This work was supported by a strategic award from the Wellcome Trust for the Centre for Immunity, Infection and Evolution (http://ciie.bio.ed.ac.uk; grant reference no. 095831). PFV was supported by a Branco Weiss fellowship (https://brancoweissfellowship.org/) and a Chancellor&#8217;s Fellowship (School of Biological Sciences, University of Edinburgh); JASJ was supported by a NERC E3 DTP PhD studentship.

1. Maintain Flies
<ol>
	<li>Maintain flies in 23 mL plastic vials containing 7 mL of freshly made Lewis medium (modified from reference31; 1 L triple distilled H2O, 6.1 g agar, 93.6 g brown sugar, 68 g maize, 18.7 g instant yeast, 15 mL Tegosept anti-fungal agent) in incubators at 25 &#177; 1 &#176;C, in a 12 h:12 h light:dark cycle with ~60% humidity. Plug the vials with non-absorbent cotton wool.</li>
	<li>After every 14 days, transfer 20&#8211;30 adults to a new food vial, with instant, dry yeast added to the surface, for 2&#8211;3 days to allow egg-laying to occur. After this time period, ensure that the eggs are visible on the surface of the food. Remove adult flies. 
	NOTE: This keeps flies in the vials as single generation, age-matched populations.</li>
	<li>Leave the eggs to develop. 
	NOTE: At 25 &#176;C, adult flies start to eclose from pupae on day 11 and continue over days 12&#8211;14.</li>
</ol>
2. Prepare Experimental Flies
<ol>
	<li>Collect the eggs of the parent generation in a population/embryo collection cage on a 75 mL apple-agar plate (1 L triple distilled H2O, 30 g agar, 33 g sucrose, 330 mL apple juice, 7 mL Tegosept anti-fungal agent) with a yeast paste spread (mix dry yeast with water to a peanut butter-like consistency). Add water-soaked cotton wool to the cage to provide moisture. 
	NOTE: To avoid confounding effects caused by differences in larval rearing density, it is important that experimental flies in different vials are reared in similar densities. The above step is performed to avoid confounding effects.</li>
	<li>Incubate for 24 h at 25 &#176;C in a 12 h:12 h light:dark cycle until egg-laying has occurred. If there are too few eggs after 24 h, provide a longer habituation period. Replace apple-agar plates and allow egg-laying to occur for a further 24 h.</li>
	<li>Take egg-laden apple-agar plates from the population cage. Remove the remaining yeast paste and any dead flies from the agar&#8217;s surface.</li>
	<li>Submerge the agar in 20 mL of 1x phosphate-buffered saline (PBS) and gently dislodge the eggs from the apple-agar with a fine paintbrush. While suspended in PBS, transfer the eggs to a 50 mL centrifuge tube and leave for 5 min so the eggs sink to the bottom. 
	NOTE: Most eggs are found on the outer edge of the agar.</li>
	<li>Remove by cutting the bottom 4 mm of a p1000 filtered pipette tip and use the pipette tip to draw 1 mL of solution, taken from the bottom of the 50 mL centrifuge tube. Transfer this to a 1.5 mL microcentrifuge tube and allow it to settle. 
	NOTE: When pipetting up eggs, snap-releasing the plunger is more efficient than a gentle release.</li>
	<li>Remove by cutting the bottom 4 mm of a p20 filtered pipette tip. Set the pipette to a desired volume and draw from the bottom of the microcentrifuge tube. 
	NOTE: With practice, a volume of 5 &#181;L contains roughly 100 eggs.</li>
	<li>Dispense the collected eggs onto the food and leave them to develop for the required amount of time.</li>
</ol>
3. Bacterial Culture
<ol>
	<li>To grow P. entomophila and P. aeruginosa cultures, inoculate 10 mL of Luria-Bertani (LB) broth with 100 &#181;L of a frozen bacterial stock at 30 &#176;C (P. entomophila) and 37 &#176;C (P. aeruginosa), respectively. Shake at 150 rpm overnight. Ensure that the bacterial culture reaches the saturation phase.</li>
	<li>To ensure the bacteria used for inoculating the flies are in the exponential phase and rapidly replicating, inoculate the overnight culture into a new subculture, of a desired volume, the following morning. Ensure that the pre-inoculum is 10% of the total volume of the subculture culture. 
	NOTE: Oral infection requires high . It is therefore necessary to grow a substantial volume of bacterial culture so that enough inoculation culture can be produced for the desired dose and experimental size. Calculate how much subculture is needed to produce the required infectious doses using the equation MsVs = MiVi, where M represents a culture&#8217;s optical density measured at 600 nm (OD600) value and V represents its volume. Subscript letters refer to whether the culture is used as a subculture (s) or an infectious dose (i).</li>
	<li>Grow this subculture in a 2 L conical flask in a volume such that the subculture&#8217;s surface falls (at most) just above the beginning of the flask&#8217;s slope. Do not fill above this mark as it will stunt the growth of bacteria.</li>
	<li>Ensure the bacteria in this subculture are in the exponential growth phase by measuring the OD every 30 min. 
	NOTE: This occurs after 3&#8211;5 h, where the subculture reaches an OD600 between 0.6&#8211;0.8.</li>
	<li>Pour equal volumes of this subculture across 50 mL centrifuge tubes and spin the subculture at 2,500 x g for 15 min at 4 &#176;C to pellet the bacteria. Once pelleted, remove and then spin the supernatant again at the above conditions to confirm the removal of the vast majority of bacteria. 
	NOTE: A pellet of negligible size (smaller than 1 mm in height) confirms this.</li>
	<li>Combine the bacterial pellets of the separate tubes by re-suspending them in 5 mL of subculture supernatant and recombining these solutions in a single 50 mL tube. Spin this concentrated culture at 2,500 x g for 15 min at 4 &#176;C to pellet the bacteria.</li>
