Name: determining temperature preference of mosquitoes and other ectotherms
Uploaded: 2022-09-28T13:50:00.000+00:00
Duration: 5 min 31 s
Description: Watch this Scientific Journal Video about Determining Temperature Preference of Mosquitoes and Other Ectotherms at JoVE.com

AHR acknowledges funding support from Majlis Amanah Rakyat (MARA).

NOTE: This protocol is written for BSL 1 or 2; for BSL 3 work, perform the entire protocol inside a class 3 biosafety cabinet (glove box).
1. Insect preparation
<ol>
	<li>Prepare two empty mosquito cages (17.5 cm x 17.5 cm x 17.5 cm) with 12 cm of sleeve openings (Figure 1). Before proceeding with the experiments, ensure there are no holes or other damage to the mosquito cages.</li>
	<li>Using a mechanical aspirator (a simple pooter with a collection chamber), transfer 30 insects (e.g., Aedes aegypti mosquitoes; here, females 3-5 days post-emergence were used), to a separate cage for easier handling and disposal after the experiment. 
	NOTE: A total of 30 insects per experiment is suggested as it is easy to manage and count without a high risk of mosquito escape. The number of insects used can be adjusted to fit the objective of the experiment.</li>
</ol>
2. Two-chamber apparatus setup
<ol>
	<li>Set the incubators to the desired temperatures, as per the incubator manufacturer&#39;s instructions.</li>
	<li>Allow the incubators to heat up and stabilize at the specific temperatures, which is &#60;30 min for temperatures in the range of 25-30 &#176;C. Check the air temperature in the incubator with a temperature probe, to ensure the incubator is set up to the intended temperature.</li>
	<li>Place an empty mosquito cage in each incubator (Figure 2A).</li>
	<li>Feed the sleeves of the cage through the front hole of the incubator. Prepare an openable cover (flap) with duct tape and place it over the hole in the acrylic tube (Figure 2B).</li>
	<li>Insert the acrylic tube into the sleeve of one cage on top of the incubator hole. The diameter of the tube is larger than the hole in the front of the incubators so that it completely covers the hole.</li>
	<li>Tighten the mesh of the sleeve around the tube with a rubber band or reusable cable tie (Figure 2C). Ensure that the acrylic tube is not loose and dangling between theincubators; if it is, pull the cage sleeves to remove excess material between the cage and rubber band.</li>
	<li>Place both incubators facing each other and repeat steps 2.5 and 2.6 with the sleeve of the other incubator. Both cages are now securely linked through the acrylic tube (Figure 2D).</li>
</ol>
3. Mosquito insertion
<ol>
	<li>Open the duct tape flap for mosquito insertion. Place a funnel into the hole. Empty the insects into the funnel that has been placed in the acrylic tube. 
	NOTE: If desired/required: for mosquitoes, use a CO2 pen to knock out all mosquitoes before placing them in the funnel21; for Drosophila, use ice to knock insects down22.</li>
	<li>Remove the funnel and cover the hole in the tube with the duct tape flap. Leave for 30 min for insects to select the preferred chamber. 
	&#8203;NOTE: If CO2 or ice was used, lightly tap the tube bridge to wake the insects after a couple of minutes.</li>
</ol>
4. Mosquito counting
<ol>
	<li>After 30 min, visually observe and write down how many insects were found in the bridge (the acrylic tube).</li>
	<li>Tap/blow the insects in the bridge to either side of the incubator. Record to deduct from the total number of insects later on. 
	NOTE: Knock out all 30 insects in the apparatus by releasing CO2 into the bridge (use CO2 for all insects as ice will not knock down insects in the cages). Also, take note of the number of insects in the bridge which fly to either side of the incubator.</li>
	<li>Pinch and close the sleeves from the acrylic tube on both sides, quickly fasten with a knot to close the cages, and ensure that the rubber band is still intact to prevent any insects from escaping.</li>
	<li>Remove the cages from the incubator, and visually count the insects in each cage (deduct the number of insects from the bridge if needed).</li>
	<li>Repeat step 4.4 with the other cage. Make sure that the numbers from the two incubators and the bridge add up to 30 (or the number of insects used, if different).</li>
	<li>If the numbers do not add up to the total number of insects used in step 1.2, look for the remaining insects in the cage sleeve.</li>
</ol>
5. Replication
<ol>
	<li>When performing experiments, be sure to account for possible external biases, such as light direction, ambient smells, etc. For example, by reversing the cages, incubator orientation and combinations between replicates.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The study describes a new method to observe the temperature preference in mosquitoes. In this method, mosquitoes are released into a tube that is connected to two incubators with independently controllable temperatures. In this manner, the mosquitoes are allowed to freely choose between two temperatures without disrupting their natural behaviors and mechanism of expressing this choice (e.g., flying).
Our first representative experiment used the mosquito optimal temperature of 27 &#176;C in both chambers. During the repetitions of this experiment, mosquitoes were observed to be freely flying between both cages for the entire 30 min, and in all replicates, there were near equal numbers in each of the two chambers. This confirmed the experimental intention of allowing the mosquitoes the ability to freely choose between cages while exhibiting their natural behaviors (flying). Conversely, the second representative experiment utilized the attractive optimal temperature of 27 &#176;C in one chamber and a sub-optimal and hence repelling temperature of 30 &#176;C in the second chamber. As expected, mosquitoes consistently selected the optimal temperature chamber at high significance, even when we swapped the incubators to avoid bias.
We also tested the setup for a different insect, D. melanogaster (fruit flies), representing another ectotherm model organism. One chamber was set to the optimum temperature of D. melanogaster, 25 &#176;C, and the other was set to 3 &#176;C higher, 28 &#176;C. Similar to mosquitoes, fruit flies also favored their optimal temperature and avoided the warmer chamber. This demonstrates that the protocol is suitable for a range of ectotherms.
Description of critical steps in the protocol 
