The authors thank Bryan T. Vinyard of the USDA, Agricultural Research Service, Henry A. Wallace Beltsville Agricultural Research Center, Biometrical Consulting Service, Beltsville, MD for statistical analyses. This work was supported in part by a grant to J.C.D. from the Deployed War Fighter Protection (DWFP) Research Program funded by the Department of Defense through the Armed Forces Pest Management Board (AFPMB).

1. Rearing Ae. aegypti adults
<ol>
	<li>Hatch eggs in approximately &#190; inch water in shallow tray. Overcrowding will reduce the size of adults.</li>
	<li>Rear larvae at 25 &#176;C (12-hL:12-hD) and feed with ground fish food. 
	NOTE: Overfeeding may reduce survival rates.</li>
	<li>Remove pupae individually by Pasteur pipette daily and transfer to plastic dishes (9 cm &#215; 5.5 cm) inside small containment buckets with fine mesh lids, thus establishing 24 hr&#160;age groupings.</li>
	<li>Feed adult mosquitoes 10% sucrose solution by cotton ball placed on the screen walls of a cage housing the mosquitoes in an environmental chamber at 27 &#176;C and 70% relative humidity under the same photoperiod as larvae.</li>
	<li>For electrophysiology, use 5-10 day old animals. In general, larger animals will produce better results. For RNA isolation, use animals that are within a single 24 hr&#160;age grouping in the 5-10 day old range.</li>
</ol>
2. Preparation of Chemicals
<ol>
	<li>Determine a suitable concentration of electrolyte (e.g. 1-10 mM) that elicits minimal activity from selected sensory neurons. 10 mM NaCl or 1mM KCl are reasonable electrolytes to test baseline activity when beginning an experiment.</li>
	<li>Dissolve each experimental chemical in appropriate solvent (if not water, use ethanol or DMSO) that has little or no effect on spike activity when compared to electrolyte alone. Final concentrations of 10% ethanol or 1% DMSO are reasonable starting concentrations.</li>
	<li>Select appropriate (e.g. quinine for bitter or sucrose for sweet) control chemicals that are well-described feeding deterrents or stimulants in insects.</li>
</ol>
3. Electrophysiology (tip recording8; Figure 1)
<ol>
	<li>Form glass electrodes by mechanically pulling glass capillaries. Optimize heat and pull settings to pull glass to form appropriately shaped recording electrodes. Carefully break off end of glass electrode using forceps under a dissecting microscope to create electrode tip opening that is wide enough to envelope target sensillum, but small enough to avoid contact with neighboring structures.</li>
	<li>Under a microscope, visually identify by morphology and position the target sensilla/sensillum to ensure repeatability of tip recording. Prep animals to allow free access to sensilla from angle of electrode entry.</li>
	<li>Cold anesthetize adult insects in freezer at -20 &#176;C. Avoid damage to peripheral neurons by overexposure to cold temperatures. Cut narrow strips of cellophane tape with razor blade on glass slide. Immobilize whole insect on a glass slide using narrow strips.</li>
	<li>Insert a chemically sharpened tungsten wire into the dorsal thorax or eye to serve as the indifferent (ground) electrode.</li>
	<li>Fill glass electrode made in step 3.1 with a stimulating solution of interest. Using an inserted syringe, fill capillary from pulled end to blunt end. Flick out all air bubbles, holding pulled end down. Insert a silver wire (chloriding optional) into the glass electrode made in step 3.1 to serve as recording and stimulating electrode.</li>
	<li>Connect electrodes to a preamplifier that is designed for recordings from contact chemoreceptive sensilla in insects. This preamplifier (300 Hz-3 kHz bandpass filter) will reduce the settling time to capture neuronal activity as close to stimulus introduction as possible. Collect, store and analyze electrical signals using a microcomputer equipped with spike recording software. 
	NOTE: Grounding the recording setup properly may drastically improve signal-to-noise ratio.</li>
	<li>Stimulate sensilla with a control solution (electrolyte only) to ensure functionality. Count spikes starting 200 msec following the stimulus artifact for all preparations.</li>
	<li>Randomize the order of test chemicals for all experiments, except dose&#8211;response studies, to eliminate bias. For dose&#8211;response studies, expose sensilla to increasing concentrations of the experimental chemical. Allow a minimum of 3 min&#160;between stimulations to ensure adequate recovery. 
	NOTE: This requirement may vary depending on insect and sensilla. Threshold is defined as that concentration for which standard error does not overlap with the standard error for the lowest concentration tested.</li>