	<li>Remove the supernatant and re-suspend the final bacteria pellet in 5% sucrose water solution. Check the OD and adjust to the desired infectious dose (OD600 = 100 for P. entomophila8 and OD600 = 25 for P. aeruginosa16,28), by re-suspending the pellets in 5% sucrose water solution to the required volume. 
	NOTE: The amount of 5% sucrose water solution to be added can be calculated using the equation in step 3.2.1 (MsVs = MiVi).</li>
</ol>
4. Orally Infecting Flies
<ol>
	<li>To ensure oral infection, starve the flies for 2&#8211;4 h before exposure to bacteria by transferring the flies to standard agar vials (1 L triple distilled H2O, 20 g agar, 84 g brown sugar, 7 mL Tegosept anti-fungal agent).</li>
	<li>Prepare infection vials while flies are being starved. Make a Pseudomonas infection vial by pipetting 500 &#181;L of standard sugar agar into the lid of a 7 mL sample tube and leave it to dry. Place a disc of filter paper in the lid and pipette 100 &#181;L of bacterial culture directly onto the filter disc. For control infections, replace the bacterial culture with the same volume of 5% sucrose water solution on the filter paper.</li>
	<li>Add single flies to the sample tube and leave for 18&#8211;24 h.</li>
	<li>To confirm oral infection, first surface-sterilize the flies immediately after bacterial exposure, by placing them in 100 &#181;L of 70% ethanol for 20&#8211;30 s. Remove the ethanol and add 100 &#181;L of triple distilled water for 20&#8211;30 s before removing the water. Add 100 &#181;L of 1x PBS and homogenize the fly.</li>
	<li>Transfer the homogenate to the top row of a 96-well plate and add 90 &#181;L of 1x PBS to every well below.</li>
	<li>Serially dilute this sample to distinguish a range of CFU values. Take 10 &#181;L of the homogenate in the top well and add this to the well below. Repeat this step with the second well, transferring 10 &#181;L to the third well, and so on, for as many serial dilutions as required. 
	NOTE: It important that new pipette tips are used for each set of dilutions.</li>
	<li>Plate the serial dilutions on an LB nutrient agar plate in 5 &#181;L droplets, to ensure all droplets remain discrete.</li>
	<li>Incubate the LB Agar plates overnight at 30 &#176;C and 37 &#176;C for P. entomophila and P. aeruginosa, respectively and count visible CFUs. 
	NOTE: While Drosophila gut microbes require distinct anaerobic growth conditions, selective medium, for example Pseudomonas Isolation Medium (PIM), may be used to make sure only Pseudomonas CFUs are counted.</li>
	<li>Calculate the number of CFUs per fly by counting the number of colonies present at the serial dilution where 10&#8211;60 CFUs are clearly visible. Then multiply by the dilution factor present to calculate the number of bacteria per fly.</li>
	<li>Perform statistical analysis. Where necessary, transform the CFUs per fly to a normal distribution. Do this by log-transformation. Once transformed, use Generalized Linear Models (GLMs)30,31,32 to test how treatment groups differ in CFUs per fly (using commonly available statistical software packages such as R33). 
	NOTE: The remaining fly homogenate can be used for measuring gene expression through quantitative reverse transcription PCR (RT-qPCR) analysis. Fix the homogenate in 50 &#181;L of RNA isolation reagent, extract RNA, and quantify specific immune gene titers by RT-qPCR (see e.g., Gupta and Vale16&#160;for a detailed protocol). The expression of specific immune gene transcripts should be normalized to the transcript levels of a housekeeping gene (i.e., rp49) and expressed as a fold change relative to control flies using the 2&#8722;&#916;&#916;Ct method31,32,33,34.</li>
</ol>
5. Recording Survivorship Following Infection
<ol>
	<li>Infect flies orally as described in step 4.2.</li>
	<li>Transfer the infected or control flies from their respective infection vials into standard Lewis vials and keep in an incubator at 25 &#176;C in a 12 h:12 h light-dark cycle (or desired conditions). Keep flies until they are dead.</li>
	<li>Count the number of living or dead flies in each vial every day, or as often as required.</li>
	<li>Transfer the flies to new vials every 5 days to avoid the flies getting stuck in the food.</li>
	<li>Present these data as Kaplan-Meier (KM) survival curves or Mean &#177; SE proportional survival plots. To analyze the effect of several factors and/or their respective interactions with one another use a statistical package (for example, the package &#8220;survival&#8221; in R33) to run a survival analysis such as the Cox Proportional Hazards model35.</li>
</ol>
6. Measuring Bacterial Load
<ol>
	<li>At the desired time point, transfer a single infected fly to a sterile 1.5 mL microcentrifuge tube.</li>
	<li>Surface sterilize the flies as described in step 4.4.</li>
	<li>Homogenize the fly and quantify the bacterial load using the protocol described in steps 4.5&#8211;4.10.</li>
</ol>
7. Measure Bacterial Shedding
<ol>
	<li>Measure the shedding alongside the internal load.</li>
	<li>After infection, transfer single flies to 1.5 mL microcentrifuge tubes containing ~50 &#181;L of Lewis medium for 24 h.</li>
	<li>Remove the flies for the internal load measurement (see step 6) and wash the tubes with 100 &#181;L of 1x PBS by vortexing heavily for 3 s.</li>
	<li>Measure the CFUs in this wash by plating on LB nutrient agar using the same protocol as described in steps 4.6&#8211;4.8.</li>
	<li>After infection, transfer single flies to 1.5 mL microcentrifuge tubes containing ~50 &#181;L of Lewis medium for 24 h.</li>
	<li>Transfer the flies to new microcentrifuge tubes containing ~50 &#181;L of Lewis medium for a further 24 h. Wash the contaminated tubes with 100 &#181;L of 1x PBS by vortexing heavily for 3 s.</li>
	<li>Measure the CFUs in this wash by plating on LB nutrient agar using the same protocol described in steps 4.6&#8211;4.8.</li>
	<li>Repeat steps 7.2 and 7.3 and record fly mortality at every transfer.</li>
</ol>

