Name: laboratory rearing of stable flies and other muscoid diptera
Uploaded: 2018-08-03T12:30:00.000+00:00
Duration: 7 min
Description: Watch this Scientific Journal Video about Laboratory Rearing of Stable Flies and Other Muscoid Diptera at JoVE.com

Nous tenons à remercier Anthony Weinhold et les nombreux étudiants qui ont travaillé avec nous pendant des années pour le support technique ainsi que des suggestions pour améliorer notre insecte procédures d’élevage.

1. oeuf Collection (Figure 1)
<ol><li>Préparer une tasse mentionnÃ, placez un bout de tissu en ~ 500 mL bécher rempli d’eau tiède (environ 40 ° C). Chevaucher les côtés de la coupe et fixez avec un élastique. Plier l’extrémité libre de la toile de retour sur le dessus de la coupe.</li>
	<li>Place coupe mentionnÃ en cage de mouches de 8 à 10 jours de vieux pour la femelle gravide ~ 2 h stable vole sera pondent leurs oeufs sur la toile.</li>
	<li>Retirer mentionnÃ tasse de cage et rinçage oeufs hors de la toile mentionnÃ dans une petite casserole avec pissette de lavage.</li>
</ol>2. préparer la moule d’élevage larvaire
<ol><li>
Préparer le support (quantités pour 1 pan d’élevage)
	<ol>
<li>Combiner le son de blé (500 g), de copeaux de bois (200 g) et de farine de poisson (115 g) dans la casserole en plastique plat 10 L et bien mélanger.</li>
		<li>Ajouter le bicarbonate d’ammonium (50 g) à l’eau (~ 25 ° C, 1 600 mL) et mélanger jusqu'à dissolution.</li>
		<li>Ajouter la solution de l’eau aux ingrédients secs et remuer pour n’assurer aucune matière unwetted. Niveau moyen à pan, mais ne pas compresser.</li>
	</ol>
</li>
	<li>
Ajouter les oeufs de mouche stables
	<ol>
<li>Faire un sillon peu profond au milieu de la longueur de la casserole.</li>
		<li>Déposer 1 mL d’oeufs de mouche stables (~ 8 000 oeufs) dans le sillon avec la pipette de l’oeuf (Figure 2). NOTE : Pipette oeuf fait en coupant une pipette graduée en plastique au point désiré, insérant 100 mesh écran et pipette collant ensemble.</li>
		<li>Couvrir les oeufs d’une mince couche de médium pour empêcher la dessiccation.</li>
		<li>Couvrir la casserole avec taie d’oreiller, fermer avec un élastique et une étiquette. Posez les poêlons individuels en salle larvaire (23 ± 2 ° C, 30 à 50 % d’humidité relative et 12:12 [DL] h photopériode). Remarque : Le développement larvaire prend 10 à 14 jours suivant la date de la ponte.</li>
	</ol>
</li>
</ol>3. traitement nymphe, les larves se déplacent jusqu’au bord du milieu 7 à 9 jours après la ponte et Pupariate par jours 13 ou 14
<ol><li>Scoop puparia, principalement située sur les bords de la casserole sous la croûte qui se forme sur la surface du milieu, dehors et placé dans une casserole propre.</li>
	<li>Remplissez la cuve, avec puparia, entre ½ et ¾ pleine d’eau. Briser les agrégats de puparia et moyennes. Les composants bran et protéines de blé du milieu évier tout en puparia > 1 jour vieux flotteur19.</li>
	<li>Enlever les matières flottantes, puparia et certaines composantes moyennes avec une passoire et laver à travers une série de tamis (#5, #7, #12, #20) pour éliminer le milieu reste. Le tamis #12 recueille le puparia et le #20 garde des déchets solides de pénétrer dans le drain. Lavage puparia entre tamis avec un pulvérisateur de l’évier.</li>
	<li>Rincez puparia, dans le tamis #12, dans une casserole propre. Remplir le bac à ½ plein d’eau. Versez puparia flottant dans une passoire et laisser les excès d’eau s’écouler.</li>
	<li>Transfert égouttés puparia à écran de séchage avec ventilateur et laisser à sec (Figure 3).</li>