</ol>
4. RNA Isolation and Sequencing (Figure 2)
<ol>
	<li>Prepare the tightly fitting RNAse-free pestles by cooling them on dry ice before tissue disruption.Prepare accompanying RNase-free pestles that fit snap cap tubes tightly. Keep collection tubes closed unless actively dissecting to avoid excess condensation.</li>
	<li>Using filtered aspirator, place 30 to 40 cold-anesthetized mosquitoes (15 sec&#160;in -20 &#176;C freezer) on cold stage and sort by sex. Place animals dorsal side down, grasping wings to not damage taste appendages.</li>
	<li>Using two pairs of fine forceps, dissect paired labella or tarsi from males or females under a dissecting microscope and limit inclusion of adjacent tissues. 
	NOTE: 500 paired labella yields approximately 800-1,000 nanograms of total mRNA. 400 individual tarsi (5 segments each) yield 1,200-1,800 nanograms of total mRNA.</li>
	<li>With one pair of gasping-forceps, lightly grasp mosquito proboscis just proximal to labella exposing inner stylets. Using other pair of collection-forceps, remove labella and add tissue to side of cold collection tube.</li>
	<li>With one pair of forceps, lightly grasp mosquito leg at junction of tibia and first tarsal segment. Using other pair of forceps, remove tarsi and add tissue to side of cold collection tube. 
	NOTE: Take into account the cell types at the interface of wanted and unwanted tissue. While it is important to limit the inclusion of neighboring tissue, it is equally important to not omit portions of desired tissue. The final sample must accurately represent the entire organ of interest. Create a precise boundary with the grasping forceps such that collection forceps can precisely liberate by scraping or grabbing only the desired tissue.</li>
	<li>Periodically, spin down collection tube briefly in 0 &#176;C centrifuge at 9,000 x g&#160;to keep tissue at bottom of tube. If moisture accumulates in collection tubes during dissection, add 50uL of cold Trizol reagent to cover collected tissue.</li>
	<li>Using anti-smudge lab marker, clearly label one RNase-free 1.5 ml&#160;snap cap tube for each tissue type for collection. Using 1.5 ml&#160;tube rack and liquid nitrogen, freeze then grind tissue into fine powder (fine pieces if in Trizol). Repeat once. Bring total volume of Trizol to 1 ml&#160;and resuspend ground tissue by vortexing.</li>
	<li>Add 200 ul&#160;of chloroform and mix thoroughly. After 5 min&#160;at room temperature, spin down in centrifuge at 4 &#176;C and 11,000 x g&#160;for 15 min. Carefully add aqueous layer directly to a genomic DNA filter column. Spin down at 11,000 x g&#160;for 5 min. Add equal volume of RNase-free 70% ethanol to flow-through and mix by pipet. Transfer liquid to an RNA collection column and continue RNA extraction per the instructions provided.</li>
	<li>Quantify and assess the purity of total RNA on a precision spectrophotometer and proceed with any standard RNA sequencing method.</li>
</ol>
5. Quantitative RT-PCR Validation of RNA Sequencing (Figure 3)
<ol>
	<li>Select as many genes as possible to compare relative gene expression levels over a dynamic range, both in predicted sequence abundance and presumed chemosensory gene function.</li>
	<li>Design primer pairs for each target gene to amplify a specific 100-180 basepair PCR product. Exclude non-specific gDNA amplification by ensuring at least one primer per set spans an intron boundary. Alternatively, use DNase treatment to reduce genomic contamination.</li>
	<li>Collect insect tissue as described in previous section. Calculate how much tissue is required to provide enough RNA to complete all qRT-PCR reactions. 
	NOTE: Biological triplicate and technical triplicate reactions will provide maximum confidence in results.</li>
	<li>Calculate relative gene quantification as Xtarget-(Ct[target]-Ct[reference]) for each target gene and average for each replicate, both biological and technical. Use housekeeping gene(s) to normalize Ct values between biological replicates and to root the Y-axis scale in comparisons of relative expression.</li>
	<li>Assess the similarity of RNA-seq and qRT-PCR datasets using the regression-based, bootstrap equivalence procedure9, iteratively applied, to the data from each tissue. 
	NOTE: This procedure identifies the smallest &#8220;region of similarity&#8221; that can be constructed around the observed RNA-seq and qRT-PCR data, and allow the relative gene expression as measured by qRT-PCR to be considered statistically equivalent to the measurement of relative gene expression by RNA-seq.</li>
</ol>

