Funding for this project was made possible through University of Wisconsin-Madison internal grants. Sincerest thanks to Kevin Bachhuber of Bachhuber Consulting Inc. for access to his unpublished guide for commercial cricket rearing and to Michael Bartlett Smith for his assistance in refining and troubleshooting these methods.

1. Preparing the oviposition substrate
NOTE: Coconut coir is an ideal oviposition substrate for G. bimaculatus. For detailed methods on how to separate coir from compressed coir brick and a note on respiratory safety, see Supplemental Materials step 1.1.
<ol>
	<li>Wash hands with soap and water.</li>
	<li>Tare a clean container on a balance and weigh a mass of dry coconut coir approximately the size of a human fist.</li>
	<li>Place coir into a sealable, clean container, which can accommodate expansion up to 6x the original volume.</li>
	<li>With clean hands, gently break up clumps of coir from the piece removed from the larger block.</li>
	<li>Using a 50 mL graduated cylinder, measure the correct volume of deionized (DI) water to achieve a 5:1 ratio by mass of five parts water to one part dry coir.</li>
	<li>Add the measured DI water slowly, evenly hydrating all coir particles. Manually macerate clumps to ensure even hydration.</li>
	<li>Re-tare the container in which the coir was previously weighed.</li>
	<li>Weigh out 75 g of wetted coir.</li>
	<li>Transfer 75 g of the wetted coir into a 100 mm x 15 mm Petri dish using a clean plastic spoon to ensure that the coir is evenly spread around the bottom of the dish and that there are no clumps.</li>
	<li>Label the side of the Petri dish with lab tape with a label denoting natal colony and date of the egg collection event.</li>
	<li>Measure an additional 45 mL of DI water in a graduated cylinder.</li>
	<li>Add water evenly over the surface of the packed coir in the Petri dish to ensure even hydration. Ensure that the coir is saturated to the point that water pools approximately &#188; the way up the sides of the container.</li>
	<li>Once the Petri dish has been packed, seal the remaining wetted coir in an airtight vessel for storage at -20 &#176;C. 
	NOTE: Following the methods prescribed by this article, G. bimaculatus individuals will reach sexual maturity after an average of 58 days post oviposition.</li>
	<li>Collecting eggs
	<ol>
		<li>Place the hydrated oviposition substrate into cages containing the desired parental stocks of crickets as far from the feed as possible due to the potential for crickets to mechanically transfer granules of feed onto the oviposition substrate.</li>
		<li>Document the date and time. 
		NOTE: The standardized working density for reproductive cricket colonies reared following these methods is n = 150 adult individuals. At that density, a 24 h oviposition window will yield between 800 and 1,500 eggs depending on the colony age, prior oviposition effort, and parent cage sex ratio.</li>
		<li>Place a small autoclavable trash receptacle on the work surface to account for the containment risk posed by handling and cleaning egg-rich oviposition substrates.</li>
		<li>Place a clean, empty 29.3 L plastic cage on the bench next to the trash receptacle to serve as the recipient cage for the egg-rich oviposition substrate.</li>
		<li>Place the 29.3 L cage containing the parent cricket stocks and oviposition substrate on the opposite side of the trash receptacle from the empty cage.</li>
		<li>After 24 h, remove the oviposition substrate from the parent cricket cage and locate it over autoclavable waste vessel.</li>
		<li>Inspect the top of the oviposition substrate for any particles of frass or feed that the crickets may have kicked onto the surface of the coir. 
		NOTE: Any substance, which is not egg mass or coir, can cause mold to form on the substrate during incubation.</li>
		<li>Remove coir contaminants into the waste vessel with a clean scoopula or plastic spoon.</li>
		<li>Place the plastic spoon into the waste receptacle.</li>
		<li>Place the cleaned oviposition substrate in the clean 29.3L cage.</li>
		<li>Place the cage in an incubator set to 27 &#176;C at 60% relative humidity on a 12 h D/12 h L photoperiod.</li>
		<li>Return the cage containing the breeding stock to the original location and clear all items from the work surface.</li>
		<li>Place the waste receptacle into an in-facility freezer dedicated to the storage of items potentially contaminated with cricket eggs.</li>
		<li>Sanitize the work surface with 10% bleach solution and let it sit for 60 s.</li>