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L. <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 Statistical Analysis of Failure Time Data. , (2002).</li><li>. Drosophila melanogaster, drawing.SVG Available from: <a target="_blank" href="https://commons.wikimedia.org/wiki/File:Drosophila-drawing.svg">https://commons.wikimedia.org/wiki/File:Drosophila-drawing.svg</a> (2007)</li><li>. Example of a survival curve estimated by Kaplan-Meier method including 95% confidence limits Available from: <a target="_blank" href="https://commons.wikimedia.org/wiki/File:Kaplan-Meier-sample-plot.svg">https://commons.wikimedia.org/wiki/File:Kaplan-Meier-sample-plot.svg</a> (2011)</li><li>Susi, H., Vale, P. F., Laine, A. -. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Host+Genotype+and+Coinfection+Modify+the+Relationship+of+within+and+between+Host+Transmission.">Host Genotype and Coinfection Modify the Relationship of within and between Host Transmission.</a> Am Nat. 186 (2), 252-263 (2015).</li><li>Fellous, S., Duncan, A. B., Quillery, E., Vale, P. F., Kaltz, O. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+influence+on+disease+spread+following+arrival+of+infected+carriers.">Genetic influence on disease spread following arrival of infected carriers.</a> Ecol Lett. 15 (3), 186-192 (2012).</li></ol>

We present a protocol for reliably orally infecting D. melanogaster with bacterial pathogens. We focus on P. aeruginosa and P. entomophila, but this protocol can easily be adapted to enable infection of other bacterial species, e.g., Serratia marcescens7. Key aspects of this protocol will vary between bacterial species. Accordingly, the most efficient infectious dose, corresponding virulence, and host genotype susceptibility should all be considered and ideally tested in pilot studies. Exposing flies to bacterial cultures of a range of optical densities and measuring their infectious dose and survival is an appropriate starting point when working with new bacterial species or fly lines.
Protocol steps such as fly starvation prior to feeding and re-suspending bacterial pellets in 5% sucrose solution are commonplace in oral infection and increase the reliability of bacterial infection during exposure7,8,9,10. However, it is important to note that during exposure, flies essentially live on a surface of bacterial culture. In the process of walking on this culture, bacteria will become lodged on the fly&#39;s surface, especially on the cuticle or around the bristles24. These epicuticular bacteria, do not reflect a successful enteric infection but would still be detected by the fly homogenization and plating. To reduce the potential for false positives, it is essential to surface sterilize flies through immersion in 70% ethanol for up to 1 min.
When considering bacterial shedding rates, oral infection is essential. The number of pathogens a host releases into the environment is often difficult to measure and the internal load is often taken as a proxy for the severity of infection and therefore transmission26,27. Measuring bacterial load alongside bacterial shedding allows an examination of the relationship between these two important components of disease severity and spread38. One limitation of the method presented is that assaying the internal bacterial load of flies requires destructive sampling. This makes it difficult to investigate longitudinal trends of pathogen growth and clearance within the same individual. However, it is possible to overcome this limitation by destructively sampling cohorts of individuals at different stages of infection, under the assumption that the average microbe load in each cohort reflects the longitudinal pathogen dynamics within any given individual. Bacterial shedding does not suffer from the same limitations, and we offer examples of how shedding can be quantified in a cross-sectional sample, or longitudinally to investigate how shedding changes within an individual over time.
Many host and pathogen traits jointly determine an individual&#39;s propensity to transmit disease25,26,39. While the significance of these traits likely varies between host-pathogen systems, shedding is likely a major determinant of fecal-oral transmission. The ability to measure bacterial shedding opens the opportunity to test this assumption. Having characterized host-pathogen dynamics in a desired panel of fly lines, experimenters could orally infect individuals, and place them alongside uninfected, susceptible hosts during their infectious periods. These &#39;recipient&#39; flies could then be assayed for internal bacterial load at various time points as a way of directly measuring transmission.