<ol><li>Klun, J. A., Khrimian, A., Debboun, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Repellent+and+deterrent+effects+of+SS220%2C+picaridin%2C+and+DEET+suppress+human+blood+feeding+by+Aedes+aegypti%2C+Anopheles+stephensi%2C+and+Phlebotomus+papatasi%5BTitle%5D)+AND+%22J.+Med.+Entomol%22%5BJournal%5D)">Repellent and deterrent effects of SS220, picaridin, and DEET suppress human blood feeding by Aedes aegypti, Anopheles stephensi, and Phlebotomus papatasi.</a> J. Med. Entomol. 43 (1), 34-39 (2006).</li>
<li>Dickens, J. C., Bohbot, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mini+review%3A+Mode+of+action+of+mosquito+repellents.Pestic%5BTitle%5D)+AND+%22Biochem.+Physiol%22%5BJournal%5D)">Mini review: Mode of action of mosquito repellents.Pestic.</a> Biochem. Physiol. 106 (3), 149-155 (2013).</li>
<li>Pitts, R. J., Rinker, D. C., Jones, P. L., Rokas, A., Zwiebel, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transcriptome+profiling+of+chemosensory+appendages+in+the+malaria+vector+Anopheles+gambiae+reveals+tissue-+and+sex-specific+signatures+of+odor+coding%5BTitle%5D)+AND+%22BMC+Genomics%22%5BJournal%5D)">Transcriptome profiling of chemosensory appendages in the malaria vector Anopheles gambiae reveals tissue- and sex-specific signatures of odor coding.</a> BMC Genomics. 12 (271), (2011).</li>
<li>Zhou, X., Slone, J. D., Rokas, A., Berger, S. L., Liebig,, et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Phylogenetic+and+transcriptomic+analysis+of+chemosensory+receptors+in+a+pair+of+divergent+ant+species+reveals+sex-specific+signatures+of+odor+coding%5BTitle%5D)+AND+%22PLOS+Genet%22%5BJournal%5D)">Phylogenetic and transcriptomic analysis of chemosensory receptors in a pair of divergent ant species reveals sex-specific signatures of odor coding.</a> PLOS Genet. 8 (8), e1002930(2012).</li>
<li>Shiao, M., Fan, W., Fang, S., Lu, M. J., Kondo, R., Li, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transcriptional+profiling+of+adult+Drosophila+antenna+by+high-throughput+sequencing%5BTitle%5D)+AND+%22Zoological+Studies%22%5BJournal%5D)">Transcriptional profiling of adult Drosophila antenna by high-throughput sequencing.</a> Zoological Studies. 52 (42), (2013).</li>
<li>Bohbot, J., Pitts, R. J., Kwon, H. W., Rützler, M., Robertson, H. M., Zwiebel, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+characterization+of+the+Aedes+aegypti+odorant+receptor+gene+family%5BTitle%5D)+AND+%22Insect+Mol.+Biol%22%5BJournal%5D)">Molecular characterization of the Aedes aegypti odorant receptor gene family.</a> Insect Mol. Biol. 16 (5), 525-537 (2007).</li>
<li>Kain, P., Boyle, S. M., Tharadra, S. K., Guda, T., Pham, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Odour+receptors+and+neurons+for+DEET+and+new+insect+repellents%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Odour receptors and neurons for DEET and new insect repellents.</a> Nature. 502 (7472), 507-512 (2013).</li>
<li>Hodgson, E. S., Lettvin, J. Y., Roeder, K. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+physiology+of+a+primary+chemoreceptor+unit%5BTitle%5D)+AND+%22Science%22%5BJournal%5D)">The physiology of a primary chemoreceptor unit.</a> Science. 122 (3166), 417-418 (1955).</li>
<li>Robinson, A. P., Duursma, R. A., Marshall, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+regression-based+equivalence+test+for+model+validation%3A+shifting+the+burden+of+proof%5BTitle%5D)+AND+%22Tree+Physiol%22%5BJournal%5D)">A regression-based equivalence test for model validation: shifting the burden of proof.</a> Tree Physiol. 25 (7), 903-913 (2005).</li>
<li>Mortazavi, A., Williams, B. A., McCue, K., Schaeffer, L., Wold, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mapping+and+quantifying+mammalian+transcriptomes+by+RNA-seq%5BTitle%5D)+AND+%22Nat.+Methods%22%5BJournal%5D)">Mapping and quantifying mammalian transcriptomes by RNA-seq.</a> Nat. Methods. 5 (7), 621-628 (2008).</li>
<li>Trapnell, C., Williams, B. A., Pertea, G., Mortazavi, A., Kwan, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Transcript+assembly+and+quantification+by+RNA-seq+reveals+unannotated+transcripts+and+isoform+switching+during+cell+differentiation%5BTitle%5D)+AND+%22Nat.+Biotechnol%22%5BJournal%5D)">Transcript assembly and quantification by RNA-seq reveals unannotated transcripts and isoform switching during cell differentiation.</a> Nat. Biotechnol. 28 (5), 511-515 (2010).</li>
<li>Sanford, J. L., Shields, V. D. C., Dickens, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Gustatory+receptor+neuron+responds+to+DEET+and+other+insect+repellents+in+the+yellow-fever+mosquito%2C+Aedes+aegypti%5BTitle%5D)+AND+%22Naturwiss%22%5BJournal%5D)">Gustatory receptor neuron responds to DEET and other insect repellents in the yellow-fever mosquito, Aedes aegypti.</a> Naturwiss. 100 (3), 269-273 (2013).</li>
<li>Sparks, J. T., Vinyard, B. T., Dickens, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Gustatory+receptor+expression+in+the+labella+and+tarsi+of+Aedes+aegypti%5BTitle%5D)+AND+%22Insect+Biochem.+Mol.+Biol%22%5BJournal%5D)">Gustatory receptor expression in the labella and tarsi of Aedes aegypti.</a> Insect Biochem. Mol. Biol. 43 (12), 1161-1171 (2013).</li>
<li>Kent, L. B., Walden, K. K. O., Robertson, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+Gr+family+of+candidate+gustatory+and+olfactory+receptors+in+the+yellow-fever+mosquito+Aedes+aegypti%5BTitle%5D)+AND+%22Chem.+Senses%22%5BJournal%5D)">The Gr family of candidate gustatory and olfactory receptors in the yellow-fever mosquito Aedes aegypti.</a> Chem. Senses. 33 (1), 79-93 (2008).</li>
<li>Ramsköld, D., Wang, E. T., Burge, C. B., Sandberg, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+abundance+of+ubiquitously+expressed+genes+revealed+by+tissue+transcriptome+sequence+data%5BTitle%5D)+AND+%22PLoS+Comput.+Biol%22%5BJournal%5D)">An abundance of ubiquitously expressed genes revealed by tissue transcriptome sequence data.</a> PLoS Comput. Biol. 5 (12), e1000598(2009).</li>
<li>Wagner, G. P., Kin, K., Lynch, V. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((A+model+based+criterion+for+gene+expression+calls+using+RNA-seq+data%5BTitle%5D)+AND+%22Theory+Biosci%22%5BJournal%5D)">A model based criterion for gene expression calls using RNA-seq data.</a> Theory Biosci. 132 (3), 159-164 (2013).</li>
<li>Hart, T., Komori, H. K., LaMere, S., Podshivalova, K., Salomon, D. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Finding+active+genes+in+deep+RNA-seq+gene+expression+studies%5BTitle%5D)+AND+%22BMC+Genomics%22%5BJournal%5D)">Finding active genes in deep RNA-seq gene expression studies.</a> BMC Genomics. 14 (778), (2013).</li>
<li>Isono, K., Morita, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+and+cellular+designs+of+insect+taste+receptor+system%5BTitle%5D)+AND+%22Front.+Cellu.+Neurosci%22%5BJournal%5D)">Molecular and cellular designs of insect taste receptor system.</a> Front. Cellu. Neurosci. 4 (20), 1-16 (2010).</li>
</ol>

