We are grateful to T. Huber and T.P. Sakmar for helpful discussions. This work was supported by grants from the US National Institutes of Health to L.B.V. and the Belgian-American Educational Foundation and the Revson Foundation to M.L.

1. Equipment and Reagents
Odor dilutions:
<ul>
<li>Paraffin oil (Sigma, CAS number: 8012-95-1)</li>
<li>Odors of the highest purity available which elicits attraction in Drosophila larvae1. In this video article, animals were tested with isoamyl acetate(Sigma, CAS number: 123-92-2). </li>
<li>1.5ml glass vials with Teflon cap (Agilent Technologies)</li>
<li>Digital measuring scale for precise dilution by weight</li>
</ul>
Assay arena set-up:
<ul>
<li>Agarose (Promega)</li>
<li>Lids of 96-well Microplate (BD Falcon, catalogue number: 353071). Our behavioral arena consists of three stacked lids.</li>
<li>Single-channel 1-20 &mu;l pipetter or multi-channel pipetter to load odorant droplet(s) into the top lid of the stack. To set up the multiple source assay, we use 5-50 &mu;l Pipet-Lite multi-channel pipetter from Rainin.</li>
</ul>
Preparation of Drosophila larvae for behavioral tests:
<ul>
<li>15% (w/w) sucrose solution</li>
<li>Regular Petri dishes (BD Falcon) to wash and store larvae</li>
<li>Paint-brush (4-5mm bristle length) to manipulate and transfer larvae</li>
</ul>
Recording of larval motion:
<ul>
<li>Small pins with a flat head for transferring larvae into arena</li>
<li>Forceps or paintbrush to remove larvae after recording</li>
<li>Illuminated stage with a homogeneous non-heating light source, "Slim Edge" Light Pad manufactured by Logan Electric</li>
<li>Cabinet to surround the behavioral arena and control light conditions</li>
<li>Video tracking system for recording of animal movement. We are using the Ethovision software (Noldus, The Netherlands).</li>
<li>CCD camera and frame grabber video card (EureCard Piccolo Basic) purchased with Ethovision software</li>
<li>Thermometer and humidity recorder to monitor the atmospheric conditions during the behavioral tests (optional)</li>
<li>Matlab (The Mathworks) for customized data analysis (optional)</li>
</ul>
2. Behavioral arena
The behavioral arena consists of three stacked 96-well dish lids. The bottom lid is used to isolate the rest of the system from the light pad and reduce convection in the arena. The second lid, covered with 25ml of a 3% agarose gel, serves as a stage for the larvae to walk on. The top lid holds the odor droplets which remain suspended by surface tension.
Odor droplets are pipetted directly into the wells of the 96-well dish lid. A single or multiple odor sources can be used to generate gradients with distinct, verifiable profiles2. To test for anosmia and chemotaxis defects in larvae, we propose to use the following two assays.
2.1. Single odor source assay: a single odor droplet laid in the well #E7 (standard 96-well plate reference system).
<ul>
<li>Stimulus: 10&mu;l of odor diluted in paraffin oil</li>
<li>Initial conditions: a single larva is introduced under the odor source (or in its close neighborhood).</li>
<li>Basic observations: For attractive stimuli (e.g. isoamyl acetate), wildtype larvae accumulate underneath the source. Anosmic animals or animals subjected to olfactory defects quickly wander away from the droplet.</li>
<li>Duration of a trial: 3 min. The motion of a given animal is recorded until the trial time has elapsed or as soon as the animal contacts the wall of the plate.</li>
<li>Possible metrics for behavioral quantification: percentage of time spent by an animal in a small circular zone centered on the odor source; mean distance to the odor source; time course of distance to odor source, etc. </li>
</ul>
This assay efficiently tests for basic anosmia in mutant larvae. It can also be used to reliably determine the detection threshold for particular odors2.
2.2. Multiple odor source assay
Several odor droplets are laid in the well of the top lid. In the present video article, we set up gradients along the central row E. A total of six droplets were introduced in wells #E2, #E4, #E6, #E8, #E10 and #E12 (standard 96-well plate reference system). In contrast with the Petri dish and single odor source assays, the multiple odor source assay allows qualitative control of the gradient geometry established along the length and width of the arena. This assay was used to characterize the sensory mechanisms allowing Drosophila larvae to chemotax2.
