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 &#215; 45 &#215; 45 cm, 18 &#215; 16 mesh aluminum screen, stockinette access</td>
		</tr>
		<tr>
			<td>9 &#215; 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 liter plastic dish pans</td>
			<td>Rubbermaid, Saratoga Springs, NY</td>
			<td>FG2951ARWHT</td>
			<td>Larval pans</td>
		</tr>
		<tr>
			<td>Stockinette, Cotton, 12&#34; X 25 yd 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

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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

Microbial Communities in Nature and Laboratory - Interview

Protocol for Mosquito Rearing (A. gambiae)

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night cycle. For this procedure, you'll need mosquito cages, 10% sterile sucrose solution, paper towels, beaker, whatman filter paper, glass feeders, human blood and serum, water bath, parafilm, distilled water, clean plastic trays, mosquito food (described below), mosquito net to cover the trays, vacuum, and a collection chamber to collect adults.

Protocol for Mosquito Rearing A. gambiae

This video illustrates the general techniques used to rear Anopheles gambiae in the laboratory. The methods for caring for laboratory mosquitoes are demonstrated through all stages of the organism's life cycle from larvae to pupae to blood-feeding adults.

This protocol describes mosquito rearing in the insectary. The insectary rooms are maintained at 28&deg;C and ~80% humidity, with a 12 hr. day/night ...

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite isolated from each other, with largely independent histories and scientific communities. However, the interface between these usually disparate fields is the developmental programs underlying acquisition of functional electrical signaling properties and differentiation of functional chemical synapses during the final phases of neural circuit formation. This interface is a critically important area for investigation. In Drosophila, these phases of functional development occur during a period of &lt;8 hours (at 25&deg;C) during the last third of embryogenesis. This late developmental period was long considered intractable to investigation owing to the deposition of a tough, impermeable epidermal cuticle. A breakthrough advance was the application of water-polymerizing surgical glue that can be locally applied to the cuticle to enable controlled dissection of late-stage embryos. With a dorsal longitudinal incision, the embryo can be laid flat, exposing the ventral nerve cord and body wall musculature to experimental investigation. This system has been heavily used to isolate and characterize genetic mutants that impair embryonic synapse formation, and thus reveal the molecular mechanisms governing the specification and differentiation of synapse connections and functional synaptic signaling properties.

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

This technique exposes the Drosophila embryonic neuromusculature for immunohistochemistry or electrophysiological recording. It is useful for studying early events in neuromuscular development or performing electrophysiology in mutants that cannot hatch.

Drosophila is a premier genetic model for the study of both embryonic development and functional neuroscience. Traditionally, these fields are quite ...

DNA Stable-Isotope Probing (DNA-SIP)

DNA stable-isotope probing (DNA-SIP) is a powerful technique for identifying active microorganisms that assimilate particular carbon substrates and nutrients into cellular biomass. As such, this cultivation-independent technique has been an important methodology for assigning metabolic function to the diverse communities inhabiting a wide range of terrestrial and aquatic environments. Following the incubation of an environmental sample with stable-isotope labelled compounds, extracted nucleic acid is subjected to density gradient ultracentrifugation and subsequent gradient fractionation to separate nucleic acids of differing densities. Purification of DNA from cesium chloride retrieves labelled and unlabelled DNA for subsequent molecular characterization (e.g. fingerprinting, microarrays, clone libraries, metagenomics). This JoVE video protocol provides visual step-by-step explanations of the protocol for density gradient ultracentrifugation, gradient fractionation and recovery of labelled DNA. The protocol also includes sample SIP data and highlights important tips and cautions that must be considered to ensure a successful DNA-SIP analysis.

DNA Stable-Isotope Probing DNA-SIP

DNA stable-isotope probing is a cultivation-independent method to identify and characterize active communities of microorganisms that are capable of utilizing specific substrates. Assimilation of substrate enriched in heavy isotope leads to incorporation of labelled atoms into microbial biomass. Density gradient ultracentrifugation retrieves labelled DNA for downstream molecular analyses.