The authors have nothing to disclose.

The most challenging aspect of recording action potentials from gustatory sensilla is deciding which responses are &#8220;normal.&#8221; When employing single gustatory sensillum tip recording the first time for a given insect species, the total number and sensitivities of the gustatory receptor neurons (GRNs) are likely unknown. Many preliminary recordings are first required to decide the range and concentrations of chemicals to test. In this instance, we began with the observation that a single GRN responded to low con...

Compounds like DEET, Picaridin, Citronellal and IR3535 have been shown to effectively repel mosquitoes, including the important disease vector Aedes aegypti1,2. We record action potentials from sensory neurons associated with specific gustatory sensilla to determine the cells involved with mosquito repellency. Coupled with downstream sequencing of expressed genes in these tissues, we may identify the genes most likely mediating the responses of these cells in order to screen new compounds for improved repellency capabilities.
RNA-seq is a powerful tool, quickly becoming standard for tracking temporal and spatial changes in gene expression. RNA-seq analyses of insect chemosensory appendages and organs have been used to uncover molecular receptors in several insect species3-5, greatly improving on conventional PCR-based searches gene by gene6. Insects represent the most diverse animal class, presenting many opportunities to study the connection between genes and unique phenotypes. RNA-seq technology can be employed on any living insect tissue. Likewise, electrophysiological recording from sensory cells within uniporous gustatory sensilla can be achieved in many different insect species. The pairing of these two techniques allows researchers to narrow the set genes involved in an observed chemosensory phenotype. Different species will present specific challenges, but may inform the connection between chemosensory receptor genes and a chemosensory adaptation. The size and morphology of chemosensory sensilla is variable and may require extensive troubleshooting when recording action potentials to reduce noise and identify repeatable signals. Dissections of chemosensory organs may be trivial or delicate and time consuming, depending on morphology and size of the insect. Recovery of high-quality RNA may require some troubleshooting as well, such as avoiding certain pigments during tissue collection.
While demonstrating the effects of repellent compounds through behavioral trials is direct and informative, this approach is time intensive and broad with respect to mechanism of action. Electrophysiology coupled with RNA-seq allows for more specific analyses of what drives avoidance behaviors in insects. Once the &#8220;toolkit&#8221; of chemical discrimination has been identified in an insect species, more specific attempts to improve on known repellents are possible. Receptors and associated proteins in sensory cells responsible for these behaviors may be expressed heterologously for direct chemical screening. Furthermore, molecular modeling can predict which chemicals will elicit strong responses from these receptors7.
The snapshot of all active genes in a narrow set of chemosensory tissues may also be useful in identifying similar genes in other species. Using sequence homology and expression similarities, researchers may form sets of molecular receptors most likely mediating responses to repellents that are broadly effective on insects. We present the following protocol to aid researchers in deconstructing insect chemosensory pathways and to persuade more to delve into the neuroethology of non-model and economically important insects.