The main critical step in the protocol is insect handling, as it generates the possibility of insects escaping. This can be prevented by determining that there are no holes large enough for escape in the cages used, that the rubber bands/cable ties used to secure the mesh sleeves to the bridge are tight, and that the cover for the insect insertion hole on the bridge is securely attached and sealed.
It is also crucial to ensure insects do not escape before or after the experiment, particularly when the insects are required for downstream experimentation or later time points for various temperature choices. This can be done by anesthetizing the insects before placing them in the acrylic bridge (using ice for Drosophila and CO2 for mosquitoes) and releasing CO2 into the bridge to knock down the insects after the experiments, prior to calculating. The usage of CO2 is ideal for mosquitoes since it won&#39;t affect the behavioral results21. In flies, exposure to CO2 can alter their flying behaviour23, hence it is recommended to use ice22.
Counting of insects is also a critical step, to ensure the numbers of insects are equal before and after the experiment for accurate results. In order to do this, we recommend the usage of a CO2 pen once the experiment is completed to knock down the insects that are located in the bridge. This will help move the insects to either side of the chamber, hence reducing the number of escapees. We also highlight in the protocol that insects can be caught in the sleeves of the cages during cage separation; therefore, ensure these are checked thoroughly during counting.
Potential modifications and troubleshooting of the technique 
The main difficulty with this technique is the flexible mesh of the cage sleeves resulting in gaps or hiding places and hence insect escape or trapping. There are some potential modifications, if needed, to improve the technique. We suggest using two or more rubber bands to ensure that the bridge is secured properly in between the chambers without leaving any potential space for the insects (loose mesh creates a hiding space for insects). We also advise particular care to pull the mesh sleeve taut, as described in step 2.6, when assembling the apparatus.
Small form factor incubators are usually heated only (i.e., have no active cooling), as was the case for the incubators used here. Consequently, using temperatures around or below the ambient room temperature will require the experiment to be performed in a cold room to ensure that the temperatures set for the incubators will go as low as desired.
In addition, this setup can also be used for BSL 3, where a class three biosafety cabinet (glove box) is needed. In this case, the glove box needs to be big enough to fit the entire apparatus. The experiment described in this protocol is ideal for experiments in a glove box because everything required will be contained within the glovebox and, importantly, the possibility of insects escaping is minimal.
Finally, there is enough space in the incubators to add external light or a humidity source without affecting the insects in the cages. Depending on the insect species or experimental design, an LED lamp with 1 cm thickness can be easily placed on top of the cage inside one or both incubators. Providing light to both and offering a temperature choice can be a more realistic protocol for some photosensitive experimental designs, or only providing light (or humidity) to one chamber is a possible modification to the protocol to assess light/humidity choice.
Advantages of this technique in the context of dual choice temperature preference assays 
The method described here presents an alternative to the traditional temperature gradient method described in previous studies10,13,14,16. In most of these studies, a large horizontal aluminum block with a thermal gradient is used, while the mechanism of generating this gradient varies, including heating/cooling blocks, water baths, etc. In these instances, the temperature gradient is produced on the surface of the aluminum block (rather than the air temperature in a cage). Consequently, most (but not all) alternative techniques do restrict the flight ability of insects more than this protocol. Here, insects can fly relatively freely between cages, allowing for a more realistic expression of natural behaviors in choice. It would even be possible to scale-up this experimental apparatus using larger cages and incubators, for example, for larger insects.
In addition to the natural behavior advantage, we also demonstrate very high temperature uniformity within the two chambers, enabling simple scoring and a clear selection of two large single temperature chambers. The use of a binary large chamber design such as this may reduce noise in the data, where, for example on a gradient apparatus, any incidental movement of the insects will alter position on the gradient and hence their perceived temperature preference.
The technique described here is also very simple and low cost. This technique does not need extra appliances to set the temperatures (i.e., a water bath10 and/or a hot plate11,12,13,14,15), no specialist equipment besides a cut acrylic tube and drilled holes, and no camera18,19 or sophisticated software19 for analysis. Such components used in other techniques can be expensive and/or require significant expertise and testing to begin experiments.
This technique can also be replicated with different devices that use batteries if there is no external power supply, making the system ideal to conduct experiments in the field. Furthermore, the same apparatus could be slightly modified to study other binary choice preference situations, such as light versus dark, high/low humidity, etc., either in the laboratory or field.
The full-sized apparatus in the protocol is significantly smaller than temperature gradient setups, allowing for an easier fit inside a BSL 3 glovebox as described above. Further, the insects are easier to contain, as they can be knocked down with CO2 at the end of the experiment, and the cages can be quickly resealed after separation from the bridge. These containment advantages are ideal for BSL 3 work.
We do however acknowledge that our apparatus only allows for a binary decision rather than a free choice along a gradient, which, depending on the application, may require additional runs to identify optimal temperatures.

