We would like to thank Anthony Weinhold and the numerous students who have worked with us over the years for technical support as well as suggestions for improving our insect rearing procedures.

1. Egg Collection (Figure 1)
<ol>
	<li>Prepare egging cup, place one end of cloth in ~500 mL beaker filled with warm (~40 &#176;C) water. Overlap the sides of the cup and fix with a rubber band. Fold the loose end of the cloth back over the top of the cup.</li>
	<li>Place egging cup in cage of 8&#8211;10 day old flies for ~2 h. Gravid female stable flies will oviposit on the cloth.</li>
	<li>Remove egging cup from cage and rinse eggs off of the egging cloth into a small pan with squeeze wash bottle.</li>
</ol>
2. Prepare Larval Rearing Pan
<ol>
	<li>Prepare medium (quantities for 1 rearing pan)
	<ol>
		<li>Combine wheat bran (500 g), wood shavings (200 g) and fish meal (115 g) in 10 L plastic dish pan and mix well.</li>
		<li>Add ammonium bicarbonate (50 g) to water (~25 &#176;C, 1600 mL) and mix until dissolved.</li>
		<li>Add water solution to dry ingredients and mix to ensure no unwetted material. Level medium in pan but do not compress.</li>
	</ol>
	</li>
	<li>Add stable fly eggs
	<ol>
		<li>Make a shallow furrow in medium the length of the pan.</li>
		<li>Deposit 1 mL of stable fly eggs (~8,000 eggs) in furrow with egg pipette (Figure 2). 
		NOTE: Egg pipette made by cutting a plastic graduated pipette at desired point, inserting 100 mesh screen and gluing pipette back together.</li>
		<li>Cover eggs with a thin layer of medium to prevent desiccation.</li>
		<li>Cover pan with pillow case, close with a rubber band and label. Place pans in larval room (23 &#177; 2 &#176;C, 30&#8211;50% RH and 12:12 [L:D] h photoperiod). 
		NOTE: Larval development takes 10&#8211;14 days from the time of oviposition.</li>
	</ol>
	</li>
</ol>
3. Pupal Processing, Larvae Move to The Edge of The Medium 7&#8211;9 Days After Oviposition and Pupariate by Days 13 or 14
<ol>
	<li>Scoop puparia, primarily located around the edges of the pan under the crust that forms on the surface of the medium, out and placed in a clean pan.</li>
	<li>Fill pan, with puparia, &#189; to &#190; full of water. Break up clumps of puparia and medium. Wheat bran and protein components of the medium sink while puparia &#62;1 day old float19.</li>
	<li>Remove floating materials, puparia and some medium components with a colander and wash through a series of sieves (#5, #7, #12, #20) to remove remaining medium. The #12 sieve collects the puparia and the #20 keeps solid waste from entering the drain. Wash puparia between sieves with a sink sprayer.</li>
	<li>Rinse puparia, in the #12 sieve, into a clean pan. Fill pan to &#189; full of water. Pour floating puparia into a colander and allow excess water to drain.</li>
	<li>Transfer drained puparia to drying screen with fan and leave until dry (Figure 3).</li>
</ol>
4. Pupal Shelf, an Alternative Method for Collecting Puparia Is with a Pupal Shelf18
NOTE: The shelf is made from a piece of plastic cut from the end of a 10 L dish pan (10.2 cm tall x 10.2 cm wide x 31.9 cm long).
<ol>
	<li>Prepare medium as described in 2.1. Taper medium in pan from ~2.5 cm deep on one end to ~7.5 cm deep on the other.</li>
	<li>Place shelf on medium in the shallow end of the rearing pan and tape to prevent larvae from crawling alongside the shelf (Figure 4).
	<ol>
		<li>Saturate sponge (~14.5 x 9.0 x 4.5 cm3, regenerated cellulose) and wrap it in a water soaked cloth (30.5 x 42 cm2, cotton) and place on the shelf about one half inch from the medium.</li>
	</ol>
	</li>
	<li>Add eggs as described in 2.2 and proceed as before.
	<ol>
		<li>Check pans daily to ensure that the sponge remains moist.</li>
	</ol>
	</li>
	<li>Rinse puparia from the shelf, sponge and cloth into a clean dish pan and pour into colander 14 days after oviposition.</li>
</ol>
5. Quality Control
<ol>
	<li>Weigh all of the puparia produced in pan (total wt.).</li>
	<li>Isolate 100 puparia, weigh and placed in a 9 cm Petri dish.</li>
	<li>Count and sex adults ~5&#8211;8 days later to determine emergence rate and sex ratio. Store Petri dishes with emerged adults in a freezer for counting at a later date if needed. Flies are sexed by the shape of their eyes and width of the fronto-orbital plate20 or genitalia under low magnification.</li>
	<li>Record total weight, weight of 100 puparia, and number of emerged adult males and females.</li>
</ol>
6. Preparing Blood
<ol>
	<li>Collect fresh bovine blood from a local abattoir in 19 L buckets containing 70 g of sodium citrate tribasic dihydrate in 500 mL of water. Stir blood vigorously for 5&#8211;10 min and returned to the laboratory.</li>
	<li>Strain blood through a colander to remove clots and partitioned into 2 L containers, label with the date of collection and store in a freezer (-20 &#176;C).</li>
	<li>Remove container from freezer and place in refrigerator 2 days before needed. Blood can be used for ~2 weeks once thawed and stored in the freezer for up to 1 year.</li>
</ol>
7. Adult Maintenance
<ol>
	<li>Prepare cages, line cage bottom with butcher paper to facilitate cleaning.</li>
	<li>Place 50 g of dry stable fly pupae (~3,500) in each cage. Adults eclose a few hours to a couple of days after pupal processing.</li>
	<li>Place a fresh blood soaked feminine napkin on top of each cage for feeding. Begin feeding within 24 h of adult eclosion and repeat daily until a day prior to oviposition (9&#8211;10 days after emergence).</li>
	<li>After oviposition (section 1), place cage in freezer for 4&#8211;8 h and then clean by removing the paper, rinsing frass from the sides and floor with hot water and scrubbing all surfaces with laundry detergent and chlorine bleach. Rinse cages with hot water and allow to dry.</li>
</ol>