<table><tbody><tr><td>Glass capillary</td><td>A-M Systems</td><td>628000</td><td>Standard, 1.5 mm X 0.86 mm, 4&quot;</td></tr><tr><td>Silver wire</td><td>A-M Systems</td><td>7875</td><td>.015&quot; bare</td></tr><tr><td>Tungsten wire</td><td>Alfa Products</td><td>369</td><td>0.127 mm diameter</td></tr><tr><td>Fine forceps</td><td>Fine Science Tools</td><td>11252</td><td>#5SF Inox</td></tr><tr><td>Refridgerated stage</td><td>BioQuip Products</td><td>1429</td><td>Chill Table</td></tr><tr><td>Preamplifier</td><td>Syntech</td><td>Taste Probe</td><td>preamplifier</td></tr><tr><td>Software for electrophysiology</td><td>Syntech</td><td>Autospike</td><td>software for electrophysiology</td></tr><tr><td>TetraMin fish food</td><td>Tetra</td><td>Tropical fish food granules</td><td>fish food ground to fine powder</td></tr><tr><td>TRIzol</td><td>Life Technologies</td><td>15596-026</td><td>RNA isolation reagent</td></tr><tr><td>RNeasy Plus Mini Kit</td><td>Qiagen</td><td>74136</td><td>includes gDNA eliminator and RNeasy spin columns</td></tr><tr><td>Nanodrop spectrophotometer</td><td>Nano Drop Products</td><td>ND-1000</td><td>tabletop spectrometer</td></tr><tr><td>R statistics</td><td>freeware (created by Robert Gentleman and Ross Ihaka)</td><td>www.r-project.org</td><td>Use the lm function of the stats package and the equiv.boot function of the equivalence package in the R computing environment.</td></tr><tr><td>1.5 ml tube rack</td><td>Evergreen</td><td>240-6388-030</td><td>Pour liquid nitrogen into a few empty wells to freeze and grind tissue.</td></tr><tr><td>1.5 ml collection tubes with pestle</td><td>Grainger</td><td>6HAX6</td><td>RNase free</td></tr><tr><td>Centrifuge</td><td>Thermo Scientific</td><td>11178160</td><td>Spin down frozen tissue to keep at bottom of 1.5 ml tube.</td></tr><tr><td>Primer-BLAST Primer Designing tool</td><td>NCBI</td><td>n/a</td><td></td></tr></tbody></table>

physiological recordings and rna sequencing of the gustatory appendages of the yellow-fever mosquito aedes aegypti

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of these disease vectors. We have recently identified a bitter sensing neuron in the labellum of female Aedes aegypti that responds to DEET and other repellents, as well as bitter quinine, through direct electrophysiological investigation. These gustatory receptor neuron responses prompted our sequencing of total mRNA from both male and female labella and tarsi samples to elucidate the putative chemoreception genes expressed in these contact chemoreception tissues. Samples of tarsi were divided into pro-, meso- and metathoracic subtypes for both sexes. We then validated our dataset by conducting qRT-PCR on the same tissue samples and used statistical methods to compare results between the two methods. Studies addressing molecular function may now target specific genes to determine those involved in repellent perception by mosquitoes. These receptor pathways may be used to screen novel repellents towards disruption of host-seeking behavior to curb the spread of harmful viruses.

The trace recordings of action potentials from Ae. aegypti gustatory sensilla (Figure 1) demonstrate the effectiveness of direct stimulation with a range of chemicals. This technique can be used to quantify responses to any stimulating chemical by counting spikes of a given amplitude and duration over a reasonable time range (generally less than 500 ms). Trace recordings must be readily reproducible under a given set of experimental conditions. Otherwise, the observed physiological responses may...

Watch this Scientific Journal Video about Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti at JoVE.com

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Electrophysiological recording of action potentials from sensory neurons of mosquitoes provides investigators a glimpse into the chemical perception of ...

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JoVE Journal

Biology

8.1K Views.  United States Department of Agriculture. Using two methods to estimate gene expression in the major gustatory appendages of Aedes aegypti, we have identified the set of genes putatively underlying the neuronal responses to bitter and repulsive compounds, as determined by electrophysiological examination.

Video: Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed in sensory hairs called sensilla, which cover the surface of olfactory organs. The surface of each sensillum is covered with tiny pores, through which odorants pass and dissolve in a fluid called sensillum lymph, which bathes the sensory dendrites of the OSNs housed in a given sensillum. The OSN dendrites express odorant receptor (OR) proteins, which in insects function as odor-gated ion channels4, 5. The interaction of odorants with ORs either increases or decreases the basal firing rate of the OSN. This neuronal activity in the form of action potentials embodies the first representation of the quality, intensity, and temporal characteristics of the odorant6, 7.
Given the easy access to these sensory hairs, it is possible to perform extracellular recordings from single OSNs by introducing a recording electrode into the sensillum lymph, while the reference electrode is placed in the lymph of the eye or body of the insect. In Drosophila, sensilla house between one and four OSNs, but each OSN typically displays a characteristic spike amplitude. Spike sorting techniques make it possible to assign spiking responses to individual OSNs. This single sensillum recording (SSR) technique monitors the difference in potential between the sensillum lymph and the reference electrode as electrical spikes that are generated by the receptor activity on OSNs1, 2, 8. Changes in the number of spikes in response to the odorant represent the cellular basis of odor coding in insects. Here, we describe the preparation method currently used in our lab to perform SSR on Drosophila melanogaster and Anopheles gambiae, and show representative traces induced by the odorants in a sensillum-specific manner.