<ul>
<li>Stimulus: 10&mu;l odor droplets are laid in alternating wells along a row. This arrangement leads to the creation of a gradient centered selected row. To establish a homogenous gradient along the width of the arena, several rows can be loaded with multiple sources.</li>
<li>Initial conditions: a single larva is introduced under the odor source between wells #E3 and #E4.</li>
<li>Basic phenotypes: anosmic larvae wander at random from the starting point. For attractive stimuli (e.g. isoamyl acetate), animals which are able to smell navigate along the direction of increasing odor concentration. The accuracy with which an animal ascends the odor trail is correlated with its ability to sense and process the odorant stimulus. Tortuous paths are often caused by sensory defects. Once an animal has reached the point of highest concentration in the trial, it remains in the vicinity of this position.</li>
<li>Duration of a trial: 3 min maximum (longer tracking durations tend to introduce noise as larvae lose interest and eventually abandon the gradient). For the gradients illustrated in this video article, the motion of an individual larva was recorded until the animal reached the neighborhood of the highest concentration source (well #E12). The recording was stopped as soon as the animal reached the target area or contacted any wall of the arena.</li>
<li>Possible metrics for behavioral quantification: percentage of time spent by an animal under the odorant line (blue rectangle); mean heading with respect to the local direction of the odor gradient; a combined chemotaxis score measuring the global tendency of an animal to follow the odorant line2.</li>
</ul>
3. Gradient profile verification
Before testing larvae in a particular gradient, it is important to determine its stability over time. On this basis, the experimenter can determine an adequate duration for each trial and the number of larvae that can be tracked sequentially in the same arena. To test the stability of the gradient, we have developed a method based on IR spectroscopy2. This method is beyond the scope of this protocol and will not be elaborated further.
4. Protocol for the preparation of an experiment
4.1. Preparation of agarose plate and odor dilutions:
<ol>
<li>1. Pour 25ml of 3% agarose onto the flat top of a 96-well dish lid; allow the agarose to cool down and solidify (~ 30 min). Important: Ensure that the 96-well lids are on a flat surface before pouring the agarose as the presence of a slope on the solidified agarose gel may affect the behavior of the larvae. When pouring the gel, remove any bubbles that form on the surface. </li>
<li>2. Dilute odors to desired concentrations in paraffin oil using a digital scale to accurately measure the amounts of paraffin oil and odor in each dilution.</li>
</ol>
Notes: When low odor concentrations are used, it is preferable to prepare a set of source concentrations based on serial dilutions. Start, for instance, with a 1.0M solution and perform a 1:2 dilution to obtain a solution 0.5M. By proceeding recurrently, one obtains a dilution series following the geometric progression 1.0M, 0.5M, 0.25M, 0.12M, 0.06M, 0.03M, 0.015M.
Important: Since many organic odors react with plastic, odor dilutions must be stored in glass vials with Teflon caps. Ideally, odors should be prepared fresh before each experiment to reduce fluctuations in actual odor concentration across experiments. This is especially true when using highly volatile chemicals.
4.2. Larvae preparation:
<ol>
<li>1. Prepare 15% sucrose solution in distilled water. Notes: A 15% sucrose solution is denser than larvae and will cause them to be buoyed to the surface of the solution. However, undissolved fly food chunks have a greater density than the solution and rapidly sink to the bottom. Thus, using this sucrose solution provides an efficient means of separating larvae from the food. Sucrose solution is an excellent medium for biological contaminants, if the solution becomes cloudy it can be filter sterilized to remove the contamination. </li>