<ol>
	<li>Cook, D. F., Dadour, I. R., Keals, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+fly,+house+fly+(Diptera:+Muscidae),+and+other+nuisance+fly+development+in+poultry+litter+associated+with+horticultural+crop+production.">Stable fly, house fly (Diptera: Muscidae), and other nuisance fly development in poultry litter associated with horticultural crop production.</a> J. Econ. Entomol. 92 (6), 1352-1357 (1999).</li><li>Dominghetti, T. F., de Barros, A. T., Soares, C. O., Can&#231;ado, P. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stomoxys+calcitrans+(Diptera:+Muscidae)+outbreaks:+current+situation+and+future+outlook+with+emphasis+on+Brazil.">Stomoxys calcitrans (Diptera: Muscidae) outbreaks: current situation and future outlook with emphasis on Brazil.</a> Rev. Bras. Parasitol. Vet. 24 (4), 387-395 (2015).</li><li>Sol&#243;rzano, J. -. A., Guilles, J., Bravo, O., Vargas, C., Gomez-Bonilla, Y., Bingham, G., Taylor, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biology+and+trapping+of+stable+flies+(Diptera:+Muscidae)+developing+in+pineapple+residues+(Ananas+comosus)+in+Costa+Rica.">Biology and trapping of stable flies (Diptera: Muscidae) developing in pineapple residues (Ananas comosus) in Costa Rica.</a> J. Insect Sci. 15 (1), 145 (2015).</li><li>Berry, I. L., Stage, D. A., Campbell, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Populations+and+economic+impacts+of+stable+flies+on+cattle.">Populations and economic impacts of stable flies on cattle.</a> Trans. Am. Soc. Agric. Eng. 26, 873-877 (1983).</li><li>Taylor, D. B., Vreysen, J., Hendrichs, R., Cardoso Pereira, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Area-wide+management+of+stable+flies.">Area-wide management of stable flies.</a> Area-wide management of insect pests. , (2017).</li><li>Bishopp, F. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+stable+fly+(Stomoxys+calcitrans+L.),+an+important+live+stock+pest.">The stable fly (Stomoxys calcitrans L.), an important live stock pest.</a> J.Econ. Entomol. 6 (1), 112-126 (1913).</li><li>Hogsette, J. A., Ruff, J. P., Jones, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+fly+biology+and+control+in+northwest+Florida.">Stable fly biology and control in northwest Florida.</a> J. Agric. Entomol. 4 (1), 1-11 (1987).</li><li>Lysyk, T., Kalischuk-Tymensen, L., Selinger, L., Lancaster, R., Wever, L., Cheng, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+stable+fly+larvae+(Diptera:+Muscidae)+on+an+egg+yolk+medium.">Rearing stable fly larvae (Diptera: Muscidae) on an egg yolk medium.</a> J. Med. Entomol. 38, 382-388 (1999).</li><li>Romero, A., Broce, A., Zurek, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+bacteria+in+the+oviposition+behaviour+and+larval+development+of+stable+flies.">Role of bacteria in the oviposition behaviour and larval development of stable flies.</a> Med. Vet. Entomol. 20, 115-121 (2006).</li><li>Glaser, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+flies+for+experimental+purposes+with+biological+notes.">Rearing flies for experimental purposes with biological notes.</a> J. Econ. Entomol. 17 (4), 486-496 (1924).</li><li>Melvin, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Physiological+studies+on+the+effect+of+flies+and+fly+sprays+on+cattle.">Physiological studies on the effect of flies and fly sprays on cattle.</a> J. Econ. Entomol. 25 (6), 1151-1164 (1932).</li><li>Doty, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convenient+method+of+rearing+the+stable+fly.">Convenient method of rearing the stable fly.</a> J. Econ. Entomol. 30 (2), 367-369 (1937).</li><li>Bridges, A. C., Spates, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Larval+medium+for+the+stable+fly+Stomoxys+calcitrans+(L.).">Larval medium for the stable fly Stomoxys calcitrans (L.).