		<li>Wipe the work surface dry with a clean paper towel. Open the freezer and dispose of the paper towel in the waste receptacle.</li>
	</ol>
	</li>
	<li>Misting and monitoring egg substrates daily 
	NOTE: For methods used to calibrate the volume of mist delivered by a spray bottle, see Supplemental Materials step 1.2.
	<ol>
		<li>Position a spray bottle over the oviposition substrate so that the water expressed is evenly distributed over the surface of the substrate.</li>
		<li>Perform the number of pump actuations calculated in the Supplemental Materials step 1.2 for each oviposition substrate daily for 11 consecutive days.</li>
		<li>Check each oviposition substrate daily, monitoring for filamentous mold growth on the surface of the coir.</li>
		<li>If fungal growth is observed, use a clean spoon or scoopula to remove spots of surface mold.</li>
		<li>Dispose of the tool and the removed coir in the autoclavable waste container stored in the facility freezer. 
		NOTE: It is unclear whether the mold adversely impacts cricket development.</li>
		<li>At day 11 post oviposition, begin looking closely at the substrate for juvenile crickets. 
		NOTE: At 27 &#176;C, the eggs of G. bimaculatus require 11-13 days to hatch.</li>
	</ol>
	</li>
	<li>Setting up natal cages
	<ol>
		<li>Select two unused 30.8 cm x 30.8 cm (12 inches x 12 inches) commercial egg carton flats. With a utility knife or strong shears, cut these into six separate 10.1 cm (4&#34;)-wide strips of equal size. Brush off the cut edges with hands to remove dangling particles of cardboard.</li>
		<li>Place the six individual 10.1 cm x 30.8 cm (4 inches x 12 inches) pieces of carton vertically into the bottom of the cage with the longer axis of the carton spanning the narrower horizontal axis of a 29.3L cage. Place a seventh piece of carton flat across the top of the six upright pieces.</li>
		<li>Select three pieces of rough brown paper towel approximately 25 cm x 25cm. Fold each in half. Place two such that they cover the top of the proximal side of the carton structure. Place one over the carton stack on the distal side.</li>
		<li>At day 11 post oviposition, move the oviposition substrate into the proximal righthand corner of the cage.</li>
	</ol>
	</li>
	<li>Caring for early instars 
	NOTE: On day 14 post oviposition, most viable eggs will have hatched and early-stage cricket nymphs will require feed and water. Young crickets are unable to break the surface tension of water droplets and can drown if water is pooled in their environment. However, they are also sensitive to desiccation. Providing a consistent relative humidity of around 60% during this stage of development is important to ensuring survival.
	<ol>
		<li>When a hatch is observed, mist the paper towels placed over the top of the cartons in step 1.4.3 until they are wetted but not actively shedding water.</li>
		<li>Thoroughly wipe both sides of a 100 mm Petri dish lid with 70% ethanol and allow it to dry. Use it as the receptacle in which cricket feed will be delivered. 
		NOTE: First-instar crickets require smaller feed particles than crickets at subsequent stages of development. This finer feed should be administered to crickets for the first 20 days post emergence.</li>
		<li>Scoop 50 g of the feed into a 60-watt single-serving blender and mill at 10,000 rpm for 1 min.</li>
		<li>Measure 1 g of the feed and shake it onto the Petri dish lid in the cage. Using the clean end of a spoon or scoopula, spread the feed out as uniformly as possible over the bottom of the dish.</li>
		<li>Replace the feed every 2 days. Use the end of the spoon to brush crickets from the feed dish before removing it. Discard the old feed in the autoclavable waste container.</li>
		<li>Monitor for mold growth on feed. If the feed begins to appear white or greenish, discard the Petri dish and feed immediately.</li>
		<li>At 14 days post oviposition, use a 2.54 cm (1 inch) paintbrush to remove crickets clinging to the natal coir substrate by brushing all crickets from the coir surface and sides of the Petri dish into the cage.</li>
		<li>Place the removed oviposition substrate into the autoclavable waste receptacle and store it in the freezer until autoclaving.</li>
		<li>Replace the natal substrate with a fresh coir dish for hydration prepared following steps 1.1.5-1.1.9.</li>
		<li>Use a DI water wash bottle to add water until the surface of the coir glistens but is not pooling. 
		&#8203;NOTE: Cricket density strongly impacts growth performance in G. bimaculatus13. Maintaining the breeding stock at excessive density risks introducing unwanted crowding-induced epigenetic effects into experiments in which their progeny is used9. Crickets must be &#34;thinned&#34; from the high densities that emerge from the oviposition substrates and distributed into densities that adhere to the standard of 0.005 crickets/cm3 of space.</li>
	</ol>
	</li>
</ol>
2. Caring for instars three to adult
<ol>
	<li>Setting up cages 
	NOTE: For details on the technique to construct the screened lid, see Supplemental Materials step 1.3.1.
	<ol>
		<li>Repeat step 1.4.1.</li>
		<li>Install five cut egg carton pieces in the distal end of the cage so the egg-shape concavities are facing outward. Ensure that the short ends are about the sides of the cage, the long side sits flat against the bottom, and there is approximately 3 cm of space between each piece of carton.</li>
		<li>Place the last cut carton piece atop the upright carton pieces such as the roof on a house as shown in Supplemental Figure S1.</li>
		<li>Follow steps 1.1.8-1.1.13 to prepare the hydration substrates.</li>
		<li>Place the hydration substrate in the proximal right-hand corner of the cricket cage as shown in Supplemental Figure S2.</li>
		<li>Use a wash bottle to add 6-10 mL of DI water until the coir surface appears wet and reflective, but the coir is not fully submerged. 
		NOTE: The surface should appear lightly dimpled, with surface tension causing the water to follow the contours of the coir.</li>
		<li>Invert the lid of a 100 mm Petri dish and place it on the proximal left-hand side of the cage. Add 2-3 g of standard cricket feed as shown in Supplemental Figure S2.</li>
	</ol>
	</li>
	<li>Modifying colony density 
	NOTE: Conduct this step at 20 days post hatching or when the crickets reach an average mass of 0.01 g.
	<ol>
		<li>On the work surface, place a large container that can accommodate the floor plan of three 29.3 L cages standing side-by-side. 
		NOTE: This is secondary containment and will control crickets that escape during their transfer from one colony to another.</li>
		<li>Remove the original colony from the rearing rack and place it on the work surface.</li>
		<li>To the right side of the original colony, place an empty cage of the same size.</li>
		<li>To the right side of the empty cage, place a cage that has been set up according to steps 2.1.1-2.1.7.</li>
		<li>Check to make sure that crickets are not clinging to the underside of the screened lid of the cage containing the crickets. If observed, tap the top of the cage to dislodge them.</li>
		<li>Open the screened lid of the cage containing the crickets.</li>
		<li>In one gentle, smooth motion, transfer the &#34;roof&#34; carton and all the crickets adhering to its contours into the middle cage.</li>
		<li>Once the carton is inside the middle cage, gently agitate the cardboard against the sides to dislodge all the crickets.</li>
		<li>Visually inspect that all crickets have been shaken free before returning the carton piece to the proximal side of the cage of origin to allow the remaining crickets to adhere to the carton.</li>
		<li>Repeat steps 2.2.8-2.2.9 with all the carton pieces in the cage, working sequentially from the front to the back of the original cage until all the crickets have been transferred into the middle cage.</li>
		<li>Gently tilt the middle cage containing the crickets so that all the crickets contained are directed toward the bottom corner.</li>
		<li>Lift the cage containing the crickets over the recipient cage.</li>
		<li>Slowly tilt the donor cage so that the crickets begin to gain purchase on the sides and can move in a controlled manner out of the mass in the bottom corner as shown in Supplemental Figure S3.</li>
		<li>If crickets advance too quickly, adjust the angle at which the middle cage is held, causing the crickets to fall back.</li>
		<li>While the cage is tilted, use a 2.54 cm (1 inch) paintbrush to direct the crickets into the recipient cage, counting each until the total count is equal 150 individuals. Use the brush to deter those advancing too swiftly for an accurate count.</li>
		<li>Label the newly stocked cricket cage with date, parental stock, and the number of crickets contained.</li>
		<li>Humanely euthanize excess crickets still in the bottom of the middle and original containers by placing the entire container into a freezer at -20 &#176;C for a minimum of 30 min.</li>
		<li>Inspect the cage of origin to ensure that all crickets have been transferred.</li>
		<li>Position the cricket cages on plant-growing racks 25 cm below light hoods containing full-spectrum fluorescent lights set to a residential-grade outdoor timer programmed to maintain a 12 h L/D photoperiod. See Supplemental Figure S4.</li>
		<li>Transfer all frass, dishes of spent hydration substrate and feed, paper towel, exuviae, and dead crickets remaining in the cage of origin into the autoclavable waste receptacle.</li>
		<li>Unless the waste is to be autoclaved immediately, store it in an in-facility freezer at -20 &#176;C.</li>
		<li>Inspect the floor, worker apparel, secondary containment cage, and work surface for escaped crickets.</li>
		<li>Sanitize the work surface and secondary containment cage with 10% bleach solution, discarding the paper towels into the autoclavable waste receptacle.</li>
	</ol>
	</li>
	<li>Feeding and watering
	<ol>
		<li>Open the airtight feed storage container and fill an empty 100 mL sample cup with cricket feed. Reach into each colony and deposit a quarter of the feed held in the cup into the Petri dish lid holding the feed.</li>
		<li>To water crickets, prepare a Petri dish of coir following steps 1.1.9-1.1.13.</li>
		<li>Increase the feed rate commensurate to the consumption rate to ensure ad libitum feed availability. 
		NOTE: Cricket feed demand changes throughout development.</li>
	</ol>
	</li>
	<li>Transferring crickets to clean cages
	<ol>
		<li>Transfer the crickets to clean cages every 2 weeks. Replicate the arrangement of the cages from steps 2.2.1-2.2.4 
		NOTE: Significant amounts of frass will have accumulated in the concavities of the egg cartons.</li>
		<li>Transfer crickets to clean cages, following steps 2.2.5-2.2.23.</li>
		<li>Gently maneuver the cartons so that the majority of frass falls into the cage of origin while allowing crickets to cling to the carton during repetition of steps 2.2.7-2.2.10.</li>
		<li>Use a brush or plastic spoon to encourage any crickets trapped in the frass to move from the middle cage into a clean cage.</li>
		<li>Inspect the cage of origin to ensure that all crickets have been transferred.</li>
		<li>Transfer all frass, dishes of spent hydration substrate and feed, paper towel, exuviae, and dead crickets remaining in the cage of origin into the autoclavable waste receptacle.</li>
		<li>Unless the waste is to be autoclaved immediately, store it in an in-facility freezer at -20 &#176;C.</li>
		<li>Inspect the floor, worker apparel, secondary containment cage, and work surface for escaped crickets.</li>
		<li>Sanitize the work surface and secondary containment cage with 10% bleach solution, discarding paper towels in autoclavable waste receptacle.</li>
	</ol>
	</li>
	<li>Setting up experimental cages 
	NOTE: Experimental cages are plastic containers that house fewer crickets. Their setup is identical to the 29.3 L cages that contain breeding stocks but rely on 7.1 L containers stocked with smaller dishes of water, feed, and contain a reduced egg carton surface area.
	<ol>
		<li>Place six 10.1 cm x 15.4 cm (4 inch x 6 inch) carton pieces into the distal end of each experimental cage with the long axes of the cartons spanning the width of the narrow dimension of the cage and the short axes of the cartons oriented toward the lid and the floor.</li>
		<li>Pack 10 g of the hydrated coir working mix into a Petri dish.</li>
		<li>Use a wash bottle containing DI water to add approximately 15 mL of DI water, or until a meniscus forms at the surface of the coir.</li>
		<li>Invert the lid of a 60 mm x 15 mm Petri dish to hold the feed. 
		NOTE: The feeding rate may vary throughout the duration of the trial. For the experimental cage cricket randomization and stocking procedure, see Supplemental Materials step 1.4.</li>
	</ol>
	</li>
	<li>Terminating insects
	<ol>
		<li>When the crickets are no longer needed for breeding or experimental use, follow step 2.2.17.</li>
		<li>When the crickets are dead, remove the cage from the freezer. Remove the lid and transfer all the contained materials into an autoclavable waste receptacle. Transfer the waste back into the freezer until autoclaving.</li>
		<li>Immerse the empty cage into 10% bleach solution and allow it to sit for a minimum of 5 min.</li>
		<li>Triple-rinse the empty cage with cold tap water to remove the bleach residue, with particular attention to channels at the bottom of the container.</li>
	</ol>
	</li>
</ol>

