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

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

Testing the Physiological Barriers to Viral Transmission in Aphids Using Microinjection

Potato loafroll virus (PLRV), from the family Luteoviridae infects solanaceous plants. It is transmitted by aphids, primarily, the green peach aphid. When an uninfected aphid feeds on an infected plant it contracts the virus through the plant phloem. Once ingested, the virus must pass from the insect gut to the hemolymph (the insect  blood ) and then must pass through the salivary gland, in order to be transmitted back to a new plant. An aphid may take up different viruses when munching on a plant, however only a small fraction will pass through the gut and salivary gland, the two main barriers for transmission to infect more plants. In the lab, we use physalis plants to study PLRV transmission. In this host, symptoms are characterized by stunting and interveinal chlorosis (yellowing of the leaves between the veins with the veins remaining green). The video that we present demonstrates a method for performing aphid microinjection on insects that do not vector PLVR viruses and tests whether the gut is preventing viral transmission.
The video that we present demonstrates a method for performing Aphid microinjection on insects that do not vector PLVR viruses and tests whether the gut or salivary gland is preventing viral transmission.

Aphids are effective transmitters of plant viruses. Aphid microinjection of virus, the procedure we will show you today, is a technique allowing researchers to inject virus directly into the hemocoel of the aphid, bypassing the gut, one of the 2 major barriers for virus transmission in a circulative manner. The same technique is also used to inject dsRNA for RNAi.

Potato loafroll virus (PLRV), from the family Luteoviridae infects solanaceous plants. It is transmitted by aphids, primarily, the green peach aphid. When ...

Biology

Detached Leaf Assays to Simplify Gene Expression Studies in Potato During Infestation by Chewing Insect Manduca sexta

The multitrophic nature of gene expression studies of insect herbivory demands large numbers of biological replicates, creating the need for simpler, more streamlined herbivory protocols. Perturbations of chewing insects are usually studied in whole plant systems. While this whole organism strategy is popular, it is not necessary if similar observations can be replicated in a single detached leaf. The assumption is that basic elements required for signal transduction are present within the leaf itself. In the case of early events in signal transduction, cells need only to receive the signal from the perturbation and transmit that signal to neighboring cells which are assayed for gene expression.
The proposed method simply changes the timing of the detachment. In whole plant experiments, larvae are confined to a single leaf which is eventually detached from the plant and assayed for gene expression. If the order of excision is reversed, from last in whole plant studies, to first in the detached study, the feeding experiment is simplified.
Solanum tuberosum var. Kennebec is propagated by nodal transfer in a simple tissue culture medium and transferred to soil for further growth if desired. Leaves are excised from the parent plant and relocated to Petri dishes where the feeding assay is conducted with the larval stages of M. sexta. Damaged leaf tissue is assayed for the expression of relatively early events in signal transduction. Gene expression analysis identified infestation-specific Cys2-His2 (C2H2) transcription factors, confirming the success of using detached leaves in early response studies. The method is easier to perform than whole plant infestations and uses less space.

The presented method creates natural herbivore damaged plant tissue through the application of Manduca sexta larvae to detached leaves of potato. The plant tissue is assayed for expression of six transcription factor homologs involved in early responses to insect herbivory.

The multitrophic nature of gene expression studies of insect herbivory demands large numbers of biological replicates, creating the need for simpler, more ...

Environment

Analysis of Gene Expression in Emerald Ash Borer (Agrilus planipennis) Using Quantitative Real Time-PCR

Emerald ash borer (EAB, Agrilus planipennis) is an exotic invasive pest, which has killed millions of ash trees (Fraxinus spp) in North America. EAB continues to spread rapidly and attacks ash trees of different ages, from saplings to mature trees. However, to date very little or no molecular knowledge exists for EAB. We are interested in deciphering the molecular-based physiological processes at the tissue level that aid EAB in successful colonization of ash trees. In this report we show the effective use of quantitative real-time PCR (qRT-PCR) to ascertain mRNA levels in different larval tissues (including midgut, fat bodies and cuticle) and different developmental stages (including 1st-, 2nd-, 3rd-, 4th-instars, prepupae and adults) of EAB. As an example, a peritrophin gene (herein named, AP-PERI1) is exemplified as the gene of interest and a ribosomal protein (AP-RP1) as the internal control. Peritrophins are important components of the peritrophic membrane/matrix (PM), which is the lining of the insect gut. The PM has diverse functions including digestion and mechanical protection to the midgut epithelium.

Analysis of Gene Expression in Emerald Ash Borer Agrilus planipennis Using Quantitative Real Time-PCR

Quantitative real-time PCR (qRT-PCR) is an effective tool to diagnose mRNA levels in different insect tissues and developmental stages. In this report we show the use of qRT-PCR to ascertain mRNA levels in different larval tissues and developmental stages of the invasive insect species, emerald ash borer.

Emerald ash borer (EAB, Agrilus planipennis) is an exotic invasive pest, which has killed millions of ash trees (Fraxinus spp) in North America. EAB ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...