<li>Obtain a vial with 6-day-old larvae or larvae at the desired developmental stage. Notes: All our experiments are carried out with 6-day old larvae (middle of third instar developmental stage); larvae at this developmental stage are large enough for ease of detection with our tracking software, but also highly active. </li>
<li>Pour sucrose into the food vial containing the larvae.</li>
<li>Using a spatula, break up and dissolve the fly food gently to release the larvae. Released larvae will float up to the surface.</li>
<li>Allow a few minutes for the fly food chunks to sink to the bottom of the vial; then, pour the sucrose and larvae into a Petri dish. If pre-selection of the larvae is necessary, animals can be collected by pouring the sucrose through a filter. Larvae can then be transferred to a dish for selection with a paintbrush.</li>
<li>Leave the larvae for approximately 30 minutes in the sucrose solution before proceeding with behavioral tests to allow the animals to habituate to the sucrose solution. As larvae do not appear to feed on sucrose, the animals will experience a state of starvation which improves the chemotaxis performance.</li>
<li>The lid of the Petri dish can be filled with water (or sucrose) and used as a receptacle for discarded animals after behavioral tests.</li>
</ol>
5. Protocol to perform behavioral recordings
5.1. Arena set-up:
<ol>
<li>1. Apply odor droplets to the underside (welled side) of a fresh 96-well dish lid. Important: To avoid contamination from different odors and odor concentrations, a fresh lid must be used for each loading set-up. None of the lids containing odor sources are recycled after the experiment. Depending on the odor concentration range used, odorant gradients have been found to be stable only for 10 to 20 minutes in a closed system. It is therefore important to minimize the time between gradient set-up and the actual beginning of the behavioral test. </li>
<li>Invert the lid and stack it above a second lid coated with the 3% agarose layer. Important: Invert the lid carefully to prevent odor droplets from spreading to adjacent wells. </li>
<li>Upon inversion of the top lid on the agarose surface, allow the odor to diffuse for 30 seconds before introducing the larva.</li>
</ol>
5.2. Loading larvae:
<ol>
<li>Lift the top lid containing the odorant droplets just enough to allow loading of the larva.</li>
<li>Transfer a larva from the sucrose solution to the agarose layer. The starting position of the animal depends on the assay being used: For the single odor source assay, larvae are released under the odorant droplet. For the multiple odor source assay, larvae are released between wells #E3 and #E4. Important: Orient the newly introduced animals in a consistent manner. We orient our animals in the direction of the gradient, ensuring that the animal body and head face the highest concentration. Animals introduced in an orientation opposite to that of the gradient initiate a searching behavior. In our view, this initial search period represents an unnecessary source of noise. Controlling the head orientation reduces the number of animals which contact the arena wall during their exploratory behavior. This practice does not prime the direction of motion of the animal or significantly increase the number of anosmic animals chemotaxing along the odorant line by chance (i.e. false-positive in smell tests). </li>
<li>Replace the top lid and start the recording promptly. Notes: Healthy larvae start crawling immediately upon their introduction in the arena and may cover a distance of 1cm within the first 10 seconds. If the experimenter is slow at initiating the recording, initial datapoints will be lost. For this reason we recommend that the experimenter becomes well acquainted with the software commands to initiate recording before attempting to run an experiment. </li>
<li>Allow the larva to behave for the duration of the recording (typically 3 minutes). Notes: Recordings are terminated before the end of the 3 min if the larva hits the arena wall or reaches a target area (such as in the gradient assay). </li>
