Special thanks to previous B. coprophila stock keepers (Jacob E. Bliss, Paula Bonazinga, Anne W. Kerrebrock, Ingrid M. Mercer, Heidi S. Smith) and research personnel (especially Robert Baird, Michael S. Foulk, Donna Kubai, John M. Urban, Yutaka Yamamoto) for fine-tuning the husbandry protocols. Initial instructions on care of B. coprophila were provided by Helen V. Crouse, Natalia Gabrusewycz-Garcia, Reba M. Goodman, Charles W. Metz, and Ellen Rasch. With gratitude to Yukiko Yamashita and Anne W. Kerrebrock for taking on the Bradysia (Sciara) stock center. Much appreciation to the following people for their helpful preparation of figures: Brian Wiegmann (Figure 1), John M. Urban (Figure 4 upper panel), Laura Ross (Figure 4 lower panel), Yutaka Yamamoto (Figure 5 left panel), Leo Kadota (Figure 7 and Figure 8). Many thanks to Ava Filiss and the Brown University Multidisciplinary Laboratory for their help with photography and filming. Thanks to Robert Baird for comments on this manuscript. Our research and maintenance of B. coprophila has been supported by NIH and NSF, including the most recent support from NIH GM121455 to S.A.G. Further details about B. coprophila are available on the Bradysia (Sciara) Stock Center websites (https://sites.brown.edu/sciara/ and https://sciara.wi.mit.edu) that are currently being constructed.

Bradysia coprophila Lab Protocols and Chromosomal Research

The author has no conflicts of interest to declare.

The protocols presented here for the husbandry of B. coprophila will be useful for scientists who wish to raise this organism in their laboratories for experiments to delve into its unique biological features. The initial description of the feeding method using yeast and mushroom powder sprinkled onto an agar base to maintain B. coprophila29 was used in the Metz laboratory to rear 14 different species of Sciarid flies5. Subsequently, it was observed that the addition of nettle and/or spinach powder further increased the vitality of B. coprophila (Gabrusewycz-Garcia, personal communication). These methods have been successful for the maintenance of related species within the Sciaridae family, including Bradysia impatiens and Lycoriella ingenua that are currently in culture&#160;(Robert Baird, personal communication).
Other methods (such as the alternative feeding methods described below) have been tried to raise B. coprophila, but the protocols described here have been optimized to have the most favorable ratio of larvae per surface area of agar to obtain the most fertile fat adults and minimize the growth of mold. To scale up, mass mating can be done in glass vials as described in protocol 2 above. Alternatively, a few (2-4) adult females can be placed together with twice as many adult males in a flask such as that used to raise Drosophila (disposable 6 oz. = 177.4 mL square bottom Drosophila polypropylene bottle). In both cases, the researcher must be fully confident that the flask contains only all female-producer or all male-producer mothers.
Only feed the larvae since the pupae and adults do not eat. Do not feed the vial if larvae have turned into pupae (a sign of this is when the first early-to-emerge adult flies appear). Once the adults eclose, put the vials into a cooler incubator (e.g., 16 &#176;C) if available as this will allow the adults to live longer. Feed three times per week (e.g., Monday, Wednesday, Friday), increasing the amount of food given per vial as the larvae grow older. Feed generously, and you will be rewarded with fat fertile adults. However, if you feed too much, white mold will appear and that is a sign to reduce the amount of food you are depositing in a vial. Further, if you feed too much, a thick pad of food will develop on top of the agar and make it harder for the adults to emerge (you can remove the pad with a forceps, but be careful to not take larvae out with the pad-it is best to not have to do this at all). Vials with few larvae (marked &#34;few&#34;) need less food. If you feed too little, the larvae will climb the walls of the vial in search of food. Underfed larvae result in small adults that are less fertile.
Alternative feeding methods 
A variety of methods have been attempted to feed larvae just once during the larval stage rather than 3 times per week.&#160;B. coprophila will not grow on Drosophila style food. John Urban (personal communication) tried mixing B. coprophila food in with the agar, but too much mold grew. He found that adding two mold inhibitors (tegosept and propionic acid) in combination and separately, trying several different concentrations, were all toxic to B. coprophila at levels that inhibit mold. The agar should be pH 6-7 (neutral) as B. coprophila gets sick at an acidic pH (as with propianic acid). Alternatively, to obviate thrice weekly feeding, he tried using a spatula or syringe without a needle to dispense a thick yeast paste (Red Star active dry yeast mixed with a little distilled water to moisten it) as a dollop on top of the agar in each vial one week after mating (i.e., about the time the larvae will begin to emerge).
Another method to avoid thrice weekly feeding is to add a living culture of fungi to each vial.&#160;Bath and Sponsler37 reported that&#160;a slanted agar surface with Sabouraud&#39;s medium should be streaked with a fungal culture of the genera Chaetoconidia (best) or else Baplosporangia or Xllescheria. The fungus was grown several days to one week before B. coprophila was introduced. No feeding was needed after this. A variant of this method was also employed by Ellen Rasch (personal communication). In our hands, the vials were too wet with this method and the larvae drowned, but it could be tried again to optimize the number of larvae relative to the vials with live fungi.
Arthur Forer (personal communication) has had some success in rearing B. coprophila the same way as crane flies38. With this approach, pupae were reared on moist papier m&#226;ch&#233;. Subsequently, the adults were mated and the eggs were deposited on fresh moist papier m&#226;ch&#233;. The resulting larvae were kept on papier m&#226;ch&#233; in petri dishes and fed with powdered nettle leaves twice a week. Pupae were put into a cage to repeat the cycle.
Yukiko Yamashita (personal communication) has tried without success to rear B. coprophila on soil, mimicking the conditions where they are found in nature in potted plants and greenhouses with high humidity. However, mold can become a problem when the humidity level is raised. Nonetheless, moist soil has been used with success to raise Pseudolycoriella (previously Bradysia) hygida larvae in plastic boxes with moist soil; they are fed decomposed Ilex paraguariensis leaves, supplemented in late larval life with 1.2% yeast extract, 1.4% cornstarch, 0.8% oatmeal flour, 1.2% agar12. Similarly, the moist soil can be replaced by moist peat moss with crushed kidney beans to raise Sciarid flies39,40.
Still other methods have been employed to maintain laboratory cultures of Bradysia: (i) autoclaved potato to which yeast and dried blood fertilizer are added41; (ii) manure42,43,44 to which dried blood can be added45; (iii) plastic containers with cotton pads and dampened paper towels with ground soybeans46.
Mites 
Mites can be transferred from Drosophila to B. coprophila. To minimize this, it is best to keep B. coprophila in a separate incubator or room, not close to Drosophila stocks. Moreover, do any B. coprophila maintenance work early in the day before handling Drosophila. Mites can also be transferred to B. coprophila from houseplants, so do not keep plants in the same room as B. coprophila. If mites invade the vials, they can be seen as small white spherical organisms crawling on the body of B. coprophila. Chemical treatments that work to destroy mites in Drosophila cannot be used for B. coprophila as the chemicals kill B. coprophila (B. coprophila is also sensitive to organic fumes such as phenol). The only treatment to rid B. coprophila stocks of mites is to manually collect the embryos on an agar plate, examine each for the absence of mites, and then transfer them to fresh agar vials using a fine paintbrush. Cotton-filled gauze plugs and cellulose acetate foam flugs (as used for Drosophila polypropylene vials) both help to prevent the entry of mites into the vials.
Usefulness of the husbandry protocols 
The protocols described here will enable the growing community of scientists to rear B. coprophila as a new/old emerging model organism to study its unique biological features. New laboratory groups are encouraged to join the growing community to maintain and investigate the unique biological features of Bradysia (Sciara).

