We thank Ellie McCabe for assistance with fly rearing. This work is supported by United States Department of Agriculture National Institute of Food and Agriculture Hatch Project grant 1010996 and National Science Foundation grant OIA-1826689 to N.M.T.

1. High-throughput CTmin assay
<ol>
	<li>Assembling the jacketed column (Figure 1A, Figure 2)
	<ol>
		<li>Cut the widest (7 cm x 6.35 cm x 0.3 cm) and narrowest (5.7 cm x 5.1 cm x 0.3 cm) acrylic tubes to equal lengths (31.5 cm) with a hacksaw (Figure 2A). These two tubes will be the outer and innermost walls of the jacketed column.</li>
		<li>Cut two rings (2 cm wide) from the midsized (6.35 cm x 5.7 cm x 0.3 cm) acrylic tube with a hacksaw (Figure 2A). These two rings will be the spacers between the inner and outermost tubes, creating a space between the two long acrylic tubes for fluid to flow.</li>
		<li>Carefully drill two holes in the outer (widest) acrylic tube, one hole at the top and one at the bottom. Ensure that each hole is 3.5 cm from the end of the tube. Drill the holes on opposite sides of the tube (Figure 2B).</li>
		<li>To reduce cracking, place tape on the tube over the spot of the future hole and drill very slowly on the lowest torque setting of the drill.</li>
		<li>Using the threading tap, thread both holes so that the hose adapters can be screwed into the two holes of the outer tube. To reduce cracking, use lubricant, and thread slowly by hand.</li>
		<li>Slide the two spacers onto the inner jacket, one at each end (bottom and top). Leave a small space (0.5 cm) between the spacer and the end of the inner jacket (Figure 2B).</li>
		<li>Weld the spacers into place using acrylic cement.</li>
		<li>After the cement on the inner tube and spacers sets, slide this construct into the larger outer tube with the holes. Ensure that the outer and inner tubes are flush on both ends. The spacers will be 0.5 cm from the end, forming small trenches on both ends of the column (Figure 2C).</li>
		<li>Weld the outer tube to the spacers using acrylic cement, using adjustable steel clamps to hold the apparatus together. Wait for the cement to set.</li>
		<li>Thread the hose adapters into the holes of the outer tube now secured to the spacers and inner tube. 
		NOTE: The adapters should only thread into the outer tube and not into the open space between the inner and outermost tubes. If the hose adapters thread too far in, shorten them to the appropriate length with a hacksaw.</li>
		<li>Seal the hose adapters into their threads on the outer tube with silicone sealant.</li>
		<li>Fill the two trenches between the inner and outermost tubes at both ends of the jacketed column with silicone sealant.</li>
		<li>To test the column, attach 0.6 cm diameter tubing to the hose adapters. Connect the adapter at the bottom of the column to a water source with tubing, and the adapter at the top of the column to a drain with a different piece of tubing.</li>
		<li>Run water through the apparatus from the bottom to the top and check for leaks. If there are leaks, identify where they are coming from and seal with silicone.</li>
	</ol>
	</li>
	<li>Setting up the jacketed column and Drosophila Funnel Monitor (DFM)
	<ol>
		<li>Secure the jacketed column to a retort stand with a three-prong retort clamp. Align the column vertically with one end open to the ceiling and the other open to the lab bench (Figure 1B).</li>
		<li>Connect the fluid input and output from a temperature-controlled refrigerated bath to the adapter nozzles of the column with 0.6 cm diameter plastic tubing (Figure 1B). Connect the fluid input to the nozzle at the bottom of the column and the fluid output to the nozzle at the top of the column.</li>
		<li>Cut two 3 cm thick circular foam insulating plugs (the same circumference as the opening of the innermost compartment of the column). Ensure the plugs fit snugly and seal the innermost column when inserted at both ends (Figure 1B).</li>
		<li>Pierce a hole through the center of one of the plugs and thread the bare end of a thermocouple through the hole about 5 cm and secure with tape. Plug the other end of the thermocouple into a thermocouple data logger.</li>
		<li>Connect the thermocouple data logger to the computer.</li>
		<li>Wedge two pieces of plastic gutter guard (5 cm x 7 cm, with ~0.5 cm diameter openings) inside the column to function as perching material. Place one piece of guard 2/3rds from the top of the column and the other 1/3rd from the top of the column (Figure 1B).</li>
		<li>Secure the bottom plug (without a thermocouple) and the top plug (with a thermocouple). When the plug is inserted at the top of the column, ensure that the thermocouple does not touch the sides of the column.</li>
		<li>Adjust the height of the column on the retort stand so there is a 25 cm distance between the bottom of the column and the bench top.</li>
		<li>Secure a retort ring (5 cm diameter) to the retort stand 5 cm below the bottom of the column and rotate the ring off to the side of the column.</li>
		<li>Set the DFM directly on the retort ring (Figure 1B). Connect all the electronic components: the power supply, power supply interface, and the computer according to the manufacturer&#39;s protocol.</li>
		<li>Once the components are connected, follow the manufacturer&#39;s protocol to finish the setup of the DFM and DFM software.</li>
	</ol>
	</li>
	<li>CTmin assay
	<ol>
		<li>Turn the input and output valves of the fluid bath to the open positions.</li>
		<li>Push the power button to turn on the temperature-controlled fluid bath and then press the play button to run a program raising and maintaining the temperature of the bath to 25 &#176;C. Give the fluid bath and column 5-10 min to reach and maintain 25 &#176;C.</li>
		<li>Remove the plug at the top of the column and replace it with a funnel (5.08 cm diameter; see Figure 1C).</li>
		<li>Tap flies from their food vial into the column.</li>
		<li>Remove the funnel and replace it with the top plug quickly, careful not to let flies escape. Give the flies 5 min to settle, occasionally tapping the bottom plug to encourage the flies to climb.</li>
		<li>Press the start button on the fluid bath and begin the CTmin ramping program (25 &#176;C for 5 min; 25 &#176;C to 10 &#176;C at 0.5 &#176;C/min; 10 &#176;C for 2 min; then 10 &#176;C to -10 &#176;C at 0.25 &#176;C/min). 
		NOTE: Other variations of this CTmin ramping protocol can be used depending on the research question (e.g., comparisons of the effects of different ramping rates on CTmin28).</li>
		<li>Click open the thermocouple recording software on the computer and then click the Record icon to begin recording the temperature inside the column every second for the duration of the assay. Ensure that each temperature recording includes a time stamp specific to the second, so that temperature data can later be merged with data from the DFM.</li>
		<li>Add 5 mL of 90% ethanol to a 15 mL conical centrifuge tube and place it in a rack below the column.</li>
		<li>Occasionally, tap the bottom plug of the column to entice any flies on the bottom to climb. Most flies will be on a perch or near the top of the column by 15 &#176;C.</li>
		<li>At 15 &#176;C, remove the bottom plug and collect any flies still on the bottom plug in the ethanol. Count and note that these flies were collected at 15 &#176;C but their CTmin is unknown. 