<li>At the end of the recording, promptly lift the lid and remove the larva with forceps or a paintbrush. Place the tracked animals in the Petri dish lid containing water (or sucrose). Important: Recorded animals are not reused. The water dish is merely a convenient receptacle to separate recorded and un-recorded animals while running the experiments. </li>
<li>If tracking multiple animals under an arena set-up, promptly load the next larva and start the recording as shown above. Important: The number of trials performed in the same arena set-up should be decided on the basis of the gradient stability which should be verified before performing any behavioral experiment2. </li>
<li>Once all the larvae to be tested under the arena set-up have been tracked and removed from the arena, discard the top lid and agarose layer. The intermediate lid (i.e. the agarose lid) can be recycled. Note: We advise Ethovision users to choose the &lsquo;background subtraction' detection method.</li>
</ol>
6. Sample experiments demonstrated in the video article
6.1. Single odor source assay with 1.0M of isoamyl acetate.
We blindly tested ten Or83b-/- mutants and ten wild type (W1118) larvae to each of which a number had been assigned. The tracking experimenter did not receive any information about the genotype of each animal. After each recording, a predictive phenotype was assigned to each animal tested: smellers, if accumulation was observed under the source for the entire trial; non-smellers, if the path quickly left the neighborhood of the odor source. After having recorded the behavior of 20 animals, paths were grouped according to phenotype. The result of this grouping was then compared with the actual genotype of the larvae: all assignments made were found to be correct.
6.2. Multiple odor source assay with exponential and linear gradients of isoamyl acetate.
In this experiment, we compared the performances of wild type larvae chemotaxing along a linear and an exponential gradient. Chemotactic behaviors were recorded for two gradients set up with the same highest concentration of 0.5M. As explained in the article, the odor sources of the exponential gradient ranged between 0.015 and 0.5M. The sources of the linear gradient ranged between 0.25 and 0.5M (arithmetic progression with common difference 0.05M).
The linear gradient represented a more challenging chemotaxis environment as the local difference in concentration was constant and relatively small, and the concentration range narrow (0.25M). In contrast, the concentration range of the exponential gradient was larger with increasing local differences in concentration along the trail.
Fifteen larvae were recorded for each of the two gradient geometries and the paths superimposed in one graph. A visual inspection of the two graphs shows that wild type larvae were able to reliably ascend the gradient for both geometries but that the amount of meandering was greater for the linear gradient condition. This suggests that the signal present along the gradient length is more reliably detected for an exponential shape than for a linear one.
6.3. Multiple odor source assay with isoamyl acetate gradient in a "roof" geometry.
Here we verify that larvae introduced in an odor gradient are sensitive to local changes in odor concentration rather than just the presence or absence of odor. An isoamyl gradient was established by the application of odor droplets with the following concentrations: 0.06, 0.12, 0.25, 0.5, 0.25, 0.12M. The ensuing gradient geometry resembled an asymmetrical "roof" peaking at 0.5M.
Larvae were released between the 0.06M and 0.12M odor droplets. Our hypothesis was that if the larvae were merely computing the presence of odor they would follow the entire gradient without a particular preference for the concentration maximum. In contrast, if larvae are sensitive to local changes in concentration they would accumulate underneath the 0.5M droplet. Fifteen larvae were recorded, and it was found that following a short exploratory period all larvae accumulated underneath the higher concentration well. This suggests that local changes in odor concentration are computed and inform the chemotactic behavior of Drosophila larvae.