</a> Southwest. Entomol. 8 (1), 6-10 (1983).</li><li>Hogsette, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+diets+for+production+of+house+flies+and+stable+flies+(Diptera:+Muscidae)+in+the+laboratory.">New diets for production of house flies and stable flies (Diptera: Muscidae) in the laboratory.</a> J. Econ. Entomol. 85 (6), 2291-2294 (1992).</li><li>McGregor, W. S., Dreiss, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rearing+stable+flies+in+the+laboratory.">Rearing stable flies in the laboratory.</a> J. Econ. Entomol. 48 (3), 327-328 (1955).</li><li>Goodhue, L. D., Cantrel, K. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+vermiculite+in+medium+for+stable+fly+larvae.">The use of vermiculite in medium for stable fly larvae.</a> J. Econ. Entomol. 51 (2), 250 (1958).</li><li>Friesen, K., Berkebile, D. R., Zhu, J. J., Taylor, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Augmenting+laboratory+rearing+of+stable+fly+(Diptera:+Muscidae)+larvae+with+ammoniacal+salts.">Augmenting laboratory rearing of stable fly (Diptera: Muscidae) larvae with ammoniacal salts.</a> J. Insect Sci. 17 (1), 1-6 (2017).</li><li>Berkebile, D. R., Weinhold, A. P., Taylor, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+method+for+collecting+clean+stable+fly+(Diptera:Muscidae)+pupae+of+known+age.">A new method for collecting clean stable fly (Diptera:Muscidae) pupae of known age.</a> Southwest. Entomol. 34 (4), 469-476 (2009).</li><li>Champlain, R. A., Fisk, F. W., Dowdy, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Some+improvements+in+rearing+stable+flies.">Some improvements in rearing stable flies.</a> J. Econ. Entomol. 47 (5), 940-941 (1954).</li><li>Zumpt, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The stomoxyine biting flies of the world. , (1973).</li><li>Wienhold, B. J., Taylor, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Substrate+properties+of+stable+fly+(Diptera:+Muscidae)+developmental+sites+associated+with+round+bale+hay+feeding+sites+in+eastern+Nebraska.">Substrate properties of stable fly (Diptera: Muscidae) developmental sites associated with round bale hay feeding sites in eastern Nebraska.</a> Environ. Entomol. 41 (2), 213-221 (2012).</li><li>Friesen, K., Berkebile, D. R., Wienhold, B. J., Durso, L., Zhu, J., Taylor, D. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Environmental+parameters+associated+with+stable+fly+(Diptera:+Muscidae)+development+at+hay+feeding+sites.">Environmental parameters associated with stable fly (Diptera: Muscidae) development at hay feeding sites.</a> Environ. Entomol. 45 (3), 570-576 (2016).</li><li>Albuquerque, T. A., Zurek, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+changes+in+the+bacterial+community+of+animal+feces+and+their+correlation+with+stable+fly+oviposition,+larval+development,+and+adult+fitness.">Temporal changes in the bacterial community of animal feces and their correlation with stable fly oviposition, larval development, and adult fitness.</a> Front. Microbiol. 5 (590), 1-9 (2014).</li><li>Bailey, D. L., Whitfield, T. L., LaBrecque, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Laboratory+biology+and+techniques+for+mass+producing+the+stable+fly,+Stomoxys+calcitrans+(L.)+(Diptera:+Muscidae).">Laboratory biology and techniques for mass producing the stable fly, Stomoxys calcitrans (L.) (Diptera: Muscidae).</a> J. Med. Entomol. 12 (2), 189-193 (1975).</li><li>Smith, J. P., Hall, R. D., Thomas, G. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Field+studies+on+mortality+of+the+immature+stages+of+the+stable+fly+(Diptera:+Muscidae).">Field studies on mortality of the immature stages of the stable fly (Diptera: Muscidae).</a> Environ. Entomol. 14 (6), 881-890 (1985).</li></ol>