A complete understanding of biological principles requires the study of many diverse organisms spanning the Tree of Life. Although a broad range of organisms were described through the end of the 19th century, by the middle of the 20th century experimental studies became restricted to a handful of less than a dozen model organisms. With the advent of the genomic era and the goal to sequence genomes from all species in the Tree of Life1, we are now in a position to expand the types of organisms being used for laboratory experiments and to reap the advantage of their diversity. Such expansion of emerging model organisms for experiments has a prerequisite of being able to maintain them in the laboratory. Here, protocols are described for rearing one such emerging new/old model organism.
The majority of animal life on Earth is accounted for by four super-radiations of Insects2. Within the Insects, there are about 158,000 species of Diptera (true flies)3, with about 3000 species in the family Sciaridae (black fungus gnats)4. The fruitfly Drosophila is the most thoroughly studied of the Dipteran flies. The lower Dipteran fly (Nematocera), Bradysia (previously called Sciara) coprophila, diverged 200 million years ago from Drosophila, which is a "higher Dipteran" fly (Brachycera). Therefore, B. coprophila is in a favorable taxonomic position for comparative studies with D. melanogaster (Figure 1). Moreover, B. coprophila has many unique biological features that are worthy of study in their own right5,6,7. Many of these features disobey the rule of DNA constancy in which all cells of an organism have the same DNA content. In B. coprophila, (i) the paternal genome is eliminated on a monopolar spindle in male meiosis I; (ii) there is non-disjunction of the X dyad in male meiosis II; (iii) germ line-limited (L) chromosomes are eliminated from the soma; and (iv) one or two X chromosomes are eliminated in the early embryo depending on the sex of the individual. Chromosome imprinting to distinguish maternal from paternal homologs was first discovered in B. coprophila and is at play for many of these chromosome elimination events. In addition to chromosome elimination, another bypass of DNA constancy occurs via developmentally regulated, locus-specific DNA amplification at the DNA puff loci in larval salivary gland polytene chromosomes. Studies of these unique features require laboratory maintenance of B.coprophila; details of its husbandry are presented here to facilitate such studies.
<img alt="Insect phylogenetic tree diagram; illustrates evolutionary relationships in Diptera and Hymenoptera." class="xfigimg" src="/files/ftp_upload/66751/66751fig01v3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: Phylogeny of Bradysia (Sciara) coprophila. Popular model organisms are indicated in blue font and their taxonomic order in red font. Bradysia and other Sciarid fungus gnats as well as mosquitoes are lower dipteran flies (formerly, suborder Nematocera), whereas Drosophila species are higher dipteran flies (suborder: Brachysera). Information on the left side of the figure is from Misof et al.33; information on the right side is from Bertone et al.34 and Wiegmann et al.2. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Previously, the genus Sciara had the largest number (700) of species for any eukaryote, prompting Steffan to subdivide them8. Subsequently, Shin proposed that the Sciaridae family be subdivided into the subfamily Sciarinae (with six genera including Sciara, Trichosia, and Leptosciarella), the subfamiliy Megalosphyinae (including the genus Bradysia) and three other groups (including Pseudolycoriella)9. The phylogeny of the Sciaridae has been studied further by several groups in recent years9,10,11. Over the past several decades, the names of many organisms in the Sciaridae family have changed12. Although most of the literature spanning more than a century refers to the organism that we study as Sciara coprophila, its current taxonomic name is now Bradysia coprophila (syn. Bradysia tilicola and other synonyms)10. They are found worldwide and are commonly known as fungus gnats since they eat mushrooms and other fungi. They were first described in 1804 by Meigen13 in Europe and subsequently by Johannsen14,15 in North America. B. coprophila was collected at Cold Spring Harbor Laboratory and laboratory stocks were established by Charles Metz in the early 1900s when he was a graduate student at Columbia University with Thomas Hunt Morgan. Thus, the current stocks reflect a century of inbreeding. Similarly, the biology of B. coprophila was further elucidated by decades of cytogenetic studies by Helen Crouse (who did her Ph.D. work with Barbara McClintock).
In the 1930s, Bradysia (Sciara) competed with Drosophila melanogaster as a model system for genetic studies. Despite its many unique biological features, B. coprophila was eclipsed by D. melanogaster as a popular model organism since radiation-induced phenotypic mutations were needed for genetic studies and were easier to achieve in the latter, even though B. coprophila is only slightly more resistant to gamma irradiation than D. melanogaster16. In the modern era of genomics, this is no longer a concern. Since the genome sequence17,18,19 (Urban, Gerbi, and Spradling, data not shown) and methods for transformation20,21 (Yamamoto and Gerbi, data not shown) for B. coprophila have recently become available, the time is now ripe to utilize it as a new/old emerging model system, as seen by the growing community of scientists that have adopted it for their research. This article describes procedures for its laboratory maintenance.
B. coprophila lacks a Y chromosome, and the sex of the offspring is determined by the mother. Females that have the X' ("X-prime") chromosome with a long paracentric inversion will have only daughters, whereas females that are homozygous for the standard (non-inverted) X chromosome will have only sons5 (Figure 2). Sequence information is available for the X' chromosome19, but the molecular mechanism remains to be elucidated on how the X' chromosome determines that offspring will be females. Males never have the X' chromosome, and after fertilization, females are X'X (heterozygous for the X') or XX. Adult X'X females can be distinguished from XX females by phenotypic markers on the wing (Figure 3). X'X females (who will have only daughters) can be recognized by the dominant Wavy (W) wing marker on the X' (as in the HoLo2 stock)22. Alternatively, XX females (who will have only sons) can be recognized by the recessive petite (p) wing marker on the X as in the 91S stock23. In this case, X'Xp females will have full-length (not petite) wings and will have only daughters. The stock 6980 carries a recessive marker on the X chromosome for swollen (sw) veins24, as well as the dominant marker Wavy on the X', allowing two markers for selection for crosses. The degree of expression of Wavy can vary and seems weaker in overcrowded vials where food is limiting or if the temperature becomes too warm. The Wavy wing phenotype is exceptionally strong if larvae are kept in the cold room (4°-8 °C) instead of the usual 21 °C. Although the recessive petite wing marker is not variable and is very easy to identify, 91S stocks are used less frequently since they are less healthy than the HoLo2 stock. The B. coprophila mating schemes are presented here (Figure 2), and described in detail for the HoLo2, 7298, and W14 stocks (Supplemental File 1), the 91S stock (Supplemental File 1), the 6980 stock (Supplemental File 1), and the translocation stocks (Supplemental File 1). The translocation stocks are no longer extant; they were reciprocal translocations of heterochromomeres (H1, H2, and H3) on the short arm of the X that contains the ribosomal RNA genes25,26,27.