Single Sensillum Recordings in the Insects Drosophila melanogaster and Anopheles gambiae

Electrophysiological responses of olfactory sensory neurons to odorants can be measured in insects using single sensillum recordings. In this video article we will demonstrate how to perform single sensillum recordings in the antennae of the vinegar fly (Drosophila melanogaster) and the maxillary palps of the malaria mosquito (Anopheles gambiae).

The sense of smell is essential for insects to find foods, mates, predators, and oviposition sites3. Insect olfactory sensory neurons (OSNs) are enclosed ...

Neuroscience

Dissection of Midgut and Salivary Glands from Ae. aegypti Mosquitoes

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol demonstrates dissection techniques for removal of the midgut and salivary glands from Aedes aegypti mosquitoes. 



The mosquito midgut and salivary glands are key entry and exit points for vector pathogens like Plasmodium falciparum and the dengue virus. This video demonstrates the dissection techniques for removing the midgut and salivary glands from Aedes aegypti mosquitoes.

The mosquito midgut and salivary glands are key entry and exit points for pathogens such as Plasmodium parasites and Dengue viruses. This video protocol ...

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the researcher to measure gustatory responses directly and quantitatively, reflecting the sensory input that the insect nervous system receives from taste stimuli in its environment. This protocol outlines all key steps in performing this technique. The critical steps in assembling an electrophysiology rig, such as selection of necessary equipment and a suitable environment for recording, are delineated. We also describe how to prepare for recording by making appropriate reference and recording electrodes, and tastant solutions. We describe in detail the method used for preparing the insect by insertion of a glass reference electrode into the fly in order to immobilize the proboscis. We show traces of the electrical impulses fired by taste neurons in response to a sugar and a bitter compound. Aspects of the protocol are technically challenging and we include an extensive description of some common technical challenges that may be encountered, such as lack of signal or excessive noise in the system, and potential solutions. The technique has limitations, such as the inability to deliver temporally complex stimuli, observe background firing immediately prior to stimulus delivery, or use water-insoluble taste compounds conveniently. Despite these limitations, this technique (including minor variations referenced in the protocol) is a standard, broadly accepted procedure for recording Drosophila neuronal responses to taste compounds.

Electrophysiological Recording From Drosophila Labellar Taste Sensilla

This protocol describes extracellular recording of the action potential responses fired by labellar taste neurons in Drosophila.

The peripheral taste response of insects can be powerfully investigated with electrophysiological techniques. The method described here allows the ...

New Methods to Study Gustatory Coding

The sense of taste allows animals to detect chemicals in the environment, giving rise to behaviors critical for survival. When Gustatory Receptor Neurons (GRNs) detect tastant molecules, they encode information about the identity and concentration of the tastant as patterns of electrical activity that then propagate to follower neurons in the brain. These patterns constitute internal representations of the tastant, which then allow the animal to select actions and form memories. The use of relatively simple animal models has been a powerful tool to study basic principles in sensory coding. Here, we propose three new methods to study gustatory coding using the moth Manduca sexta. First, we present a dissection procedure for exposing the maxillary nerves and the subesophageal zone (SEZ), allowing recording of the activity of GRNs from their axons. Second, we describe the use of extracellular electrodes to record the activity of multiple GRNs by placing tetrode wires directly into the maxillary nerve. Third, we present a new system for delivering and monitoring, with high temporal precision, pulses of different tastants. These methods allow the characterization of neuronal responses in vivo directly from GRNs before, during and after tastants are delivered. We provide examples of voltage traces recorded from multiple GRNs, and present an example of how a spike sorting technique can be applied to the data to identify the responses of individual neurons. Finally, to validate our recording approach, we compare extracellular recordings obtained from GRNs with tetrodes to intracellular recordings obtained with sharp glass electrodes.

We present three new methods to study gustatory coding. Using a simple animal, the moth Manduca sexta (Manduca), we describe a dissection protocol, the use of extracellular tetrodes to record the activity of multiple gustatory receptor neurons, and a system for delivering and monitoring precisely timed pulses of tastants.

The sense of taste allows animals to detect chemicals in the environment, giving rise to behaviors critical for survival. When Gustatory Receptor Neurons ...