		NOTE: The temperature at which the bottom plug is removed should be decided before the assay and based on the predicted CTmin of the test species or treatment. For this assay, 15 &#176;C was chosen based on the CTmin of these particular DGRP lines found in preliminary assays.</li>
		<li>Place a 75 mm outer diameter glass funnel into the DFM. Adjust the retort ring, DFM, and funnel so that they are under the column. Ensure that the lip of the funnel completely seals the bottom of the column (Figure 1D).</li>
		<li>Insert the bottom of the funnel into the 15 mL collection tube (Figure 1D).</li>
		<li>Open the DFM software on the computer by clicking the Software icon. The software will immediately start recording the time/date at which flies reach their CTmin. Flies that reach their CTmin lose neuromuscular function and fall from their perches, and thereafter through the DFM.</li>
		<li>Monitor whether all the flies have reached their CTmin as the temperature decreases by checking the top plug and perches to see if any flies are still perched (i.e., still maintaining neuromuscular function). The trial ends when all the flies have reached their CTmin.</li>
		<li>At the end of the trial, adjust the DFM and funnel away from the column opening. Some flies may reach their CTmin but remain stuck in the column (i.e. wedged in a perch or dangling by a single tarsal hook). Open the top plug and remove these flies. The CTmin of these flies is unknown.</li>
		<li>Combine the .txt output files from the thermocouple recording software (i.e., temperature, date, and time) and the DFM software (i.e., number of flies through the funnel, date, and time) using the Merge command in RStudio. Merge the two files based on the shared date/time variable.</li>
	</ol>
	</li>
</ol>
2. High-throughput heat KD assay
<ol>
	<li>Apparatus assembly and preparation
	<ol>
		<li>With an adhesive, fix the steel woven wire mesh (~1.5 mm aperture) to the bottom of a 96 well no-bottom plate.</li>
		<li>Attach magnets to the opposite sides of the bottom of a 96 well no-bottom plate with a hot glue gun and hot glue (Figure 3).</li>
		<li>To craft a custom septum lid with adhesive film designed for 96 well plates, stick two films sticky sides together to form a ridged plastic sheet.</li>
		<li>Place the plastic sheets over the 96 well plate and use tape to adhere it to all four sides of the plate. Over the opening to each well on the plate, cut an &#39;x&#39; in the plastic sheet with a box cutter (i.e., 96 total x&#39;s).</li>
		<li>Anesthetize flies with CO2 and load them individually into each well of the modified 96 well no-bottom plate with an aspirator and septum lid. Remove the septum lid from the 96 well plate while the flies are anesthetized with CO2 and replace it with a tight-fitting clear lid.</li>
		<li>Place the 96 well no-bottom plate loaded with flies and with a clear tight-fitting lid on food. Ensure the flies have at least 48 h between CO2 anesthetization and the start of the assay (steps 2.2.1-2.2.5). 
		NOTE: The bottom of the modified 96 well no-bottom plates is made of mesh, so flies anaesthetized with CO2 can be loaded and left on food for at least 48 h before a trial begins. Any plastic container &#62;8.5 cm wide x 13 cm long that is at least 2 cm deep to accommodate a 1 cm deep layer of food can be used.</li>
		<li>Fix a webcam to the bottom of the inside of a temperature-controlled incubator with tape. Point the camera directly up (Figure 4). Secure an incubator shelf about 10 cm above the camera. 
		NOTE: The webcam points up and records the 96 well plate from below to ensure as much of the well surface is in view as possible (e.g., not blocked out from view by the well walls of the plate). When the flies reach their KDT they fall to the bottom of the well; in this case, the side closest to the webcam, and are therefore in view regardless of how far their well is from the center of view.</li>
		<li>Connect the webcam to a computer.</li>
		<li>With tape, attach a white sheet of paper (8.5 cm x 13 cm; the exact area of the bottom of the 96 well plate) to the bottom of the shelf (Figure 4). Ensure that the paper fills the entire frame when viewed through the webcam.</li>
		<li>Place a light source in the incubator. Use paper or other materials to dampen the lighting and minimize glare. 
		NOTE Step 2.1.10 is specific to each incubator because lighting and reflections vary among incubators. The goal is to have sufficient lighting in the incubator to provide a good contrast between the flies in each well and the white sheet of paper behind the plate when viewed with the webcam.</li>
	</ol>
	</li>
	<li>Performing the heat KD assay
	<ol>
		<li>Set the incubator to 37.5 &#176;C and wait about 30 min to give the incubator time to reach and maintain the desired temperature. The exact temperature will depend on the insect being assessed and any other time considerations.</li>
		<li>Place the 96 well plate inverted in the incubator, such that the bottom of the plate (mesh side) is against the white paper on the bottom of the tray (Figure 4). Take note of the orientation of the wells (column and row names) on the tray and in the frame of the webcam. Colored tape along the sides of the 96 well plate and edges of the white piece of paper can verify the orientation (Figure 4). 
		NOTE: Ensure that the incubator temperature is consistent with the temperature inside the 96 well plate by recording the temperature inside the plate with a thermocouple during a test trial of the heat KD assay. It is also prudent to check that there is negligible variation in temperature between wells of the 96 well plate with multiple thermocouples before conducting the heat KD assay.</li>
		<li>Close the incubator door.</li>
		<li>Click Record on the video recording software.</li>
		<li>After 2 h, check the recording to see that all flies have reached their final resting spot and stopped moving. Once all flies have stopped moving, click Stop on the video recording software. For the genotypes tested here, reared at 25 &#176;C, most flies reach their KDT by 60 min at 37.5 &#176;C (also see J&#248;rgensen et al.9).</li>
		<li>Dispose of the flies.</li>
		<li>Use the custom Python scripts (Supplementary Coding Files 1-3) to approximate the time in the video when flies reach their heat KDT. 
		NOTE: Step 2.2.7 is optional. To speed up video analysis, a set of custom Python scripts were developed to measure changes in pixel density over time in each well (see Supplementary Coding File). When the flies stop moving, the pixel density is constant, and a plot of these data can be used to locate the approximate time in the video when flies are knocked down. While it may be possible to use this script to automate data analysis, currently slight imperfections in the video lead to minor discrepancies between changes in pixel density and the true KD time.</li>
		<li>Click open the video file and record the KDT of each fly in each well. The most consistent measure of heat KDT between trials and observers is recording the time at which a fly reaches its final resting spot.</li>
		<li>Track the video in reverse, focusing on a single well, and noting the time at which the fly first moves off its final resting spot. Repeat this process for each well.</li>
	</ol>
	</li>
</ol>