<img alt="Genetic inheritance diagram showing sex determination process in germline and zygote development." class="xfigimg" src="/files/ftp_upload/66751/66751fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Mating scheme for B. coprophila. This organism has no Y chromosome (the male soma has a single X); mothers determine the sex of their offspring. XX mothers have only sons and X'X females have only daughters. The X' chromosome has a long paracentric inversion when compared to the X chromosome. The paternal or maternal lineage of the X (or X') chromosome is denoted by blue or red, respectively, in this figure. Sperm are haploid for the autosomes but have two copies of the X chromosome due to non-disjunction in meiosis II. The somatic lineage of early embryos eliminates one or two copies of the paternally derived X if they will be female or male, respectively. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Insect morphology comparison, seven positions, labeled A-G, with wing structures, specimen analysis." class="xfigimg" src="/files/ftp_upload/66751/66751fig03.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Wing phenotypes of B. coprophila. Adult female flies are shown with various wing phenotypes: (A) straight-wing (XX), (B) Wavy wing (X'WX), (C) extreme Wavy wing (X'WX) phenotype that has a shriveled appearance after storing larvae in the cold room, (D) petite wing (XpXp) that is vestigial-like, (E) straight wing with wildtype (XX) and not swollen veins, (F) straight wing with swollen veins (XswXsw) where small bubbles (black arrows) appear on the upper edge of the wing and/or near the tip of both wings, (G) extreme example of swollen where a blister (white arrow) occurs on one or both wings. Males lack the X' chromosome and therefore will never have Wavy wings, but they have petite or swollen wings in the 91S stock or 6980 stock, respectively. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
The goal in stock maintenance is to perform crosses where half the crosses are from female-producing mothers and half the crosses from male-producing mothers to have equal numbers of female and male adults in the next generation for subsequent crosses. However, this also involves planning since the life cycle for males is shorter than for females and adult males emerge up to a week before adult females. Nature accommodates for this asynchrony between the sexes by having the male embryos emerge as larvae 1-2 days after the female larvae from a cross on the same date. However, to ensure that male and female adults are available at the same time for laboratory crosses, female development can be somewhat speeded up by leaving vials with female larvae at room temperature rather than at 21 °C or placing vials with male larvae at slightly cooler temperatures (e.g., 16 °C). Another route that is more foolproof is to do crosses with female-producer mothers on Monday and crosses with male-producer mothers on Friday of the same week. The easiest route, which is what we employ, is to perform crosses with female-producer and male-producer mothers on the same day each week and perform crosses on that day in each consecutive week. In that approach, adult females from a cross on week 1 can be mated with adult males that emerged from a cross on week 2.
The life cycle for female B. coprophila is 5 weeks when raised at 21 °C (Table 1). The length of their life cycle is somewhat longer at cooler temperatures or if they are underfed. The life cycle of male B. coprophila is ~4-4.5 weeks since they pupate 0.5-1 week before females. The end of each larval instar is marked by the shedding of the cuticle, which is triggered by a burst in the level of the steroid hormone ecdysone. Unlike D. melanogaster, which has three larval instars, B. coprophila has four larval instars.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Developmental Stage</td>
			<td>Days post mating (dpm)</td>
			<td>Length of stage (days)</td>
		</tr>
		<tr>
			<td>Egg laid</td>
			<td>1-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Embryo</td>
			<td>1-2 to 7-8</td>
			<td>~7 days</td>
		</tr>
		<tr>
			<td>Larva </td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Larval instars 1, 2, and 3</td>
			<td>7-8 to 16-19</td>
			<td>~10</td>
		</tr>
		<tr>
			<td>4th larval instar pre-eyespot</td>
			<td>16-19 to 21-24</td>
			<td>5</td>
		</tr>
		<tr>
			<td>4th larval instar eye-spot stage</td>
			<td>21-24 to 25-28</td>
			<td>4</td>
		</tr>
		<tr>
			<td>Pupa </td>
			<td>25-28 to 30-33</td>
			<td>5</td>
		</tr>
		<tr>
			<td>Adult</td>
			<td colspan="2">lives 1-2 days at 21 °C if mated or lives 2-3 weeks at 16 °C if not mated. </td>
		</tr>
	</tbody>
</table>
Table 1: Life cycle of female B. coprophila at 21 °C.
B. coprophila can be kept anywhere in the range of 15 °C-25 °C, with development progressing more slowly at cooler temperatures. This insect prefers a humid environment (being found in the soil of houseplants or mushroom beds), so we keep a beaker with deionized water in the incubator. B. coprophila can be kept at room temperature in a metal bread box with a loosely fitting lid and containing a beaker of water, but they go into heat shock at 37 °C28, which is a danger in hot climates. Michael Ashburner and others have tried with little success to store D. melanogaster in the cold to reduce the time needed for stockkeeping. In contrast, a major advantage of B. coprophila is that vials with mid-stage larvae can be stored for up to 3 months on an open shelf in the cold room (4-8 °C) with minimal care of only feeding once a month. They develop extremely slowly in the cold up to the pupal stage and will emerge as fertile adults when the vials are brought back to 21 °C. Presumably, this mimics their overwintering in the wild. This cold-induced developmental stalling might be comparable to that seen after gamma irradiation of mid-stage B. coprophila larvae16, but developmental stalling is not seen in late-stage larvae that have passed the point of no return for their normal developmental progression.

