sample size is large enough, omit this step.</li>
		<li>Add 200 &#181;L of cold 100% molecular-grade ethanol to the supernatant. Invert tubes to mix and incubate at room temperature for at least 15 min. 
		NOTE: The protocol can be paused here, and samples can be stored at -20 &#176;C up to 24 h.</li>
		<li>Centrifuge at 13,000 x g for 15 min at room temperature. Remove ethanol by pipetting.</li>
	</ol>
	</li>
	<li>DNA wash and elution
	<ol>
		<li>Add 200 &#181;L of cold 70% molecular-grade ethanol. Then flick the tube to resuspend and wash the pellet.</li>
		<li>Centrifuge at 13,000 x g for 5 min. While visualizing the pellet, carefully remove the ethanol by pipetting and add 200 &#181;L of cold 100% molecular-grade ethanol.</li>
		<li>Centrifuge at 13,000 x g for 5 min. While visualizing the pellet, carefully remove ethanol by pipetting. 
		NOTE: Repeat the 100-70-100 ethanol wash if necessary.</li>
		<li>Air dry the pellets for 5&#8211;10 min with the tube laying horizontally open on a tissue paper.</li>
		<li>Resuspend the DNA pellet in 80 &#181;L of low TE (10 mM Tris-HCl, 0.1 mM EDTA).</li>
		<li>Quantify the resuspended DNA using a spectrophotometer.</li>
		<li>Store at 4 &#176;C.</li>
	</ol>
	</li>
</ol>
4. Microsatellite PCR and Sequencing for Aphid Genotyping
<ol>
	<li>Order the appropriate F and R primers for microsatellite sequencing (Table 120). 
	NOTE: Reverse primer sequences should be modified with 5&#39;-6-FAM or 5&#39;-5-HEX fluorescent labels to allow for multiplexed samples for microsatellite sequencing.</li>
	<li>Perform PCR with single aphid DNA samples (described in Section 3) and fluorescently labeled microsatellite primers.
	<ol>
		<li>Mix PCR reactions according to the manufacturer&#39;s protocol (0.2 &#181;M each F/R primer, 2.5 mM MgCl2, 50&#8211;200 ng DNA template).</li>
		<li>Use the following thermocycler settings: initial denaturation at 94 &#176;C for 4 min, 35 cycles of 94 &#176;C for 30 s, 58 &#176;C for 35 s, 72 &#176;C for 45 s, and a final elongation step at 72 &#176;C for 10 min.</li>
	</ol>
	</li>
	<li>Combine PCR samples with different fluorescent tags to reduce the number of samples sequenced and sequence the microsatellite samples at a genotyping facility.</li>
	<li>Analyze the .fsa raw sample files using microsatellite analysis software.</li>
</ol>
5. RNA Extraction
<ol>
	<li>Collect aphids&#39; samples for RNA extraction in 1.5 mL RNase/ DNase-free tubes and immediate freeze in liquid nitrogen. 
	NOTE: If the following steps are not performed immediately, the samples can be stored at -80 &#176;C.</li>
	<li>Tissue homogenization
	<ol>
		<li>With the sterile pestle in the tube with aphid, freeze in liquid nitrogen for 10&#8211;15 seconds, until the sample stop sizzling. Crush the aphid well with the pestle as described in step 3.2. 
		NOTE: Optimal tissue disintegration is achieved when the aphid is positioned between the pestle and side of the tube.</li>
		<li>In the fume hood, add 800 &#181;L of guanidinium thiocyanate-phenol-chloroform extraction reagent to the sample (1&#8211;5 adult aphids). Homogenize samples with the pestle and dispose of the pestle.</li>
	</ol>
	</li>
	<li>Phase separation 
	NOTE: All steps should be performed in a fume hood.
	<ol>
		<li>Incubate the homogenized samples for 5 min at room temperature. Add 160 &#181;L of chloroform to sample. Shake vigorously by hand for 15 s.</li>
		<li>Incubate for 2-3 min at room temperature. Centrifuge for 15 min at 12,000 x g at 4 &#176;C. 
		NOTE: Following centrifugation, the mixture separates into 3 layers: a lower, red phenol-chloroform phase, an interphase and a colorless upper aqueous phase. The RNA remains exclusively in the aqueous phase. The volume of the aqueous will be ~480 &#181;L.</li>
	</ol>
	</li>
	<li>RNA precipitation
	<ol>
		<li>In a fume hood, transfer the aqueous phase to a fresh, RNase-free tube. Do not disturb the intermediate phase.</li>
		<li>Precipitate the RNA by adding 400 &#181;L of isopropanol and incubate the sample at -20 &#176;C for 10 min. 
		NOTE: The protocol can be paused here, and samples can be stored at -20 &#176;C up to 24 h.</li>
		<li>Centrifuge the sample for 10 min at 12,000 x g at 4 &#176;C.</li>
	</ol>
	</li>
	<li>RNA wash and elution
	<ol>
		<li>Remove the supernatant; watch for the RNA pellet.</li>
		<li>Wash the RNA pellet with 1 mL of 75% ethanol in DEPC-treated water. Mix by slow vortexing. Centrifuge for 5 min at 7,500 x g at 4 &#176;C.</li>
		<li>Repeat steps 5.5.1&#8211;5.5.2 to help remove phenol contaminants.</li>
		<li>Remove the supernatant and air dry the pellet for 5-10 min with tube laying horizontally open on a sterile bench. Do not let the RNA pellet dry completely.</li>
		<li>Dissolve the RNA pellet in 30 &#181;L of RNase-free or DEPC-treated water. Gently pipette up and down to mix. Incubate at 55&#8211;60 &#176;C for 10&#8211;15 min.</li>
	</ol>
	</li>
</ol>
6. RNAseq De Novo Transcriptome Assembly, Annotation, and Differential Expression Analysis
<ol>
	<li>Analyze RNA sample concentration and quality using a chip-based capillary electrophoretic system. 