Single Sensillum Recordings for Locust Palp Sensilla Basiconica

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially for the detection of non-volatile chemical cues through sensilla chaetica (previously named terminal sensilla or crested sensilla). There is now increasing evidence that these palps also have an olfactory function. An odorant receptor (LmigOR2) and an odorant-binding protein (LmigOBP1) have been localized in the neurons and accessory cells, respectively, in the sensilla basiconica of the palps. Single sensillum recording (SSR) is used for recording the responses of odorant receptor neurons, which is an effective method for screening active ligands on specific odorant receptors. SSR is used in functional studies of odorant receptors in palp sensilla. The structure of the sensilla basiconica located on the dome of the palps differs somewhat from the structure of those on the antennae. Therefore, when performing an SSR elicited by odorants, some specific advice may be helpful for obtaining optimum results. In this paper, a detailed and highly effective protocol for an SSR from insect palp sensilla basiconica is introduced.

This paper describes a detailed and highly effective protocol for single sensillum recordings from the sensilla basiconica on the palps of insect mouthparts.

The palps of locust mouthparts are considered to be conventional gustatory organs that play an important role in a locust&#39;s food selection, especially ...

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In the laboratory, researchers can deliver animal-derived and artificial blood meals to mosquitoes via membrane feeders, which allow for manipulation of meal composition. Here, we present methods for feeding blood and artificial blood meals to Aedes aegypti mosquitoes and quantifying the volume consumed by individual females.
Targeted feeding and quantification of artificial/blood meals have broad uses, including testing the effects of meal components on mosquito behavior and physiology, delivering pharmacological compounds without injection, and infecting mosquitoes with specific pathogens. Adding fluorescein dye to the meal prior to feeding allows for subsequent meal size quantification. The meal volume consumed by mosquitoes can be measured either by weight, if the females are to be used later for behavioral experiments, or by homogenizing individual females in 96-well plates and measuring fluorescence levels using a plate reader as an endpoint assay. Meal size quantification can be used to determine whether changing the meal components alters the meal volume ingested or if meal consumption differs between mosquito strains. Precise meal size quantification is also critical for downstream assays, such as those measuring effects on host attraction or fecundity. The methods presented here can be further adapted to track meal digestion over the course of days or to include multiple distinguishable markers added to different meals (like nectar and blood) to quantify the consumption of each meal by a single mosquito.
These methods allow researchers to singlehandedly perform high-throughput measurements to compare the meal volume consumed by hundreds of individual mosquitoes. These tools will therefore be broadly useful to the community of mosquito researchers for answering diverse biological questions.

Feeding and Quantifying Animal-Derived Blood and Artificial Meals in Aedes aegypti Mosquitoes

The goal of this protocol is to deliver animal-derived and artificial blood meals to Aedes aegypti mosquitoes through an artificial membrane feeder and precisely quantify the volume of meal ingested.

Females of certain mosquito species can spread diseases while biting vertebrate hosts to obtain protein-rich blood meals required for egg development. In ...

Immunology and Infection

Heterologous Expression and Functional Analysis of Aedes aegypti Odorant Receptors to Human Odors in Xenopus Oocytes

The mosquito Aedes aegypti (Linnaeus), a vector of many important human diseases including yellow fever, dengue fever and Zika fever, shows a strong preference for human hosts over other warm-blooded animals for blood meals. Olfactory cues play a critical role for mosquitoes as they explore their environment and seek a human host to obtain blood meals, thus transmitting human diseases. Odorant receptors (ORs) expressed in the olfactory sensory neurons are known to be responsible for the interaction of mosquito vectors with human odors. To gain deeper insights into Ae. aegypti&#8217;s olfactory physiology and investigate their interactions with humans at the molecular level, we used an optimized protocol of Xenopus Oocytes heterologous expression to functionally analyze Ae. aegypti odorant receptors in response to human odors. Three example experiments are presented: 1) Cloning and synthesizing cRNAs of ORs and odorant receptor co-receptor&#160;(Orco) from four to six days old Ae. aegypti antennae; 2) Microinjection and expression of ORs and Orco in Xenopus oocytes; and 3) Whole-cell current recording from Xenopus oocytes expressing mosquito ORs/Orco with a two-electrode voltage-clamp. These optimized procedures provide a new way for researchers to investigate human odor reception in Aedes mosquitoes and reveal the underlying mechanisms governing their host-seeking activity at a molecular level.

Heterologous Expression and Functional Analysis of Aedes aegypti Odorant Receptors to Human Odors in Xenopus Oocytes

A protocol is presented that functionally characterizes mosquito ORs in response to human odors using a Xenopus oocyte expression system coupled with a two-electrode voltage clamp, providing a powerful new technique for exploring the responses of mosquitoes ORs to exposure to human odors.

The mosquito Aedes aegypti (Linnaeus), a vector of many important human diseases including yellow fever, dengue fever and Zika fever, shows a strong ...

