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 Sincerest thanks to Kevin Bachhuber of Bachhuber Consulting Inc. for access to his unpublished guide for commercial cricket rearing and to Michael Bartlett Smith for his assistance in refining and troubleshooting these methods.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.

<table><tbody><tr><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></tr><tr><td>3-tier/12-tray Grow Light Stand</td><td>Fischer Scientific</td><td>NC1938548</td><td></td></tr><tr><td>50-gal (189.27L) tote bin with lid</td><td>Sterilite</td><td>#14796603</td><td>Used as secondary containment when handling crickets</td></tr><tr><td>50 mL polypropylene graduated cylinder</td><td>Fischer Scientific</td><td>S95171</td><td></td></tr><tr><td>7.5-qt (7.1 L) snap-lid tote bin with lid</td><td>HOMZ</td><td>3410CLBL</td><td>Used to house exprimental stock</td></tr><tr><td>Accuris 500 g x 0.01 g Balance</td><td>Manufactured by Accuris, a subsidieary of Benchmark Scientific</td><td>W3300-500</td><td>Purchased from Dot Scientific through University of Wisconsin system purchasing service &quot;ShopUW+&quot;</td></tr><tr><td>Ace Premier 1 Inch Flat Chip Brush</td><td>Ace Hardware&amp;nbsp;</td><td>#1803261</td><td></td></tr><tr><td>Bel-Art SP Scienceware deionized water wash bottle</td><td>Fischer Scientific</td><td>03-421-160&amp;nbsp;</td><td></td></tr><tr><td>Bright aluminum window screen&amp;nbsp;</td><td>Phifer</td><td>UNSPSC# 11162108</td><td>Mesh size 18 x 16&quot;</td></tr><tr><td>Clear Disposable Plastic Portion Cups 5.5 oz w/ lids</td><td>Wal-Mart</td><td>N/A</td><td></td></tr><tr><td>Deionized water</td><td></td><td></td><td></td></tr><tr><td>Diablo 4-4/8&quot; x 13 TPI Ultra Fine Finish Bi-Metal Jigsaw Blade</td><td>Home Depot</td><td>#313114935</td><td></td></tr><tr><td>Egg Filler Flats-Paper, 12 x 12&quot;</td><td>Uline</td><td>S-5189</td><td></td></tr><tr><td>Fisherbrand Petri Dishes with Clear Lid 100 x 15mm</td><td>Fischer Scientific</td><td>FB0875714</td><td></td></tr><tr><td>Fisherbrand Petri Dishes with Clear Lid 60 x 15mm</td><td>Fischer Scientific</td><td>FB0875713A</td><td></td></tr><tr><td>Georgia-Pacific Envision Brown Paper Towels</td><td>Home Depot</td><td>#205675843</td><td></td></tr><tr><td>Infinity Tough Guy high performance hot-melt glue sticks</td><td>Infinity Bond</td><td>Infinity IM-Tough-Guy-12</td><td></td></tr><tr><td>Mazuri Cricket Diet</td><td>Land O&#039; Lakes International</td><td>SKU#&amp;nbsp; 3002219-105</td><td></td></tr><tr><td>Stanley TimeIt Twin 2-outlet Grounded Mechanical 24 Hour Timer</td><td>Wal-Mart</td><td>N/A</td><td></td></tr><tr><td>Vermont Organics Reclamation Soil 11 lb Coir Block</td><td>Home Depot</td><td>#300679904</td><td></td></tr></tbody></table>

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

Biology

3.1K Views.  University of Wisconsin-Madison. Deze technieken tonen praktische en beproefde methoden voor het fokken van Gryllus bimaculatus, een belangrijke mini-diersoort die wordt gekweekt voor diervoeder en menselijk voedsel en een steeds belangrijker alternatief modelorganisme. Deze laboratoriummethoden voor de cultuur van Gryllus bimaculatus zijn afgeleid van technieken die gebruikelijk zijn in de Noord-Amerikaanse cricketlandbouwindustrie en vertegenwoordigen een facsimile op laboratoriumschaal van cricketlandbouwactiviteiten. Om ...

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

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

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

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

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

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

Genetics

The Insect Galleria mellonella as a Powerful Infection Model to Investigate Bacterial Pathogenesis

The study of bacterial virulence often requires a suitable animal model. Mammalian models of infection are costly and may raise ethical issues. The use of insects as infection models provides a valuable alternative. Compared to other non-vertebrate model hosts such as nematodes, insects have a relatively advanced system of antimicrobial defenses and are thus more likely to produce information relevant to the mammalian infection process. Like mammals, insects possess a complex innate immune system1. Cells in the hemolymph are capable of phagocytosing or encapsulating microbial invaders, and humoral responses include the inducible production of lysozyme and small antibacterial peptides2,3. In addition, analogies are found between the epithelial cells of insect larval midguts and intestinal cells of mammalian digestive systems. Finally, several basic components essential for the bacterial infection process such as cell adhesion, resistance to antimicrobial peptides, tissue degradation and adaptation to oxidative stress are likely to be important in both insects and mammals1. Thus, insects are polyvalent tools for the identification and characterization of microbial virulence factors involved in mammalian infections.
Larvae of the greater wax moth Galleria mellonella have been shown to provide a useful insight into the pathogenesis of a wide range of microbial infections including mammalian fungal (Fusarium oxysporum, Aspergillus fumigatus, Candida albicans) and bacterial pathogens, such as Staphylococcus aureus, Proteus vulgaris, Serratia marcescens Pseudomonas aeruginosa, Listeria monocytogenes or Enterococcus faecalis4-7. Regardless of the bacterial species, results obtained with Galleria larvae infected by direct injection through the cuticle consistently correlate with those of similar mammalian studies: bacterial strains that are attenuated in mammalian models demonstrate lower virulence in Galleria, and strains causing severe human infections are also highly virulent in the Galleria model8-11. Oral infection of Galleria is much less used and additional compounds, like specific toxins, are needed to reach mortality.
G. mellonella larvae present several technical advantages: they are relatively large (last instar larvae before pupation are about 2 cm long and weight 250 mg), thus enabling the injection of defined doses of bacteria; they can be reared at various temperatures (20 &deg;C to 30 &deg;C) and infection studies can be conducted between 15 &deg;C to above 37 &deg;C12,13, allowing experiments that mimic a mammalian environment. In addition, insect rearing is easy and relatively cheap. Infection of the larvae allows monitoring bacterial virulence by several means, including calculation of LD5014, measurement of bacterial survival15,16 and examination of the infection process17. Here, we describe the rearing of the insects, covering all life stages of G. mellonella. We provide a detailed protocol of infection by two routes of inoculation: oral and intra haemocoelic. The bacterial model used in this protocol is Bacillus cereus, a Gram positive pathogen implicated in gastrointestinal as well as in other severe local or systemic opportunistic infections18,19.