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 ...

Immunology and Infection

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant defense response to microbial elicitors and to wounding caused by chewing insects, eliciting systemic calcium signals in plants. However, the role of calcium in vivo during biotic stress is still unclear. This protocol describes the use of a genetically-encoded calcium sensor to detect calcium signals in plants during feeding by a hemipteran pest. Hemipterans such as aphids pierce a small number of cells with specialized, elongated sucking mouthparts, making them the ideal tool to study calcium dynamics when a plant is faced with a biotic stress, which is distinct from a wounding response. In addition, fluorescent biosensors are revolutionizing the measurement of signaling molecules in vivo in both animals and plants. Expressing a GFP-based calcium biosensor, GCaMP3, in the model plant Arabidopsis thaliana allows for the real-time imaging of plant calcium dynamics during insect feeding, with a high spatial and temporal resolution. A repeatable and robust assay has been developed using the fluorescence microscopy of detached GCaMP3 leaves, allowing for the continuous measurement of cytosolic calcium dynamics before, during, and after insect feeding. This reveals a highly-localized rapid calcium elevation around the aphid feeding site that occurs within a few minutes. The protocol can be adapted to other biotic stresses, such as additional insect species, while the use of Arabidopsis thaliana allows for the rapid generation of mutants to facilitate the molecular analysis of the phenomenon.

Real-time In Vivo Recording of Arabidopsis Calcium Signals During Insect Feeding Using a Fluorescent Biosensor

This protocol outlines a simple method for analyzing calcium signals in plants generated by feeding hemipteran insects, such as aphids. Arabidopsis thaliana transformed with the GFP calcium biosensor GCaMP3 allow for the real-time in vivo imaging of calcium dynamics with a high temporal and spatial resolution.

Calcium ions are predicted to be key signaling entities during biotic interactions, with calcium signaling forming an established part of the plant ...

Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain into an axenic insect is a powerful method to verify gut microbial function and elucidate the mechanisms underlying gut microbe-host interactions. Administering antibiotics or sterilizing egg surfaces are two commonly used methods to remove gut bacteria from insects. However, in addition to the potential adverse effects of antibiotics on insects, previous studies indicated that feeding antibiotics could not eliminate gut bacteria. Thus, germ-free artificial diets are generally employed to maintain axenic insects, which is a tedious and labor-intensive process that cannot fully resemble nutritional components in natural food. Described here is an efficient and simple protocol to prepare and maintain axenic larvae of a leaf beetle&#160;(Plagiodera versicolora). Specifically, surfaces of the beetle eggs were sterilized, following which germ-free poplar leaves were used to rear axenic larvae. The axenic status of the insects was further confirmed via culture-dependent and culture-independent assays. Collectively, by combining egg disinfection and germ-free cultivation, an efficient and convenient method was developed to obtain axenic P. versicolora, providing a readily transferable tool for other leaf-eating insects.

To obtain an axenic insect, its egg surface is sterilized, and the hatched larva is subsequently reared using axenic leaves. This method provides an efficient way for axenic insect preparation without administering antibiotics or developing an artificial diet, which can also be applied to other leaf-eating insects.

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain ...

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human food. Outside its native range, culturing G. bimaculatus is feasible due to its dietary plasticity, rapid reproductive cycle, lack of diapause requirement, tolerance for high-density rearing, and robustness against pathogens. Thus, G. bimaculatus can be&#160;a versatile model for studies of insect physiology, behavior, embryology, or genetics.
Cultural parameters, such as stocking density, within-cage refugia, photoperiod, temperature, relative humidity, and diet, all impact cricket growth, behavior, and gene expression and should be standardized. In the burgeoning literature on farming insects for human consumption, these crickets are frequently employed to evaluate candidate feed admixtures derived from crop residues, food-processing byproducts, and other low-cost waste streams.
To support ongoing experiments evaluating G. bimaculatus growth performance and nutritional quality in response to variable feed substrates, a comprehensive set of standard protocols for breeding, upkeep, handling, measurement, and euthanasia in the laboratory was developed and is presented here. An industry-standard cricket feed has proven nutritionally adequate and functionally appropriate for the long-term maintenance of cricket breeding stocks, as well as for use as an experimental control feed. Rearing these crickets at a density of 0.005 crickets/cm3 in screen-topped 29.3 L polyethylene cages at an average temperature of 27 &#176;C on a 12 light (L)/12 dark (D) photoperiod, with moistened coconut coir serving both as hydration source and oviposition medium has successfully sustained healthy crickets over a 2-year span. Following these methods, crickets in a controlled experiment yielded an average mass of 0.724 g 0.190 g at harvest, with 89% survivorship and 68.2% sexual maturation between stocking (22 days) and harvest (65 days).

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

This paper outlines basic methods to standardize important factors such as density, feed availability, hydration source, and environmental controls for the long-term rearing of laboratory cultures of the edible cricket, Gryllus bimaculatus.

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

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

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Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology