Name: administering and detecting protein marks on arthropods for dispersal research
Uploaded: 2016-01-28T15:00:00.000+00:00
Duration: 10 min 30 s
Description: Watch this Scientific Journal Video about Administering and Detecting Protein Marks on Arthropods for Dispersal Research at JoVE.com

Funding was provided by USDA CRIS 5347-22620-021-00D and, in part, by the Agriculture and Food Research Initiative Competitive Grant no. 2011-67009-30141 from the USDA National Institute of Food and Agriculture. We are grateful for the technical support of Johanna Nassif. We also thank Paul Baker, David Horton, Diego Nieto, and Frances Sivakoff for providing some of the pictures used in Figure 3.

1. Internal Mark, Retention, and Detection Procedure
<ol>
	<li>Internal marking procedure
	<ol>
		<li>Collect insects of interest (n &#8776; 100 individuals) from a laboratory colony reared on an artificial diet or from the field and divide into two clean rearing containers.</li>
		<li>Place a regular 20 ml diet packet (unmarked negative control treatment) into one of the containers. Supplement a second 20 ml artificial diet packet with 1.0 ml of a 1.0 mg/ml chicken IgG/IgY solution, mix thoroughly, and place it in the other container. 
		Note: If the insect of interest does not have an artificial diet, the mark can be placed on or in whatever diet they are normally sustained on (e.g., green bean, moth eggs, water, sugar, honey).</li>
		<li>Allow the insects to resume their feeding activity for 24 hr.</li>
	</ol>
	</li>
	<li>Internal mark retention procedure
	<ol>
		<li>Remove the diet packets from each container and supply the insects with another food source to sustain them over the duration of the experiment (e.g., green beans).</li>
		<li>Collect a cohort (n = 20) of insects daily (or at any desired time interval) from each treatment and freeze immediately at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Sandwich ELISA procedure
	<ol>
		<li>Add 50 &#181;l of rabbit anti-chicken IgY primary antibody diluted 1:500 in Tris buffered saline (TBS) to each well of a clean ELISA microplate and incubate for a minimum of 1 hr at RT (note: the microplates can also be incubated O/N at 4 &#176;C).</li>
		<li>Discard the antibody from the plate and block each well with 300 &#181;l of a 1.0% non-fat dry milk solution for 30 min.</li>
		<li>While waiting for the previous incubation steps, place frozen insects individually in 1.6 ml microcentrifuge tubes with 1.0 ml of TBS.</li>
		<li>Grind each insect with a clean pestle until thoroughly homogenized and add a 100 &#181;l of each sample to an individual well of the ELISA plate. Allow the samples to incubate at RT for 1 hr.</li>
		<li>Add 100 &#181;l of negative (TBS only) and positive (egg whites in TBS) controls to wells in the first column of each ELISA plate. Also, add 100 &#181;l of negative insect controls (unmarked insects) to the 8 wells in the last column of each plate (Figure 1). Allow the control samples to incubate at RT for 1 hr. Note that the template provided in Figure 1 is the template we use for our standardized assays. The placement of the samples in the wells is up to the discretion of the researcher.</li>
		<li>Wash each well three times with phosphate buffered saline (PBS)-tween and then add 50 &#181;l of rabbit anti-chicken IgG/IgY conjugated with horseradish peroxidase diluted 1:10,000 in 1% milk solution and incubate for 1 hr at RT.</li>
		<li>Wash each well three times with PBS-tween and add 50 &#181;l of tetramethylbenzidine (TMB) substrate to each well.</li>
		<li>After 10 min, read the wells with a microplate spectrophotometer set at a wavelength of 650 nm.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/53693/53693fig1.jpg" /> 
Figure 1. A schematic diagram showing the standardized well designations used for a typical 96 well ELISA microplate. Each plate contains 7 wells dedicated to Tris-buffered saline negative controls (TBS), a well dedicated to a positive protein control (Pos Ctrl), 80 individual wells dedicated to recaptured insects (Sample 1&#8230;80), and 8 wells dedicated to unmarked (negative) insect controls (Neg Ctrl 1&#8230;8). <a href="https://www.jove.com/files/ftp_upload/53693/53693fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. External Mark, Retention, and Detection Procedure
<ol>
	<li>External marking procedure
	<ol>
		<li>Collect insects of interest (n &#8776; 100 individuals) from a laboratory colony or from the field and place into two 1.0 L plastic containers with a 2.5 cm diameter hole punched out of the side of each container.</li>
		<li>Spray the insects in the first container with 1.0 ml of dH2O (negative control treatment) using a medical nebulizer. Spray the insects in the second container with 1.0 ml of a 5.0% solution of chicken egg whites using a medical nebulizer.</li>
		<li>Attach the hose of the nebulizer to a standard laboratory air outlet and then insert the mouth of the nebulizer into the 2.5 cm hole of the container. Then, turn on the air outlet gradually until the nebulizer produces a vapor.</li>
		<li>Continue spraying (marking) until the egg whites solution has been dispensed (approx. 2-5 min).</li>
		<li>Remove the nebulizer and place a cork into the hole of the plastic container. Allow the insects to stand at RT until the solution has dried completely.</li>
	</ol>
	</li>
	<li>External mark retention procedure
	<ol>
		<li>Place each treatment of the dry insects into a separate clean rearing container to avoid further exposure to the mark.</li>
		<li>Add food and water to the clean containers to sustain the insects over the duration of the study.</li>
		<li>Collect a cohort (n = 20) of insects from each treatment daily and freeze immediately at -20 &#176;C.</li>
	</ol>
	</li>
	<li>Indirect ELISA procedure
	<ol>
		<li>Place the frozen insects individually into 1.6 ml microcentrifuge tubes and add 1.0 ml of TBS.</li>
		<li>Soak the samples for 1 hr on an orbital shaker set at 120 rpm.</li>
		<li>Add negative and positive controls as described in Step 1.3.5.</li>
		<li>After soaking, pipette a 100 &#181;l aliquot from each sample into one of the 80 designated sample wells on a new 96 well ELISA plate as shown in Figure 1. Allow the samples to incubate for 1 hr at RT (note: the microplates can also be incubated O/N at 4 &#176;C).</li>
		<li>Wash the wells three times with PBS-tween and then add 300 &#181;l of phosphate buffered saline containing 1% bovine serum albumin (PBS-BSA) to each well.</li>
		<li>After 30 min at RT wash twice with PBS-tween.</li>
		<li>Add 50 &#181;l of rabbit anti-egg albumin primary antibody diluted 1:8,000 in PBS-BSA-Silwet (0.05%) to each well and incubate for 1 hr at RT.</li>
		<li>Wash each well three times with PBS-tween and then add 50 &#181;l of goat anti-rabbit IgG conjugated with horseradish peroxidase diluted 1:2,000 in PBS-BSA-Silwet (0.05%) for 1 hr at RT.</li>
		<li>Wash each well three times with PBS-tween and add 50 &#181;l of TMB substrate to each well.</li>
		<li>After 10 min, read the wells with a microplate spectrophotometer set at a wavelength of 650 nm.</li>
	</ol>
	</li>
</ol>
3. Data Analysis&#160;
<ol>
	<li>Calculate the mean absorbance value of the 8 negative control insects on each ELISA plate and then calculate the standard deviation based on the pooled negative controls from all of the ELISA plates of a given study12.</li>
	<li>Add six times the pooled standard deviation to the mean obtained for each ELISA plate to obtain an ELISA positive threshold value. Any insect sample yielding an absorbance value equal to or greater than this value is deemed to be positive for the presence of the protein mark.</li>
</ol>

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The arthropod protein immunomarking procedure was first described almost a quarter of a century ago9. Since then, the procedure has been adapted to study the dispersal patterns of a wide variety of arthropods using both internally and externally administered IgG/IgYs. These proteins have proven steadfast markers for the wide variety of insect species tested thus far. However, as mentioned above, the major limitation for using IgG/IgYs is that they are very expensive. Consequently, IgG/IgYs are only useful for ...

Tracking the movement of arthropod pests, natural enemies (parasitoids and predators), and pollinators in nature is essential for better understanding how to improve ecosystem services. The key component for most types of dispersal research is having a reliable method to tag the arthropod(s) of interest. A variety of materials (e.g., paints, dyes, colored dusts, tags, rare elements, proteins) have been used to mark arthropods to assess their population dynamics, dispersal capabilities, feeding behaviors, and other ecological interactions1,2.
The appropriateness of a marker used for any given dispersal research will be dependent on the type of study being conducted. There are three broad categorizations for marking arthropods: (1) mark-release-recapture (MRR), (2) mark-capture, and (3) self-mark-capture. For mark-release-recapture research, the investigator typically marks the arthropods collectively in the laboratory and releases them at a central point in the field. The arthropods are then recaptured at various spatial and temporal intervals using different collection devices (e.g., sweep net, vacuum, sticky trap)3,4,5. The recaptured specimens are then examined for the specific mark to distinguish released from native individuals. For mark-capture research, the investigator usually applies the mark directly in the field using spray equipment (e.g., backpack sprayer, boom and nozzle sprayer). The best markers for mark-capture research are inexpensive and easily applied to the arthropod&#39;s habitat. For self-mark-capture research, the investigator usually applies marks to an arthropod bait6,7 or nest entrance8. In turn, the arthropod marks itself internally by devouring the marked bait or externally by &#34;brushing&#34; up against the mark as it exits the nest.
As mentioned above, many types of markers have been used to tag a variety of arthropod species. However, very few are useful for all three of these dispersal research categories. The development of the protein immunomarking procedure was a major breakthrough for marking insects. Immunomarking puts a protein label on arthropods either internally or externally which, in turn, is detected by an anti-protein specific enzyme-linked immunosorbent assay (ELISA). The first such protein markers used were rabbit immunoglobulin (IgG) and chicken IgG/IgY9,10. They proved to be very effective marks for MRR and self-mark-capture research (see discussion). Unfortunately, IgG/IgY proteins are costly and are therefore not practical for mark-capture research and most types of self-mark-capture research. Subsequently, second-generation protein detection ELISAs were developed for proteins contained in chicken egg whites (albumin), cow&#39;s milk (casein) and soy milk (trypsin inhibitor protein). Each assay is highly sensitive, specific, and, most importantly, uses proteins that are much less expensive than the IgG/IgY proteins11. These proteins have proven effective for MRR, mark-capture, and self-mark-capture research (see discussion).
In this article, we describe and demonstrate how to conduct protein mark laboratory retention studies. Such studies are the first phase of research needed for any type of field dispersal study. Specifically, it is critical that investigators know how long the mark will be retained on the targeted arthropod species prior to embarking on field dispersal studies. Here, we describe and demonstrate how to internally and externally mark insects for MRR, mark-capture, and self-mark-capture type field studies. We then demonstrate how to detect the presence of the marks with indirect and sandwich ELISAs.

<table><tbody><tr><td>Food storage container (for insect rearing)</td><td>Rubbermaid/Tupperware</td><td>Various sizes</td><td>Various manufacturers &amp;amp; vendors. Mesh fabric is glued in a window</td></tr><tr><td>Small volume nebulizer (Micro Mist)</td><td>Teleflex-Hudson RCI</td><td>1881</td><td>Available online and in medical supply shops. Other brands will work as well.</td></tr><tr><td>Microcentrifuge tubes, 1.6 ml w/cap</td><td>Continental Lab Products</td><td>4445.X</td><td>Any similar type will work. We prefer brands that have easy to open lids.</td></tr><tr><td>Styrofoam storage box for microcentrifuge tubes (5x20 grid)</td><td>RPI Corp.</td><td>145746</td><td>This design is nice because it doesn&#039;t crowd the tubes.</td></tr><tr><td>Dispenser, repeater</td><td>Eppendorf</td><td>4982000322</td><td>Anything similar will help speed up the dispensing.</td></tr><tr><td>Pestles, plastic disposable tissue grinders, 1.5 ml</td><td>Kimble Chase</td><td>749521-1500</td><td>Available with various vendors. Pestles can be cleaned and autoclaved for reuse.</td></tr><tr><td>BD Falcon ELISA plates, 96-well (flat bottom)</td><td>BD</td><td>351172</td><td></td></tr><tr><td>Ovation pipette, manual (20-200 &amp;micro;l)</td><td>VistaLab</td><td>1057-0200 or 1070-0200</td><td>Any similar type will work. This model is ergonomic.</td></tr><tr><td>Reservoirs, sterile reagent</td><td>VistaLab</td><td>4054-1000</td><td>Anything similar will work.</td></tr><tr><td>Electronic pipette, 8-channel (50-1,200 &amp;micro;l) (Biohit eLine)</td><td>Sartorius</td><td>730391</td><td>If you had to pick one electronic model, this size is most useful.</td></tr><tr><td>Electronic pipette, 8-channel (10-300 &amp;micro;l) (Biohit eLine)</td><td>Sartorius</td><td>730361</td><td>Anything similar will work.</td></tr><tr><td>Manifold, multiwell plate washer (8-position, straight)</td><td>Sigma-Aldrich</td><td>Z369802&amp;nbsp;</td><td>Anything similar will work for manual plate washing.</td></tr><tr><td>Microplate reader with absorbance detection</td><td>Molecular Devices</td><td>SpectraMax 250</td><td>Anything similar will work.</td></tr><tr><td>Chicken IgG/IgY</td><td>United States Biological</td><td>I1903-15R</td><td>Also available from other sources</td></tr><tr><td>Egg whites</td><td>Grocery store</td><td>&amp;ldquo;All Whites&amp;rdquo;, &amp;ldquo;Egg Beaters&amp;rdquo;, &lt;em&gt;etc.&lt;/em&gt;</td><td>Mix 5 ml with 95 ml dH&lt;sub&gt;2&lt;/sub&gt;O for 5% solution</td></tr><tr><td>Tris (Trizma Base)</td><td>Sigma-Aldrich</td><td>T1503</td><td>2.42 g/L to make TBS</td></tr><tr><td>Sodium chloride (NaCl)</td><td>Sigma-Aldrich</td><td>S9888</td><td>29.22 g/L to make TBS and 8.0&amp;nbsp;g&amp;#8288;/&amp;#8288;L for PBS</td></tr><tr><td>Sodium phosphate dibasic (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>S0876</td><td>1.14 g/L for PBS</td></tr><tr><td>Potassium phosphate monobasic (KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Sigma-Aldrich</td><td>P5655</td><td>0.2 g/L for PBS</td></tr><tr><td>Potassium chloride (KCl)</td><td>Sigma-Aldrich</td><td>P9541</td><td>0.2 g/L for PBS</td></tr><tr><td>Tween 20, EIA grade</td><td>Sigma-Aldrich</td><td>P1379</td><td>Add 0.5 ml per L to PBS after salts are dissolved.</td></tr><tr><td>PBS with 1% BSA</td><td>Sigma-Aldrich</td><td>P3688</td><td>Mix 1 packet in 1 L dH&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Anti-Chicken IgY (IgG) (whole molecule) antibody produced in rabbit (1:500)</td><td>Sigma-Aldrich</td><td>C6409</td><td>Mix 100 &amp;micro;l in 50 ml TBS</td></tr><tr><td>Dry Milk, Powdered or Fresh Milk (1%)</td><td>Grocery store</td><td></td><td>Mix 10 g with 1 L dH&lt;sub&gt;2&lt;/sub&gt;O for 1% solution</td></tr><tr><td>Anti-Chicken IgY (IgG) (whole molecule)-Peroxidase antibody produced in rabbit (1:10,000)</td><td>Sigma-Aldrich</td><td>A9046</td><td>Mix 100 &amp;micro;l in 1 L of 1% milk</td></tr><tr><td>Anti-Chicken Egg Albumin antibody produced in rabbit (1:8,000)</td><td>Sigma-Aldrich</td><td>C6534</td><td>Dilute 125 &amp;micro;l in 1 L PBS-BSA</td></tr><tr><td>Goat Anti-Rabbit IgG Peroxidase Conjugate (1:2,000)</td><td>Sigma-Aldrich</td><td>A6154</td><td>Dilute 0.5 ml in 1 L PBS-BSA, then add 1.3&amp;nbsp;ml Silwet</td></tr><tr><td>Vac-In-Stuff (Silwet L-77) silicon-polyether copolymer</td><td>Lehle Seeds</td><td>VIS-01</td><td>Add as last ingredient</td></tr><tr><td>TMB Microwell One Component Peroxidase Substrate</td><td>SurModics</td><td>TMBW-1000-01</td><td></td></tr></tbody></table>

administering and detecting protein marks on arthropods for dispersal research

Monitoring arthropod movement is often required to better understand associated population dynamics, dispersal patterns, host plant preferences, and other ecological interactions. Arthropods are usually tracked in nature by tagging them with a unique mark and then re-collecting them over time and space to determine their dispersal capabilities. In addition to actual physical tags, such as colored dust or paint, various types of proteins have proven very effective for marking arthropods for ecological research. Proteins can be administered internally and/or externally. The proteins can then be detected on recaptured arthropods with a protein-specific enzyme-linked immunosorbent assay (ELISA). Here we describe protocols for externally and internally tagging arthropods with protein. Two simple experimental examples are demonstrated: (1) an internal protein mark introduced to an insect by providing a protein-enriched diet and (2) an external protein mark topically applied to an insect using a medical nebulizer. We then relate a step-by-step guide of the sandwich and indirect ELISA methods used to detect protein marks on the insects. In this demonstration, various aspects of the acquisition and detection of protein markers on arthropods for mark-release-recapture, mark-capture, and self-mark-capture types of research are discussed, along with the various ways that the immunomarking procedure has been adapted to suit a wide variety of research objectives.

Internal marking:
The results of the internal mark retention test are shown in Figure 2A. The calculated ELISA critical threshold value was 0.054. Overall (all four sample dates combined), the insects treated without protein yielded consistently low ELISA values (x&#772; = 0.038 &#177; 0.002, n = 80). Conversely, all of the insects fed the protein enriched diet yielded consistently stro...

Watch this Scientific Journal Video about Administering and Detecting Protein Marks on Arthropods for Dispersal Research at JoVE.com

Administering and Detecting Protein Marks on Arthropods for Dispersal Research

Proteins are often used to mark arthropods for dispersal research. Methods for administering internal and external protein marks on arthropods are demonstrated. Protein mark detection techniques, either through indirect or sandwich enzyme-linked immunosorbent assays (ELISAs), are also illustrated.

Monitoring arthropod movement is often required to better understand associated population dynamics, dispersal patterns, host plant preferences, and other ...

administering-detecting-protein-marks-on-arthropods-for-dispersal

Research

JoVE Journal

Environment

7.3K Views.  Arid-Land Agricultural Research Center. Proteins are often used to mark arthropods for dispersal research. Methods for administering internal and external protein marks on arthropods are demonstrated. Protein mark detection techniques, either through indirect or sandwich enzyme-linked immunosorbent assays (ELISAs), are also illustrated. 

Video: Administering and Detecting Protein Marks on Arthropods for Dispersal Research

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes. The majority of them belong to the superfamily of G protein-coupled receptors (GPCRs). We have focused on characterizing arthropod kinin receptors from the tick and mosquito. Arthropod kinins are multifunctional neuropeptides with myotropic, diuretic, and neurotransmitter function. Here, a method for systematic analyses of structure-activity relationships of insect kinins on two heterologous kinin receptor-expressing systems is described. We provide important information relevant to the development of biostable kinin analogs with the potential to disrupt the diuretic, myotropic, and/or digestive processes in ticks and mosquitoes.
The kinin receptors from the southern cattle tick, Boophilus microplus (Canestrini), and the mosquito Aedes aegypti (Linnaeus), were stably expressed in the mammalian cell line CHO-K1. Functional analyses of these receptors were completed using a calcium bioluminescence plate assay that measures intracellular bioluminescence to determine cytoplasmic calcium levels upon peptide application to these recombinant cells. This method takes advantage of the aequorin protein, a photoprotein isolated from luminescent jellyfish. We transiently transfected the aequorin plasmid (mtAEQ/pcDNA1) in cell lines that stably expressed the kinin receptors. These cells were then treated with the cofactor coelenterazine, which complexes with intracellular aequorin. This bond breaks in the presence of calcium, emitting luminescence levels indicative of the calcium concentration. As the kinin receptor signals through the release of intracellular calcium, the intensity of the signal is related to the potency of the peptide.
This protocol is a synthesis of several previously described protocols with modifications; it presents step-by-step instructions for the stable expression of GPCRs in a mammalian cell line through functional plate assays (Staubly et al., 2002 and Stables et al., 1997). Using this methodology, we were able to establish stable cell lines expressing the mosquito and the tick kinin receptors, compare the potency of three mosquito kinins, identify critical amino acid positions for the ligand-receptor interaction, and perform semi-throughput screening of a peptide library. Because insect kinins are susceptible to fast enzymatic degradation by endogenous peptidases, they are severely limited in use as tools for pest control or endocrinological studies. Therefore, we also tested kinin analogs containing amino isobutyric acid (Aib) to enhance their potency and biostability. This peptidase-resistant analog represents an important lead in the development of biostable insect kinin analogs and may aid in the development of neuropeptide-based arthropod control strategies.

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito Aedes aegypti and Tick Rhipicephalus microplus G Protein-coupled Receptors

This protocol provides instructions for clonal-cell line selection and a calcium bioluminescence assay to analyze the structure-activity relationships of synthesized arthropod neuropeptides on their cognate GPCRs. This assay can be used for receptor deorphanization and structure-activity relationship studies for synthetic analog design and peptide/drug-lead discovery.

Arthropod hormone receptors are potential targets for novel pesticides as they regulate many essential physiological and behavioral processes.  The ...

Immunology and Infection

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells named wing imaginal discs formed during embryonic development. In the larval stages the imaginal discs grow, increasing its number of cells and forming monolayered epithelial&#160;structures. Inside the pupal case, the imaginal discs bud out and fold into bilayers along a line that becomes the future margin of the wing. During this process, the longitudinal primodia veins originate vein cells on the prospective dorsal and ventral surfaces of the wing. During the pupal stage the stripes of vein cells of each surface communicate in order to generate tight tubes; at the same time, the cross-veins begin their formation.
With the help of appropriate molecular markers, it is possible to identify the major elements composing the wing during its development. For this reason, the ability to accurately detect transcripts or proteins in this structure is critical for studying their abundance and localization related to the development process of the wing.
The procedure described here focuses on manipulating pupal wings, providing detailed instructions on how to dissect the wing during the pupal stage. The dissection of pupal tissue is more difficult to perform than their counterparts in third instar larvae. This is why this approach was developed, to obtain rapid and efficient high quality samples. Details of how to immunostain and mount these wing samples, to allow the visualization of proteins or cell components, are provided in the protocol. With little expertise it is possible to collect 8-10 high quality pupal wings in a short amount of time.

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

The ability to accurately detect transcripts or proteins in Drosophila tissues is critical for studying their abundance and localization related to the development process. Here is the description of a straightforward procedure to dissect pupal wings. These wings can be used as samples in several techniques (immunohistochemistry, PCR assay, etc.).

Wing development in Drosophila melanogaster is an ideal model for studying morphogenesis at tissue level. These appendages develop from a group of cells ...

Developmental Biology

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Drosophila flight muscle is a powerful model to study diverse processes such as transcriptional regulation, alternative splicing, metabolism, and mechanobiology, which all influence muscle development and myofibrillogenesis. Omics data, such as those generated by mass spectrometry or deep sequencing, can provide important mechanistic insights into these biological processes. For such approaches, it is beneficial to analyze tissue-specific samples to increase both selectivity and specificity of the omics fingerprints. Here we present a protocol for dissection of fluorescent-labeled flight muscle from live pupae to generate highly enriched muscle samples for omics applications. We first describe how to dissect flight muscles at early pupal stages (&#60;48 h after puparium formation [APF]), when the muscles are discernable by green fluorescent protein (GFP) labeling. We then describe how to dissect muscles from late pupae (&#62;48 h APF) or adults, when muscles are distinguishable under a dissecting microscope. The accompanying video protocol will make these technically demanding dissections more widely accessible to the muscle and Drosophila research communities. For RNA applications, we assay the quantity and quality of RNA that can be isolated at different time points and with different approaches. We further show that Bruno1 (Bru1) is necessary for a temporal shift in myosin heavy chain (Mhc) splicing, demonstrating that dissected muscles can be used for mRNA-Seq, mass spectrometry, and reverse transcription polymerase chain reaction (RT-PCR) applications. This dissection protocol will help promote tissue-specific omics analyses and can be generally applied to study multiple biological aspects of myogenesis.

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

Drosophila flight muscle is a powerful model to study transcriptional regulation, alternative splicing, metabolism, and mechanobiology. We present a protocol for dissection of fluorescent-labeled flight muscle from live pupae to generate highly enriched samples ideal for proteomics and deep-sequencing. These samples can offer important mechanistic insights into diverse aspects of muscle development.

Drosophila flight muscle is a powerful model to study diverse processes such as transcriptional regulation, alternative splicing, metabolism, and ...

Genetics

A Method for Quantifying Foliage-Dwelling Arthropods

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimates of density requires a method with high detection probability and a known sampling area. While most described methods provide a qualitative or semi-quantitative estimate adequate for describing species presence, richness, and diversity, few provide an adequately consistent detection probability and known or consistent sampling areas to provide an index or estimate with adequate precision to detect differences in abundance across environmental, spatial, or temporal variables. We describe how to quantify leaf-dwelling arthropods by sealing the leaves and end of branches in a bag, clipping and freezing the bagged material, and rinsing the previously frozen material in water to separate arthropods from the substrate and quantify them. As we demonstrate, this method can be used at a landscape scale to quantify leaf-dwelling arthropods with adequate precision to test for and describe how spatial, temporal, environmental, and ecological variables influence arthropod richness and abundance. This method allowed us to detect differences in density, richness, and diversity of leaf-dwelling arthropods among 5 genera of trees commonly found in southeastern deciduous forests.

We describe how to quantify leaf dwelling arthropods by sealing the leaves and end of branches in a bag, clipping and freezing the bagged material, and rinsing the previously frozen material in water to separate arthropods from the substrate for quantification.

Terrestrial arthropods play an important role in our environment. Quantifying arthropods in a way that allows for a precise index or estimates of density ...

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila express gustatory receptors1,2, ion channels3-6, and ionotropic receptors7 that are involved in the detection of volatile and non-volatile sensory cues. These neurons directly contact tastants such as food, noxious substances and pheromones and therefore influence many complex behaviors such as feeding, egg-laying and mating. Electrode recordings and calcium imaging have been widely used in insects to quantify the neuronal responses evoked by these tastants. However, electrophysiology requires specialized equipment and obtaining measurements from a single taste sensillum can be technically challenging depending on the cell-type, size, and position. In addition, single neuron resolution in Drosophila can be difficult to achieve since taste sensilla house more than one type of chemosensory neuron. The live calcium imaging method described here allows responses of single gustatory neurons in live flies to be measured. This method is especially suitable for imaging neuronal responses to lipid pheromones and other ligand types that have low solubility in water-based solvents.

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

The forelegs and proboscis of Drosophila contain a rich repertoire of gustatory sensory neurons. Here, we present a method using calcium imaging to measure physiological responses from sensory neurons in the foreleg and proboscis of live flies upon exogenous application of a gustatory pheromone.

Unlike mammals, insects such as Drosophila have multiple taste organs. The chemosensory neurons on the legs, proboscis, wings and ovipositor of Drosophila ...

SwarmSight: Real-time Tracking of Insect Antenna Movements and Proboscis Extension Reflex Using a Common Preparation and Conventional Hardware

Many scientifically and agriculturally important insects use antennae to detect the presence of volatile chemical compounds and extend their proboscis during feeding. The ability to rapidly obtain high-resolution measurements of natural antenna and proboscis movements and assess how they change in response to chemical, developmental, and genetic manipulations can aid the understanding of insect behavior. By extending our previous work on assessing aggregate insect swarm or animal group movements from natural and laboratory videos using the video analysis software SwarmSight, we developed a novel, free, and open-source software module, SwarmSight Appendage Tracking (SwarmSight.org) for frame-by-frame tracking of insect antenna and proboscis positions from conventional web camera videos using conventional computers. The software processes frames about 120 times faster than humans, performs at better than human accuracy, and, using 30 frames per second (fps) videos, can capture antennal dynamics up to 15 Hz. The software was used to track the antennal response of honey bees to two odors and found significant mean antennal retractions away from the odor source about 1 s after odor presentation. We observed antenna position density heat map cluster formation and cluster and mean angle dependence on odor concentration.

This protocol describes steps for using the novel software, SwarmSight, for frame-by-frame tracking of insect antenna and proboscis positions from conventional web camera videos using conventional computers. The free, open-source software processes frames about 120 times faster than humans and performs at better than human accuracy.

Many scientifically and agriculturally important insects use antennae to detect the presence of volatile chemical compounds and extend their proboscis ...

Neuroscience

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host plant.1,2 When a host plant is an important agronomic commodity feeding damage by insect pests can inflict serious economic losses to growers. Accordingly, kairomones can be used as attractants to lure or confuse these insects and, thus, offer an environmentally friendly alternative to pesticides for insect control.3 Unfortunately, plants can emit a vast number volatiles with varying compositions and ratios of emissions dependent upon the phenology of the commodity or the time of day. This makes identification of biologically active components or blends of volatile components an arduous process. To help identify the bioactive components of host plant volatile emissions we employ the laboratory-based screening bioassay electroantennography (EAG). EAG is an effective tool to evaluate and record electrophysiologically the olfactory responses of an insect via their antennal receptors. The EAG screening process can help reduce the number of volatiles tested to identify promising bioactive components. However, EAG bioassays only provide information about activation of receptors. It does not provide information about the type of insect behavior the compound elicits; which could be as an attractant, repellent or other type of behavioral response. Volatiles eliciting a significant response by EAG, relative to an appropriate positive control, are typically taken on to further testing of behavioral responses of the insect pest. The experimental design presented will detail the methodology employed to screen almond-based host plant volatiles4,5 by measurement of the electrophysiological antennal responses of an adult insect pest navel orangeworm (Amyelois transitella) to single components and simple blends of components via EAG bioassay. The method utilizes two excised antennae placed across a "fork" electrode holder. The protocol demonstrated here presents a rapid, high-throughput standardized method for screening volatiles. Each volatile is at a set, constant amount as to standardize the stimulus level and thus allow antennal responses to be indicative of the relative chemoreceptivity. The negative control helps eliminate the electrophysiological response to both residual solvent and mechanical force of the puff. The positive control (in this instance acetophenone) is a single compound that has elicited a consistent response from male and female navel orangeworm (NOW) moth. An additional semiochemical standard that provides consistent response and is used for bioassay studies with the male NOW moth is (Z,Z)-11,13-hexdecadienal, an aldehyde component from the female-produced sex pheromone.6-8

A method to rapidly screen host plant volatiles by measurement of the electrophysiological response of adult navel orangeworm (Amyelois transitella) antennae to single components and blends via electroantennographic analysis is demonstrated.

Plant volatiles play an important role in plant-insect interactions. Herbivorous insects use plant volatiles, known as kairomones, to locate their host ...

Biology

Monitoring On-Target Signaling Responses in Larval Zebrafish - Z-REX Unmasks Precise Mechanisms of Electrophilic Drugs and Metabolites

Reactive metabolites and related electrophilic drugs are among the most challenging small molecules to study. Conventional approaches to deconstruct the mode of action (MOA) of such molecules leverage bulk treatment of experimental specimens with an excess of a specific reactive species. In this approach, the high reactivity of electrophiles renders non-discriminate labeling of the proteome in a time- and context-dependent manner; redox-sensitive proteins and processes can also be indirectly and often irreversibly affected. Against such a backdrop of innumerable potential targets and indirect secondary effects, linking phenotype to specific target engagement remains a complex task. Zebrafish targeting reactive electrophiles and oxidants (Z-REX)-an on-demand reactive-electrophile delivery platform adapted for use in larval zebrafish-is designed to deliver electrophiles to a specific protein of interest (POI) in otherwise unperturbed live fish embryos. Key features of this technique include a low level of invasiveness, along with dosage-, chemotype-, and spatiotemporally-controlled precision electrophile delivery. Thus, in conjunction with a unique suite of controls, this technique sidesteps off-target effects and systemic toxicity, otherwise observed following uncontrolled bulk exposure of animals to reactive electrophiles and pleiotropic electrophilic drugs. Leveraging Z-REX, researchers can establish a foothold in the understanding of how individual stress responses and signaling outputs are altered as a result of specific reactive ligand engagement with a specific POI, under near-physiologic conditions in intact living animals.