The authors have nothing to disclose.

Stable flies can be found developing in a broad variety of substrates in nature and can be reared in many types of media in the laboratory. We have used wood shavings and vermiculite as bulking agents. Vermiculite worked well, but made separating puparia from rearing medium difficult and was expensive (~$0.60/pan). Possibly because of the added rigor of separating puparia from the medium, eclosion was also lower with vermiculite, 57% vs. 75% for wood shavings. Cottonwood shavings were comparable to vermiculite, but at ti...

Stable flies, Stomoxys calcitrans (L.), are hematophagous flies whose painful bites disrupt the grazing behavior of livestock, cause pain and suffering to companion animals and disrupt human recreational activities worldwide. Immature stable flies develop in fermenting vegetative matter, often contaminated with animal waste. Changing agronomic practices and crops have produced serious outbreaks of stable flies in crop residues, vegetables in Australia1, sugar cane in Brazil2, and pineapple in Costa Rica3. Although just 14 stable flies per animal are considered to be the economic threshold4, observations of more than 2,000 flies per animal have been made during recent outbreaks5. Such infestation levels reduce host productivity to near zero and can cause mortality6. As a result of agronomically associated outbreaks, stable flies are receiving renewed interest and demand for laboratory colonies has increased dramatically.
As for all holometamorphic insects, stable flies obtain all of the nutrients required for growth during the immature or larval stage. Therefore, an important component of a rearing system is the larval diet or substrate. Stable fly larvae have been observed developing in a broad range of substrates in the field7 and they are dependent upon the microbial community of the substrate8,9. Natural larval substrates are primarily composed of decomposing or fermenting vegetative materials often contaminated with nitrogenous wastes.
For laboratory rearing, stable fly larval substrates are usually composed of a vegetative material and an added nitrogen source. Numerous materials have been used for stable fly larval diets. The first larval diets mimicked natural substrates and included fermenting oat straw and horse or cow manure10,11. Carbohydrate sources include wheat bran12,13,14, alfalfa meal12,13,14 and a commercial formulation developed by Chemical Specialties Manufacturers Association (CSMA, 33% wheat bran, 27% alfalfa meal, 40% brewer&#39;s yeast granules)13,14,15,16. Nitrogen sources include yeast suspension12, fish meal and ammonium bicarbonate17. Inert bulking materials are often included in diets including oat hulls12, bagasse13, vermiculite16, wood chips13,18 and pelleted peanut hulls14.
A primary objective of laboratory rearing is to produce a product that is as physiologically similar to &#34;wild type&#34; as possible in order that laboratory experiments will produce results reflecting those of field populations. This requires that in-breeding and selection be minimized to maintain genetic diversity and nutritional resources be comparable to those in the field. Secondary objectives are to minimize labor and expenses. A major component of minimizing expenses is the use of locally available diet components. The stable fly rearing system presented was developed to meet these objectives.