<ol>
	<li>Fishilevich, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotaxis+behavior+mediated+by+single+larval+olfactory+neurons+in+Drosophila.">Chemotaxis behavior mediated by single larval olfactory neurons in Drosophila.</a> Curr Biol. 15, 2086-2096 (2005).</li><li>Louis, M., Huber, T., Benton, R., Sakmar, T. P., Vosshall, L. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bilateral+olfactory+sensory+input+enhances+chemotaxis+behavior.">Bilateral olfactory sensory input enhances chemotaxis behavior.</a> Nature Neuroscience advance online publication. , (2007).</li></ol>

The authors have nothing to disclose.

high-resolution measurement of odor-driven behavior in drosophila larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Watch this Scientific Journal Video about High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae at JoVE.com

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

high-resolution-measurement-odor-driven-behavior-drosophila

Research

JoVE Journal

Biology

10.7K visualizações.  Rockefeller University. Neste artigo de vídeo, descrevemos um método novo que permite a construção de gradientes de odores com geometrias estável e controlável. Brevemente ilustrar como esses gradientes podem ser usados ​​para triagem de defeitos olfativa (anosmia total e parcial) e estudar características mais sutis de comportamento de quimiotaxia.

Vídeo: High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Dissection of Imaginal Discs from 3rd Instar Drosophila Larvae

This protocol demonstrates the dissection technique used for isolating imaginal discs from drosophila larvae at the 3rd instar stage. Methods for fixing the tissue after saying and removing the wing, leg, and eye discs are demonstrated directly on a microscope slide for subsequent visualization.

Dissecação de discos imaginais de 3 larvas Drosophila Instar

Biologia

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

De alta resolução de vídeo de Rastreamento Locomotion em Adult Drosophila melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Na imagem in vivo de larvas Drosophila Intact no Sub-celular Resolução

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

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.

Medição do Tempo de vida em<em&gt; Drosophila melanogaster</em

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

Force Measurement During Contraction to Assess Muscle Function in Zebrafish Larvae

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional measures of muscle health. In this video article, we demonstrate a method to measure force generation during contraction of zebrafish larval trunk muscle. Force measurements are accomplished by placing an anesthetized larva into a chamber filled with a salt solution. The anterior end of the larva is tied to a force transducer and the posterior end of the larva is tied to a length controller. An isometric twitch contraction is elicited by electric field stimulation and the force response is recorded for analysis. Force generation during contraction provides a measure of overall muscle health and specifically provides a measure of muscle function. Although we describe this technique for use with wild-type larvae, this method can be used with genetically modified larvae or with larvae treated with drugs or toxicants, to characterize muscle disease models and evaluate treatments, or to study muscle development, injury, or chemical toxicity.

Force measurements can be used to demonstrate changes in muscle function due to development, injury, disease, treatment or chemical toxicity. In this video, we demonstrate a method to measure force during a maximal contraction of zebrafish larval trunk muscle.

Medição de força durante a contração para avaliar a função muscular em larvas do peixe

Zebrafish larvae provide models of muscle development, muscle disease and muscle-related chemical toxicity, but related studies often lack functional ...

Design and Analysis of Temperature Preference Behavior and its Circadian Rhythm in Drosophila

The circadian clock regulates many aspects of life, including sleep, locomotor activity, and body temperature (BTR) rhythms1,2. We recently identified a novel Drosophila circadian output, called the temperature preference rhythm (TPR), in which the preferred temperature in flies rises during the day and falls during the night 3. Surprisingly, the TPR and locomotor activity are controlled through distinct circadian neurons3. Drosophila locomotor activity is a well&#160;known circadian behavioral output and has provided strong contributions to the discovery of many conserved mammalian circadian clock genes and mechanisms4. Therefore, understanding TPR will lead to the identification of hitherto unknown molecular and cellular circadian mechanisms. Here, we describe how to perform and analyze the TPR assay. This technique not only allows for dissecting the molecular and neural mechanisms of TPR, but also provides new insights into the fundamental mechanisms of the brain functions that integrate different environmental signals and regulate animal behaviors. Furthermore, our recently published data suggest that the fly TPR shares features with the mammalian BTR3. Drosophila are ectotherms, in which the body temperature is typically behaviorally regulated. Therefore, TPR is a strategy used to generate a rhythmic body temperature in these flies5-8. We believe that further exploration of Drosophila TPR will facilitate the characterization of the mechanisms underlying body temperature control in animals.