<table><tbody><tr><td>Agar (bacteriological)</td><td>U.S. Biological</td><td>A0930</td><td>https://www.usbio.net;&amp;nbsp;</td></tr><tr><td>CO&lt;sub&gt;2&lt;/sub&gt; FlyStuff Foot Pedal</td><td>Genesee Scientific</td><td>59-121</td><td></td></tr><tr><td>CO2 FlyStuff Blowgun</td><td>Genesee Scientific</td><td>54-104</td><td></td></tr><tr><td>CO2 FlyStuff UltimaterFlypad</td><td>Genesee Scientific</td><td>59-172</td><td>https://www.geneseesci.com</td></tr><tr><td>Ether fume hood</td><td>Labconco&amp;nbsp;</td><td>3955220</td><td>Sits on top of lab bench</td></tr><tr><td>&amp;nbsp; &amp;nbsp;</td><td>&amp;nbsp; &amp;nbsp;</td><td>&amp;nbsp; &amp;nbsp;</td><td>Filter replacement&amp;nbsp; cat # 6961300</td></tr><tr><td>Food: Brewer&amp;rsquo;s Yeast Powder</td><td>Solgar</td><td>&amp;nbsp; &amp;nbsp;</td><td>Obtain from Amazon or health food store</td></tr><tr><td>&amp;nbsp; &amp;nbsp;</td><td>&amp;nbsp; &amp;nbsp;</td><td>&amp;nbsp; &amp;nbsp;</td><td>https://www.solgar.com;</td></tr><tr><td>Food: Nettle Powder&amp;nbsp; (pesticide free)</td><td>Starwest Botanicals</td><td>209460-51</td><td></td></tr><tr><td>Food: Shitake Mushrooms (pesticide free)</td><td>Starwest Botanicals</td><td>202127-5</td><td>https://www.starwest-botanicals.com;</td></tr><tr><td>Food: Spinach Powder ( pesticide free)</td><td>Starwest Botanicals</td><td>209583-5</td><td></td></tr><tr><td>Food: Straw (pesticide free )</td><td>Starwest Botanicals</td><td>209465-3</td><td></td></tr><tr><td>Jar: clear glass, polypropylene lid</td><td>Fisher Scientific:</td><td>FB02911765</td><td>73 mm dia, 89 mm ht (240 ml)&lt;br/&gt; https://www.fishersci.com;</td></tr><tr><td>Needle Probe, wooden handle</td><td>US Geo Supply Inc</td><td>SKU: 4190</td><td>5.75&amp;rdquo; long probe,&lt;br/&gt; stainless steel needle&lt;br/&gt; https://usgeosupply.com; (970)-434-3708</td></tr><tr><td>Vials: glass, preferred:</td><td>Wilmad LabGlass</td><td></td><td></td></tr><tr><td>Wilmad-glass custom vials</td><td></td><td></td><td>28-33 mm inner dia, 33 mm outer dia, 9.5 cm ht&lt;br/&gt; Wilmad: https://www.SP-WilmadLabglass.com</td></tr><tr><td>Vials: glass (cheaper and ok)&amp;nbsp;</td><td>Fisher Scientific</td><td>03-339-26H</td><td>29 mm outer dia, 9.5 cm h&lt;br/&gt; &amp;nbsp;https://www.fishersci.com;&amp;nbsp;</td></tr><tr><td>Vials: glass (a bit narrow)&amp;nbsp;</td><td>Genesee Scientific&amp;nbsp;</td><td>32-201</td><td>24.5 mm outer dia,9.5 cm h&lt;br/&gt; thttps://www.geneseesci.com</td></tr><tr><td>Vials: polypropylene&amp;nbsp;</td><td>Genesee Scientific</td><td>32-114</td><td>28.5 mm outer dia,9.5 cm ht</td></tr><tr><td>Vial Plugs</td><td></td><td></td><td></td></tr><tr><td>roll of non-absorbent cotton</td><td>Fisher Scientific</td><td>22-456881</td><td></td></tr><tr><td>cheesecloth</td><td>Fisher Scientific</td><td>22-055053</td><td>https://www.fishersci.com;</td></tr></tbody></table>