The fruit fly (also known as the vinegar fly), D. melanogaster, has been extensively used as a model organism for infection and immunity against a variety of pathogens1,2. This work has offered fundamental insights into the physiological consequences of infection and was also pioneering in unraveling the molecular pathways underlying the host immune response against parasitoid, bacterial, fungal, and viral infections. This knowledge is not only useful to understand the innate immune response of insects and other invertebrates, but because many of the immune mechanisms are evolutionarily conserved between insects and mammals, Drosophila has also spurred the discovery of major immune mechanisms in mammals, including humans3.
Most work on Drosophila infection and immunity has focused on systemic infections, using inoculation methods that deliver pathogens directly into the body of the insect by pricking or injection4,5,6. The advantage of these methods in allowing the delivery of a controlled infectious dose is clear and supported by a large body of work on systemic infections. However, many naturally occurring bacterial pathogens of D. melanogaster are acquired through feeding on decomposing organic matter where gut immunocompetence plays a significant role in host defence7,8,9,10,11,12,13,14,15. Experiments that employ systemic infections bypass these defenses, and, therefore, provide an altogether different picture of how insects mount defenses against natural pathogens. This is especially relevant if the aim of the work is to test predictions about the ecology and evolution of infection, where the use of natural pathogens and routes of infection is important16,17. Recent work has highlighted how the route taken by pathogens significantly affects disease outcome18,19, elicits distinct immune pathways20,21, can determine the protective effect of inherited endosymbionts16, and may even play an important role in the evolution of host defenses17.
Another reason to employ oral routes of infection is that it allows the investigation of the variation in pathogen transmission by measuring bacterial shedding during fecal excretion following oral infection22,23,24. Understanding the sources of host heterogeneity in disease transmission is challenging in natural populations25,26, but measuring components of transmission, such as pathogen shedding, under controlled laboratory conditions offers a useful alternative approach27. By feeding flies bacteria and measuring bacterial shedding under a variety of genetic and environmental contexts in controlled experimental conditions, it is possible to identify sources of variation in transmission among hosts.
Here, we describe a protocol for orally infecting D. melanogaster with bacterial pathogens, and for quantifying the bacterial growth and shedding that follows (Figure 1). We describe this protocol on two Pseudomonas bacteria: a virulent strain of the opportunistic pathogen P. aeruginosa (PA14), and a less virulent strain of the natural fly pathogen P. entomophila. Pseudomonads are common gram-negative bacteria with a broad host range, infecting insects, nematodes, plants, and vertebrates, and are found in most environments4,6. Enteric infection of Drosophila by P. aeruginosa and P. entomophila results in pathology to intestinal epithelia12,13,14,15,28. While we focus on these two bacterial pathogens, the methods described here can in principle be applied to any bacterial pathogen of interest with minor modifications. Following oral exposure, we measure post-infection survival, and measure the microbe load within individual flies and the viable microbes shed into the environment, expressed in colony forming units (CFUs). Finally, because gut immunocompetence results from a combination of epithelial barrier and humoral responses, we also measure the survival of fly lines where these defenses are disrupted. Specifically, Drosocrystallin (Dcy) mutants have been previously shown to be more susceptible to oral bacterial infection due to a depleted peritrophic matrix in the gut29. We also measure survival in a Relish (Rel) mutant which is impeded from producing antimicrobial peptides against Gram-negative bacteria via the IMD pathway30.