Organisms can live and reproduce only within their thermal tolerance range. As environmental temperature varies due to seasons changing and global warming, species must adapt and respond accordingly to ensure their survival. This includes ectotherms, where the body temperature is in equilibrium with the environment1. Hence, each insect has their own optimal environmental temperature range which they seek to remain within2.
Temperature is one of the important factors used to predict the distribution and range of insects3,4,5, observing pathogen-insect relationships6,7 and the effect of external factors on the fitness of ectotherms such as their adult lifespan, fecundity, and feeding rate8,9.
Previous studies have investigated the preferred temperature of ectotherms with different setups. The most common is using a large aluminum block either with a cooled or heated water bath10, an ice bath and programmable heater element11, cold and hot plates12,13, thermal regulator plates14,15, or a heat pack and ice pack16 at either end to create a temperature gradient. Additionally, other studies have also used a temperature gradient incubator to study the growth of selected bacteria17 and mounted an aluminum rod on a thermoelectric device (heated and cooled at the ends) to observe the thermal preference of Drosophila melanogaster18,19.
However, the alternative methodology proposed here has significant advantages for certain insect applications. Firstly, other solutions require complete construction from scratch with basic materials, including aluminum sheets, constructing acrylic chambers for the insects, and often a camera setup and specialist software; this can be expensive and time-consuming to set up. Secondly, many alternative apparatuses rely on a temperature gradient on a surface (as opposed to air temperature). Consequently, the chamber in which the insects are studied is often very narrow (e.g., 24 cm long gradients with only 2 cm width and 1 cm depth16), which may prevent natural behaviors, such as flight, which are essential for the normal mobility of insects and hence imperative in selecting a preferred temperature. Some studies do measure the air temperature; however, the scoring of choice still involves counting the number of mosquitoes landing on the Peltier elements as opposed to insects flying freely in the cages20.
In this study, we describe a simpler setup, which uses minimally modified standard equipment and provides insects with sufficient room to fly and navigate relatively unhindered in a standard-sized colony maintenance cage. Further, rather than relying on a gradient, the protocol utilizes two relatively large-sized sections of consistent internal temperature, allowing for natural roaming of the insects at their preferred temperature and a simple binary scoring. Hence, the apparatus and protocol described here provide a low-cost and simple means of studying mosquito temperature preference in a less obstructive and more realistic setting.
The protocol involves preparation of the insects before the experiment followed by the two-chamber apparatus setup. Further steps include placing insects in the apparatus to allow the choice of temperature and scoring of results. For an illustration of the method here, we chose the optimal (standard rearing) temperature of the insects, 27 &#176;C for Aedes aegypti, 25 &#176;C for Drosophila melanogaster, and a higher repelling temperature for both species of insects, 30 &#176;C and 28 &#176;C, respectively. Insects are given 30 min to select a preferred chamber. This time was found to be sufficient, and a longer duration did not change the results; however, this may be extended depending on species/temperature/other variables as needed.