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.

The protocols described here have led to proven success in raising B. coprophila. When recently eclosed fat adults are chosen for mating (Figure 4), over 90% of the crosses can be fertile and yield offspring. The fertility success does vary with different stocks (Table 2). Stock 7298 (X' chromosome with Wavy marker) was the healthiest of the stocks but went through a period of decline, apparently due to activation of DNA mobile elements creating genome rearrangements32. The HoLo2 stock represents a healthy strain derived from 7298, where apparently the genome rearrangements have stabilized, and it has replaced the parental 7298 stock in the stock center. The HoLo2 stock is the one that was used to sequence the B. coprophila genome and is the one used most widely by various laboratory groups. Recently, CRISPR mutagenesis of HoLo2 flies has been used to create the W14 stock with a white eye phenotype to be used for transformation with fluorescent eye markers (Yamamoto and Gerbi, data not shown). The W14 strain is exceptionally robust. The 6980 stock (Wavy wing and swollen vein markers) is somewhat less robust and the 91S stock (petite wing marker) is even less robust.
Successful crosses result in embryos (Figure 5). Embryos undergo elimination of the imprinted paternal X chromosomes at the 7th to 9th cleavage division. In addition, germ-line limited L chromosomes are eliminated from the somatic lineage in embryos at the 5th to 6th cleavage division. The embryos emerge as larvae, which should not be confused with mold that also may be present (Figure 6). Eyespots (anlage to the adult eyes) appear at the second half of the 4th larval instar (Figure 7). The size of the eyespots provides a convenient phenotypic marker for the onset and progression of DNA puff amplification, which is one of only two known examples of naturally occurring developmentally regulated site-specific intrachromosomal DNA (gene) amplification. Subsequently, pupae develop, and the amount of pigment filling their eyes can serve as a developmental marker for meiosis I and II in spermatogenesis (Figure 8) with its unique chromosome behaviors in these divisions.
<img alt="Three insect phenotypes: XwX female, XX female, XO male, shown in genetic variation study diagram." class="xfigimg" src="/files/ftp_upload/66751/66751fig04.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Mating of adult B. coprophila flies. The upper panel shows the three types of adult flies in the HoLo2 stock: female-producing mothers with Wavy wings (X'WX adult females), male-producing mothers with straight wings (XX adult females), and males with straight wings (X0 adult males). Note the pointed ovipositor at the posterior end of the female flies and the hook-shaped clasper at the posterior end of the male flies. The lower panel shows a male and female mating, where the male clasper has grabbed the female's ovipositor. The sperm will be stored in the spermatheca of the female and will fertilize the eggs as they are discharged to the outside. The length of adults is 2.0 mm (males), 2.5 mm (females). <a href="https://www.jove.com/files/ftp_upload/66751/66751fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Drosophila embryogenesis stages, nuclei visualization, developmental process, microscopy images." class="xfigimg" src="/files/ftp_upload/66751/66751fig05v2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: B. coprophila embryos. The upper left panel is a view of embryos using standard light in a dissecting microscope; the nuclei in the syncytial cytoplasm appear as black dots. The panel on the right uses fluorescence microscopy to visualize the propidium iodide-stained nuclei of embryos. The embryos have an average length of 200 microns and an average width of 150 microns. The nuclei for the germ cells cluster at the posterior pole of the embryo as seen for cycles 6 (embryo tilted forward), and 7-9, after which they are interspersed with somatic nuclei. L chromosome elimination in the somatic lineage occurs in cleavage division 5 or 6; X chromosome elimination in the somatic lineage occurs in the 7th, 8th, or 9th cleavage division. Cellularization occurs during interphase of cycle 11. The table at the left shows the average length of time for each division cycle at 22 °C. The table on the left and panel on the right are adapted with permission from de Saint Phalle and Sullivan35. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Fungal infection process; insect cadaver, spore structures, microscopy image, entomopathogenic study." class="xfigimg" src="/files/ftp_upload/66751/66751fig06.jpg" data-altupdatedat="1747911167000" data-alt="Figure 6"> 
Figure 6: B. coprophila embryos emerge as larvae. A cluster of embryos (blue thick arrow) is seen near the ovipositor of the adult female who died after egg-laying. A week after egg-laying, the embryos become young larvae, several of whom are indicated by the black arrows. The newly emerged larvae have a black jaw at the anterior end and a translucent body. They move on top of the agar surface and should not be confused with mold that does not move. The inset shows some mold, with a white filament (red arrow) and a black spore at its tip (green arrow), and is very slightly smaller than the newly emerged larvae. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Larval development stages diagram, highlighting eye formation through microscopic images." class="xfigimg" src="/files/ftp_upload/66751/66751fig07.jpg" data-altupdatedat="1747911167000" data-alt="Figure 7"> 
Figure 7: Eyespot stages of B. coprophila larvae. Eyespots form at the anterior of the larva, just behind the jaw, and are composed of pigment granules that increase in number. The eyespots are the anlage to the adult eye. The upper panel shows larvae that were visualized with a dissecting microscope; the lower panel is a magnified view of the eyespots using a phase contrast microscope to visualize a larva on a microscope slide with a drop of distilled water and a coverslip lightly floating on top. The nomenclature of the eyespot stages is according to Gabrusewycz-Garcia30 where the number of granules is counted in the longest row (e.g., 12) and the number of additional rows excluding the longest row is noted (e.g., 6 for eyespot stage 12x6). Initiation of site-specific DNA amplification in the salivary gland polytene chromosomes begins at eyespot stage 10x5 and is completed at 14x7 when there is a burst of transcription at the locus and expansion of the DNA puffs31. At the subsequent edge-eye/dropped jaw stage, the eyespot granules begin to merge, and they move laterally away from the midline; the length of the larval body shortens. Further, the DNA puffs condense at this stage. It takes about one day at 21 °C to traverse each eyespot stage. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Pupal development stages in insects; images A-D showing progressive metamorphosis stages." class="xfigimg" src="/files/ftp_upload/66751/66751fig08.jpg" data-altupdatedat="1747911167000" data-alt="Figure 8"> 
Figure 8: Development of B. coprophila pupae. During pupation, all larval tissues histolyze except for the nervous system and are replaced by adult tissues that arise by cell divisions of the imaginal discs. The body color changes from white to tan to brown to black. Pigment gradually fills in the pupal eye; meiosis I and II occur in male pupae with eyes that are 1/4 to 1/2 filled with pigment36. Scale bar = 1 mm. <a href="https://www.jove.com/files/ftp_upload/66751/66751fig08large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Stock name</td>
			<td>Markers </td>
			<td>Fertility rate </td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>7298</td>
			<td>Wavy (W) wing</td>
			<td>~75%</td>
			<td></td>
		</tr>
		<tr>
			<td>HoLo2</td>
			<td>Wavy (W) wing</td>
			<td>~90%</td>
			<td>derived from 7298</td>
		</tr>
		<tr>
			<td>W14</td>
			<td>Wavy (W) wing; White eyes</td>
			<td>~95%</td>
			<td>derived from HoLo2</td>
		</tr>
		<tr>
			<td>6980</td>
			<td>Wavy (W) wing; swollen (sw) veins</td>
			<td>~65%</td>
			<td></td>
		</tr>
		<tr>
			<td>91S</td>
			<td>weak Wavy (W) wing; petite (p) wings</td>
			<td>~50%</td>
			<td>the Wavy marker was introduced in a cross to rescue 91S</td>
		</tr>
	</tbody>
</table>
Table 2: Stocks of Bradysia (Sciara) coprophila. Table 1 of Gerbi6 lists these markers and others that are no longer extant. Five translocations (T1, T23, T29, T32, T70) at the centromere end of the X27 are summarized in Figure 8 of Gerbi6 but are no longer extant.
Supplemental File 1: HoLo2 (and 7298 and W14) crosses, 91S cross and rescue, 6980 cross, translocation crosses. <a href="https://www.jove.com/files/ftp_upload/66751/Supplemental%20Materials.zip" target="_blank">Please click here to download this File.</a>