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The authors have no conflicts of interest to declare.

The simplicity of this approach to cricket rearing can benefit a range of research areas and represents a generic template for successful cricket husbandry, easily adaptable to a variety of experimental needs. Compared to several other studies of G. bimaculatus, the individual body adult size is smaller and maturation is slower14, which we attribute to sub-optimal rearing temperature imposed on us by circumstance. The methods described above have been used and refined over the course of 2...

As typified by the iconic insect, the fruit fly Drosophila melanogaster, the use of insects as laboratory model organisms provides distinct advantages for studies in genetics, toxicology, and physiology1. The small size of insects reduces the space needed for cultures and the amount of feed and consumable materials required. Many insects&#160;reproduce quickly making them uniquely suited to the creation of specialized genetic lines and studies requiring the evaluation of multiple successive generations.
Many studies focus on holometabolous insects such as Drosophila, which exhibit complete metamorphosis and pupation. However, other models are available, including Gryllus bimaculatus (De Geer),&#160;the two-spotted field cricket. G. bimaculatus is a paurometabolous insect that undergoes between 7 and 11 nymphal instars before reaching sexual maturity2. This cricket displays a wide range of behaviors related to sexual selection, including stridulation, territorial displays, and mate-guarding3. Immature crickets are unlike the larvae of holometabolous insect species in that they, similar to many paurometabolous juveniles, are able to regenerate lost and damaged limbs during ecdysis4. Additionally, the fully sequenced genome of G. bimaculatus was published in 20215. These characteristics make these crickets appealing as a target for basic research.
Two-spotted field crickets are widely reared for human food and animal feed. The scale of these operations is often much larger than for laboratory research6,7. Despite the difference in scale, the challenges faced by researchers overlap greatly with those encountered by commercial cricket farmers. These considerations converge in the context of lab-based research aiming to improve edible insect production. As the edible insect industry continues to evolve and grow, optimizing feed inputs and myriad other aspects of production is a primary goal8. Laboratory studies demonstrating measured improvements in rearing efficiency, survivorship, or generation time in these crickets have the potential to help increase the profitability of cricket farming operations long-term.
Standardized rearing protocols enable closer comparison between studies investigating rearing optimization. To-date, few in-depth protocols for rearing G. bimaculatus in the laboratory have been published. An ideal protocol would reflect conditions encountered in real-world cricket farming operations, while maintaining the strictly controlled conditions necessary to accurately measure changes in growth performance arising from experimental treatments and highlighting risk mitigation strategies. The methods described in this paper were developed based on published protocols, techniques, and apparatus used to rear a variety of cricket species at a broad range of laboratory and commercial production scales2,9,10,11,12. These methods are also informed by several non-peer reviewed sources, including unpublished technical bulletins and personal communication with commercial cricket farmers in North America. This protocol was developed with the intention of facilitating the establishment of laboratory cultures of G. bimaculatus specifically for use in trials related to insect agriculture.

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			<td>31-qt (29.3 L) Snap-lid tote bin with lid</td>
			<td>HOMZ</td>
			<td>3430CLBL</td>
			<td>Used to house breeding stock</td>
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			<td>Fischer Scientific</td>
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			<td>50-gal (189.27L) tote bin with lid</td>
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			<td>Used as secondary containment when handling crickets</td>
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			<td>7.5-qt (7.1 L) snap-lid tote bin with lid</td>
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			<td>Manufactured by Accuris, a subsidieary of Benchmark Scientific</td>
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			<td>Wal-Mart</td>
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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).

Data demonstrating successful cricket rearing from hatching to 65 days old were collected during a September 2021 feed trial. Crickets were grown from eggs following steps 1.1.1-2.6.1 of these protocols, and six replicate cages were stocked with 24 random 22-day-old (third instar) crickets following step 2.7 above. Crickets were then reared in ambient room conditions; however, due to a malfunctioning facility air handling unit, the average room temperature was 25 &#177; 1 &#176;C at 20% relative humidity rather than the ...

Watch this Scientific Journal Video about Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology at JoVE.com

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.

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 ...

maintaining-laboratory-cultures-gryllus-bimaculatus-versatile

Research

JoVE Journal

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3.1K Views.  University of Wisconsin-Madison. 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.

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

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, ...

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.

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.