<table><tbody><tr><td>Acrylic tube (Bridge)</td><td>Perspex</td><td>900 mm OD</td><td>Size (Length x diameter): 8 cm x 9 cm; 1 cm bigger than the size of the hole in front of the incubator. Size of the hole on top: 1.6 cm</td></tr><tr><td>Carbon dioxide (CO&lt;sub&gt;2&lt;/sub&gt;) inflator</td><td>Peaken</td><td>B08HM2BDDB</td><td>Any CO&lt;sub&gt;2 &lt;/sub&gt;pen will work&amp;nbsp;</td></tr><tr><td>Digital Incubator (&amp;times;2)</td><td>VWR&amp;nbsp;</td><td>VWR INCU-Line 1L 10 (390-0384)</td><td>Size of hole in front of incubator: 8 cm diameter. Holes need to be position in the center and have the same exact position on both incubators to allow alignment of bridge.This should be pre-drilled using a standard 8 cm &amp;lsquo;holesaw&amp;rsquo; drill bit. Incubator must be just large enough to contain one mosquito cage.&amp;nbsp;</td></tr><tr><td>Mechanical aspirator (for mosquitoes)&amp;nbsp;</td><td>Watkins and Doncaster</td><td>E710</td><td>Ideal barrel size 50 x 28 mm and tube diameter 9mm.</td></tr><tr><td>Mosquito cage (&amp;times;3; two for the experiments, one for storing insects)</td><td>BugDorm</td><td>BD4S1515</td><td>Size: 17.5 cm x 17.5 cm x 17.5 cm with 12 cm sleeve opening. Mesh material : Knitted nylon</td></tr><tr><td>Plastic funnel&amp;nbsp;</td><td></td><td></td><td>Diameter of opening = 5 cm&lt;br/&gt; Length of funnel = 5 cm&lt;br/&gt; Diameter of aperture = 1 cm</td></tr><tr><td>Plastic Pocket Pooter (for Drosophila or small insects)</td><td>Watkins and Doncaster</td><td>E714</td><td>Manual/mouth aspirated&amp;nbsp;</td></tr><tr><td>Rubber band or Reusable cable tie</td><td></td><td></td><td>Either, depending on preference.</td></tr><tr><td>Temperature probe</td><td>Eidyer</td><td>B07J4T1VQZ</td><td>Any thermometer with at least 100 cm narrow wire probe</td></tr></tbody></table>

determining temperature preference of mosquitoes and other ectotherms

Most insects and other ectotherms have a relatively narrow optimal temperature window, and deviation from their optima can have significant effects on their fitness, as well as other characteristics. Consequently, many such ectotherms seek out their optimal temperature range. Although temperature preferences of mosquitoes and other insects have been well studied, the traditional experimental setup is performed using a temperature gradient on an aluminum surface in a highly enclosed space. In some cases, this equipment restricts many natural behaviors, such as flying, which may be important in preference selection.
The objective of this study is to observe insect preference for air temperature by using a two-chamber apparatus with sufficient room for flight. The two chambers consist of independent temperature-controlled incubators, each with a large aperture. The incubators are connected by these apertures using a short acrylic bridge. Inside the incubators are two netted cages, linked via the apertures and bridge, allowing the insects to freely fly between the different conditions. The acrylic bridge also acts as a temperature gradient between the two incubators.
Due to the spacious area in the cage and easy construction, this method can be used to study any small ectotherm and/or any manipulation which may alter temperature preference including sensory organ manipulation, diet, gut flora, and endosymbiont presence at biosafety levels 1 or 2 (BSL 1 or 2). Additionally, the apparatus can be used for the study of pathogen infection using further containment (e.g., inside of a biosafety cabinet) at BSL 3.