	NOTE: A chip-based capillary electrophoresis system is the preferable method of choice than analyzing with a spectrophotometer because it provides a more accurate and sensitive measure of RNA concentration and quality.
	<ol>
		<li>If samples are of suitable quality (&#8805; 250 ng total, RIN (RNA Integrity Number) &#8805; 5), perform RNA sequencing. 
		NOTE: Importantly, because this sequencing data will be used for both expression profiling and de novo transcriptome assembly, more read depth will result in a higher quality transcriptome. For a reasonably comprehensive assembly using Illumina sequencing technology, 100&#8211;200 million 100 bp, paired end reads would be a recommended starting point. Total mRNA library preparation and RNA sequencing were performed by a sequencing facility.</li>
	</ol>
	</li>
	<li>Check the quality of reads using Fast QC22.</li>
	<li>Combine all sample reads and assemble the transcriptome de novo using Trinity23,24 (Trimmomatic quality filtering enabled).</li>
	<li>Refine the assembly
	<ol>
		<li>Use Transdecoder25 to identify open reading frames (ORFs) that are a minimum of 100 amino acids in length.</li>
		<li>Perform homology searches for the translated ORFs against Pfam26 and UniProt27 databases using BLASTP28 and HMMER29, respectively.</li>
		<li>Remove bacterial transcripts (any translated sequence whose best BLAST hit was to a bacterial gene with a bit score of over 300 and a minimum amino acid sequence identity of 50%).</li>
		<li>Collapse any complete, translated ORFs that are at least 99% identical at the amino acid level using CD-HIT30.</li>
		<li>Collapse the remaining, incomplete ORFs that are at least 95% identical at the nucleotide level using CD-HIT30.</li>
		<li>Assign the remaining nucleotide sequences with unique, species-specific identifiers (e.g., APHNE 0001).</li>
	</ol>
	</li>
	<li>Assess the completeness of the refined assembly, using BUSCO (http://busco.ezlab.org/) and the Arthropoda gene dataset31.</li>
	<li>Transcriptome annotation
	<ol>
		<li>First, annotate the refined transcriptome using HMMER against the Pfam database26,29.</li>
		<li>Second, annotate the transcriptome using BLASTP against the UniProt database27,28.</li>
		<li>Third, annotate the transcriptome using BLASTP against the coding sequences of selected insects with published, annotated genomes.</li>
		<li>Last, annotate the transcriptome using BLASTP against the pea aphid protein database only.</li>
		<li>Use Trinotate to generate GO annotations from UniProt accessions.</li>
		<li>Use Trinotate to organize all the annotation results into a SQLite database and generate an annotation report.</li>
	</ol>
	</li>
	<li>Differential expression analysis 
	NOTE: Using the refined transcriptome as a reference, align and quantify each library separately.
	<ol>
		<li>Use Trimmomatic to quality-filter and trim the original read files32. 
		Note: If performing this step after a Trinity assembly, one may instead use the Trimmomatic output from that step.</li>
		<li>Perform local alignments for each sample using Bowtie233.</li>
		<li>Extract the read counts from each sample individually using SAMtools34.</li>
		<li>Calculate the differential expression between samples of interest using DESeq2 with the default parameters and a parametric fit35.</li>
	</ol>
	</li>
</ol>
7. qPCR Verification of Differentially Expressed Genes
NOTE: If users are interested in differentially expressed genes from their RNAseq experiments, the following protocol can be used to verify patterns of differential expression.
<ol>
	<li>Generate RNA samples as described above (step 5).</li>
	<li>Quantitate RNA extractions using a spectrophotometer to check for the quality and obtain the concentration.</li>
	<li>Synthesize cDNA samples using a commercially available kit as per the manufacturer&#39;s recommendation.</li>
	<li>Determine the primer efficiencies for genes of interest to ensure accurate two-fold PCR amplification.
	<ol>
		<li>Based on original RNA concentrations, perform serial dilutions (101) to obtain 3 cDNA concentrations.</li>
		<li>Using a quantitative PCR master mix, mix triplicate qPCR reactions according to the manufacturer&#39;s protocol using three primer concentrations (e.g., 100 nM, 200 nM, 300 nM) with three serially diluted cDNA concentrations (e.g., 0.1 ng/&#181;L, 10 ng/&#181;L, 100 ng/&#181;L).</li>
		<li>For each target gene, calculate the slope (m) of the line created using the mean Ct values for each sample as the dependent variables and the log (cDNA concentration) as the independent variables (three points total).</li>
		<li>Use the following equation to calculate the primer efficiency (E) where m is the slope calculated in 7.4.3: 
		E = 10^(-1/m) 
		NOTE: Primer efficiencies between 90-110% are suitable for analyses. This process ensures equal amplification of all genes included in the calculations.</li>
	</ol>
	</li>
	<li>Use the &#916;&#916;Ct method with a housekeeping gene to quantify the differential expression for genes of interest36.</li>
</ol>