Read this Scientific Article Protocol about Author Spotlight: Exploring Bradysia coprophila's Unique Biology: A Guide to Laboratory Maintenance at JoVE.com

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 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. ...

laboratory-maintenance-lower-dipteran-fly-bradysia-sciara-coprophila

Research

JoVE Journal

Biology

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

Video: Author Spotlight: Exploring Bradysia coprophila's Unique Biology – A Guide to Laboratory Maintenance

Vibrio cholerae: Model Organism to Study Bacterial Pathogenesis - Interview

I am Es.In MindUP, we study biology of human pathogens in non hosts environments. Many human pathogens after exiting their human host have to survive in the environment before infecting another host. So our goal is to better understand processes that control survival of human pathogens in the environment.And our goal is to then better understand transmission dynamics of infectious agents. So we are using Vibrio cholera, the cost of agent of diarrhea, disease, cholera as a model organism. Vibrio cholera is a facultative human pathogen and it has an aquatic lifecycle.Therefore, it provides us great opportunity to study the survival of human pathogen in a natural environment. There are many tools to study this organism. Its genome is completely sequenced and there are micro array representing every gene in the genome.And furthermore, there are many tools that we can manipulate the genome of this organism. My research program takes advantage of these tools and integrates them to study aquatic lifecycle of cholera. So in our experiments, we try to simulate the aquatic environments that organism lives in.And then we would like to know how does this organism so senses fluctuations in its environment with respect to nutrient load, temperature, salinity, or other biological stressors such as pages and tzo? So in aquatic environment, RIA cholera is found either in a free living form or in a surface attached form. And the surface attached forms are known as biofilms, which are simply composed of microorganisms and their extra polymeric substances, which can be exo polysaccharide and proteins and sometimes DNA.And this matrix acts as a glue and hold the organism together. And when grown in biofilm mode, these organisms are actually resistant to fluctuating parameters of their environments and they can be better protected from, for example, phage and proteasome probations. And these are factors that are controlling abundance of microorganisms in the environment.Some of the recent techno technological advance that helped us in our studies are the completion of the genome sequence of the organism and development of microray technology. Using these technologies, we can determine at the whole genome level how this organism responds to major fluctuations in its environment such as nutrient deprivation or changes in temperature. Or one of our major goal is to integrate our whole genome expression studies to a network analysis.So we would like to identify genetic regulatory network that controls response of to fluctuating environmental parameters. We would like to be able to model behavior of organism in the environment. And also we still do not know when in the environment with whom Vibra Cide interacts with.We do not know much about ecology of this organism, and we also do not know relevance of the genes and processes we identified to be important for survival of this organism in the laboratory to the real world. No are the genes that we identified express when Ria cholera is present in natural environment. We also would like to be able to identify and culture organisms that Vibrio Cora interacts with because in the environment, Vibrio Cora is a part of the complex microbial community.So I think understanding biology of Vibrio Cora in natural environment would be a, it's a challenging and it's critical for us to understand biology of the organism. Within the next five years, I hope that we would be able to better characterize op polysaccharide components of biofilms. For example, in Vibrio cholera, we still do not know structure of the op polysaccharide that is holding the cells together.We also do not know all the components of biofilm metrics. Eventually I think it be great to be able to quantify components of the biofilm metrics in natural biofilms of these organisms. I think another major challenge is to study, be OC in natural environment.And I hope that we would be able to take advantage of comparative genomic studies and combine whole genomic expression profiling comparative genomics to better understand evolution of IBR cholera as a pathogen.