</ol>4. plateau nymphe, une méthode Alternative pour la collecte Puparia est une étagère Pupal18
NOTE : Le plateau est issu d’un morceau de coupe en plastique de l’extrémité d’un pan de plat 10 L (10,2 cm de hauteur x 10,2 cm de large x 31,9 cm de long).
<ol><li>Préparer le milieu comme décrit en 2.1. Support conique dans le moule de ~2.5 cm profond sur un bout de ~7.5 cm profond sur l’autre.</li>
	<li>
Poser le plateau sur le milieu dans la partie peu profonde de la moule d’élevage et ruban pour éviter les larves de ramper à côté de l’étagère (Figure 4).
	<ol>
<li>Saturer l’éponge (~14.5 x 9,0 x 4,5 cm3, cellulose régénérée) et l’envelopper dans un chiffon gorgés d’eau (30,5 x 42 cm2, coton) et la place sur le plateau environ un demi pouce du milieu.</li>
	</ol>
</li>
	<li>
Ajouter les oeufs comme indiqué au point 2.2 et procédez comme ci-dessus.
	<ol>
<li>Vérifier quotidiennement les casseroles pour vous assurer que l’éponge reste humide.</li>
	</ol>
</li>
	<li>Rinçage puparia du plateau, éponge et chiffon dans un plat propre casserole et versez dans une passoire 14 jours après la ponte.</li>
</ol>5. contrôle de la qualité
<ol><li>Peser tous le puparia produite dans casserole (poids total).</li>
	<li>Isoler puparia 100, peser et placé dans une boîte de Pétri de 9 cm.</li>
	<li>Nombre et sexe adultes ~ 5 – 8 jours plus tard afin de déterminer le taux d’émergence et sex-ratio. Stocker Pétri avec des adultes a émergé dans un congélateur pour le comptage à une date ultérieure si nécessaire. Les mouches sont sexés par la forme de leurs yeux et de la largeur de la plaque fronto-orbital20 ou sous faible grossissement, les organes génitaux.</li>
	<li>Poids total record, poids de 100 puparia et le nombre de levées mâles et femelles adultes.</li>
</ol>6. préparation de sang
<ol><li>Recueillir le sang de bovin fraîche d’un abattoir local dans des seaux de 19 L contenant 70 g de citrate de sodium dihydraté tribasique dans 500 mL d’eau. Remuez vigoureusement pendant 5 à 10 min de sang et retournés au laboratoire.</li>
	<li>Souche sang à travers une passoire pour éliminer des caillots et partitionné en conteneurs de 2 L, étiquette avec la date de la collection et le magasin dans un congélateur (-20 ° C).</li>
	<li>Conteneur de retirer du congélateur et le placer dans le réfrigérateur 2 jours avant nécessaire. Sang peut être utilisé pour environ 2 semaines une fois décongelés et stockés au congélateur jusqu'à 1 an.</li>
</ol>7. adult Maintenance
<ol><li>Préparer les cages, fond de cage de ligne avec du papier de boucher pour faciliter le nettoyage.</li>
	<li>Placer 50 g des pupes de mouches à secs stables (~ 3 500) dans chaque cage. Eclose adultes quelques heures à quelques jours après la transformation pupale.</li>
	<li>Place un sang frais imbibés serviette féminine sur le dessus de chaque cage pour se nourrir. Commence à se nourrir dans les 24h d’eclosion adulte et répéter tous les jours jusqu'à un jour avant la ponte (9 à 10 jours après la levée).</li>
	<li>Après la ponte (section 1), placez la cage dans le congélateur pendant 4 à 8 h et puis nettoyer en retirant le papier, le rinçage des excréments sur les côtés et le plancher avec de l’eau chaude et frotter toutes les surfaces avec un détergent à lessive et eau de Javel. Rincer les cages à l’eau chaude et laisser pour sécher.</li>
</ol>

<ol>
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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. 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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. 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Les auteurs n’ont rien à divulguer.