<table>
	<tbody>
		<tr>
			<td>Vials</td>
			<td>Sarstedt Ltd.</td>
			<td>58.490</td>
			<td>Polystyrene flat base tube 75mm x 23.5mm, 23ml</td>
		</tr>
		<tr>
			<td>Agar</td>
			<td>Sigma-Aldrich</td>
			<td>A7002</td>
			<td>Agar, ash 2.0-4.5%</td>
		</tr>
		<tr>
			<td>Brown sugar</td>
			<td>Bidvest</td>
			<td>66032</td>
			<td>Light brown soft sugar</td>
		</tr>
		<tr>
			<td>Maize</td>
			<td>Dove&#39;s Farm</td>
			<td></td>
			<td>Organic maize flour</td>
		</tr>
		<tr>
			<td>Fermipan yeast</td>
			<td>Bidvest</td>
			<td>96360</td>
			<td>Dry, instant yeast</td>
		</tr>
		<tr>
			<td>Methyl-4-hydroxybenzoate</td>
			<td>Sigma-Aldrich</td>
			<td>H5501</td>
			<td>&#62;=99.0%, crystalline</td>
		</tr>
		<tr>
			<td>Sucrose</td>
			<td>Sigma-Aldrich</td>
			<td>84097</td>
			<td>Sucrose BioUltra, for molecular biology, &#62;=99.5% (HPLC)</td>
		</tr>
		<tr>
			<td>Petri dishes</td>
			<td>Fisher Scientific UK Ltd.</td>
			<td>15788517</td>
			<td>X600 Petri Dish 90 X 16.2MM Sterile Triple Vent</td>
		</tr>
		<tr>
			<td>Falcon tubes (50ml)</td>
			<td>Greiner Bio-one Inc</td>
			<td>210261</td>
			<td>Centrifuge tube, 50ml, skirted, bagged, sterile</td>
		</tr>
		<tr>
			<td>Eppendorf tube (0.5ml)</td>
			<td>Sarstedt Ltd.</td>
			<td>72.699</td>
			<td>Sarstedt Micro Tube Conical Base Push Cap 0.5ml Standard Neutral</td>
		</tr>
		<tr>
			<td>Eppendorf tube (1.5ml)</td>
			<td>Sarstedt Ltd.</td>
			<td>72.690.001</td>
			<td>Sarstedt Micro Tube Conical Base Push Cap 1.5ml Standard Neutral</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>VWR International Ltd.</td>
			<td>20821.33</td>
			<td>ETHANOL ABSOLUTE ANALAR NP ACS/R.PE - Analytical Grade</td>
		</tr>
		<tr>
			<td>2 L Conical Flask</td>
			<td>VWR International Ltd.</td>
			<td>214-0038</td>
			<td>Narrow neck, 2 L Erlenmeyer flask</td>
		</tr>
		<tr>
			<td>Sterile filter paper</td>
			<td>Fisher Scientific UK Ltd.</td>
			<td>1001-020</td>
			<td>Plain circle and sheets; Particle Retention: greater than11um; Filtration speed: 150 herzberg; Air flow: 10.5s/100mL/in2; Medium porosity; Smooth surface; Grade 1; Type: circle; Dia: 20mm</td>
		</tr>
		<tr>
			<td>Bijou sample container</td>
			<td>Fisher Scientific UK Ltd.</td>
			<td>129A</td>
			<td>7ml polystyrene sample container</td>
		</tr>
		<tr>
			<td>Pseudomonas isolation agar</td>
			<td>Sigma-Aldrich</td>
			<td>17208</td>
			<td>Contains agar 13.6 g/L, magnesium chloride 1.4 g/L, peptic digest of animal tissue 20g/L, potassium sulfate 10g/L, triclosan 0.025g/L</td>
		</tr>
		<tr>
			<td>LB broth, Miller</td>
			<td>Fisher Scientific UK Ltd.</td>
			<td>BP1426</td>
			<td>Contains 10g tryptone, 5g yeast extract, 10g sodium chloride per litre</td>
		</tr>
		<tr>
			<td>Large Embryo Collection Cages</td>
			<td>Scientific Laboratory Supplies Ltd.</td>
			<td>59-101</td>
			<td>Flystuff- fits 100mm petri dish</td>
		</tr>
		<tr>
			<td>TRI reagent solution</td>
			<td>Life Technologies</td>
			<td>AM9738</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well microplate</td>
			<td>Scientific Laboratory Supplies Ltd.</td>
			<td>353072</td>