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer tissue with minor or no modification, allowing the dietary routing between different trophic levels. However, the FA trophic marker approach is still hampered by the limited knowledge in lipid metabolism of the soil fauna. This study used entirely labelled palmitic acid (13C16:0, 99 atom%) as a tracer in fatty acid metabolism pathways of two widespread soil Collembola, Protaphorura fimata and Heteromurus nitidus. In order to investigate the fate and metabolic modifications of this precursor, a method of isotopologue profiling is presented, performed by mass spectrometry using single ion monitoring. Moreover, the upstream laboratory feeding experiment is described, as well as the extraction and methylation of dominant lipid fractions (neutral lipids, phospholipids) and the related formula and calculations. Isotopologue profiling does not only yield the overall 13C enrichment in fatty acids derived from the 13C labeled precursor but also produces the pattern of isotopologues exceeding the mass of the parent ion (i.e., the FA molecular ion M+) of each labeled FA by one or more mass units (M+1, M+2, M+3, etc.). This knowledge allows conclusions on the ratio of dietary routing of an entirely consumed FA in comparison to de novo biosynthesis. The isotopologue profiling is suggested as a useful tool for evaluation of fatty acid metabolism in soil animals to disentangle trophic interactions.

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

The fatty acid trophic marker approach, i.e., the assimilation of fatty acids as entire molecule and transfer into consumer tissue with no or minor modification, is hampered by knowledge gaps in fatty acid metabolism of small soil invertebrates. Isotopologue profiling is provided as a valuable tool to disentangle trophic interactions.

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer ...

Biochemistry

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly relevant as a model for the studies of osmoregulatory mechanisms because of its extreme tolerance to low salinity. It is also widely used as a model of settling biology, in particular in relation to antifouling research. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae for studies. A protocol for the all-year-round culturing of B. improvisus has been developed and a detailed description of all steps in the production line is outlined (i.e., the establishment of adult cultures on panels, the collection and rearing of barnacle larvae, and the administration of feed for adults and larvae). The description also provides guidance on troubleshooting and discusses critical parameters (e.g., the removal of contamination, the production of high-quality feed, the manpower needed, and the importance of high-quality seawater). Each batch from the culturing system maximally yields roughly 12,000 nauplii and can deliver four batches in a week, so up to almost 50,000 larvae per week can be produced. The method used to culture B. improvisus is, probably, to a large extent also applicable to other marine invertebrates with free-swimminglarvae. Protocols are presented for the dissection of various tissues from adults as well as the production of high-quality RNA for studies on gene expression. It is also described how cultured adults and reared cyprids can be utilized in a wide array of experimental designs for examining gene expression in relation to external factors. The use of cultured barnacles in gene expression is illustrated with studies of possible osmoregulatory roles of Na+/K+ ATPase and aquaporins.

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

The barnacle Balanus (Amphibalanus) improvisus is a model for studying osmoregulation and antifouling. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae. Here, a protocol for the all-year-round culturing of B. improvisus is described, including the production of larvae. The use of cultured barnacles in gene expression studies is illustrated.

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly ...

Environment

Microinjection of Medaka Embryos for use as a Model Genetic Organism

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1, as in zebrafish and the mouse. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions.
To take advantage of medaka fish whose embryos are transparent and develop externally, microinjection is an essential technique to inject cell-tracers for labeling cells, mRNAs or anti-sense oligonucleotides for over-expressing and knocking-down genes of interest, and DNAs for making transgenic lines.

Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows ...

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and behavioural research. As a vertebrate, zebrafish share considerable genetic sequence similarity with humans and are being used as an animal model for various human disease conditions. The advantages of zebrafish in comparison to other common vertebrate models include high fecundity, low maintenance cost, transparent embryos, and rapid development. Due to the spur of interest in zebrafish research, the need to establish and maintain a productive zebrafish housing facility is also increasing. Although literature is available for the maintenance of a zebrafish laboratory, a concise video protocol is lacking. This video illustrates the protocol for regular housing, feeding, breeding and raising of zebrafish larvae. This process will help researchers to understand the natural behaviour and optimal conditions of zebrafish husbandry and hence troubleshoot experimental issues that originate from the fish husbandry conditions. This protocol will be of immense help to researchers planning to establish a zebrafish laboratory, and also to graduate students who are intending to use zebrafish as an animal model.

Regular Care and Maintenance of a Zebrafish Danio rerio Laboratory: An Introduction

This protocol outlines regular maintenance and care to maintain optimal conditions for zebrafish husbandry. The video illustrates the protocol for system maintenance, regular housing, feeding, breeding, and raising of zebrafish larvae.

This protocol describes regular care and maintenance of a zebrafish laboratory. Zebrafish are now gaining popularity in genetics, pharmacological and ...