In order to test the efficacy and effectiveness of this experimental setup, 30 mosquitoes were tested with the same temperature in both incubators in four replicates (Figure 3). When both chambers were set to the mosquito optimal temperature of 27 &#176;C, there was no significant difference between chamber preference (P = 0.342; Wilcoxon signed-rank test). However, when one chamber was set to the attractive optimal temperature of 27 &#176;C and the other chamber to a sub-optimal temperature of 30 &#176;C, mosquitoes consistently demonstrated active preference toward their optima (P = 0.029; Wilcoxon signed-rank test; mean value of 78.2% and 21.8% for 27 &#176;C and 30 &#176;C, respectively). We also tested using Drosophila to determine the applicability with another ectotherm model and similar results were observed.
Temperature uniformity within cages 
Figure 4 shows the temperature uniformity of the two-chamber apparatus. Once assembled, the two sides were set to 27 &#176;C and 30 &#176;C and allowed to equilibrate as per the instructions given here. All parts of the incubator and bridge are within 0.4 &#176;C of the central temperature, except (consistently) for one corner. Note, that the front bottom left-hand corner (as viewed from the front) is a consistent hot spot at both 27 &#176;C and 30 &#176;C. This is likely due to the electronics of the incubator controls being situated just beneath that section of the incubator, rather than the manipulations performed; hence, it is likely incubator model-specific. This demonstrates that the manipulation and addition to the incubator have minimal effect on the temperature uniformity. Furthermore, the bridge temperature was intermediate between the two chambers, ensuring that insects are not confronted with a temperature trough that they would have to fly through.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64356/64356fig01.jpg" /> 
Figure 1: Description of the mosquito cage. Mosquito cage (17.5 cm x 17.5 cm x 17.5 cm) with 12 cm sleeve openings. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64356/64356fig01large.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/64356/64356fig02.jpg" /> 
Figure 2: Pictures and diagram of the apparatus during setup. (A) Empty insect cage placed in the incubator. (B) Acrylic tube with an openable cover (flap) made from duct tape. (C)&#160;Side view of the setup with a schematic diagram. The mesh of the sleeve was tightened around the acrylic tube with a rubber band. For these experiments, 3-5 day old, mated, female Ae. aegypti mosquitoes were used. (D)&#160;Complete setup. Two incubators facing each other are connected by an acrylic tube. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64356/64356fig02large.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/64356/64356fig03.jpg" /> 
Figure 3: Temperature preference in insects. The two-chamber apparatus was assembled as per the instructions. Insects were inserted as per the protocol and left for 30 min to select their preferred chamber (temperature) and then counted. Black points represent individual replicates, and blue represents the mean. (A) Both incubators were set to the same temperature (27 &#176;C) and the temperature preference of Ae. aegypti was observed. (B) Incubators were set to different temperatures (27 &#176;C vs. 30 &#176;C) and the temperature preference of Ae. Aegypti was observed. (C) Incubators were set to different temperatures (25 &#176;C vs. 28 &#176;C) and temperature preference of D. melanogaster was observed. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64356/64356fig03large.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/64356/64356fig04v2.jpg" /> 
Figure 4: Temperature uniformity within the chambers and bridge. As described, two incubators, two cages, and the bridge were assembled as per the instructions. The temperature was adjusted to 27 &#176;C on both incubators and 30 &#176;C in the center. A temperature probe was used to measure the temperature in the center of the cage, all eight corners of the incubator, and inside of the bridge. The temperatures measured are shown here. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/64356/64356fig04largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Determining Temperature Preference of Mosquitoes and Other Ectotherms at JoVE.com

Determining Temperature Preference of Mosquitoes and Other Ectotherms

Insects have an optimal environmental temperature range which they seek to remain within, and many external and internal factors can alter this preference. Here, we describe a cost-effective and simple method to study temperature choice, which allows insects to freely exhibit their natural behaviors.

Most insects and other ectotherms have a relatively narrow optimal temperature window, and deviation from their optima can have significant effects on ...

determining-temperature-preference-of-mosquitoes-and-other-ectotherms

Nanyang Technological University

Research

JoVE Journal

Biology

2.1K Views.  University of Liverpool. Deze opstelling met tweekamerapparatuur wordt gebruikt om de temperatuurvoorkeur van insecten te bestuderen en kan worden aangepast om andere binaire keuzevoorkeursituaties te bestuderen, zoals licht versus donker of hoge versus lage luchtvochtigheid. De belangrijkste voordelen van deze techniek zijn de eenvoudige opstelling, het ruime gebied waarmee de insecten hun vliegende aard kunnen vertonen bij het kiezen van hun voorkeurstemperatuur en gemakkelijk scoren zonder externe apparatuur. Het ...