<table><tbody><tr><td>Diamalt</td><td>Premier Malt Products, Inc., Saddle Brook, NJ</td><td>2540</td><td></td></tr><tr><td>CSMA Fly media</td><td>Purina Animal Nutrition, Arden Hills, MN</td><td>5S6Z</td><td></td></tr><tr><td>Thin Maxi Pad</td><td>The Tranzonic Co., Cleveland, OH, USA</td><td>5001M</td><td></td></tr><tr><td>Calf Manna</td><td>MannaPro, Chesterfield, MO, USA</td><td>Manna Pro</td><td></td></tr><tr><td>Ammonium Bicarbonate</td><td>Spectrum Chemical Manufacturing Corp, Gardena, CA</td><td>A1125</td><td></td></tr><tr><td>Wheat bran, Coarse</td><td>Siemer Milling Company, Teutopolis, IL</td><td></td><td></td></tr><tr><td>Wood shavings</td><td>Tractor Supply Company, Brentwood, TN</td><td>502770699</td><td></td></tr><tr><td>Fishmeal</td><td>Consumer Supply Distributing, North Sioux City, SD</td><td>F1550</td><td></td></tr><tr><td>Adult cages</td><td>All Aluminum Window Company, Lincoln, Ne</td><td>Custom</td><td>45 &amp;times; 45 &amp;times; 45 cm, 18 &amp;times; 16 mesh aluminum screen, stockinette access</td></tr><tr><td>9 &amp;times; 28 cm black cotton cloth</td><td>Robert Kaufman Fabrics, Los Angeles, CA</td><td>K040-114</td><td>Egging cloth</td></tr><tr><td>10 L plastic dish pans</td><td>Rubbermaid, Saratoga Springs, NY</td><td>FG2951ARWHT</td><td>Larval pans</td></tr><tr><td>Stockinette, Cotton, 12 inch x 25 yard roll</td><td>Tex-Care Medical Company, Burlington, NC</td><td>91311-225</td><td></td></tr></tbody></table>

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.

Larvae pupariate 10&#8211;14 days and adults emerge 14&#8211;16 days after oviposition. Generation time, egg to egg, is ~24 days. Rearing data for May 2013 to January 2017 with three different bulking agents and two colonies are presented in Figure 5. Cottonwood gave the best yield, 3867 &#177;1442 (<img alt="Equation" src="/files/ftp_upload/57341/57341eq1.jpg" /> &#177; standard deviation) pupae weighing 12.5 &#177;1.6 mg with 74 &#177;19% eclosion producing...

Watch this Scientific Journal Video about Laboratory Rearing of Stable Flies and Other Muscoid Diptera at JoVE.com

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

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 ...

laboratory-rearing-of-stable-flies-and-other-muscoid-diptera

Research

JoVE Journal

Biology

8.5K Views.  ARS-USDA. A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Video: Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is a model for understanding microbial interactions with animal hosts, facilitated by approaches to rear large sample sizes of Drosophila under microorganism-free (axenic) conditions, or with defined microbial communities (gnotobiotic). This work outlines a method for collection of Drosophila embryos, hypochlorite dechorionation and sterilization, and transfer to sterile diet. Sterilized embryos are transferred to sterile diet in 50 ml centrifuge tubes, and developing larvae and adults remain free of any exogenous microbes until the vials are opened. Alternatively, flies with a defined microbiota can be reared by inoculating sterile diet and embryos with microbial species of interest. We describe the introduction of 4 bacterial species to establish a representative gnotobiotic microbiota in Drosophila. Finally, we describe approaches for confirming bacterial community composition, including testing if axenic Drosophila remain bacteria-free into adulthood.

Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

A method for rearing Drosophila melanogaster under axenic and gnotobiotic conditions is presented. Fly embryos are dechorionated in sodium hypochlorite, transferred aseptically to sterile diet, and reared in closed containers. Inoculating diet and embryos with bacteria leads to gnotobiotic associations, and bacterial presence is confirmed by plating whole-body Drosophila homogenates.

