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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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 &#8211; 5.08 cm to 7.62 cm</td>
			<td>Any</td>
			<td>Any</td>
			<td></td>
		</tr>
		<tr>
			<td>Clear acrylic tubing &#8211; 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 &#8211; 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 &#8211; 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) &#8211; &#188; 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&#34; 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&#34; 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&#34; 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 ...

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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

Choice and No-Choice Assays for Testing the Resistance of A. thaliana to Chewing Insects

Larvae of the small white cabbage butterfly are a pest in agricultural settings. This caterpillar species feeds from plants in the cabbage family, which include many crops such as cabbage, broccoli, Brussel sprouts etc. Rearing of the insects takes place on cabbage plants in the greenhouse. At least two cages are needed for the rearing of Pieris rapae. One for the larvae and the other to contain the adults, the butterflies. In order to investigate the role of plant hormones and toxic plant chemicals in resistance to this insect pest, we demonstrate two experiments. First, determination of the role of jasmonic acid (JA - a plant hormone often indicated in resistance to insects) in resistance to the chewing insect Pieris rapae. Caterpillar growth can be compared on wild-type and mutant plants impaired in production of JA. This experiment is considered "No Choice", because larvae are forced to subsist on a single plant which synthesizes or is deficient in JA. Second, we demonstrate an experiment that investigates the role of glucosinolates, which are used as oviposition (egg-laying) signals. Here, we use WT and mutant Arabidopsis impaired in glucosinolate production in a "Choice" experiment in which female butterflies are allowed to choose to lay their eggs on plants of either genotype. This video demonstrates the experimental setup for both assays as well as representative results.

Plant resistance to chewing insect herbivores can be tested in several ways. Here, we demonstrate how to set-up a choice and a no-choice experiment with the model plant Arabidopsis thaliana to identify resistance against the pest species Pieris rapae.

Larvae of the small white cabbage butterfly are a pest in agricultural settings. This caterpillar species feeds from plants in the cabbage family, which ...

High-Throughput Measurement and Classification of Organic P in Environmental Samples

Many types of organic phosphorus (P) molecules exist in environmental samples1. Traditional P measurements do not detect these organic P compounds since they do not react with colorimetric reagents2,3. Enzymatic hydrolysis (EH) is an emerging method for accurately characterizing organic P forms in environmental samples4,5. This method is only trumped in accuracy by Phosphorus-31 Nuclear Magnetic Resonance Spectroscopy (31P-NMR) -a method that is expensive and requires specialized technical training6. We have adapted an enzymatic hydrolysis method capable of measuring three classes of phosphorus (monoester P, diester P and inorganic P) to a microplate reader system7. This method provides researchers with a fast, accurate, affordable and user-friendly means to measure P species in soils, sediments, manures and, if concentrated, aquatic samples. This is the only high-throughput method for measuring the forms and enzyme-lability of organic P that can be performed in a standard laboratory. The resulting data provides insight to scientists studying system nutrient content and eutrophication potential.

This protocol describes a high-throughput method of enzymatic hydrolysis that utilizes a microplate reader to measure and classify soil phosphorus as P monoesters, P diesters and inorganic P. Up to 96 samples can be measured at one time in a standard laboratory.

Many types of organic phosphorus (P) molecules exist in environmental samples1.  Traditional P measurements do not detect these organic P compounds since ...

High-throughput Yeast Plasmid Overexpression Screen

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, including those with direct relevance to human disease. Because of its short generation time and well-characterized genome, a major experimental advantage of the yeast model system is the ability to perform genetic screens to identify genes and pathways that are involved in a given process. Over the last thirty years such genetic screens have been used to elucidate the cell cycle, secretory pathway, and many more highly conserved aspects of eukaryotic cell biology 1-5. In the last few years, several genomewide libraries of yeast strains and plasmids have been generated 6-10. These collections now allow for the systematic interrogation of gene function using gain- and loss-of-function approaches 11-16. Here we provide a detailed protocol for the use of a high-throughput yeast transformation protocol with a liquid handling robot to perform a plasmid overexpression screen, using an arrayed library of 5,500 yeast plasmids. We have been using these screens to identify genetic modifiers of toxicity associated with the accumulation of aggregation-prone human neurodegenerative disease proteins. The methods presented here are readily adaptable to the study of other cellular phenotypes of interest.