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L., Abbot, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insect+adaptations+toward+plant+toxins+in+milkweed-herbivores+systems+-+a+review.">Insect adaptations toward plant toxins in milkweed-herbivores systems - a review.</a> Entomologia Experimentalis et Applicata. 58, 579-610 (2018).</li><li>Rothschild, M., von Euw, J., Reichstein, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+glycosides+in+the+oleander+aphid,+Aphis+nerii.">Cardiac glycosides in the oleander aphid, Aphis nerii.</a> Journal of Insect Physiology. 16, 1141-1145 (1970).</li><li>Harrison, J. S., Mondor, E. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+an+invasive+aphid+'superclone':+extremely+low+genetic+diversity+in+oleander+aphid+(Aphis+nerii)+populations+in+the+southern+United+States.">Evidence for an invasive aphid 'superclone': extremely low genetic diversity in oleander aphid (Aphis nerii) populations in the southern United States.</a> PLoS ONE. 6, 17524 (2011).</li><li>Mooney, K. A., Halitschke, R., Kessler, A., Agrawal, A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evolutionary+trade-offs+in+plants+mediate+the+strength+of+trophic+cascades.">Evolutionary trade-offs in plants mediate the strength of trophic cascades.</a> Science. 327, 1642-1644 (2010).</li><li>Grabherr, M. G., 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=Full-length+transcriptome+assembly+from+RNA-Seq+data+without+a+reference+genome.">Full-length transcriptome assembly from RNA-Seq data without a reference genome.</a> Nature Biotechnology. 29, 644-652 (2011).</li><li>Haas, B. J., 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=De+novo+transcript+sequence+reconstruction+from+RNA-seq+using+the+Trinity+platform+for+reference+generation+and+analysis.">De novo transcript sequence reconstruction from RNA-seq using the Trinity platform for reference generation and analysis.</a> Nature Protocols. 8, 1494-1512 (2013).</li><li>Finn, R. 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=The+Pfam+protein+families+database:+towards+a+more+sustainable+future.">The Pfam protein families database: towards a more sustainable future.</a> Nucleic Acids Research. 44, 279-285 (2016).</li><li>The UniProt Consortium. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=UniProt:+the+universal+protein+knowledgebase.">UniProt: the universal protein knowledgebase.</a> Nucleic Acids Research. 45, 158-169 (2017).</li><li>Johnson, 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=NCBI+BLAST:+a+better+web+interface.">NCBI BLAST: a better web interface.</a> Nucleic Acids Research. 1 (36), 5-9 (2008).</li><li>Fu, L., Niu, B., Zhu, Z., Wu, S., Li, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=CD-HIT:+accelerated+for+clustering+the+next+generation+sequencing+data.">CD-HIT: accelerated for clustering the next generation sequencing data.</a> Bioinformatics. 28 (23), 3150-3152 (2012).</li><li>Sim&#227;o, F. A., Waterhouse, R. M., Ioannidis, P., Kriventseva, E. V., Zdobnov, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=BUSCO:+assessing+genome+assembly+and+annotation+completeness+with+single-copy+orthologs.">BUSCO: assessing genome assembly and annotation completeness with single-copy orthologs.</a> Bioinformatics. 31, 3210-3212 (2015).</li><li>Bolger, A. M., Lohse, M., Usadel, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Trimmomatic:+a+flexible+trimmer+for+Illumina+sequence+data.">Trimmomatic: a flexible trimmer for Illumina sequence data.</a> Bioinformatics. 30, 2114-2120 (2014).</li><li>Langmead, B., Salzberg, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fast+gapped-read+alignment+with+Bowtie+2.">Fast gapped-read alignment with Bowtie 2.</a> Nature Methods. 9, 357-359 (2012).</li><li>Li, H., 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=The+Sequence+Alignment/Map+format+and+SAMtools.">The Sequence Alignment/Map format and SAMtools.</a> Bioinformatics. 25, 2078-2079 (2009).</li><li>Love, M. I., Huber, W., Anders, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Moderated+estimation+of+fold+change+and+dispersion+for+RNA-seq+data+with+DESeq2.">Moderated estimation of fold change and dispersion for RNA-seq data with DESeq2.</a> Genome Biology. 15, 31 (2014).</li><li>Rieu, I., Powers, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+quantitative+RT-PCR:+design,+calculations,+and+statistics.">Real-time quantitative RT-PCR: design, calculations, and statistics.</a> Plant Cell. 21, 1031-1033 (2009).</li><li>Malcolm, S. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+defence+in+chewing+and+sucking+insect+herbivores:+plant-derived+cardenolides+in+the+monarch+butterfly+and+oleander+aphid.">Chemical defence in chewing and sucking insect herbivores: plant-derived cardenolides in the monarch butterfly and oleander aphid.</a> Chemoecology. 1, 12-21 (1990).</li><li>Agrawal, A. A., Underwood, N., Stinchcombe, J. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intraspecific+variation+in+the+strength+of+density+dependence+in+aphid+populations.">Intraspecific variation in the strength of density dependence in aphid populations.</a> Ecological Entomology. 29, 521-526 (2004).</li><li>Zehnder, C. B., Hunter, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+comparison+of+maternal+effects+and+current+environment+on+vital+rates+of+Aphis+nerii,+the+milkweed-oleander+aphid.">A comparison of maternal effects and current environment on vital rates of Aphis nerii, the milkweed-oleander aphid.</a> Ecological Entomology. 32, 172-180 (2007).</li><li>Hartbauer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Collective+defense+of+Aphis+nerii+and+Uroleucon+hypochoeridis+(Homoptera,+Aphididae)+against+natural+enemies.">Collective defense of Aphis nerii and Uroleucon hypochoeridis (Homoptera, Aphididae) against natural enemies.</a> PLoS ONE. 5, 10417 (2010).</li></ol>