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G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Critical+thermal+limits+affected+differently+by+developmental+and+adult+thermal+fluctuations.">Critical thermal limits affected differently by developmental and adult thermal fluctuations.</a> Journal of Experimental Biology. 220 (23), 4471-4478 (2017).</li><li>Hoffmann, A. A., Hallas, R., Anderson, A. R., Telonis-Scott, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evidence+for+a+robust+sex-specific+trade-off+between+cold+resistance+and+starvation+resistance+in+Drosophila+melanogaster.">Evidence for a robust sex-specific trade-off between cold resistance and starvation resistance in Drosophila melanogaster.</a> Journal of Evolutionary Biology. 18 (4), 804-810 (2005).</li><li>Kelty, J. D., Lee, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+cold-hardening+of+Drosophila+melanogaster+(Diptera:+Drosophilidae)+during+ecologically+based+thermoperiodic+cycles.">Rapid cold-hardening of Drosophila melanogaster (Diptera: Drosophilidae) during ecologically based thermoperiodic cycles.</a> Journal of Experimental Biology. 204 (9), 1659-1666 (2001).</li><li>Sinclair, B. J., Vernon, P., Klok, C. J., Chown, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Insects+at+low+temperatures:+An+ecological+perspective.">Insects at low temperatures: An ecological perspective.</a> Trends in Ecology and Evolution. 18 (5), 257-262 (2003).</li></ol>