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

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

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

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

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

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to study the functions and evolution of bacterial symbionts in their host. As most symbionts cannot be cultured in vitro, methods to isolate an adequate quantity of bacteria for genome sequencing are very important. In the whitefly Bemisia tabaci, a number of endosymbionts have been identified and are predicted to be of importance in the development and reproduction of the pests through multiple approaches. However, the mechanism underpinning the associations remains largely unknown. The obstacle partially comes from the fact that the endosymbionts in whitefly, mostly restrained in bacteriocytes, are hard to separate from the host cells. Here we report a step-by-step protocol for the identification, extraction and purification of endosymbionts from the whitefly B. tabaci mainly by dissection and filtration. Endosymbiont samples prepared by this method, although still a mixture of different endosymbiont species, are suitable for subsequent genome sequencing and analysis of the possible roles of endosymbionts in B. tabaci. This method may also be used to isolate endosymbionts from other insects.

Methods for the Extraction of Endosymbionts from the Whitefly Bemisia tabaci

Here, we present a protocol to isolate endosymbionts from the whitefly Bemisia tabaci through dissection and filtration. After amplification, the DNA samples are suitable for subsequent sequencing and study of the mutualism between endosymbionts and whitefly.

Bacterial symbionts form an intimate relationship with their hosts and confer advantages to the hosts in most cases. Genomic information is critical to ...

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ecologically-important endpoints, though they are rarely considered in toxicological studies. D. melanogaster is emerging as a toxicology model for the behavioral, neurological, and genetic impacts of toxicants, to name a few. More importantly, species in the genus Drosophila can be utilized as a model system for an integrative framework approach to incorporate emergent properties and answer ecologically-relevant questions in&#160;toxicology research. The aim of this paper is to provide a protocol for exposing species in the genus Drosophila to pollutants to be used as a model system for a range of phenotypic outputs and ecologically-relevant questions. More specifically, this protocol can be used to 1) link multiple biological levels of organization and understand the impact of toxicants on both individual- and population-level fitness; 2) test the impact of toxicants at different stages of developmental exposure; 3) test multigenerational and evolutionary implications of pollutants; and 4) test multiple contaminants and stressors simultaneously.

Experimental Protocol for Using Drosophila As an Invertebrate Model System for Toxicity Testing in the Laboratory

In this paper, we provide a detailed protocol for exposing species in the genus Drosophila to pollutants with the goal of studying the impact of exposure on a range of phenotypic outputs at different developmental stages and for more than one generation.

Emergent properties and external factors (population-level and ecosystem-level interactions, in particular) play important roles in mediating ...

A Versatile Model of Hard Tick Infestation on Laboratory Rabbits

The use of live animals in tick research is crucial for a variety of experimental purposes including the maintenance of hard tick colonies in the laboratory. In ticks, all developmental stages (except egg) are hematophagous, and acquiring a blood-meal when attached to their vertebrate hosts is essential for the successful completion of their life cycle. Here we demonstrate a simple method that uses easily openable capsules for feeding of hard ticks on rabbits. The advantages of the proposed method include its simplicity, short duration and most importantly versatile adjustment to the needs of specific experimental requirements. The method makes possible the use of multiple chambers (of various sizes) on the same animal, which permits feeding of multiple stages or different experimental groups while reducing the overall animal requirement. The non-irritating and easily accessible materials used minimizes discomfort to the animals, which can be easily recovered from an experiment and offered for adoption or reused if the ethical protocol allows it.

We have developed a simple and versatile system to feed hard ticks on laboratory rabbits. Our non-laborious protocol uses easily accessible materials and can be adjusted depending on the requirements of the various experimental settings. The method allows comfortable monitoring and/or sampling of ticks during the entire feeding period.

The use of live animals in tick research is crucial for a variety of experimental purposes including the maintenance of hard tick colonies in the ...