DNA stable-isotope probing (DNA-SIP) is a powerful technique for identifying active microorganisms that assimilate particular carbon substrates and ...

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 ...

Aseptic Laboratory Techniques: Plating Methods

Microorganisms are present on all inanimate surfaces creating ubiquitous sources of possible contamination in the laboratory. Experimental success relies on the ability of a scientist to sterilize work surfaces and equipment as well as prevent contact of sterile instruments and solutions with non-sterile surfaces. Here we present the steps for several plating methods routinely used in the laboratory to isolate, propagate, or enumerate microorganisms such as bacteria and phage. All five methods incorporate aseptic technique, or procedures that maintain the sterility of experimental materials. Procedures described include (1) streak-plating bacterial cultures to isolate single colonies, (2) pour-plating and (3) spread-plating to enumerate viable bacterial colonies, (4) soft agar overlays to isolate phage and enumerate plaques, and (5) replica-plating to transfer cells from one plate to another in an identical spatial pattern. These procedures can be performed at the laboratory bench, provided they involve non-pathogenic strains of microorganisms (Biosafety Level 1, BSL-1). If working with BSL-2 organisms, then these manipulations must take place in a biosafety cabinet. Consult the most current edition of the Biosafety in Microbiological and Biomedical Laboratories (BMBL) as well as Material Safety Data Sheets (MSDS) for Infectious Substances to determine the biohazard classification as well as the safety precautions and containment facilities required for the microorganism in question. Bacterial strains and phage stocks can be obtained from research investigators, companies, and collections maintained by particular organizations such as the American Type Culture Collection (ATCC). It is recommended that non-pathogenic strains be used when learning the various plating methods. By following the procedures described in this protocol, students should be able to: 
 
&#9679; Perform plating procedures without contaminating media. 
&#9679; Isolate single bacterial colonies by the streak-plating method. 
&#9679; Use pour-plating and spread-plating methods to determine the concentration of bacteria. 
&#9679; Perform soft agar overlays when working with phage. 
&#9679; Transfer bacterial cells from one plate to another using the replica-plating procedure. 
&#9679; Given an experimental task, select the appropriate plating method.

When working with media and reagents used to culture microorganisms, aseptic technique must be practiced to ensure contamination is minimized. A variety of plating methods are routinely used to isolate, propagate, or enumerate bacteria and phage, all of which incorporate procedures that maintain the sterility of experimental materials.

Microorganisms are present on all inanimate surfaces creating ubiquitous sources of possible contamination in the laboratory. Experimental success relies ...

Simple Homemade Tools to Handle Fruit Flies&#8212;Drosophila melanogaster

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in practice, as adult flies fly. Demonstrated here is how to make some simple and cost-effective tools to address difficult issues in the handling of Drosophila. Holes in foam stoppers are made and pipette tips or funnels are inserted into the holes. Flies then move only in one direction into the pipette tip/funnel assemblage, allowing efficient control of the transfer of adult Drosophila into or out of a vial. Existing protocols have been modified for cool-anesthetizing flies by chilling in crushed ice and transferring them onto a cold, hard icepack surface. The icepack is covered with a piece of medical gauze that keeps immobilized flies from the condensed water when examined under a stereomicroscope. The flies are finally euthanized for counting and sorting or discarded by microwaving. A bottle-shaped cage has also been developed for collecting eggs, as well as a labor-saving device and accompanying protocol for cleaning glass culture vials.

Described here is the use of several homemade tools to transfer, chill, and kill adult Drosophila, as well as to clean glass culture vials and collect eggs. These tools are easy to make and are rather efficient in handling Drosophila.

The fruit fly, Drosophila melanogaster, is widely used both in biological research and biology education. Handling adult flies is common but difficult in ...