The authors have nothing to disclose.

It has long been recognized that the aposematic A. nerii can provide insights into the patterns and mechanisms of resistance to plant defenses and particularly chemical sequestration18,37. A number of genomic resources have recently emerged for A. nerii10, offering new opportunities for ecological and functional genomic studies that use A. nerii as a model. We outline basic protocols in aphid and plant culture, a...

Aphids are small, hemimetabolous insects that colonize on diverse plant families worldwide. They are distinctive for several features, most notably their complex life cycles involving cyclical parthenogenesis and discrete polyphenisms, and their obligate nutritional symbioses with bacterial or yeast endosymbionts that supply nutrients missing from their diet of plant sap1. While most aphids are host plant specialists, some generalist species are important crop pests, inflicting considerable economic damage on crops either directly or via the pathogens and viruses they vector2. The publication of the first aphid genome in 2010, the pea aphid Acyrthosiphon pisum3, marked an important milestone in the study of aphid biology because it provided the genomic resources for addressing questions about the insect&#39;s adaptations to the herbivorous lifestyles, including those that might lead to a better control strategies4. Since that time, additional genomic resources have accumulated with the publication of an annotated genome for the soybean aphid Aphis glycines5, and publicly-available whole genome resources for another three-aphid species (Myzus cerasi (black cherry aphid), Myzus persicae (peach-potato aphid), Rhopalosiphum padi (bird cherry-oat aphid)6. Valuable de novo transcriptomic resources are available as well for a number of other aphid species (e.g.,Aphis gossypii (cotton aphid)7, Sitobion avenae (grain aphid)8, Cinara pinitabulaeformis (pine aphid)9, Aphis nerii (milkweed-oleander aphid)10).
Aphids have also made lasting contributions to our understanding of the plant-insect interactions and the ecology of the life on plants11. One area where aphids have made particularly important contributions is in our understanding of the chemical ecology of the host plant interactions. Herbivorous insects express diverse adaptations for overcoming plant defenses, and some even co-opt plant defenses for their own benefit12,13,14. For example, the milkweed-oleander aphid, Aphis nerii, is a bright yellow, invasive aphid found in temperate and tropical regions worldwide that colonizes on plants in the milkweed family (Apocynaceae). Plants in the family Apocynaceae have evolved diverse chemical defenses, including milky latex and cardiac glycosides known as cardenolides, that bind the cation carrier Na,K-ATPase and are effective deterrents to generalist herbivores15,16. Milkweed specialists express various modes of resistance to cardenolides, and some selectively or passively accumulate or modify cardenolides in their tissues as a means to deter predation or for other benefits17. A. nerii is thought to sequester cardenolides in this way, although the mechanisms and functional benefits remain unclear10,18.
Given the genomic resources at hand, A. nerii provides an excellent experimental model for the study of the molecular and genetic mechanisms involved in the chemo-ecological interactions between toxic host plants and their specialist herbivores. It is worth noting that, while some of the earliest studies of A. nerii focused on sequestration of cardenolides19, since that time, studies of A. nerii have provided insights into a broad set of evolutionary and ecological questions, including the genetic structure of invasive insects20 and the interplay between bottom-up and top-down regulation on the herbivore density21. A. nerii is thus a good candidate as an experimental model for an especially broad set of studies of the insect-plant interactions. Critical to the success of any study with A. nerii is the careful culture of aphid populations, which includes the culture of plants on which the aphids depend, as well as an efficient generation of high-quality -omic data. Our goal is to guide the reader through both. Outlined below are methods for the generation and maintenance of the plant and aphid cultures in the greenhouse and laboratory, DNA and RNA extractions, microsatellite analysis, de novo transcriptome assembly and annotation, transcriptome differential expression analysis, and qPCR verification of differentially expressed genes. While these methods are written for A. nerii, the general culturing, extraction, and analysis methods can extend to a variety of aphid species.