Les mouches stables se trouve dans une large variété de substrats dans la nature et peuvent être élevés dans de nombreux types de médias dans le laboratoire. Nous avons utilisé des copeaux de bois et de la vermiculite comme agents gonflants. La vermiculite a bien fonctionné, mais fait séparation puparia d’élevage moyennement difficile et était cher (~$0.60/pan). Peut-être à cause de la rigueur supplémentaire de séparation puparia du milieu, eclosion était également plus faible avec la vermiculite, 57 %...

Les mouches stables, Stomoxys calcitrans (L.), sont des mouches hématophages dont les piqûres douloureuses perturbent le comportement de pâturage du bétail, causent des douleurs et souffrances aux animaux de compagnie et perturbent les activités récréatives humaines dans le monde entier. Immatures mouches stables se développent dans la fermentation de matière végétale, souvent contaminé par des déchets animaux. Cultures et modification des pratiques agronomiques ont produit des graves éclosions de mouches stables dans des résidus de cultures, légumes en Australie1, canne à sucre au Brésil2et ananas au Costa Rica,3. Bien que seulement 14 mouches stables par animal sont considérés comme le seuil économique4, observations de plus de 2 000 mouches par animal ont été faites au cours de récentes éclosions5. Ces niveaux d’infestation réduire la productivité hôte à près de zéro et peut causer la mortalité6. À la suite de flambées associées sur le plan agronomique, mouches stables reçoivent le regain d’intérêt et demande des colonies de laboratoire a augmenté de façon spectaculaire.
En ce qui concerne tous les holometamorphic insectes, mouches stables obtenir tous les éléments nutritifs nécessaires à sa croissance durant le stade immature ou larvaire. Par conséquent, un élément important d’un système d’élevage est l’alimentation larvaire ou le substrat. Les larves de mouches stables ont été observés dans un large éventail de substrats dans la zone7 et ils dépendent de la communauté microbienne du substrat8,9. Substrats naturels de larves sont principalement composées de décomposition ou de fermentation matériel végétatif souvent contaminés par des déchets azotés.
Pour l’élevage de laboratoire, des substrats de larves mouches stables sont généralement composées d’un matériau végétal et une source d’azote supplémentaire. Nombreux matériaux ont été utilisés pour un regime de larve mouche stable. La première alimentation larvaire a imité substrats naturels et inclus la fermentation paille d’avoine et de10,du fumier de cheval ou vache11. Sources de glucides incluent blé bran12,13,14, luzerne repas12,13,14 , et une formulation commerciale développée par Chemical Specialties Manufacturers Association (CSMA, 33 % du son de blé, farine de luzerne de 27 %, 40 % de levure granulés)13,14,15,16. Sources d’azote comprennent de suspension de levure12, de farine de poisson et de bicarbonate d’ammonium17. Gonflement des matériaux inertes figurent souvent dans les régimes dont avoine coques12, bagasse13, vermiculite16, copeaux de bois13,18 et granulés coques d’arachides14.
Des principaux objectifs de l’élevage de laboratoire sont de produire un produit qui est aussi physiologiquement similaire à « type sauvage » que possible, afin que des expériences en laboratoire produira des résultats reflétant celles des populations naturelles. Cela nécessite que l’élevage et la sélection être minimisé pour maintenir génétique, diversité et ressources nutritionnelles être comparables à ceux dans le domaine. Les objectifs secondaires sont à minimiser le travail et les dépenses. Une composante majeure de la réduction des dépenses est l’utilisation de composants d’alimentation disponibles localement. La mouche stable SYSTEME présenté d’élevage a été développée afin d’atteindre ces objectifs.