The authors have nothing to disclose.

The two methods detailed above generate high-throughput data of ecologically relevant metrics for upper and lower thermal limits. These protocols build upon previously established methodologies common to research on insect thermal limits (summarized in Sinclair et al.)6. Both protocols can be completed in a short amount of time (~2 h each), produce data sets with large sample sizes, do not sacrifice repeatability or accuracy, and minimize experimenter error by eliminating manual data record...

Thermal limits of insects 
Variation in environmental conditions, including temperature, is a major factor influencing the performance, fitness, survival, and geographic distribution of organisms1,2. Upper and lower thermal limits determine the theoretical range of environments an organism can tolerate, and, therefore, these limits are important predictors of plant and animal distributions, especially in the face of climate change3,4. Thus, protocols to accurately measure thermal limits are important tools for ecologists, physiologists, evolutionary biologists, and conservation biologists, among others.
As the most abundant and diverse terrestrial animals, insects are frequently used for measurements of thermal limits.&#160;Critical thermal maxima (CTmax) and critical thermal minima (CTmin) are commonly used to assess intra- and interspecific variation in thermal tolerance5,6,7. While CTmax and CTmin can be measured for multiple phenotypes, including growth, reproductive output, and behavior, they are most commonly applied to locomotor function5,6,7. Thus, CTmax (also called heat knockdown temperature) and CTmin are often defined as the high and low temperatures at which insects lose motor function and are unable to remain upright5,6,7,8,9,10,11. CTmin coincides with the onset of chill coma, a reversible paralysis brought on by cold temperatures6. While paralysis at the thermal limits is often reversible, continued exposure to these temperatures leads to ecological death5.
Common methods for measuring thermal limits 
A variety of apparatuses have been used to measure thermal limits (summarized in Sinclair et al.)6. Briefly, insects are heated or cooled in incubators12,13, containers submerged in fluid baths11,14,15,16, aluminum blocks10,17, or jacketed containers18, and monitored until locomotion ceases. To monitor insects during the assay, the most common method is direct observation, in which individuals are continuously monitored in real time or retrospectively with recorded video6,9,10,11,15,17. While direct observation methods have minimal equipment requirements, they are labor-intensive and limit throughput. Alternatively, insects can be observed indirectly by collecting individuals at discrete times as they fall from perches6,19,20,21 or using activity monitors13.
Indirect methods for measuring thermal limits are generally higher-throughput and potentially less error prone than direct observation methods. The most common method for indirect monitoring uses a jacketed temperature-controlled column6,8,19,20,21. Insects are placed inside a column with perches, and the temperature of the inner chamber is controlled by pumping fluid from a temperature-controlled fluid bath through the jacketed lining of the column. Individuals that reach their thermal limit fall from their perch and are collected at discrete temperatures or time intervals. While this method works well for CTmin, it has been found unsuitable for CTmax, because flies voluntarily walk out of the bottom of the column when the temperature increases. The new method described here circumvents this issue by individually containing flies during automated measurements.
In addition to the method of observation, two types of temperature regimes are commonly used to assess upper thermal limits. Dynamic assays consist of gradually increasing temperature until motor function is lost; that temperature is the dynamic CTmax7,8,9,13. In contrast, static assays consist of a constant stressful temperature until motor function is lost; that time point is the heat knockdown time (heat KDT), also called the static CTmax (sCTmax) in a recent paper by J&#248;rgensen et al.7,8,9,16,22. Although CTmax and heat knockdown assays (heat KD assays) produce metrics with different units, mathematical modeling of the two traits indicates they give comparable information on heat tolerance and are both ecologically relevant8,9. Dynamic assays yield a temperature that can be compared to environmental conditions, and they are preferable when there are large differences in heat tolerance, such as comparisons between species with widely different thermal niches. However, due to the high Q10 for heat injury accumulation, a static assay may be preferable for detecting small effect sizes, such as intraspecific variation in heat tolerance9. Also, practically speaking, a static assay requires less sophisticated equipment than a dynamic assay.
Objective 
The objective of this paper is to formalize methods for CTmin and heat KD assays that can be used in future research to assess the thermal limits of motile insects. The protocols are adapted from previously established methodologies and are designed to be high-throughput, automated, and cost-effective. Both assays can be completed in a short amount of time (~2 h), which means that multiple experiments can be conducted in a single day, producing large amounts of data without sacrificing repeatability or accuracy. With this setup, the heat tolerance of 96 flies can be measured simultaneously, while the column for CTmin can hold more than 100 flies, provided there is adequate surface area for perching.
The high-throughput method for observing CTmin modifies the common jacketed column methodology with the addition of an infrared sensor to automatically count flies. The use of an infrared sensor for counting was first proposed by Shuman et al. in 199623 but has not been widely adopted. The addition of the infrared sensor allows for the generation of continuous data rather than collecting data at discrete intervals. This protocol also minimizes experimenter error by eliminating manual data entry and the need to manually switch collection tubes below the jacked column at discrete time points.
The high-throughput method for recording heat KDT is modified from two previous studies of heat tolerance in insects10,12. Individual flies are stored in a 96 well plate in a temperature-controlled incubator and video is recorded. This protocol minimizes experimenter bias in determining heat KDT because experiments can be reviewed and verified by playing back the recording. This protocol also provides a set of custom Python scripts that can be used to speed up video analysis. The use of individual wells eliminates interference that can occur when other individuals move around or fall over, which can be a problem when groups of individuals are observed in the same arena10,17. Furthermore, the temperature-controlled incubator provides a stable temperature across all 96 wells, unlike the temperature gradient sometimes observed across a temperature-controlled aluminum block10. Also note that the 96 well recording method can be adapted to measure dynamic CTmax and potentially CTmin (see Discussion).
To demonstrate each protocol, the thermal limits of adult Drosophila melanogaster females from select lines of the Drosophila melanogaster Genetic Reference Panel (DGRP) were compared24. These lines were selected because preliminary experiments indicated significant differences in thermal tolerance. These assays proved to be robust methods for discriminating differences in thermal tolerance. The following two protocols, high-throughput CTmin assay (section 1) and high-throughput heat KD assay (section 2), describe the necessary actions to produce CTmin and heat KDT data for any motile insect life stage capable of fitting in the apparatuses, such as adult Drosophila. For CTmin it is also essential that the insect be able to perch. Here, each assay is demonstrated in adult Drosophila melanogaster. However, modifications may be required for other taxa or life stages6. Minor changes might include using perching material with larger openings to accommodate larger specimens in the CTmin assay or using a higher quality camera to discern the subtle KDT of a slow moving insect or life stage in the heat KD assay. This protocol does not describe methods for preparing flies, but it is important to standardize rearing protocols to ensure repeatability25 (see Garcia and Teets26 and Teets and Hahn27). The protocols provided include information on how to build and set up the apparatuses, how to record measurements, and a brief description of data analysis.