Design and Analysis of Temperature Preference Behavior and its Circadian Rhythm in Drosophila

We recently identified a novel Drosophila circadian output, temperature preference rhythm (TPR), in which the preferred temperature in flies rises during the day and falls during the night. TPR is regulated independently from another circadian output, locomotor activity. Here&#160;we describe the design and analysis of TPR in Drosophila.

Projeto e Análise do Comportamento de Preferência de Temperatura e seu Ritmo Circadiano em Drosophila

The circadian clock regulates many aspects of life, including sleep, locomotor activity, and body temperature (BTR) rhythms1,2. We recently identified a ...

Rapid and Efficient Zebrafish Genotyping Using PCR with High-resolution Melt Analysis

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic tools have been introduced, including multiple strategies for inducing mutations and generating transgenic lines. However, large-scale screening is limited by traditional genotyping methods, which are time-consuming and labor-intensive. Here we describe a technique to analyze zebrafish genotypes by PCR combined with high-resolution melting analysis (HRMA).&#160;This approach is rapid, sensitive, and inexpensive, with lower risk of contamination artifacts.&#160;Genotyping by PCR with HRMA can be used for embryos or adult fish, including in high-throughput screening protocols.

PCR combined with high-resolution melt analysis (HRMA) is demonstrated as a rapid and efficient method to genotype zebrafish.

Rápida e eficiente Zebrafish Genotipagem Usando PCR com alta resolução Análise Melt

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic ...

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial scientific task. Many disease causing genes in humans have a fly homologue, making Drosophila a good model to study signaling pathways involved in the development of different disorders. Additionally, the tractability of Drosophila simplifies genetic screens to aid in identifying novel therapeutic targets that may regulate metabolism. In order to perform such a screen a simple and fast method to identify changes in the metabolic state of flies is necessary. In general, carbon dioxide production is a good indicator of substrate oxidation and energy expenditure providing information about metabolic state. In this protocol we introduce a simple method to measure CO2 output from flies. This technique can potentially aid in the identification of genetic perturbations affecting metabolic rate.

Measurement of Metabolic Rate in Drosophila using Respirometry

Metabolic disorders are among one of the most common diseases in humans. The genetically tractable model organism D. melanogaster can be used to identify novel genes that regulate metabolism. This paper describes a relatively simple method which allows studying the metabolic rate in flies by measuring their CO2 production.

A medição da taxa metabólica em<em&gt; Drosophila</em&gt; Usando respirometria

Metabolic disorders are a frequent problem affecting human health. Therefore, understanding the mechanisms that regulate metabolism is a crucial ...

Measurement of Larval Activity in the Drosophila Activity Monitor

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This protocol repurposes an instrument often used to measure adult activity, the TriKinetics Drosophila activity monitor (MB5 Multi-Beam Activity Monitor) to study larval activity. The instrument can monitor the movements of animals in 16 individual 8 cm glass assay tubes, using 17 infrared detection beams per tube. Logging software automatically saves data to a computer, recording parameters such as number of moves, times sensors were triggered, and animals&#8217; positions within the tubes. The data can then be analyzed to represent overall locomotion and/or position preference as well as other measurements. All data are easily accessible and compatible with basic graphing and data manipulation software. This protocol will discuss how to use the apparatus, how to operate the software and how to run a larval activity assay from start to finish.

Measurement of Larval Activity in the Drosophila Activity Monitor

This report describes a method for measuring Drosophila larval activity using the TriKinetics Drosophila Activity Monitor. The device employs infrared beams to detect movements of up to 16 individual animals. Data can be analyzed to represent motion parameters including rates and the positions of the animals within the assay chambers.

Medição da Actividade larval no<em&gt; Drosophila</em&gt; Monitor de Atividade

Drosophila larvae are used in many behavioral studies, yet a simple device for measuring basic parameters of larval activity has not been available. This ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Quantificação de alta resolução Celulose cristalina Acumulação em<em&gt; Arabidopsis</em&gt; Raízes para monitorar celular específico do tecido da parede Modificações

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...