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A)

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer progression. How the chromosomal location and function of a centromere (i.e., centromere identity) are determined and participate in accurate chromosome segregation is a fundamental question. CENP-A is proposed to be the non-DNA indicator (epigenetic mark) of centromere identity, and CENP-A ubiquitylation is required for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability.
Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-CENP-A K124R mutant suggesting that ubiquitylation at a different lysine is induced because of the EYFP tagging in the CENP-A K124R mutant protein. Lysine 306 (K306) ubiquitylation in EYFP-CENP-A K124R was successfully identified, which corresponds to lysine 56 (K56) in CENP-A through mass spectrometry analysis. A caveat is discussed in the use of GFP/EYFP or the tagging of high molecular weight protein as a tool to analyze the function of a protein. Current technical limit is also discussed for the detection of ubiquitylated bands, identification of site-specific ubiquitylation(s), and visualization of ubiquitylation in living cells or a specific single cell during the whole cell cycle.
The method of mass spectrometry analysis presented here can be applied to human CENP-A protein with different tags and other centromere-kinetochore proteins. These combinatory methods consisting of several assays/analyses could be recommended for researchers who are interested in identifying functional roles of ubiquitylation.

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A EYFP-CENP-A

CENP-A ubiquitylation is an important requirement for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability. Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A) protein.

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer ...

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no prevention. Control of mosquito vectors can rely on chemicals applied to the immature and adult stages, which can contribute to the mortality of non-targets and more importantly, lead to insecticide resistance in the vector. The sterile insect technique (SIT) is a method of controlling populations of pests through the release of sterilized adult males that mate with wild females to produce non-viable offspring. This paper describes the process of producing sterile males for use in an operational SIT program for the control of Aedes aegypti mosquitoes. Outlined here are the steps used in the program including rearing and maintaining a colony, separating male and female pupae, irradiating and marking adult males, and shipping Aedes aegypti males to the release site. Also discussed are procedural caveats, program limitations, and future objectives.

Preparing Irradiated and Marked Male Aedes aegypti Mosquitoes for Release in an Operational Sterile Insect Technique Program

The sterile insect technique (SIT) is used to control specific, medically important mosquito populations that may be resistant to chemical controls. Here, we describe a method of mass rearing and preparation of sterile male mosquitoes for release in an operational SIT program targeting the Aedes aegypti mosquito.

The control of such human diseases as dengue, Zika, and chikungunya relies on the control of their vector, the Aedes aegypti mosquito, because there is no ...

Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to investigate metabolic capability. To obtain the measures of mitochondrial respiration, fresh biological samples must be used, and the entire laboratory procedure must be completed within approximately 2 h. Furthermore, multiple pieces of specialized equipment are required to perform these laboratory assays. This creates a challenge for measuring mitochondrial respiration in the tissues of wild animals living far from physiology laboratories as live tissue cannot be preserved for very long after collection in the field. Moreover, transporting live animals over long distances induces stress, which can alter mitochondrial energetics.
This manuscript introduces the Auburn University (AU) MitoMobile, a mobile mitochondrial physiology laboratory that can be taken into the field and used on-site to measure mitochondrial metabolism in tissues collected from wild animals. The basic features of the mobile laboratory and the step-by-step methods for measuring isolated mitochondrial respiration rates are presented. Additionally, the data presented validate the success of outfitting the mobile mitochondrial physiology laboratory and making mitochondrial respiration measurements. The novelty of the mobile laboratory lies in the ability to drive to the field and perform mitochondrial measurements on the tissues of animals captured on site.

We designed and constructed a mobile laboratory to measure respiration rates in isolated mitochondria of wild animals captured at field locations. Here, we describe the design and outfitting of a mobile mitochondrial laboratory and the associated laboratory protocols.

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to ...

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite the growing burden of tick-borne diseases, research on ticks has lagged behind insect disease vectors due to challenges in applying genetic transformation tools for functional studies to the unique biology of ticks. Genetic interventions have been gaining attention to reduce mosquito-borne diseases. However, the development of such interventions requires stable germline transformation by injecting embryos. Such an embryo injection technique is lacking for chelicerates, including ticks. Several factors, such as an external thick wax layer on tick embryos, hard chorion, and high intra-oval pressure, are some&#160;obstacles that previously prevented embryo injection protocol development in ticks. The present work has overcome these obstacles, and an embryo injection technique for the black-legged tick, Ixodes scapularis, is described here.&#160;This technique can be used to deliver components, such as CRISPR/Cas9, for stable germline transformations.

Embryo Injection Technique for Gene Editing in the Black-Legged Tick, Ixodes scapularis

The present protocol describes a method for injecting tick embryos. Embryo injection is the preferred technique for genetic manipulation to generate transgenic lines.

Ticks can transmit various viral, bacterial, and protozoan pathogens and are therefore considered vectors of medical and veterinary importance. Despite ...