Here we describe a plasmid overexpression screen in Saccharomyces cerevisiae, using an arrayed plasmid library and a high-throughput yeast transformation protocol with a liquid handling robot.

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, ...

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted into fuel alcohols1. The potential for enzymatic hydrolysis of cellulosic biomass to provide renewable energy has intensified efforts to engineer cellulases for economical fuel production2. Of particular interest are fungal cellulases3-8, which are already being used industrially for foods and textiles processing.
Identifying active variants among a library of mutant cellulases is critical to the engineering process; active mutants can be further tested for improved properties and/or subjected to additional mutagenesis. Efficient engineering of fungal cellulases has been hampered by a lack of genetic tools for native organisms and by difficulties in expressing the enzymes in heterologous hosts. Recently, Morikawa and coworkers developed a method for expressing in E. coli the catalytic domains of endoglucanases from H. jecorina3,9, an important industrial fungus with the capacity to secrete cellulases in large quantities. Functional E. coli expression has also been reported for cellulases from other fungi, including Macrophomina phaseolina10 and Phanerochaete chrysosporium11-12. 
We present a method for high throughput screening of fungal endoglucanase activity in E. coli. (Fig 1) This method uses the common microbial dye Congo Red (CR) to visualize enzymatic degradation of carboxymethyl cellulose (CMC) by cells growing on solid medium. The activity assay requires inexpensive reagents, minimal manipulation, and gives unambiguous results as zones of degradation (&#x201C;halos&#x201D;) at the colony site. Although a quantitative measure of enzymatic activity cannot be determined by this method, we have found that halo size correlates with total enzymatic activity in the cell. Further characterization of individual positive clones will determine , relative protein fitness. 
Traditional bacterial whole cell CMC/CR activity assays13 involve pouring agar containing CMC onto colonies, which is subject to cross-contamination, or incubating cultures in CMC agar wells, which is less amenable to large-scale experimentation. Here we report an improved protocol that modifies existing wash methods14 for cellulase activity: cells grown on CMC agar plates are removed prior to CR staining. Our protocol significantly reduces cross-contamination and is highly scalable, allowing the rapid screening of thousands of clones. In addition to H. jecorina enzymes, we have expressed and screened endoglucanase variants from the Thermoascus aurantiacus and Penicillium decumbens (shown in Figure 2), suggesting that this protocol is applicable to enzymes from a range of organisms.

High Throughput Screening of Fungal Endoglucanase Activity in Escherichia coli

We describe a low cost, high throughput method to screen for fungal endoglucanase activity in E. coli. The method relies on a simple visual readout of substrate degradation, does not require enzyme purification, and is highly scalable. This allows for the rapid screening of large libraries of enzyme variants.

Cellulase enzymes (endoglucanases, cellobiohydrolases, and &beta;-glucosidases) hydrolyze cellulose into component sugars, which in turn can be converted ...

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place over the next 5 years. For example, the sequencing of the genomes for 15 malaria mosquitospecies is currently being done using an Illumina platform3,4. This Anopheles species cluster includes both vectors and non-vectors of malaria. When the genome assemblies become available, researchers will have the unique opportunity to perform comparative analysis for inferring evolutionary changes relevant to vector ability. However, it has proven difficult to use next-generation sequencing reads to generate high-quality de novo genome assemblies5. Moreover, the existing genome assemblies for Anopheles gambiae, although obtained using the Sanger method, are gapped or fragmented4,6.
Success of comparative genomic analyses will be limited if researchers deal with numerous sequencing contigs, rather than with chromosome-based genome assemblies. Fragmented, unmapped sequences create problems for genomic analyses because: (i) unidentified gaps cause incorrect or incomplete annotation of genomic sequences; (ii) unmapped sequences lead to confusion between paralogous genes and genes from different haplotypes; and (iii) the lack of chromosome assignment and orientation of the sequencing contigs does not allow for reconstructing rearrangement phylogeny and studying chromosome evolution. Developing high-resolution physical maps for species with newly sequenced genomes is a timely and cost-effective investment that will facilitate genome annotation, evolutionary analysis, and re-sequencing of individual genomes from natural populations7,8.
Here, we present innovative approaches to chromosome preparation, fluorescent in situ hybridization (FISH), and imaging that facilitate rapid development of physical maps. Using An. gambiae as an example, we demonstrate that the development of physical chromosome maps can potentially improve genome assemblies and, thus, the quality of genomic analyses. First, we use a high-pressure method to prepare polytene chromosome spreads. This method, originally developed for Drosophila9, allows the user to visualize more details on chromosomes than the regular squashing technique10. Second, a fully automated, front-end system for FISH is used for high-throughput physical genome mapping. The automated slide staining system runs multiple assays simultaneously and dramatically reduces hands-on time11. Third, an automatic fluorescent imaging system, which includes a motorized slide stage, automatically scans and photographs labeled chromosomes after FISH12. This system is especially useful for identifying and visualizing multiple chromosomal plates on the same slide. In addition, the scanning process captures a more uniform FISH result. Overall, the automated high-throughput physical mapping protocol is more efficient than a standard manual protocol.