<table><tbody><tr><td>Sun Gro Fafard Germination Mix</td><td>Hummert International</td><td>10-0952-2</td><td></td></tr><tr><td>Sun Gro Fafard 3B/ Metro Mix</td><td>Hummert International</td><td>10-0951-2</td><td></td></tr><tr><td>2&quot; x 4&quot; Round Standard Pot</td><td>Anderson Pots</td><td>1503</td><td></td></tr><tr><td>DreamTaq DNA Polymerase</td><td>ThermoFisher Scientific</td><td>EP0701</td><td></td></tr><tr><td>Trizol</td><td>ThermoFisher Scientific</td><td>15596026</td><td></td></tr><tr><td>SuperScript&amp;reg; III First-Strand Synthesis kit</td><td>ThermoFisher Scientific</td><td>18080051</td><td></td></tr><tr><td>Power SYBR Green PCR Master Mix</td><td>ThermoFisher Scientific</td><td>4367659</td><td></td></tr></tbody></table>

maintaining biological cultures and measuring gene expression in aphis nerii: a non-model system for plant-insect interactions

Aphids are excellent experimental models for a variety of biological questions ranging from the evolution of symbioses and the development of polyphenisms to questions surrounding insect's interactions with their host plants. Genomic resources are available for several aphid species, and with advances in the next-generation sequencing, transcriptomic studies are being extended to non-model organisms that lack genomes. Furthermore, aphid cultures can be collected from the field and reared in the laboratory for the use in organismal and molecular experiments to bridge the gap between ecological and genetic studies. Last, many aphids can be maintained in the laboratory on their preferred host plants in perpetual, parthenogenic life cycles allowing for comparisons of asexually reproducing genotypes. Aphis nerii, the milkweed-oleander aphid, provides one such model to study insect interactions with toxic plants using both organismal and molecular experiments. Methods for the generation and maintenance of the plant and aphid cultures in the greenhouse and laboratory, DNA and RNA extractions, microsatellite analysis, de novo transcriptome assembly and annotation, transcriptome differential expression analysis, and qPCR verification of differentially expressed genes are outlined and discussed here.

Plant cultures: Seeds will take approximately two to four weeks, depending on the season, to grow large enough to be re-potted (Figure 1A). Re-potted seedlings will take another two to four weeks to grow to an optimal size for aphid cultures (Figure 1B).
Aphid cultures: Adult A. nerii are disti...

Watch this Scientific Journal Video about Maintaining Biological Cultures and Measuring Gene Expression in Aphis nerii: A Non-model System for Plant-insect Interactions at JoVE.com

Maintaining Biological Cultures and Measuring Gene Expression in Aphis nerii: A Non-model System for Plant-insect Interactions

The aphid Aphis nerii colonizes on highly-defended plants in the dogbane family (Apocyanaceae) and provides numerous opportunities to study plant-insect interactions. Here, we present a series of protocols for the maintenance of plant and aphid cultures, and the generation and analysis of molecular and -omic data for A. nerii.

Maintaining Biological Cultures and Measuring Gene Expression in Aphis nerii: A Non-model System for Plant-insect Interactions

Aphids are excellent experimental models for a variety of biological questions ranging from the evolution of symbioses and the development of polyphenisms ...

maintaining-biological-cultures-measuring-gene-expression-aphis-nerii

Research

JoVE Journal

Genetics

7.5K Views.  Vanderbilt University. The aphid Aphis nerii colonizes on highly-defended plants in the dogbane family (Apocyanaceae) and provides numerous opportunities to study plant-insect interactions. Here, we present a series of protocols for the maintenance of plant and aphid cultures, and the generation and analysis of molecular and -omic data for A. nerii.