<table><tbody><tr><td>ARCTIC A40 Refrigerated fluid circulator (Programable teperature ramps)</td><td>Thermo Scientific; Waltham, MA</td><td>153-5401</td><td></td></tr><tr><td>C922 Pro Stream Webcam</td><td>Logitech; Newark, CA</td><td>960-001087</td><td></td></tr><tr><td>Circular adjustable steel clamp &amp;ndash; 5.08 cm to 7.62 cm</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Clear acrylic tubing &amp;ndash; 5.7 cm x 5.1 cm x 0.3 cm</td><td>United States Plastic Corp., OH</td><td>44036</td><td></td></tr><tr><td>Clear acrylic tubing &amp;ndash; 6.35 cm x 5.7 cm x 0.3 cm</td><td>United States Plastic Corp., OH</td><td>440515</td><td></td></tr><tr><td>Clear acrylic tubing &amp;ndash; 7 cm x 6.35 cm x 0.3 cm</td><td>United States Plastic Corp., OH</td><td>44041</td><td></td></tr><tr><td>Clear silicone sealant</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Collection tube (15 ml)</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Cordless Drill</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Drosophila Funnel Monitor (DFM)</td><td>TriKinetics; Waltham, MA</td><td>DFM</td><td>Used to count the number of flies that fall through the funnel at a given time point</td></tr><tr><td>DAM data collection software</td><td>TriKinetics; Waltham, MA</td><td></td><td>Records data input from the DFM</td></tr><tr><td>Fly Storage Lid</td><td>FlySorter; Seatle, WA</td><td>FS-96LID-5PK</td><td>Used to load flies into the storage plate for the sCTmax assay</td></tr><tr><td>Fly Storage Plate</td><td>FlySorter; Seatle, WA</td><td>FS-96PLATE-5PK</td><td>Used to hold flies during in the sCTmax assay</td></tr><tr><td>Fly Food Tray</td><td>FlySorter; Seatle, WA</td><td>FS-TRAY-5PK</td><td>Used to keep flies on food after loading into the 96-well plate until the sCTmax assay</td></tr><tr><td>Glass funnel</td><td>Kimax</td><td>28950-75</td><td>75mm</td></tr><tr><td>Gutter guard</td><td>Any</td><td>Any</td><td>~0.5 cm diameter openings</td></tr><tr><td>Hacksaw</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Heratherm Thermo Scientific incubator</td><td>Thermo Scientific; Waltham, MA</td><td>OMS100</td><td></td></tr><tr><td>Hose nylon adapters (2) &amp;ndash; &amp;frac14; MNPT x 3/8</td><td>United States Plastic Corp., OH</td><td>61135</td><td></td></tr><tr><td>Hot glue gun and glue</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Light Source</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Magnets</td><td>Any</td><td>Any</td><td></td></tr><tr><td>OMEGA TC-08 Recorder and TC-08 Player Software</td><td>OMEGA; Norwalk, CT</td><td></td><td></td></tr><tr><td>OMEGA thermocouple (Type T)</td><td>OMEGA; Norwalk, CT</td><td>5LRTC-TT-K-20-36</td><td></td></tr><tr><td>Plastic funnel</td><td>Any</td><td>Any</td><td>2&quot; diameter</td></tr><tr><td>Plastic tubing - 0.6 cm diameter</td><td>United States Plastic Corp., OH</td><td>62852</td><td></td></tr><tr><td>Retort ring</td><td>Any</td><td>Any</td><td>2&quot; diameter</td></tr><tr><td>Retort stand</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Retort three-prong clamp</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Rstudio</td><td></td><td></td><td></td></tr><tr><td>Serial port connector (PSIU9)</td><td>TriKinetics; Waltham, MA</td><td>PSIU9</td><td>Intermediate connection between the DFM and computer, allows for multiple DFM connections</td></tr><tr><td>Styrofoam (2&quot; thick)</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Tape</td><td>Any</td><td>Any</td><td></td></tr><tr><td>Uninterrupted Power Supply (PS9-1)</td><td>TriKinetics; Waltham, MA</td><td>PS9-1</td><td>Power supply for the DFM and PSIU9</td></tr><tr><td>Weld-on #4 Acrylic Cement</td><td>United States Plastic Corp., OH</td><td>45737</td><td></td></tr></tbody></table>