Collection, Isolation and Enrichment of Naturally Occurring Magnetotactic Bacteria from the Environment

Magnetotactic bacteria (MTB) are aquatic microorganisms that were first notably described in 19751 from sediment samples collected in salt marshes of Massachusetts (USA). Since then MTB have been discovered in stratified water- and sediment-columns from all over the world2. One feature common to all MTB is that they contain magnetosomes, which are intracellular, membrane-bound magnetic nanocrystals of magnetite (Fe3O4) and/or greigite (Fe3S4) or both3, 4. In the Northern hemisphere, MTB are typically attracted to the south end of a bar magnet, while in the Southern hemisphere they are usually attracted to the north end of a magnet3,5. This property can be exploited when trying to isolate MTB from environmental samples.
One of the most common ways to enrich MTB is to use a clear plastic container to collect sediment and water from a natural source, such as a freshwater pond. In the Northern hemisphere, the south end of a bar magnet is placed against the outside of the container just above the sediment at the sediment-water interface. After some time, the bacteria can be removed from the inside of the container near the magnet with a pipette and then enriched further by using a capillary racetrack6 and a magnet. Once enriched, the bacteria can be placed on a microscope slide using a hanging drop method and observed in a light microscope or deposited onto a copper grid and observed using transmission electron microscopy (TEM).
Using this method, isolated MTB may be studied microscopically to determine characteristics such as swimming behavior, type and number of flagella, cell morphology of the cells, shape of the magnetic crystals, number of magnetosomes, number of magnetosome chains in each cell, composition of the nanomineral crystals, and presence of intracellular vacuoles.

We demonstrate a method to collect magnetotactic bacteria (MTB) that can be applied to natural waters. MTB can be isolated and enriched from sediment samples using a relatively simple setup that takes advantage of the bacteria's natural magnetism. Isolated MTB can then be examined in detail using both light and electron microscopy.

Magnetotactic bacteria (MTB) are aquatic microorganisms that were first notably described in 19751 from sediment samples collected in salt marshes of ...

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster. In contrast to the syncytial cleavages of Drosophila, the early cleavages of Parhyale are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

The amphipod Parhyale hawaiensis is a promising model organism for studies of crustacean embryology and comparative arthropod development and evolution. This protocol describes a method for manual removal of single blastomeres from early cleavage stage embryos of Parhyale.

The amphipod Parhyale hawaiensis is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale has emerged as a ...

Measurement of Survival Time in Brachionus Rotifers: Synchronization of Maternal Conditions

Rotifers are microscopic cosmopolitan zooplankton used as models in ecotoxicological and aging studies due to their several advantages such as short lifespan, ease of culture, and parthenogenesis that enables clonal culture. However, caution is required when measuring their survival time as it is affected by maternal age and maternal feeding conditions. Here we provide a protocol for powerful and reproducible measurement of the survival time in Brachionus rotifers following a careful synchronization of culture conditions over several generations. Empirically, poor synchronization results in early mortality and a gradual decrease in survival rate, thus resulting in weak statistical power. Indeed, under such conditions, calorie restriction (CR) failed to significantly extend the lifespan of B. plicatilis although CR-induced longevity has been demonstrated with well-synchronized rotifer samples in past and present studies. This protocol is probably useful for other invertebrate models, including the fruitfly Drosophila melanogaster and the nematode Caenorhabditis elegans, because maternal age effects have also been reported in these species.

Measurement of Survival Time in Brachionus Rotifers: Synchronization of Maternal Conditions

Rotifers are microscopic zooplankton used as models in ecotoxicological and aging studies. Here we provide a protocol for powerful and reproducible measurement of survival time in Brachionus rotifers. Synchronization of culture conditions over several generations is of particular importance because maternal condition affects life history of offspring.

Rotifers are microscopic cosmopolitan zooplankton used as models in ecotoxicological and aging studies due to their several advantages such as short ...

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.

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

Culturing Larvae of Crepidula fornicata in Laboratory and Field Conditions

Laboratory and Field Culture of Larvae of The Slipper Limpet, Crepidula fornicata

The calyptraeid gastropod mollusk, Crepidula fornicata, has been widely used for studies of larval developmental biology, physiology, and ecology. Brooded veliger larvae of this species were collected by siphoning onto a sieve after natural release by adults, distributed into the culture at a density of 200/L, and fed with Isochrysis galbana (strain T-ISO) at 1 x 105 cells/mL. Shell growth and acquisition of competence for metamorphosis were documented for sibling larvae reared in ventilated 800 mL cultures designed for equilibration to ambient air or to defined atmospheric gas mixtures. Contrasting with these laboratory culture conditions; growth and competence data were also collected for larvae reared in a 15 L flow-through ambient seawater mesocosm located in a field population of reproductive adults. Growth rates and timing of metamorphic competence in the laboratory cultures were similar to those reported in previously published studies. Larvae reared in the field mesocosm grew much faster and metamorphosed sooner than reported for any laboratory studies. Together, these methods are suited for exploring larval development under predetermined controlled conditions in the laboratory as well as under naturally occurring conditions in the field.

Author Spotlight: Bridging the Gap Between Field Observations and Lab Manipulations in Larval Ecology Research

The paper presents methods for larval culture of the gastropod Crepidula fornicata in a small-volume laboratory-scale system and in an ambient-seawater mesocosm system that can be deployed in the field.

The calyptraeid gastropod mollusk, Crepidula fornicata, has been widely used for studies of larval developmental biology, physiology, and ecology. Brooded ...

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

Podsumowanie

Przeglądaj więcej filmów

Rozdziały w tym wideo

Vibrio cholerae: Model Organism to Study Bacterial Pathogenesis - Interview

Fatty Acid ¹³C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Microinjection of Medaka Embryos for use as a Model Genetic Organism

Regular Care and Maintenance of a Zebrafish (Danio rerio) Laboratory: An Introduction

Collection, Isolation and Enrichment of Naturally Occurring Magnetotactic Bacteria from the Environment

Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis

Measurement of Survival Time in Brachionus Rotifers: Synchronization of Maternal Conditions

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

Laboratory and Field Culture of Larvae of The Slipper Limpet, Crepidula fornicata