			<td>Falcon 96 Well Clear Flat Bottom TC-Treated Polystyrene Cell Culture Microplate with Lid Sterile</td>
		</tr>
		<tr>
			<td>Pestle</td>
			<td>Fisher Scientific UK Ltd</td>
			<td>12649595</td>
			<td>Pestle, Presterilized; Axygen; Tissue grinder; Blue; Inert polypropylene construction; Fits 1.5 and 2.0mL centrifuge tubes; Individually wrapped; 100/Pk</td>
		</tr>
		<tr>
			<td>Glycerol</td>
			<td>Scientific Laboratory Supplies Ltd.</td>
			<td>CHE2068</td>
			<td>Glycerol A.R. 99.5% 2.5 L</td>
		</tr>
		<tr>
			<td>Cotton wool- non absorbent</td>
			<td>Cowens Ltd</td>
			<td>ABL</td>
			<td>Non Absorbent Large quantity 20 bags x 500</td>
		</tr>
		<tr>
			<td>Cotton wool- absorbent</td>
			<td>Cowens Ltd</td>
			<td>ABS</td>
			<td>BP small quantity 20 bags x 500</td>
		</tr>
		<tr>
			<td>Vortex</td>
			<td>Fisherbrand</td>
			<td></td>
			<td>Whirlimixer 75W 50-6-Hz 220-240V</td>
		</tr>
		<tr>
			<td>Orbital incubator</td>
			<td>Gallenkamp</td>
			<td>INR-200-010V</td>
			<td>220 V, 50 Hz, 5 A. Nominal temperature: 70&#186;C. Shaking frequency: 0...400 rpm.</td>
		</tr>
		<tr>
			<td>Absorbance Microplate Reader</td>
			<td>Biotek Instruments Ltd</td>
			<td></td>
			<td>ELx808&#8482; Absorbance Microplate Reader</td>
		</tr>
		<tr>
			<td>Gen 5 Microplate Reader and Imager Software</td>
			<td>Biotek Instruments Ltd</td>
			<td></td>
			<td>For Absorbance Microplate Reader</td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Beckman Coulter</td>
			<td>392304</td>
			<td>Allegra X-12R Benchtop Centrifuge, refridgerated 50Hz 230V</td>
		</tr>
		<tr>
			<td>Step One Plus Real Time qPCR System</td>
			<td>Thermofisher Scientific</td>
			<td>4376600</td>
			<td>Applied biosystems Step One Plus real time qPCR system</td>
		</tr>
		<tr>
			<td>Step One Software 2.3</td>
			<td>Thermofisher Scientific</td>
			<td></td>
			<td>For Stepone and SteponePlus real time qPCR systems</td>
		</tr>
		<tr>
			<td>R Statistical Software</td>
			<td>https://cran.r-project.org/</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#181;L pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>741015</td>
			<td>Easlyload gilson-style. Graduated, clear, refill, 960 pcs</td>
		</tr>
		<tr>
			<td>200 &#181;L pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>741065</td>
			<td>Easlyload gilson-style. Graduated, clear, refill, 960 pcs</td>
		</tr>
		<tr>
			<td>1000 &#181;L pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>741045</td>
			<td>Easlyload gilson-style. Graduated, clear, refill, 960 pcs</td>
		</tr>
		<tr>
			<td>20 &#181;L filtered pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>774288</td>
			<td>Filter tip gilson-style. Clear, rack blue, single packed, R/Dnase free, 10 racks of 96pcs</td>
		</tr>
		<tr>
			<td>200 &#181;L filtered pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>739288</td>
			<td>Filter tip gilson-style. Clear, rack blue, single packed, R/Dnase free, 10 racks of 96pcs</td>
		</tr>
		<tr>
			<td>1000 &#181;L filtered pipette tips</td>
			<td>Greiner Bio-one Inc</td>
			<td>740288</td>
			<td>Filter tip gilson-style. Clear, rack blue, single packed, R/Dnase free, 10 racks of 60pcs</td>
		</tr>
	</tbody>
</table>