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.

The thermal limits (i.e., CTmin and heat KDT) of females from the Drosophila melanogaster Genetic Reference Panel (DGRP) were measured to demonstrate the high-throughput data generated from the two described protocols. CTmin was assayed using the DGRP lines 714 (n = 37) and 913 (n = 45). Heat KDT was assayed and compared with the DGRP lines 189 (n = 42) and 461 (n = 42), and video files were manually analyzed. The total time of the experiments, including wat...

Watch this Scientific Journal Video about High-Throughput Assays of Critical Thermal Limits in Insects at JoVE.com

High-Throughput Assays of Critical Thermal Limits in Insects

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 ...

high-throughput-assays-of-critical-thermal-limits-in-insects

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

5.0K Views.  University of Kentucky. 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.

Video: High-Throughput Assays of Critical Thermal Limits in Insects

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.
The Drosophila larva is an established model system to study thermal nociception, a sensory response to potentially harmful temperatures that is evolutionarily conserved across species1,2. The advantages of Drosophila for such studies are the relative simplicity of its nervous system and the sophistication of the genetic techniques that can be used to dissect the molecular basis of the underlying biology4-6 In Drosophila, as in all metazoans, the response to noxious thermal stimuli generally involves a "nocifensive" aversive withdrawal to the presented stimulus7. Such stimuli are detected through free nerve endings or nociceptors and the amplitude of the organismal response depends on the number of nociceptors receiving the noxious stimulus8. In Drosophila, it is the class IV dendritic arborization sensory neurons that detect noxious thermal and mechanical stimuli9 in addition to their recently discovered role as photoreceptors10. These neurons, which have been very well studied at the developmental level, arborize over the barrier epidermal sheet and make contacts with nearly all epidermal cells11,12. The single axon of each class IV neuron projects into the ventral nerve cord of the central nervous system11 where they may connect to second-order neurons that project to the brain.
Under baseline conditions, nociceptive sensory neurons will not fire until a relatively high threshold is reached. The assays described here allow the investigator to quantify baseline behavioral responses or, presumably, the sensitization that ensues following tissue damage. Each assay provokes distinct but related locomotory behavioral responses to noxious thermal stimuli and permits the researcher to visualize and quantify various aspects of thermal nociception in Drosophila larvae. The assays can be applied to larvae of desired genotypes or to larvae raised under different environmental conditions that might impact nociception. Since thermal nociception is conserved across species, the findings gleaned from genetic dissection in Drosophila will likely inform our understanding of thermal nociception in other species, including vertebrates.