Video: Determining Temperature Preference of Mosquitoes and Other Ectotherms

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific responses to changing temperatures according to their physiological tolerance and adaptability. Drosophila flies also possess a temperature sensing system that has become fundamental to understanding the neural basis of temperature processing in ectotherms. We present here a temperature-controlled arena that permits fast and precise temperature changes with temporal and spatial control to explore the response of individual flies to changing temperatures. Individual flies are placed in the arena and exposed to pre-programmed temperature challenges, such as uniform gradual increases in temperature to determine reaction norms or spatially distributed temperatures at the same time to determine preferences. Individuals are automatically tracked, allowing the quantification of speed or location preference. This method can be used to rapidly quantify the response over a large range of temperatures to determine temperature performance curves in Drosophila or other insects of similar size. In addition, it can be used for genetic studies to quantify temperature preferences and reactions of mutants or wild-type flies. This method can help uncover the basis of thermal speciation and adaptation, as well as the neural mechanisms behind temperature processing.

An Automated Method to Determine the Performance of Drosophila in Response to Temperature Changes in Space and Time

Here we present a protocol to automatically determine the locomotor performance of Drosophila at changing temperatures using a programmable temperature-controlled arena that produces fast and accurate temperature changes in time and space.

Temperature is a ubiquitous environmental factor that affects how species distribute and behave. Different species of Drosophila fruit flies have specific ...

Behavior

Exploring Life History Choices: Using Temperature and Substrate Type as Interacting Factors for Blowfly Larval and Female Preferences

Blowflies (Diptera: Calliphoridae) present a wide range of larval lifestyles, typically classified as obligate parasitism, facultative parasitism, and complete sapro-necrophagy. Several parasitic species, both obligate and facultative, are considered to be of sanitary and economic importance, as their larvae can cause myiasis (maggot infestation in live tissue). However, it is noteworthy that the adult female plays a decisive role as she chooses the oviposition site, and, therefore, largely determines the feeding habit and developmental conditions of the larvae. In this study, two protocols are proposed to test larval feeding preference and female oviposition site preference considering two interacting factors: meat substrate type and temperature. The setups presented here allowed to test Lucilia cuprina larvae and gravid females in a four-choice assay with two temperatures (33 &plusmn; 2 &#176;C and 25 &plusmn; 2 &#176;C) and two types of meat substrates (fresh meat supplemented with blood and 5-day-old rotten meat). Larvae or gravid females can choose to burrow or lay their eggs, respectively, in either of the following: rotten meat at 25 &#176;C (simulating a necrophagous species condition), fresh meat supplemented with blood at 33 &#176;C (simulating a parasitic species condition), and two controls, rotten meat at 33 &#176;C, or fresh meat supplemented with blood at 25 &#176;C. The preference is assessed by counting the number of larvae or eggs laid in each option for each replicate. Comparing the observed results to a random distribution allowed for the estimation of the statistical significance of the preference. The results indicated that L. cuprina larvae have a strong preference for the rotten substrate at 25 &#176;C. Conversely, oviposition-site preference by females was more varied for the meat type. This methodology can be adapted to test the preference of other insect species of similar size. Other questions can also be explored by using alternative conditions.

Herein, two protocols for assessing food source and oviposition preferences in larvae and females of blowflies are detailed. These comprise four choices with two interacting factors: substrate type and temperature. The assays enable the determination of the food source preference of the larvae and the oviposition site preference for the females.

Blowflies (Diptera: Calliphoridae) present a wide range of larval lifestyles, typically classified as obligate parasitism, facultative parasitism, and ...

High-Throughput Heat Tolerance Measurement in Drosophila Using DAM2 Systems

Heat Tolerance Assays Using the Drosophila Activity Monitor System: A Guide to an Executable Application for Data Analysis

The study of heat tolerance in&#8239;Drosophila melanogaster has been of particular interest to researchers for decades, with a common approach to assessing heat tolerance being to monitor the time to knockdown (TKD) after exposure to an elevated temperature. Classically, flies are housed in individual vials and placed inside a heated water bath. TKD is then monitored manually by researchers. While very well-established, there remain problems of subjectivity and consistent application of a tangible definition of cessation of all movement, including muscular spasms, when implementing these manual assays. We have developed a high-throughput method for automating heat tolerance assays using the TriKinetics Drosophila Activity Monitors (DAM2). To accompany the DAM2 system, we have written a program and created an easy-use executable to automatically read the last time of movement from the activity data generated. This script then writes to a .csv file the time to heat paralysis (TKD) for each fly. Our data show that this automated DAM2 method is consistent and reliable. Meanwhile, activity profiles created from the activity count data are of interest. These activity profiles can be compiled and have the potential to expand heat tolerance assays to include the relatively unstudied behavioral components of heat tolerance. This protocol will describe in detail how to use the DAM2 system and the HoTDAM! software to estimate heat tolerance in D. melanogaster.