oral bacterial infection and shedding in drosophila melanogaster

The fruit fly Drosophila melanogaster is one of the best developed model systems of infection and innate immunity. While most work has focused on systemic infections, there has been a recent increase of interest in the mechanisms of gut immunocompetence to pathogens, which require methods to orally infect flies. Here we present a protocol to orally expose individual flies to an opportunistic bacterial pathogen (Pseudomonas aeruginosa) and a natural bacterial pathogen of D. melanogaster (Pseudomonas entomophila). The goal of this protocol is to provide a robust method to expose male and female flies to these pathogens. We provide representative results showing survival phenotypes, microbe loads, and bacterial shedding, which is relevant for the study of heterogeneity in pathogen transmission. Finally, we confirm that Dcy mutants (lacking the protective peritrophic matrix in the gut epithelium) and Relish mutants (lacking a functional immune deficiency (IMD) pathway), show increased susceptibility to bacterial oral infection. This protocol, therefore, describes a robust method to infect flies using the oral route of infection, which can be extended to the study of a variety genetic and environmental sources of variation in gut infection outcomes and bacterial transmission.

Here, we present illustrative results from experiments where D. melanogaster was orally infected with P. aeruginosa or P. entomophila. Figure 2 demonstrates the successful oral infection of flies following a 12 h or 24 h exposure period to bacterial cultures of OD600 = 25 and 100 for P. aeruginosa (Figure 2A) and P. entomophila (Figure 2B, C), respectively. Figure 2B illustrates the importance of using a more concentrated culture of P. entomophila, shown by the increase in bacterial load when flies are exposed to bacterial cultures of greater optical density. Male and female Oregon R (OreR) flies clear P. aeruginosa infection at the same rate (Figure 3) and shed the same number of P. aeruginosa CFUs (Figure 4A). When infected with P. entomophila however, male and female OreR flies differ in the number of bacteria shed, in a manner that changes over time (Figure 4B). Males and females die from P. aeruginosa (Figure 5A) and P. entomophila (Figure 5B) at different rates. We also see that Dcy mutants (which lack the protective peritrophic matrix in the gut epithelium) and Relish mutants (which lack a functional IMD immune pathway), show decreased survival following P. entomophila and P. aeruginosa oral infection (Figure 5C).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57676/57676fig1.jpg" /> 
Figure 1: Schematic overview of protocols for measuring survival, shedding, and internal bacterial load following oral infection in Drosophila melanogaster. An illustration of 3 potential experiments following the oral infection of D. melanogaster. Measure the &#39;survival&#39; by transferring single flies to vials and recording their infected lifespan. Measure &#39;shedding&#39; by transferring single flies to 1.5 mL microcentrifuge tubes with 50 &#181;L of Lewis medium in the cap. After 24 h in the tube, remove the fly and vortex the tube with 100 &#181;L of 1x PBS. Remove and plate this solution on LB nutrient agar to calculate the bacterial shedding. Measure the shedding in the same fly longitudinally, by transferring flies to fresh tubes with Lewis medium in the cap after 24 h, and washing and plating the now contaminated tube. A fly&#39;s &#39;internal load&#39; can be measured by taking an infected fly, surface sterilizing it, and homogenizing it before finally plating the homogenate on LB nutrient agar. This can be performed after shedding has been measured to calculate how the &#39;internal load&#39; and shedding correlate. The fly illustration used in this figure was originally drawn by B. Nuhanen36. The authors have modified it to accompany the example Kaplan-Meier curve which is taken from Wikimedia Commons37. All other illustrations are original. <a href="/files/ftp_upload/57676/57676fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57676/57676fig2.jpg" /> 
Figure 2: Infectious dose of bacteria following oral infection. (A) Infectious dose of male and female Oregon-R flies following exposure to a P. aeruginosa culture (OD600 = 25) for 12 h. The mean and SE were calculated from 3 males and 3 females. (B) Infectious dose of outcrossed wild-type females following exposure to one of four P. entomophila cultures (OD600 = 100, 75, 50, and 25) or control 5% sucrose solution for 24 h. The statistical difference of (F3,76 = 18.567, p &#60;0.001) in the infectious dose between exposure treatments is denoted by differing letters above bars. The means were calculated from 5 flies for the OD600 = 0 dose, and 18-20 for all other doses. (C) The infectious dose of male and female Oregon-R flies following exposure to P. entomophila culture (OD600 = 100) for 24 h. The mean and SE were calculated from 20 males and 20 females. <a href="/files/ftp_upload/57676/57676fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57676/57676fig3.jpg" /> 
Figure 3: Internal P. aeruginosa load in flies after oral infection. Mean &#177; SE bacterial load of male and female Oregon-R flies following oral infection with P. aeruginosa (OD600 = 25) up to 168 h post-infection. The mean and SE of each time point are calculated from 3 individuals. A fly&#39;s internal bacterial load significantly changes over time (p &#60;0.001). <a href="/files/ftp_upload/57676/57676fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/57676/57676fig4.jpg" /> 
Figure 4: Bacterial shedding following oral infection. (A) P. aeruginosa shed by the same flies used in Figure 3, up to 120 h post-infection. The mean and SE were calculated from 3 males and 3 females. (B) P. entomophila shed by male and female Oregon-R flies following oral infection with P. entomophila (OD600 = 100) up to 120 h post-infection. The mean and SE were calculated from 34 males and 38 females. For both P. aeruginosa and P. entomophila, the number of CFUs shed by a fly significantly changes over time (p &#60;0.001). <a href="/files/ftp_upload/57676/57676fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/57676/57676fig5.jpg" /> 
Figure 5: Survival of flies following bacterial oral infection. Kaplan-Meier (KM) survival curves of (A) Oregon-R male and female flies following oral infection with P. aeruginosa (OD600 = 25) or control 5% sucrose solution. The KM survival curve was calculated from 4 vials of 20 flies per treatment group. (B) OreR male and female flies after oral infection with P. entomophila (OD600 = 100). The KM survival curve was calculated from 4 single control flies and 34 infected flies for both males and females. (C) Immune mutants: Dcy (Drosocrystallin-peritrophic matrix mutant) and Rel (Relish-IMD mutant), exposed to P. entomophila (Pe), P. aeruginosa (Pa14), or a control 5% sucrose solution. All infected groups die significantly faster than the control flies (p &#60;0.001). <a href="/files/ftp_upload/57676/57676fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Oral Bacterial Infection and Shedding in Drosophila melanogaster at JoVE.com