The influence of microbes on myriad animal traits and behaviors has been increasingly recognized in recent years. The fruit fly Drosophila melanogaster is ...

Developmental Biology

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are also a useful scientific model to study motion vision and flight dynamics in a controlled laboratory setting. As the larvae develop in organically polluted water, they are useful models for investigating investment in microbial immunity. While large scale commercial breeding for agriculture already occurs, there are no standardized protocols for maintaining captive populations for scientific studies. This is important as commercial captive breeding programs focusing on mass output during peak pollination periods may fail to provide a population that is consistent, stable and robust throughout the year, as is often needed for other research purposes. Therefore, a method to establish, maintain and refresh a captive research population is required. Here, we describe the utilization of an artificial hibernation cycle, in addition to the nutritional and housing requirements, for long term maintenance of Eristalis tenax. Using these methods, we have significantly increased the health and longevity of captive populations of E. tenax compared to previous reports. We furthermore discuss small scale rearing methods and options for optimizing yields and manipulating population demographics.

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

The overall goal of these procedures is to establish, maintain and refresh a captive population of Eristalis tenax in a research setting.

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are ...

Environment

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors remains poorly understood. In Drosophila melanogaster, environmental cues, such as food availability, affect mating activity offering a tractable system to investigate the mechanisms modulating sexual behavior. In D. melanogaster, environmental cues are often sensed via the chemosensory gustatory and olfactory systems. Here, we present a method to test the effect of environmental chemical cues on mating behavior. The assay consists of a small mating arena containing food medium and a mating couple. The mating frequency for each couple is continuously monitored for 24 h. Here we present the applicability of this assay to test environmental compounds from an external source through a pressurized air system as well as manipulation of the environmental components directly in the mating arena. The use of a pressurized air system is especially useful to test the effect of very volatile compounds, while manipulating components directly in the mating arena can be of value to ascertain a compound's presence. This assay can be adapted to answer questions about the influence of genetic and environmental cues on mating behavior and fecundity as well as other male and female reproductive behaviors.

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

We demonstrate an assay to analyze the environmental and genetic cues that influence mating behavior in the fruit fly Drosophila melanogaster.

An individual's sexual drive is influenced by genotype, experience and environmental conditions. How these factors interact to modulate sexual behaviors ...

Behavior

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are delivered using a non-quantitative method. Accurate delivery of known quantities of nucleic acids or other chemicals is critical to measure the efficacy of the compound being tested and to allow reliable comparison between compounds.
Here we provide a reproducible, quantitative microinjection protocol that ensures accurate delivery of specific doses of dsRNA, reducing the mortality typically induced by injection injury. These modifications include the addition of Rhodamine B, a graduated injection needle, and an improved recovery method borrowed from Isoe and Collins. This method allows calculation of dose responses and facilitates comparisons between compounds. Versions of this method have been successfully used on three genera of mosquitoes as well as house flies to assess the reduction in fecundity resulting from gene silencing of ribosomal RNA transcripts.
This protocol provides strategies to reduce several challenges of small insect microinjection. Together, mechanical delivery of dsRNAs accompanied by visual verification, identification of effective locations for delivery, and inclusion of a post-injection recovery period ensure accurate dosing and low injury mortality. This protocol also describes an oviposition bioassay for uniform determination of effects on fecundity.

This protocol describes a microinjection methodology that we have standardized and used for several years to deliver specific quantities of nucleic acids directly to the hemolymph of mosquitoes and house flies. This protocol results in minimal injection mortality and allows dose correlated measurements of fecundity.

Synthetic dsRNAs, used to induce RNA interference, may have dose dependent phenotypic effects. These effects are difficult to define if the dsRNAs are ...

Flying Insect Detection and Classification with Inexpensive Sensors

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow for the development of many useful applications in vector and pest control for both medical and agricultural entomology. Given this, the last sixty years have seen many research efforts devoted to this task. To date, however, none of this research has had a lasting impact. In this work, we show that pseudo-acoustic optical sensors can produce superior data; that additional features, both intrinsic and extrinsic to the insect&#8217;s flight behavior, can be exploited to improve insect classification; that a Bayesian classification approach allows to efficiently learn classification models that are very robust to over-fitting, and a general classification framework allows to easily incorporate arbitrary number of features. We demonstrate the findings with large-scale experiments that dwarf all previous works combined, as measured by the number of insects and the number of species considered.