This report describes a method for measuring adult Drosophila melanogaster time to knockdown using a Drosophila Activity Monitor (DAM2) in response to an air conduction heat stressor within an incubator chamber. The DAM2 measures activity by recording individual fly movements as they cross an infrared beam. Data analysis is facilitated by a novel executable file created by the authors.

The study of heat tolerance in&#8239;Drosophila melanogaster has been of particular interest to researchers for decades, with a common approach to ...

A Temperature Gradient Assay to Determine Thermal Preferences of Drosophila Larvae

Many animals, including the fruit fly, Drosophila melanogaster, are capable of discriminating minute differences in&#160;environmental temperature, which enables them to seek out their preferred thermal landscape. To define the temperature preferences of larvae over a defined linear range, we developed an assay using a temperature gradient. To establish a single-directional gradient, two aluminum blocks are connected to independent water baths, each of which controls the temperature of individual blocks. The two blocks set the lower and upper limits of the gradient. The temperature gradient is established by placing an agarose-coated aluminum plate over the two water-controlled blocks so that the plate spans the distance between them. The ends of the aluminum plate that is set on the top of the water blocks defines the minimum and maximum temperatures, and the regions in-between the two blocks form a linear temperature gradient. The gradient assay can be applied to&#160;larvae of different ages and can be used to identify mutants that exhibit phenotypes, such as those with mutations affecting genes encoding TRP channels and opsins, which are required for temperature discrimination.

A Temperature Gradient Assay to Determine Thermal Preferences of Drosophila Larvae

Here, we present a protocol to determine the preferred environmental temperature of Drosophila larvae using a continuous thermal gradient.

Many animals, including the fruit fly, Drosophila melanogaster, are capable of discriminating minute differences in&#160;environmental temperature, which ...

Neuroscience

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.

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

Adapting Microrespirometry for Drosophila - Insights into CASK Mutant Metabolism

Measuring O2 Consumption in Drosophila melanogaster Using Coulometric Microrespirometry

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable environment. A coulometric microrespirometer consists of an airtight chamber in which O2 is consumed and the CO2 produced by the organism is removed by an absorbent medium. The resulting pressure decrease triggers electrolytic O2 production, and the amount of O2 produced is measured by recording the amount of charge used to generate it. In the present study, the method has been adapted to Drosophila melanogaster tested in small groups, with the sensitivity of the apparatus and the environmental conditions optimized for high stability. The amount of O2 consumed by wildtype flies in this apparatus is consistent with that measured by previous studies. Mass-specific O2 consumption by CASK mutants, which are smaller and known to be less active, was not different from congenic controls. However, the small size of CASK mutants resulted in a significant reduction in O2 consumption on a per-fly basis. Therefore, the microrespirometer is capable of measuring O2 consumption in D. melanogaster, can distinguish modest differences between genotypes, and adds a versatile tool for measuring metabolic rates.

Author Spotlight: Influence of Temperature on Drosophila melanogaster and Desert-Adapted Beetles 

Coulometric respirometry is ideal for measuring the metabolic rate of small organisms. When adapted for Drosophila melanogaster in the present study, measured O2 consumption was within the range reported for wildtype D. melanogaster by previous studies. Per-fly O2 consumption by CASK mutants, which are smaller and less active, was significantly lower than the wildtype.

Coulometric microrespirometry is a straightforward, inexpensive method for measuring the O2 consumption of small organisms while maintaining a stable ...

Forced Salivation As a Method to Analyze Vector Competence of Mosquitoes

Vector competence is defined as the potential of a mosquito species to transmit a mosquito-borne virus (mobovirus) to a vertebrate host. Viable virus particles are transmitted during a blood meal via the saliva of an infected mosquito. Forced salivation assays allow determining the vector potential on the basis of single mosquitoes, avoiding the use of animal experiments. The method is suitable to analyze a large number of mosquitoes in one experiment within a short period of time. Forced salivation assays were used to analyze 856 individual mosquitoes trapped in Germany, including two different Culex pipiens pipiens biotypes, Culex torrentium as well as Aedes albopictus, which were experimentally infected with Zika virus (ZIKV) and incubated at 18 &#176;C or 27 &#176;C for two and three weeks. The results indicated the lack of vector competence of the different Culex taxa for ZIKV. In contrast, Aedes albopictus was susceptible to ZIKV, but only at 27 &#176;C, with transmission rates similar to an Aedes aegypti laboratory colony tested in parallel.