Dissection of the Endolymphatic Sac from Mice

The study of mutant mouse models of human hearing and balance disorders has unraveled many structural and functional changes which may contribute to the human phenotypes. Although important progress has been done in the understanding of the development and function of the neurosensory epithelia of the cochlea and vestibula, limited knowledge is available regarding the development, cellular composition, molecular pathways and functional characteristics of the endolymphatic sac. This is, in large part, due to the difficulty of visualizing and microdissecting this tissue, which is an epithelium comprised of only one cell layer. The study presented here describes an approach to access and microdissect the endolymphatic sac from the wild-type mouse inner ear at different ages. The result of a similar dissection is shown in a pendrin-deficient mouse model of enlargement of the vestibular aqueduct. A transgenic mouse with a fluorescent endolymphatic sac is presented. This reporter mouse can be used to readily visualize the endolymphatic sac with limited dissection and determine its size. It can also be used as an educational tool to teach how to dissect the endolymphatic sac. These dissection procedures should facilitate further characterization of this understudied part of the inner ear.

This study describes how to dissect the endolymphatic sac from the inner ear of the mouse at different ages. The result of a similar dissection is shown in an Slc26a4-null mouse model of enlargement of the endolymphatic sac. A transgenic mouse with a fluorescent reporter expressed in the endolymphatic sac is presented as a model to readily visualize the endolymphatic sac, and educational tool.

The study of mutant mouse models of human hearing and balance disorders has unraveled many structural and functional changes which may contribute to the ...

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules derived from the parental cells. EVs function as intra- and inter-species communication vectors while also contributing to the interaction between bacteria and host organisms in the context of infection and colonization. Given the multitude of functions attributed to EVs in health and disease, there is a growing interest in isolating EVs for in vitro and in vivo studies. It was hypothesized that the separation of EVs based on physical properties, namely size, would facilitate the isolation of vesicles from diverse bacterial cultures. 
The isolation workflow consists of centrifugation, filtration, ultrafiltration, and size-exclusion chromatography (SEC) for the isolation of EVs from bacterial cultures. A pump-driven tangential flow filtration (TFF) step was incorporated to enhance scalability, enabling the isolation of material from liters of starting cell culture. Escherichia coli was used as a model system expressing EV-associated nanoluciferase and non-EV-associated mCherry as reporter proteins. The nanoluciferase was targeted to the EVs by fusing its N-terminus with cytolysin A. Early chromatography fractions containing 20-100 nm EVs with associated cytolysin A - nanoLuc were distinct from the later fractions containing the free proteins. The presence of EV-associated nanoluciferase was confirmed by immunogold labeling and transmission electron microscopy. This EV isolation workflow is applicable to other human gut-associated gram-negative and gram-positive bacterial species. In conclusion, combining centrifugation, filtration, ultrafiltration/TFF, and SEC enables scalable isolation of EVs from diverse bacterial species. Employing a standardized isolation workflow will facilitate comparative studies of microbial EVs across species.

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

Bacteria secrete nanometer-sized extracellular vesicles (EVs) carrying bioactive biological molecules. EV research focuses on understanding their biogenesis, role in microbe-microbe and host-microbe interactions and disease, as well as their potential therapeutic applications. A workflow for scalable isolation of EVs from various bacteria is presented to facilitate standardization of EV research.

Diverse bacterial species secrete ~20-300 nm extracellular vesicles (EVs), comprised of lipids, proteins, nucleic acids, glycans, and other molecules ...

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 ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

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Microbial Communities in Nature and Laboratory - Interview

Protocol for Mosquito Rearing (A. gambiae)

Harvesting and Preparing Drosophila Embryos for Electrophysiological Recording and Other Procedures

DNA Stable-Isotope Probing (DNA-SIP)

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

Aseptic Laboratory Techniques: Plating Methods

Simple Homemade Tools to Handle Fruit Flies—Drosophila melanogaster

Dissection of the Endolymphatic Sac from Mice

Scalable Isolation and Purification of Extracellular Vesicles from Escherichia coli and Other Bacteria

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