We proposed a system that uses inexpensive, noninvasive pseudo-acoustic optical sensors to automatically and accurately detect, count, and classify flying insects based on their flying sound.

An inexpensive, noninvasive system that could accurately classify flying insects would have important implications for entomological research, and allow ...

Bioengineering

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine parasitoids known to infect the host during the egg stage1. Second, it has a wide range of potential fruit fly hosts. Perhaps due to its life history, F. arisanus has been a successfully used for biological control of fruit flies in multiple tropical regions2-4. One impediment to the wide use of F. arisanus for fruit fly control is that it is difficult to establish a stable laboratory colony5-9. Despite this difficulty, in the 1990s USDA researchers developed a reliable method to maintain laboratory populations of F. arisanus10-12. There is significant interest in F. arisanus biology13,14, especially regarding its ability to colonize a wide variety of Tephritid hosts14-17; interest is especially driven by the alarming spread of Bactrocera fruit fly pests to new continents in the last decade18. Further research on F. arisanus and additional deployments of this species as a biological control agent will benefit from optimizations and improvements of rearing methods. In this protocol and associated video article we describe an optimized method for rearing F. arisanus based on a previously described approach12. The method we describe here allows rearing of F. arisanus in a small scale without the use of fruit, using materials available in tropical regions around the world and with relatively low manual labor requirements.

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Fopius arisanus is an egg-larval parasitoid of Tephritid fruit flies that is successfully used in biological control of these important tropical pests. We describe here an optimized protocol for rearing F. arisanus on a small scale using readily available materials.

Fopius arisanus (Sonan) is an important parasitoid of Tephritid fruit flies for at least two reasons. First, it is the one of only three opiine ...

Maintenance of a Drosophila melanogaster Population Cage

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life cycle of Drosophila enables collection of large quantities of material from embryos, larvae, pupae and adult flies, in a synchronized way, at a low economic cost. A major strategy for propagating large numbers of flies is the use of a fly population cage. This useful and common tool in the Drososphila community is an efficient way to regularly produce milligrams to tens of grams of embryos, depending on uniformity of developmental stage desired. While a population cage can be time consuming to set up, maintaining a cage over months takes much less time and enables rapid collection of biological material in a short period. This paper describes a detailed and flexible protocol for the maintenance of a Drosophila melanogaster population cage, starting with 1.5 g of harvested material from the previous cycle.

Maintenance of a Drosophila melanogaster Population Cage

This manuscript reports a detailed protocol for culturing, on a regular basis, a population of Drosophila melanogaster using a fly population cage.

Large quantities of DNA, RNA, proteins and other cellular components are often required for biochemistry and molecular biology experiments. The short life ...

Circadian Entrainment of Drosophila Melanogaster

Nearly universal among organisms, circadian rhythms coordinate biological activity to earth&#39;s orbit around the sun. To identify factors creating this rhythm and to understand the resulting outputs, entrainment of model organisms to defined circadian time-points is required. Here we detail a procedure to entrain many Drosophila to a defined circadian rhythm. Furthermore, we detail post-entrainment steps to prepare samples for immunofluorescence, nucleic acid, or protein extraction-based analysis.

Here, we detail how to synchronize Drosophila to a circadian day. This is the first, and most important step necessary for studying biological rhythms and chronobiology.

Nearly universal among organisms, circadian rhythms coordinate biological activity to earth&#39;s orbit around the sun. To identify factors creating this ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

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. ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

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Rearing the Fruit Fly Drosophila melanogaster Under Axenic and Gnotobiotic Conditions

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

A Method to Test the Effect of Environmental Cues on Mating Behavior in Drosophila melanogaster

Reproducible dsRNA Microinjection and Oviposition Bioassay in Mosquitoes and House Flies

Flying Insect Detection and Classification with Inexpensive Sensors

An Optimized Protocol for Rearing Fopius arisanus, a Parasitoid of Tephritid Fruit Flies

Maintenance of a Drosophila melanogaster Population Cage

Circadian Entrainment of Drosophila Melanogaster

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

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