Local and Global Methods of Assessing Thermal Nociception in Drosophila Larvae

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of thermal nociceptors1,2 while the second involves a wholesale (global) activation of most or all such neurons3. Together, these techniques allow visualization and quantification of the behavioral functions of Drosophila nociceptive sensory neurons.

In this article, we demonstrate assays to study thermal nociception in Drosophila larvae. One assay involves spatially-restricted (local) stimulation of ...

Neuroscience

Standardized Methods for Measuring Induction of the Heat Shock Response in Caenorhabditis elegans

The heat shock response (HSR) is a cellular stress response induced by cytosolic protein misfolding that functions to restore protein folding homeostasis, or proteostasis. Caenorhabditis elegans occupies a unique and powerful niche for HSR research because the HSR can be assessed at the molecular, cellular, and organismal levels. Therefore, changes at the molecular level can be visualized at the cellular level and their impacts on physiology can be quantitated at the organismal level. While assays for measuring the HSR are straightforward, variations in the timing, temperature, and methodology described in the literature make it challenging to compare results across studies. Furthermore, these issues act as a barrier for anyone seeking to incorporate HSR analysis into their research. Here, a series of protocols is&#160;presented for measuring induction of the HSR in a robust and reproducible manner with RT-qPCR, fluorescent reporters, and an organismal thermorecovery assay. Additionally, we show that a widely used thermotolerance assay is not dependent on the well-established master regulator of the HSR, HSF-1, and therefore should not be used for HSR research. Finally, variations in these assays found in the literature are discussed and best practices are proposed to help standardize results across the field, ultimately facilitating neurodegenerative disease, aging, and HSR research.

Standardized Methods for Measuring Induction of the Heat Shock Response in Caenorhabditis elegans

Here, standardized protocols are presented to assess induction of the heat shock response (HSR) in Caenorhabditis elegans using RT-qPCR at the molecular level, fluorescent reporters at the cellular level, and thermorecovery at the organismal level.

The heat shock response (HSR) is a cellular stress response induced by cytosolic protein misfolding that functions to restore protein folding homeostasis, ...

Environment

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 ...

Behavior

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 ...

Measurements of Physiological Stress Responses in C. Elegans

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and physiology. Thus, organisms have evolved targeted and specific stress responses dedicated to repair damage and maintain homeostasis. These mechanisms include the unfolded protein response of the endoplasmic reticulum (UPRER), the unfolded protein response of the mitochondria (UPRMT), the heat shock response (HSR), and the oxidative stress response (OxSR). The protocols presented here describe methods to detect and characterize the activation of these pathways and their physiological consequences in the nematode, C. elegans. First, the use of pathway-specific fluorescent transcriptional reporters is described for rapid cellular characterization, drug screening, or large-scale genetic screening (e.g., RNAi or mutant libraries). In addition, complementary, robust physiological assays are described, which can be used to directly assess sensitivity of animals to specific stressors, serving as functional validation of the transcriptional reporters. Together, these methods allow for rapid characterization of the cellular and physiological effects of internal and external proteotoxic perturbations.

Measurements of Physiological Stress Responses in C. Elegans

Here, we characterize cellular proteotoxic stress responses in the nematode C. elegans by measuring the activation of fluorescent transcriptional reporters and assaying sensitivity to physiological stress.

Organisms are often exposed to fluctuating environments and changes in intracellular homeostasis, which can have detrimental effects on their proteome and ...

Field-Based Thermal Physiology Assay: Cold Shock Recovery under Ambient Conditions

Ecological physiology, particularly of ectotherms, is increasingly important in this changing world as it uses measures of species and environmental traits to explore the interactions between organisms and their surroundings to better understand their survival and fitness. Traditional thermal assays are costly in terms of time, money, and equipment and are therefore often limited to small sample sizes and few species. Presented here is a novel protocol that generates detailed data on individual behavior and physiology of large, volant, terrestrial insects, using the example of butterflies. This paper describes the methods of a cold shock recovery assay that can be performed in the field under ambient environmental conditions and does not require costly laboratory equipment. This method has been used to understand the response and recovery strategy to cold shock of tropical butterflies, generating individual level data across entire butterfly communities. These methods can be employed in both remote field settings and classrooms and can be used to generate ecologically relevant physiological data and as a teaching tool.