For efficient control of mosquito borne virus transmission, the knowledge of the vector potential of respective mosquitoes is of particular interest. We describe forced salivation as a method to analyze vector competence of Aedes albopictus and three different Culex taxa for the transmission of Zika virus.

Vector competence is defined as the potential of a mosquito species to transmit a mosquito-borne virus (mobovirus) to a vertebrate host. Viable virus ...

A Blood-Free Diet to Rear Anopheline Mosquitoes

Malaria research requires large-scale breeding and production conditions for mosquitoes (Anopheles spp.) in captivity. The sustainable and reliable production of mosquitoes is currently inhibited by the supply of fresh vertebrate blood. Alternatives to blood are required to promote efficient control strategies for malaria and other vector borne diseases that are transmitted by blood feeding insects. With this in mind, artificial liquid diets were formulated as substitutes for fresh vertebrate blood. Herein we report a blood-free artificial liquid diet that delivers feeding rates similar to blood and mimics the physiological effects of a fresh vertebrate blood meal. The diet induces ovarian and egg maturation of Anopheles mosquitoes and also produces good larval survival and development of functional adults. The formulated blood-free liquid diet is an important advance towards sustainable mosquito breeding in captivity and will reduce the maintenance costs of mosquito colonies and eliminate the need for fresh vertebrate blood.

A protocol is presented for formulation of a blood-free artificial diet to feed Anopheles mosquitoes in captivity. This diet has a similar performance to vertebrate blood and triggers oogenesis and egg maturation and produces viable adult progeny.

Malaria research requires large-scale breeding and production conditions for mosquitoes (Anopheles spp.) in captivity. The sustainable and reliable ...

High-Throughput Assays of Critical Thermal Limits in Insects

Upper and lower thermal limits of plants and animals are important predictors of their performance, survival, and geographic distributions, and are essential for predicting responses to climate change. This work describes two high-throughput protocols for measuring insect thermal limits: one for assessing critical thermal minima (CTmin), and the other for assessing heat knock down time (KDT) in response to a static heat stressor. In the CTmin assay, individuals are placed in an acrylic-jacketed column, subjected to a decreasing temperature ramp, and counted as they fall from their perches using an infrared sensor. In the heat KDT assay, individuals are contained in a 96 well plate, placed in an incubator set to a stressful, hot temperature, and video recorded to determine the time at which they can no longer remain upright and move. These protocols offer advantages over commonly used techniques. Both assays are low cost and can be completed relatively quickly (~2 h). The CTmin assay reduces experimenter error and can measure a large number of individuals at once. The heat KDT protocol generates a video record of each assay and thus removes experimenter bias and the need to continuously monitor individuals in real time.

Thermal limits can predict the environments organisms tolerate, which is valuable information in the face of rapid climate change. Described here are high-throughput protocols to assess critical thermal minima and heat knockdown time in insects. Both protocols maximize the throughput and minimize the cost of the assays.

Upper and lower thermal limits of plants and animals are important predictors of their performance, survival, and geographic distributions, and are ...

A Step-by-Step Guide to Mosquito Electroantennography

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when acquiring a blood-meal. To locate a host to feed on, mosquitoes rely on a wide range of sensory cues, including visual, mechanical, thermal, and olfactory. The study details a technique, electroantennography (EAG), that allows researchers to assess whether the mosquitoes can detect individual chemicals and blends of chemicals in a concentration-dependent manner. When coupled with gas-chromatography (GC-EAG), this technique allows to expose the antennae to a full headspace/complex mixture and determines which chemicals present in the sample of interest, the mosquito can detect. This is applicable to host body odors as well as plant floral bouquets or other ecologically relevant odors (e.g., oviposition sites odorants). Here, we described a protocol that permits long durations of preparation responsiveness time and is applicable to both female and male mosquitoes from multiple genera, including Aedes, Culex, Anopheles, and Toxorhynchites mosquitoes. As olfaction plays a major part in mosquito-host interactions and mosquito biology in general, EAGs and GC-EAG can reveal compounds of interest for the development of new disease vector control strategies (e.g., baits). Complemented with behavioral assays, the valence (e.g., attractant, repellent) of each chemical can be determined.

The present article details a step-by-step protocol for successful and low noise electroantennograms in several genera of mosquitoes, including both females and males.

Female mosquitoes are the deadliest animals on earth, claiming the lives of more than 1 million people every year due to pathogens they transmit when ...

Determining Temperature Preference of Mosquitoes and Other Ectotherms

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