An Experimental and Bioinformatics Protocol for RNA-seq Analyses of Photoperiodic Diapause in the Asian Tiger Mosquito, Aedes albopictus

Photoperiodic diapause is an important adaptation that allows individuals to escape harsh seasonal environments via a series of physiological changes, most notably developmental arrest and reduced metabolism. Global gene expression profiling via RNA-Seq can provide important insights into the transcriptional mechanisms of photoperiodic diapause. The Asian tiger mosquito, Aedes albopictus, is an outstanding organism for studying the transcriptional bases of diapause due to its ease of rearing, easily induced diapause, and the genomic resources available. This manuscript presents a general experimental workflow for identifying diapause-induced transcriptional differences in A. albopictus. Rearing techniques, conditions necessary to induce diapause and non-diapause development, methods to estimate percent diapause in a population, and RNA extraction and integrity assessment for mosquitoes are documented. A workflow to process RNA-Seq data from Illumina sequencers culminates in a list of differentially expressed genes. The representative results demonstrate that this protocol can be used to effectively identify genes differentially regulated at the transcriptional level in A. albopictus due to photoperiodic differences. With modest adjustments, this workflow can be readily adapted to study the transcriptional bases of diapause or other important life history traits in other mosquitoes.

An Experimental and Bioinformatics Protocol for RNA-seq Analyses of Photoperiodic Diapause in the Asian Tiger Mosquito, Aedes albopictus

RNA-Seq analyses are becoming increasingly important for identifying the molecular underpinnings of adaptive traits in non-model organisms. Here, a protocol to identify differentially expressed genes between diapause and non-diapause Aedes albopictus mosquitoes is described, from mosquito rearing, to RNA sequencing and bioinformatics analyses of RNA-Seq data.

Photoperiodic diapause is an important adaptation that allows individuals to escape harsh seasonal environments via a series of physiological changes, ...

Fat Body Organ Culture System in Aedes Aegypti, a Vector of Zika Virus

The insect fat body plays a central role in insect metabolism and nutrient storage, mirroring functions of the liver and fat tissue in vertebrates. Insect fat body tissue is usually distributed throughout the insect body. However, it is often concentrated in the abdomen and attached to the abdominal body wall.
The mosquito fat body is the sole source of yolk proteins, which are critical for egg production. Therefore, the in vitro culture of mosquito fat body tissues represents an important system for the study of mosquito physiology, metabolism, and, ultimately, egg production. The fat body culture process begins with the preparation of solutions and reagents, including amino acid stock solutions, Aedes physiological saline salt stock solution (APS), calcium stock solution, and fat body culture medium. The process continues with fat body dissection, followed by an experimental treatment. After treatment, a variety of different analyses can be performed, including RNA sequencing (RNA-Seq), qPCR, Western blots, proteomics, and metabolomics.
In our example experiment, we demonstrate the protocol through the excision and culture of fat bodies from the yellow fever mosquito, Aedes aegypti, a principal vector of arboviruses including dengue, chikungunya, and Zika. RNA from fat bodies cultured under a physiological condition known to upregulate yolk proteins versus the control were subject to RNA-Seq analysis to demonstrate the potential utility of this procedure for investigations of gene expression.

Fat Body Organ Culture System in Aedes Aegypti, a Vector of Zika Virus

The fat body is the central metabolic organ in insects. We present a live organ culture system that enables the user to study the responses of isolated fat body tissue to various stimuli.

The insect fat body plays a central role in insect metabolism and nutrient storage, mirroring functions of the liver and fat tissue in vertebrates. Insect ...

A Step-by-Step Guide to Mosquito Electroantennography

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when acquiring a blood-meal. To locate a host to feed on, mosquitoes rely on a wide range of sensory cues, including visual, mechanical, thermal, and olfactory. The study details a technique, electroantennography (EAG), that allows researchers to assess whether the mosquitoes can detect individual chemicals and blends of chemicals in a concentration-dependent manner. When coupled with gas-chromatography (GC-EAG), this technique allows to expose the antennae to a full headspace/complex mixture and determines which chemicals present in the sample of interest, the mosquito can detect. This is applicable to host body odors as well as plant floral bouquets or other ecologically relevant odors (e.g., oviposition sites odorants). Here, we described a protocol that permits long durations of preparation responsiveness time and is applicable to both female and male mosquitoes from multiple genera, including Aedes, Culex, Anopheles, and Toxorhynchites mosquitoes. As olfaction plays a major part in mosquito-host interactions and mosquito biology in general, EAGs and GC-EAG can reveal compounds of interest for the development of new disease vector control strategies (e.g., baits). Complemented with behavioral assays, the valence (e.g., attractant, repellent) of each chemical can be determined.

The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males.

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when ...

Physiological Recordings and RNA Sequencing of the Gustatory Appendages of the Yellow-fever Mosquito Aedes aegypti

Summary

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Heterologous Expression and Functional Analysis of Aedes aegypti Odorant Receptors to Human Odors in Xenopus Oocytes

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Fat Body Organ Culture System in Aedes Aegypti, a Vector of Zika Virus

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