Here, a low-cost, accessible protocol is described to evaluate cold shock recovery of butterflies under ambient environmental conditions.

Ecological physiology, particularly of ectotherms, is increasingly important in this changing world as it uses measures of species and environmental ...

Topical Application Bioassay to Quantify Insecticide Toxicity for Mosquitoes and Fruit Flies

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. Insecticide resistance surveillance of mosquito populations is typically done through Centers for Disease Control and Prevention (CDC) bottle bioassays or World Health Organization (WHO) tube tests. However, these methods can result in a high degree of variability in mortality data due to variable insecticide contact with&#160;the insect, the relatively small numbers of organisms tested, extensive variation in mass between populations, and constantly changing environmental conditions, leading to variable outcomes. This paper presents the topical application bioassay, adapted as a high-throughput phenotypic bioassay for both mosquitoes and fruit flies, to test large numbers of insects along&#160;a range of insecticide concentrations.
This assay 1) ensures consistent treatment and insecticide contact with every organism, 2) produces highly specific dose-response curves that account for differences in average mass between strains and sexes (which is particularly important for field-collected organisms), and 3) allows for the calculation of statistically rigorous median lethal doses (LD50), which are necessary for resistance ratio comparisons-an alternative surveillance approach from diagnostic dose mortality, which is also used for larvicide resistance surveillance. This assay will be a complementary tool for accurately phenotyping mosquito populations and, as illustrated using fruit flies, is easily adaptable for use with other insects. We argue that this assay will help fill the gap between genotypic and phenotypic insecticide resistance in multiple insect species.

We describe the methodology and importance of the topical application bioassay to measure insecticide susceptibility in mosquitoes and fruit flies. The presented assay is high-throughput, utilizes insect mass-thus allowing for calculating a mass-relativized lethal dose instead of concentration-and likely has lower variability than other similar methods.

The continued use of insecticides for public health and agriculture has led to widespread insecticide resistance and hampering of control methods. ...

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on host physiology. It also fluoresces in blue when dying, providing a convenient means of pinpointing death. This property has been exploited to develop high-throughput label-free C. elegans survival assays (LFASS). These involve time-lapse fluorescence recording of worm populations set in multiwell plates, from which population median time of death can be derived. The present study adopts the LFASS approach to screen multiple microbial isolates at once for the effects on C. elegans susceptibility to severe heat and oxidative stresses. Such microbial screening pipeline, which can notably be used to prescreen probiotics, using severe stress resistance as a proxy for host health is reported here. The protocol describes how to grow both C. elegans gut microbiota isolate collections and synchronous worm populations in multiwell arrays before combining them for the assays. The example provided covers the testing of 47 bacterial isolates and one control strain on two worm strains, in two stress assays in parallel. However, the approach pipeline is readily scalable and applicable to the screening of many other modalities. Thus, it provides a versatile setup to rapidly survey a multiparametric landscape of biological and biochemical conditions that impact C. elegans health.

High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

Gut microbes may positively or negatively impact the health of their host via specific or conserved mechanisms. Caenorhabditis elegans is a convenient platform to screen for such microbes. The present protocol describes high-throughput screening of 48 bacterial isolates for impact on nematode stress resistance, used as a proxy for worm health.

With its small size, short lifespan, and easy genetics, Caenorhabditis elegans offers a convenient platform to study the impact of microbial isolates on ...

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.

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 ...

Thermal Limits Determination for Zooplankton Using a Heat Block

Thermal limits and breadth have been widely used to predict species distribution. As the global temperature continues to rise, understanding how thermal limit changes with acclimation and how it varies between life stages and populations are vital for determining the vulnerability of species to future warming. Most marine organisms have complex life cycles that include early planktonic stages. While quantifying the thermal limit of these small early developmental stages (tens to hundreds of microns) helps identify developmental bottlenecks, this process can be challenging due to the small size of target organisms, large bench space requirement, and high initial fabrication cost. Here, a setup that is geared toward small volumes (mL to tens of mL) is presented. This setup combines commercially available components to generate a stable and linear thermal gradient. Production specifications of the setup, as well as procedures to introduce and enumerate live versus dead individuals and compute lethal temperature, are also presented.

The present protocol illustrates the use of commercially available components to generate a stable and linear thermal gradient. Such gradient can then be used to determine the upper thermal limit of planktonic organisms, particularly invertebrate larvae.

Thermal limits and breadth have been widely used to predict species distribution. As the global temperature continues to rise, understanding how thermal ...

High-Throughput Assays of Critical Thermal Limits in Insects

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High-Throughput Screening of Microbial Isolates with Impact on Caenorhabditis elegans Health

Determining Temperature Preference of Mosquitoes and Other Ectotherms

Thermal Limits Determination for Zooplankton Using a Heat Block