Video: Maintaining Biological Cultures and Measuring Gene Expression in Aphis nerii: A Non-model System for Plant-insect Interactions

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. ...

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative without tools to study gene function. The development and sharing of detailed protocols for the establishment of new model systems in laboratories, and for tools to carry out functional studies, is thus crucial for leveraging the power of genomics. Coleoptera (beetles) are the largest clade of insects and occupy virtually all types of habitats on the planet. In addition to providing ideal models for fundamental research, studies of beetles can have impacts on pest control as they are often pests of households, agriculture, and food industries. Detailed protocols for rearing and maintenance of D. maculatus laboratory colonies and for carrying out dsRNA-mediated interference in D. maculatus are presented. Both embryonic and parental RNAi procedures-including apparatus set up, preparation, injection, and post-injection recovery-are described. Methods are also presented for analyzing embryonic phenotypes, including viability, patterning defects in hatched larvae, and cuticle preparations for unhatched larvae. These assays, together with in situ hybridization and immunostaining for molecular markers, make D. maculatus an accessible model system for basic and applied research. They further provide useful information for establishing procedures in other emerging insect model systems.

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Here, we present protocols for rearing an intermediate-germ beetle, Dermestes maculatus (D. maculatus) in the lab. We also share protocols for embryonic and parental RNAi and methods for analyzing embryonic phenotypes to study gene function in this species.

Advances in genomics have raised the possibility of probing biodiversity at an unprecedented scale. However, sequence alone will not be informative ...

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental studies. Like other aphids, A. pisum propagate rapidly via parthenogenetic viviparous reproduction, where the embryos develop within egg chambers in an assembly-line fashion in the ovariole. Previously we have established a robust platform of whole-mount in situ hybridization allowing detection of mRNA expression in the aphid embryos. For analyzing the expression of protein, though, established protocols for immunostaining the ovarioles of asexual viviparous aphids did not produce satisfactory results. Here we report conditions optimized for increasing tissue permeability and decreasing background staining, both of which were problems when applying established approaches. Optimizations include: (1) incubation of proteinase K (1 &#181;g/ml, 10 min), which was found essential for antibody penetration in mid- and late-stage aphid embryos; (2) replacement of normal goat serum/bovine serum albumin with a blocking reagent supplied by a Digoxigenin (DIG)-based buffer set and (3) application of methanol rather hydrogen peroxide (H2O2) for bleaching endogenous peroxidase; which significantly reduced the background staining in the aphid tissues. These critical conditions optimized for immunostaining will allow effective detection of gene products in the embryos of A. pisum and other aphids.

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental ...

Developmental Biology

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and tomato. The susceptibility of M. sexta larvae to a variety of entomopathogenic bacterial species1-5, as well as the wealth of information available regarding the insect's immune system6-8, and the pending genome sequence9 make it a good model organism for use in studying host-microbe interactions during pathogenesis. In addition, M. sexta larvae are relatively large and easy to manipulate and maintain in the laboratory relative to other susceptible insect species. Their large size also facilitates efficient tissue/hemolymph extraction for analysis of the host response to infection.
The method presented here describes the direct injection of bacteria into the hemocoel (blood cavity) of M. sexta larvae. This approach can be used to analyze and compare the virulence characteristics of various bacterial species, strains, or mutants by simply monitoring the time to insect death after injection. This method was developed to study the pathogenicity of Xenorhabdus and Photorhabdus species, which typically associate with nematode vectors as a means to gain entry into the insect. Entomopathogenic nematodes typically infect larvae via natural digestive or respiratory openings, and release their symbiotic bacterial contents into the insect hemolymph (blood) shortly thereafter10. The injection method described here bypasses the need for a nematode vector, thus uncoupling the effects of bacteria and nematode on the insect. This method allows for accurate enumeration of infectious material (cells or protein) within the inoculum, which is not possible using other existing methods for analyzing entomopathogenesis, including nicking11 and oral toxicity assays12. Also, oral toxicity assays address the virulence of secreted toxins introduced into the digestive system of larvae, whereas the direct injection method addresses the virulence of whole-cell inocula.
The utility of the direct injection method as described here is to analyze bacterial pathogenesis by monitoring insect mortality. However, this method can easily be expanded for use in studying the effects of infection on the M. sexta immune system. The insect responds to infection via both humoral and cellular responses. The humoral response includes recognition of bacterial-associated patterns and subsequent production of various antimicrobial peptides7; the expression of genes encoding these peptides can be monitored subsequent to direct infection via RNA extraction and quantitative PCR13. The cellular response to infection involves nodulation, encapsulation, and phagocytosis of infectious agents by hemocytes6. To analyze these responses, injected insects can be dissected and visualized by microscopy13, 14.