<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

Les mouches stables, Stomoxys calcitrans, sont des ravageurs importants des animaux, les humains, animaux de compagnie et la faune dans le monde entier. Durant les dernières 20 années +, changements dans les pratiques agronomiques a entraîné de graves flambées de mouches stables dans plusieurs pays. Ces flambées perturbé la production animale et humaine loisirs aboutissant à des demandes de la population pour accroître les efforts de recherche et de la gestion de cet organisme nuisible. Une procédure simple et peu coûteuse pour l’élevage des mouches stables pour des études en laboratoire est présentée. La procédure utilise des fournitures, des équipements et des composants de l’alimentation disponible localement. La procédure peut être adaptée pour l’élevage des autres mouches muscoid dont mouche faciale (Musca autumnalis), mouche des cornes (Haematobia irritans) et mouche domestique (Musca domestica). La procédure produit puparia mouche stable, avec une moyenne de 12,5 mg et environ 35 % des œufs à la survie des adultes. Environ 3000 mouches sont produites dans chaque casserole.

Les larves pupariate 10 – 14 jours et les adultes émergent de 14 à 16 jours après la ponte. Générationnel, oeuf d’oeuf, est ~ 24 jours. Données d’élevage pour mai 2013 à janvier 2017 avec trois différents agents gonflants et deux colonies sont présentées à la Figure 5. Cottonwood a donné le meilleur rendement, 3867 ±1442 (<img alt="Equation" src="/files/ftp_upload/57341/57341eq1.jpg"> </img> ± écart-type) pesant 12,5 mg 1,6 avec 74 ±19 ...

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

Une procédure pour l’élevage des mouches stables (Stomoxys calcitrans) est présentée. La procédure utilise des matériaux disponibles localement pour les composants de l’alimentation, de matériel et de fournitures.

Laboratoire d’élevage de mouches Stable et autres diptères Muscoid

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 vues.  ARS-USDA. Une procédure pour l’élevage des mouches stables (Stomoxys calcitrans) est présentée. La procédure utilise des matériaux disponibles localement pour les composants de l’alimentation, de matériel et de fournitures.

Vidéo: Laboratory Rearing of Stable Flies and Other Muscoid Diptera

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.

Élevage et entretien à long terme des syrphes Eristalis tenax pour études

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

Recherche

Environnement

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.

DsRNA reproductible Microinjection et bioessais sur la ponte chez les moustiques et les mouches domestiques

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.

Insectes volants Détection et classification des capteurs peu coûteux

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

Bio-ingénierie

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.

Un protocole optimisé pour l&#39;élevage Fopius arisanus, Un parasitoïde des mouches des fruits Tephritidae

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 ...

Biologie

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.

Le maintien d&#39;un<em&gt; Drosophila melanogaster</em&gt; 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.

Formation circadienne de Drosophila Melanogaster

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

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).

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.

Maintenir les cultures de laboratoire de Gryllus bimaculatus, un modèle orthoptère polyvalent pour l’agriculture des insectes et la physiologie des invertébrés

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

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.

Essai biologique d’application topique pour quantifier la toxicité des insecticides pour les moustiques et les mouches des fruits

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.

Maintenance en laboratoire de la mouche des diptères inférieurs Bradysia (Sciara) coprophila : un organisme modèle nouveau/ancien émergent

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

Drosophila Maintenance

Drosophila melanogaster, commonly known as fruit flies, are a frequently used model organism for life science research. Although starting a collection of these critters may seem as easy as leaving a banana on your kitchen counter for too long, a productive fly colony in the lab requires careful husbandry and maintenance.
This video demonstrates the necessary steps for maintaining a healthy fly stock. The overview begins with the preparation and storage of the yeast and sugar-containing media on which flies feed. Next, the vessels most commonly used for housing Drosophila are shown, as well as how and when to move flies between these containers. Finally, the presentation also includes examples of the ways in which housing and feeding conditions are manipulated for biological experiments.

Maintenance and Husbandry of Drosophila melanogaster

Maintien et élevage de la drosophile

Drosophila melanogaster, commonly known as fruit flies, are a frequently used model organism for life science research. Although starting a collection of ...

Laboratoire d’élevage de mouches Stable et autres diptères Muscoid

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