Zebrafish targeting reactive electrophiles and oxidants (Z-REX) is a chemical biology-based method for the investigation of reactive small-molecule signaling. This technique can be applied to live fish of different developmental stages. Here, we couple standard assays in zebrafish with Z-REX for signaling pathway analysis.

Reactive metabolites and related electrophilic drugs are among the most challenging small molecules to study. Conventional approaches to deconstruct the ...

High-Resolution Imaging and Analysis of Drosophila Eye Defects

A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It is a simple structure of repeating unit eyes, termed ommatidia, that is used to characterize genetic interactions and gene functions. Mutations that affect eye architecture can be easily detected and analyzed; hence, this system is frequently used in under-resourced institutions. Further phenotypic analysis often includes a Scanning Electron Microscope (SEM) to generate high-magnification images suitable for quantitative analysis. However, SEMs are expensive and require costly reagents; sample preparation spans days; and, often, they need full-time staff for sample preparation and instrument maintenance. This limits their utility at under-resourced institutions or during budgetary austerity. In entomology, the use of high-resolution digital imaging technology is a common practice for the identification and characterization of species. This paper describes a method that combines strategies and allows for high-resolution digital imaging of adult Drosophila structures and quantitative analysis using the open software ImageJ. The workflow is a rapid and student-friendly alternative that remedies the limitations of underfunded and under-resourced research facilities with a cost-effective and rapid approach to quantitative phenotypic analysis.

Here, we present a rapid, low-cost, workflow for high-resolution imaging of adult Drosophila eyes to quantify patterning and growth defects. We describe our protocol for sample preparation by point-mounting, high-resolution image acquisition, and image analysis.

The Drosophila compound eye is a precisely patterned tissue that has revealed molecular mechanisms and biological processes that drive morphogenesis. It ...

Non-invasive Electropenetrography to Study Mosquito Probing Behaviors

AC-DC Electropenetrography for the Study of Probing and Ingestion Behaviors of Culex tarsalis Mosquitoes on Human Hands

Mosquitoes transmit pathogens that negatively affect human and animal health. A greater understanding of their blood-feeding biology and interactions with hosts and pathogens could be exploited to develop new targets for controlling mosquito-borne diseases. Unfortunately, probing (i.e., biting) behaviors of mosquitoes are poorly understood because they occur inside host tissues. Here, a non-invasive procedure is described for using AC-DC electropenetrography (EPG) to indirectly visualize and quantify mosquito feeding behaviors by recording changes in electrical signals generated when probing and ingesting on human hands. Thin gold wires are attached to the mosquitoes using conductive silver glue and connected to the EPG instrument. The human host holds a substrate voltage probe in their hand. Probing of the host by the wired mosquito completes the electrical circuit, and electrical signals are recorded on a computer as &#34;waveforms&#34; that can then be measured and enumerated for analysis. This methodology has been used to characterize the probing and ingestion behaviors of Aedes aegypti and Culex tarsalis mosquitoes and can be applied to other mosquito species. EPG can be used to study the effects of pathogens, insecticides, and other factors on mosquito feeding behaviors.

Procedures are described for studying the probing and ingestion behaviors of Culex tarsalis&#160;mosquitoes on human hands using AC-DC electropenetrography.

Mosquitoes transmit pathogens that negatively affect human and animal health. A greater understanding of their blood-feeding biology and interactions with ...

Administering and Detecting Protein Marks on Arthropods for Dispersal Research

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Chapters in this video

A Calcium Bioluminescence Assay for Functional Analysis of Mosquito (Aedes aegypti) and Tick (Rhipicephalus microplus) G Protein-coupled Receptors

A Rapid and Efficient Method to Dissect Pupal Wings of Drosophila Suitable for Immunodetections or PCR Assays

Dissection of Drosophila melanogaster Flight Muscles for Omics Approaches

A Method for Quantifying Foliage-Dwelling Arthropods

Measuring Physiological Responses of Drosophila Sensory Neurons to Lipid Pheromones Using Live Calcium Imaging

SwarmSight: Real-time Tracking of Insect Antenna Movements and Proboscis Extension Reflex Using a Common Preparation and Conventional Hardware

Electroantennographic Bioassay as a Screening Tool for Host Plant Volatiles

Monitoring On-Target Signaling Responses in Larval Zebrafish - Z-REX Unmasks Precise Mechanisms of Electrophilic Drugs and Metabolites

A Rapid, Simple Workflow for Quantification of External Adult Drosophila Structures

AC-DC Electropenetrography for the Study of Probing and Ingestion Behaviors of Culex tarsalis Mosquitoes on Human Hands