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

The method described here utilizes direct injection of entomopathogenic bacteria into the hemocoel of Manduca sexta insect larvae. M. sexta is a commercially available and well-studied insect. Thus, this method represents a simple approach to analyzing host-bacterial interactions from the perspective of one or both partners.

Manduca sexta, commonly known as the tobacco hornworm, is considered a significant agricultural pest, feeding on solanaceous plants including tobacco and ...

Immunology and Infection

Mass-Rearing and Molecular Studies in Tortricidae Pest Insects

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural losses. To understand the biology of such pest moths, fundamental techniques have been in high demand. Here, methods for mass-rearing, observations, and molecular studies are developed using two tea tortrix, Homona magnanima and Adoxophyes honmai (Lepidoptera: Tortricidae). Insects were mass-reared with sliced artificial diet and maintained by inbreeding for over 100 generations by considering their biological characteristics. Insects have various sex dimorphisms; hence it is difficult to distinguish the sex during the developing stages, which have prevented subsequent assays. The present work highlighted that the sex of tortricids larvae could be determined by observing testes or lactic-acetic orcein staining to visualize the female-specific W chromosome. Moreover, using the sex determination methods, the present study enabled nucleic acid extractions from sex determined embryos and application toward high throughput sequencing. These tips are applicable for other pest insects and will facilitate further morphological and genetic studies.

The present protocol describes the rearing method of tortricid pest insects in the laboratories. The procedures to distinguish insects' sex and extract nucleic acids for high throughput sequencing are established using two tortricid pests.

Tortricidae (Lepidoptera), commonly known as tortrix or leafroller moths, comprises many agricultural and forestry pests, which cause serious agricultural ...

Environment

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees (Apis mellifera)

This video demonstrates novel techniques of RNA interference (RNAi) which downregulate two genes simultaneously in honey bees using double-stranded RNA (dsRNA) injections. It also presents a protocol of proboscis extension response (PER) assay for measuring gustatory perception.	
	RNAi-mediated gene knockdown is an effective technique downregulating target gene expression. This technique is usually used for single gene manipulation, but it has limitations to detect interactions and joint effects between genes. In the first part of this video, we present two strategies to simultaneously knock down two genes (called double gene knockdown). We show both strategies are able to effectively suppress two genes, vitellogenin (vg) and ultraspiracle (usp), which are in a regulatory feedback loop. This double gene knockdown approach can be used to dissect interrelationships between genes and can be readily applied in different insect species.	
	The second part of this video is a demonstration of proboscis extension response (PER) assay in honey bees after the treatment of double gene knockdown. The PER assay is a standard test for measuring gustatory perception in honey bees, which is a key predictor for how fast a honey bee's behavioral maturation is. Greater gustatory perception of nest bees indicates increased behavioral development which is often associated with an earlier age at onset of foraging and foraging specialization in pollen. In addition, PER assay can be applied to identify metabolic states of satiation or hunger in honey bees. Finally, PER assay combined with pairing different odor stimuli for conditioning the bees is also widely used for learning and memory studies in honey bees.

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees Apis mellifera

In this protocol, we describe two strategies that simultaneously suppress two genes (double gene knockdown) in honey bees. Then we present how to use the proboscis extension response (PER) assay to study the effect of double gene knockdown on honey bee gustatory perception.

This video demonstrates novel techniques of RNA  interference (RNAi) which downregulate two genes simultaneously in honey bees  using double-stranded RNA ...

Neuroscience

Culturing and Genetically Manipulating Entomopathogenic Nematodes

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic of their life cycle is the mutualistic association with the bacteria Photorhabdus and Xenorhabdus, respectively. The nematode parasites are able to locate and enter suitable insect hosts, subvert the insect immune response, and multiply efficiently to produce the next generation that will actively hunt new insect prey to infect. Due to the properties of their life cycle, entomopathogenic nematodes are popular biological control agents, which are used in combination with insecticides to control destructive agricultural insect pests. Simultaneously, these parasitic nematodes represent a research tool to analyze nematode pathogenicity and host anti-nematode responses. This research is aided by the recent development of genetic techniques and transcriptomic approaches for understanding the role of nematode secreted molecules during infection. Here, a detailed protocol on maintaining entomopathogenic nematodes and using a gene knockdown procedure is provided. These methodologies further promote the functional characterization of entomopathogenic nematode infection factors.

Entomopathogenic nematodes live in symbiosis with bacteria and together they successfully infect insects by undermining their innate immune system. To promote research on the genetic basis of nematode infection, methods for maintaining and genetically manipulating entomopathogenic nematodes are described.

Entomopathogenic nematodes in the genera Heterorhabditis and Steinernema are obligate parasites of insects that live in the soil. The main characteristic ...