High-throughput Physical Mapping of Chromosomes using Automated in situ Hybridization

Genome assemblies based on massively parallel DNA sequencing technologies are usually highly fragmented. The development of physical chromosome maps can potentially improve genome assemblies. Here, we demonstrate innovative approaches to chromosome preparation, fluorescent in situ hybridization, and imaging that significantly increase throughput of the physical map development.

Projects to obtain whole-genome sequences for 10,000 vertebrate species1 and for 5,000 insect and related arthropod species2 are expected to take place ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

High Throughput Microinjections of Sea Urchin Zygotes

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of treatment solution is loaded into a microinjection needle that is used to physically inject individual immobilized cells or embryos. Despite the need for initial training to perform this procedure for high-throughput delivery, microinjection offers maximum efficiency and reproducible delivery of a wide variety of treatment solutions (including complex mixtures of samples) into cells, eggs or embryos. Applications to microinjections include delivery of DNA constructs, mRNAs, recombinant proteins, gain of function, and loss of function reagents. Fluorescent or colorimetric dye is added to the injected solution to enable instant visualization of efficient delivery as well as a tool for reliable normalization of the amount of the delivered solution. The described method enables microinjection of 100-400 sea urchin zygotes within 10-15 min.

Microinjection is a common technique used to deliver DNA constructs, mRNAs, morpholino antisense oligonucleotides or other treatments into eggs, embryos, and cells of various species.

Microinjection into cells and embryos is a common technique that is used to study a wide range of biological processes. In this method a small amount of ...

High-throughput Titration of Luciferase-expressing Recombinant Viruses

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with viruses that do not form plaques. As an alternative to plaque assays, we have developed a high-throughput titration method that allows for the simultaneous titration of a high volume of samples in a single day. This approach involves infection of the samples with a Firefly luciferase tagged virus, transfer of the infected samples onto an appropriate permissive cell line, subsequent addition of luciferin, reading of plates in order to obtain luminescence readings, and finally the conversion from luminescence to viral titers. The assessment of cytotoxicity using a metabolic viability dye can be easily incorporated in the workflow in parallel and provide valuable information in the context of a drug screen. This technique provides a reliable, high-throughput method to determine viral titers as an alternative to a standard plaque assay.

This article presents a high-throughput luciferase expression-based method of titrating various RNA and DNA viruses using automated and manual liquid handlers.

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of energy homeostasis and obesity. The small size of zebrafish allows for assays on embryos to be conducted in a 96- or 384-well plate format, Morpholino and CRISPR based technologies promote ease of genetic manipulation, and drug treatment by bath application is viable. Moreover, zebrafish are ideal for forward genetic screens allowing for novel gene discovery. Given the relative novelty of zebrafish as a model for obesity, it is necessary to develop tools that fully exploit these benefits. Herein, we describe a method to measure energy expenditure in thousands of embryonic zebrafish simultaneously. We have developed a whole animal microplate platform in which we use 96-well plates to isolate individual fish and we assess cumulative NADH2 production using the commercially available cell culture viability reagent alamarBlue. In poikilotherms the relationship between NADH2 production and energy expenditure is tightly linked. This energy expenditure assay creates the potential to rapidly screen pharmacological or genetic manipulations that directly alter energy expenditure or alter the response to an applied drug (e.g. insulin sensitizers).

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism for the study of energy homeostasis. By utilizing a NADH2 sensitive redox indicator, alamar Blue, we have developed an assay that measures the metabolic rate of zebrafish larvae in a 96 well plate format and can be applied to drug or gene discovery.

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of ...