The Insect Galleria mellonella as a Powerful Infection Model to Investigate Bacterial Pathogenesis

Oral and intra haemocolic infection of larvae of the greater wax moth Galleria mellonella is described. This insect can be used to study virulence factors of entomopathogenic as well as mammalian opportunistic bacteria. Rearing of the insects, methods of infection and examples of in vivo analysis are described.

The study of bacterial virulence often requires a suitable animal model. Mammalian models of infection are costly and may raise ethical issues. The use of ...

Immunology and Infection

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 ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in agronomic practices resulted in serious outbreaks of stable flies in several countries. These outbreaks disrupted livestock production and human recreation resulting in public demands for increasing research and management efforts for this pest. A simple and inexpensive procedure for rearing stable flies for laboratory studies is presented. The procedure uses locally available diet components, equipment and supplies. The procedure can be adapted for rearing other muscoid flies including face fly (Musca autumnalis), horn fly (Haematobia irritans), and house fly (Musca domestica). The procedure produces stable fly puparia averaging 12.5 mg and ~35% egg to adult survival. Approximately 3000 flies are produced in each pan.

A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in ...

Establishment and Maintenance of Gnotobiotic American Cockroaches (Periplaneta americana)

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment and maintenance of gnotobiotic American cockroaches (Periplaneta americana). This approach includes built-in sterility checks for ongoing quality control. Gnotobiotic insects are defined here as cockroaches that still contain their vertically transmitted endosymbiont (Blattabacterium) but lack other microbes that normally reside on their surface and in their digestive tract. For this protocol, egg cases (oothecae) are removed from a (nonsterile) stock colony and surface sterilized. Once collected and sterilized, the oothecae are incubated at 30 &#176;C for approximately 4&#8722;6 weeks on brain-heart infusion (BHI) agar until they hatch or are removed due to contamination. Hatched nymphs are transferred to an Erlenmeyer flask containing a BHI floor, sterile water, and sterile rat food. To ensure that the nymphs are not housing microbes that are unable to grow on BHI in the given conditions, an additional quality control measure uses restriction fragment-length polymorphism (RFLP) to test for nonendosymbiotic microbes. Gnotobiotic nymphs generated using this approach can be inoculated with simple or complex microbial communities and used as a tool in gut microbiome studies.

Establishment and Maintenance of Gnotobiotic American Cockroaches Periplaneta americana

This protocol is used in establishing and maintaining gnotobiotic American cockroaches (Periplaneta americana) by surface sterilizing the egg cases (oothecae) prior to hatching. These gnotobiotic insects contain their vertically transmitted Blattabacterium endosymbionts but have axenic guts.

Gnotobiotic animals are a powerful tool for the study of controls on microbiome structure and function. Presented here is a protocol for the establishment ...

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

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

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

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

Bradysia coprophila Lab Protocols and Chromosomal Research

Laboratory Maintenance of the Lower Dipteran Fly Bradysia (Sciara) coprophila: A New/Old Emerging Model Organism

Laboratory stocks of the lower dipteran fly, Bradysia (Sciara) coprophila, have been maintained for over a century. Protocols for laboratory upkeep of B. coprophila are presented here. These protocols will be useful for the rapidly increasing number of laboratories studying B. coprophila to take advantage of its unique biological features, which include (1) a monopolar spindle in male meiosis I; (2) non-disjunction of the X dyad in male meiosis II; (3) chromosome imprinting to distinguish maternal from paternal homologs; (4) germ line-limited (L) chromosomes; (5) chromosome elimination (paternal chromosomes in male meiosis I; one to two X chromosomes in early embryos; L chromosomes from the soma in early embryos); (6) sex determination by the mother (there is no Y chromosome); and (7) developmentally regulated DNA amplification at the DNA puff loci in larval salivary gland polytene chromosomes. 
It is now possible to explore these many unique features of chromosome mechanics by using the recent advances in sequencing and assembly of the B. coprophila genome and the development of transformation methodology for genomic engineering. The growing scientific community that uses B. coprophila for research will benefit from the protocols described here for mating the flies (phenotypic markers for mothers that will have only sons or only daughters; details of mass mating for biochemical experiments), checking embryo hatch, feeding larvae, and other comments on its rearing.

Author Spotlight: Exploring Bradysia coprophila's Unique Biology &#8211; A Guide to Laboratory Maintenance

This paper outlines the laboratory maintenance (including mating and feeding) of the lower dipteran fly Bradysia (Sciara) coprophila.

Laboratory stocks of the lower dipteran fly, Bradysia (Sciara) coprophila, have been maintained for over a century. Protocols for laboratory upkeep of B. ...

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

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