Transient Expression and Cellular Localization of Recombinant Proteins in Cultured Insect Cells

Heterologous protein expression systems are used for the production of recombinant proteins, the interpretation of cellular trafficking/localization, and the determination of the biochemical function of proteins at the sub-organismal level. Although baculovirus expression systems are increasingly used for protein production in numerous biotechnological, pharmaceutical, and industrial applications, nonlytic systems that do not involve viral infection have clear benefits but are often overlooked and underutilized. Here, we describe a method for generating nonlytic expression vectors and transient recombinant protein expression. This protocol allows for the efficient cellular localization of recombinant proteins and can be used to rapidly discern protein trafficking within the cell. We show the expression of four&#160;recombinant proteins in a commercially available insect cell line, including two aquaporin proteins from the insect Bemisia tabaci, as well as subcellular marker proteins specific for the cell plasma membrane and for intracellular lysosomes. All recombinant proteins were produced as chimeras with fluorescent protein markers at their carboxyl termini, which allows for the direct detection of the recombinant proteins. The double transfection of cells with plasmids harboring constructs for the genes of interest and a known subcellular marker allows for live cell imaging and improved validation of cellular protein localization.

Nonlytic insect cell expression systems are underutilized for production, cellular trafficking/localization, and recombinant protein functional analysis. Here, we describe methods to generate expression vectors and subsequent transient protein expression in commercially available lepidopteran cell lines. The co-localization of Bemisia tabaci aquaporins with subcellular fluorescent marker proteins is also presented.

Heterologous protein expression systems are used for the production of recombinant proteins, the interpretation of cellular trafficking/localization, and ...

Biology

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, chikungunya and Zika. Wolbachia-infected mosquitoes can be released into the field to either reduce population sizes through incompatible matings or to transform populations with mosquitoes that are refractory to virus transmission. For these strategies to succeed, the mosquitoes released into the field from the laboratory must be competitive with native mosquitoes. However, maintaining mosquitoes in the laboratory can result in inbreeding, genetic drift and laboratory adaptation which can reduce their fitness in the field and may confound the results of experiments. To test the suitability of different Wolbachia infections for deployment in the field, it is necessary to maintain mosquitoes in a controlled laboratory environment across multiple generations. We describe a simple protocol for maintaining Ae. aegypti mosquitoes in the laboratory, which is suitable for both Wolbachia-infected and wild-type mosquitoes. The methods minimize laboratory adaptation and implement outcrossing to increase the relevance of experiments to field mosquitoes. Additionally, colonies are maintained under optimal conditions to maximize their fitness for open field releases.

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

Aedes aegypti mosquitoes infected with Wolbachia are being released into natural populations to suppress the transmission of arboviruses. We describe methods to rear Ae. aegypti with Wolbachia infections in the laboratory for experiments and field release, taking precautions to minimize laboratory adaptation and selection.

Aedes aegypti mosquitoes experimentally infected with Wolbachia are being utilized in programs to control the spread of arboviruses such as dengue, ...

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

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, host-symbiont interactions, speciation, and venom synthesis. Despite the availability of several molecular tools, including CRISPR/Cas9, functional genetic studies are still limited in this organism. The major limitation of applying CRISPR/Cas9 technology in N. vitripennis stems from the challenges of embryonic microinjections. Injections of embryos are particularly difficult in this organism and in general in many parasitoid wasps, due to small embryo size and the requirement of a host pupa for embryonic development. To address these challenges, Cas9 ribonucleoprotein complex delivery into female&#160;ovaries by adult injection, rather than embryonic microinjection, was optimized, resulting in both somatic and heritable germline edits. The injection procedures were optimized in pupae and female wasps using either ReMOT Control (Receptor-Mediated Ovary Transduction of Cargo) or BAPC (Branched Amphiphilic Peptide Capsules). These methods are shown to be effective alternatives to embryo injection, enabling site-specific and heritable germline mutations.

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

Here, we describe methods for efficient pupal and adult injections in Nasonia vitripennis as accessible alternatives to embryo microinjection, enabling functional analysis of genes of interest using either RNA-silencing via RNA interference (RNAi) or gene knockout via CRISPR/Cas9 genome editing.

The jewel wasp, Nasonia vitripennis, has become an efficient model system to study epigenetics of haplo-diploid sex determination, B-chromosome biology, ...

Maintaining Biological Cultures and Measuring Gene Expression in Aphis nerii: A Non-model System for Plant-insect Interactions

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

Rearing and Double-stranded RNA-mediated Gene Knockdown in the Hide Beetle, Dermestes maculatus

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

Rearing and Injection of Manduca sexta Larvae to Assess Bacterial Virulence

Mass-Rearing and Molecular Studies in Tortricidae Pest Insects

RNAi-mediated Double Gene Knockdown and Gustatory Perception Measurement in Honey Bees (Apis mellifera)

Culturing and Genetically Manipulating Entomopathogenic Nematodes

Transient Expression and Cellular Localization of Recombinant Proteins in Cultured Insect Cells

Maintaining Aedes aegypti Mosquitoes Infected with Wolbachia

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