Oral Bacterial Infection and Shedding in Drosophila melanogaster

This protocol describes methods to orally expose and infect the fruit fly Drosophila melanogaster with bacterial pathogens, and to measure the number of infectious bacteria shed following gut infection. We further describe the effect of immune mutants on fly survival following oral bacterial infection.

Oral Bacterial Infection and Shedding in Drosophila melanogaster

The fruit fly Drosophila melanogaster is one of the best developed model systems of infection and innate immunity. While most work has focused on systemic ...

oral-bacterial-infection-and-shedding-in-drosophila-melanogaster

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11.4K Views.  University of Edinburgh. This protocol describes methods to orally expose and infect the fruit fly Drosophila melanogaster with bacterial pathogens, and to measure the number of infectious bacteria shed following gut infection. We further describe the effect of immune mutants on fly survival following oral bacterial infection.

Video: Oral Bacterial Infection and Shedding in Drosophila melanogaster

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Wolbachia Bacterial Infection in Drosophila

My name is Voia Friedman. I am associate researcher at Princeton University. And today I'm going to tell you about a few protocols that I developed to trans infect, to I initially to isolate, to go back from the heli of infected flies, and later on help to inject the heli feed to unaffected flights.So basically, this is a very simple protocol to isolate him oly for basics. And you will use this device from, from mepo that contains a filter that goes on top of an E.So you basically needs to put you, we start from around a hundred flies. You anize the flies, you put them into the filter device, you can just use a funnel and that will add our other fit very easily on this device and wash it with 70%ethanol.You just take the, take the future away, you can throw away the ethanol, and then after three times your last wash you do with molecule water because you don't want have the bacteria from the, with traces of ethanol. Well, and then you just go to a new weapon door and, and then you spin down on a micro, real micro for two minutes And later on how you can inject that. He only feed do unaffected flies.So basically take a double stick tape and usually these double stick tapes have two side, one side is a sticker than the other. So leave the sticker side up because that's the side that you're gonna glue that flies on top of it. And basically you'll, the gas apparatus, if you have this light on top of it, you'll be no gas flow and the flies you wake up if they're on top of the slide.So you can just direct the flow, making a little hood, a little chamber using tape. Then you direct the gas over the flies. So then you put your, this light here with the tape and you, you have your flies.Stick the flies on the top of the tape, all of them the same orientation, and put their head towards the direction that you'll be bringing the needle from. Because then you can just over the, the wing, you make sure since they fly, you don't want to fly. If they woke up during the procedure, don't want them to go away.So make sure that you take a little tip and pressure against the, the, the wings of the fly on the tail. And it's okay if they wings get damaged. And when you get rid, when you take the flies up the injection, most likely they'll lose their wings anyway, so they'll survive.That can, that can be useful because you're not gonna lose the flies that, so he puts a mineral oil on the micrometer for the back of the needle, then the needle injection polar, and then the, you can just adjust the pressure to bring the material up to the tip of the needle. I, I did to inject Somewhere between 10 to 15 nanoliters, but as, as long, as long as I'm consistents. Okay, so one important detail there is a cushion of this experiment is that the, the fly, once the apply is being injected when you saw the needle needs to come as far as you can through the upper part of the abdomen.Because here, basically when the needle penetrates, you have a, a, a, a, a bigger space to go up and down. And basically if it's close, closer to the cuticle, it's less likely they're gonna penetrate vital organs in the flight. You, it is very likely unlikely that you're gonna punch a hole in your gut, for instance, if that is gonna kill the flight that you are performing the surgery.My name is Voia Friedman. I am associate researcher at Princeton University in the lab of Eric Hels. And today I would like to tell you about this protocol that you just saw, the demonstration.So this protocol was done because we wanted to prove that UBA is capable of crossing tissues. So UBA is one of the most successful intracellular paras on the face of the earth. Infects from 20 to 70%of insects and many particles.And also is, is very important in terms of human diseases because also infect nematodes that transmit diseases like river blindness or s and ssis. And so, and CCA is our obligatory endo in, in these worms. And there's a great, a very big interest in using CCA also as means to control parasitic diseases transmitter by mosquito and Danny and this and by mosquito such as malaria and dengue.So this, this, this protocol basically was developed to show that once CCA inside the abdominal cavity of a fly, they can cross tissues and reach the germ. So basically what you can see here is the oph of the drosophila ovary and go back once present in the he leaf it, it can cross a pink peronial. She that is around the ovary.And then they cross a muscle layer. And actually one of the first places that they, they go inside the ovary is the somatic stem cell image. And then they can reach the germline.And this bacteria is transmitted the same way that mitochondria is transmitted is through, is through the mother transmitted, through the mother cyop plants present in the egg. So of course they needs to be present into the germline. So that's why this protocol is developed.And this protocol is, can be very useful to basically traditionally the way that people would try to trans infect wbaa basically take wbaa and introduce a new insect species. It would be through doing micro injections into the, into the embryo. But those are much more complicated.And basically here we are just taking advantage of the o biology since they are capable to cross tissues and moving into the stem cell tissues from, they can pass to the, through to the germline. So that's make easier taking UBA from certain, from, from certain species of insects that are infected and they can be transmitted to uninfected species. Prob I think it's the injection itself.Basically you need to be careful to mobilize the fly as well as you can. We are using double sided stick tape and you need to penetrate the needle basically as close to the, as parallel to the fly as possible so you don't go to deep. And it's less likely that we're gonna kill the fly after they're performing the surgery.

Protocols for Oral Infection of Lepidopteran Larvae with Baculovirus

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. This video shows how lepidopteran larvae can be infected with polyhedra by droplet feeding and diet plug-based bioassays. This accompanying Springer Protocols section provides an overview of the baculovirus lifecycle and use of baculoviruses as insecticidal agents, including discussion of the pros and cons for use of baculoviruses as insecticides, and progress made in genetic enhancement of baculoviruses for improved insecticidal efficacy.

In this video, we demonstrate oral infection techniques of lepidopteran larvae with baculovirus in order to determine insecticidal efficiency.

Baculoviruses are widely used both as protein expression vectors and as insect pest control agents. This video shows how lepidopteran larvae can be ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called imaginal discs. These discs give rise to a high percentage of adult structures that are found within the adult fly. Here we describe a protocol that has been optimized to recover these discs and prepare them for analysis with antibodies, transcriptional reporters and protein traps. This procedure is best suited for thin tissues like imaginal discs, but can be easily modified for use with thicker tissues such as the larval brain and adult ovary. The written protocol and accompanying video will guide the reader/viewer through the dissection of third instar larvae, fixation of tissue, and treatment of imaginal discs with antibodies. The protocol can be used to dissect imaginal discs from younger first and second instar larvae as well. The advantage of this protocol is that it is relatively short and it has been optimized for the high quality preservation of the dissected tissue. Another advantage is that the fixation procedure that is employed works well with the overwhelming number of antibodies that recognize Drosophila proteins. In our experience, there is a very small number of sensitive antibodies that do not work well with this procedure. In these situations, the remedy appears to be to use an alternate fixation cocktail while continuing to follow the guidelines that we have set forth for the dissection steps and antibody incubations.

Dissection and Immunostaining of Imaginal Discs from Drosophila melanogaster

The adult structures of Drosophila are derived from sac-like structures called imaginal discs. Analysis of these discs provides insight into many developmental processes including tissue determination, compartment boundary establishment, cell proliferation, cell fate specification, and planar cell polarity. This protocol is used to prepare imaginal discs for light/fluorescent microscopy.

A significant portion of post-embryonic development in the fruit fly, Drosophila melanogaster, takes place within a set of sac-like structures called ...

Maintenance of a Drosophila melanogaster Population Cage

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

Maintenance of a Drosophila melanogaster Population Cage

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

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