This work was supported by a donation from Frank Leslie. Some strains were provided by the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440).

1. Maintenance and synchronization of C. elegans
<ol>
	<li>Maintain worms at 20 &#176;C on Nematode Growth Medium (NGM) plates seeded with OP50 Escherichia coli bacteria by transferring several adults to fresh plates approximately 2x per week10. Care should be taken to prevent worms from running out of food, because this can affect their physiology11.
	<ol>
		<li>Preparation of NGM plates.
		<ol>
			<li>Mix 3 g of NaCl, 2.5 g of Bacto-peptone, 20&#160;g of agar, and deionized (DI) H2O up to 1 L in a flask.</li>
			<li>Autoclave the mixture for sterilization.</li>
			<li>Allow mixture to cool to ~50 &#176;C.</li>
			<li>Add 25 mL of 1 M KH2PO4 (pH = 6), 1 mL of 1 M CaCl2, 1 mL of 1 M MgSO4, and 1 mL of cholesterol (5 mg/mL in 100% ethanol).</li>
			<li>Using sterile technique, pour the mixture into 6 cm plates to yield approximately 100 plates. Pouring plates is easier if the mixture is first transferred to a 300 mL sterile beaker.</li>
			<li>Allow 1 day to solidify at room temperature (RT) before seeding with bacteria or storing at 4 &#176;C.</li>
		</ol>
		</li>
		<li>Seeding of OP50 bacteria onto NGM plates.
		<ol>
			<li>Grow a saturated overnight OP50 bacterial culture in LB at 30 &#176;C or 37 &#176;C.</li>
			<li>Place approximately 300 &#181;L of the culture onto the center of a 6 cm NGM plate.</li>
			<li>Let plates dry at RT for 1-3 days as needed for the bacterial lawn to adhere to the plate. Plates can then be used or stored at 4 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Grow the worms synchronously either by isolating freshly laid eggs (described here)&#160;or alternatively&#160;by collecting eggs after dissolving worms with bleach.
	<ol>
		<li>Transfer approximately 10 gravid adult worms to a fresh plate using a platinum wire pick. Egg-lay synchronization works best if the adults are in the first day of adulthood.</li>
		<li>After approximately 1 h, remove the worms from the plate. This should result in 40-60 eggs per plate, depending on the conditions and the strain.</li>
	</ol>
	</li>
</ol>
2. Fluorescent imaging of HSR reporters
<ol>
	<li>Synchronize the worms (section 1.2) and maintain at 20 &#176;C until the desired developmental stage. For the AM446 (hsp-70p::gfp) and CL2070 (hsp-16.2p::gfp) fluorescent reporter strains, young adult worms that have not yet reached reproductive maturity are generated 64 h after the egg-laying synchronization. 
	NOTE: The developmental timing varies with each strain and the temperature at which the worms are raised. Both HSR reporter strains exhibit a slight developmental delay relative to N2. Importantly, the magnitude of HSR induction declines approximately 2-4x after the onset of reproductive maturity (see Discussion).</li>
	<li>Heat shock the worms by wrapping plates with paraffin film and submerging in a circulating water bath at 33 &#176;C for 1 h. A thin strip of paraffin film should be wrapped 2x around the plate to seal the edges. Do not cover the bottom of the plate or it could interfere with heat transfer. Submerge the plates upside down using a test tube rack and a lead weight. Remember to include a negative control sample (no heat shock) if necessary. 
	NOTE: If the paraffin film is not secure, then water will enter the plate and the plate should not be used for data collection.</li>
	<li>Recover the worms by removing the plates from the water bath and drying with a paper towel. Remove the paraffin film and incubate the worms at 20 &#176;C for 6-24 h. This recovery period allows sufficient time for GFP synthesis and folding before imaging.</li>
	<li>Prepare slides for imaging. Slides should be prepared fresh for each use.
	<ol>
		<li>Make a 3% agarose solution in water and heat using a microwave until the agarose is dissolved.</li>
		<li>Place a microscope slide for imaging between two other microscope slides that have a strip of laboratory tape on them to create a spacer for the agarose pad.</li>
		<li>Using a 1,000 &#181;L pipette, place a drop (~150 &#181;L) of the heated 3% agarose in the center of the microscope slide.</li>
		<li>Immediately cover the microscope slide with a blank microscope slide perpendicular to the first slide so that the top slide rests on the laboratory tape on the adjacent slides. This spreads out the drop of agarose to create a pad of uniform width.</li>
		<li>Carefully remove the top slide.</li>
	</ol>
	</li>
	<li>Immobilize the worms by using a 200 &#181;L pipette to add a small drop (~5 &#181;L) of 1 mM levamisole in M9 buffer to the center of the agarose pad. Then transfer 10 worms into the drop of levamisole using a platinum wire pick. Cover with a coverslip. Sealing the coverslip is not necessary for an upright microscope. Optionally, the worms can be aligned when they become paralyzed by spreading the levamisole off, to the outside of the agarose pad, and aligning the worms with a platinum wire pick. Alternatively, the levamisole can be soaked up using a laboratory wipe. 
	NOTE: Image as soon as possible, because prolonged incubation in levamisole could alter fluorescence.</li>
	<li>Image the worms using a fluorescence microscope. The details of image capture vary by microscope and software. 
	NOTE: To directly compare image intensities, use identical microscope settings in one imaging session. Avoid oversaturating the image.</li>
</ol>
3. Measurement of HSR gene expression using RT-qPCR
<ol>
	<li>Synchronize worms (section 1.2) and maintain at 20 &#176;C until the desired developmental stage. For N2 worms, young adult worms that have not yet reached reproductive maturity are generated 60 h after the egg-laying synchronization. 
	NOTE: The developmental timing varies with each strain and the temperature at which the worms are raised. Importantly, the magnitude of HSR induction declines approximately 2-4x after the onset of reproductive maturity (see Discussion).</li>
	<li>Heat shock worms as described in step 2.2.</li>
	<li>Take the plates out of the water bath, remove the paraffin film, and immediately collect the worms. The worms can be collected by washing the plates gently with 1 mL of M9, collecting the liquid in a microcentrifuge tube, and then removing the M9 after centrifugation at 400 x g for 1 min.</li>
	<li>Lyse the worms and purify the RNA using organic extraction.
	<ol>
		<li>Add 250 &#181;L of RNA isolation reagent (see Table of Materials).</li>
		<li>Vortex tubes by hand for 30 s.</li>
		<li>Vortex tubes for 20 min at 4 &#176;C using a microcentrifuge tube attachment (see Table of Materials).</li>
		<li>Add 50 &#181;L of chloroform.</li>
		<li>Vortex for 30 s.</li>
		<li>Incubate the samples at RT for 3 min.</li>
		<li>Centrifuge at &#8805;14,000 x g for 15 min at 4 &#176;C.</li>
		<li>Transfer the aqueous layer (i.e., top layer, ~125 &#181;L) to a new microcentrifuge tube. 
		NOTE: Avoid the organic layer and the material in the interface.</li>
		<li>Add 50 &#181;L of chloroform.</li>
		<li>Vortex for 30 s.</li>
		<li>Incubate the samples at RT for 3 min.</li>
		<li>Centrifuge at &#8805;14,000 x g for 5 min at 4 &#176;C.</li>
		<li>Transfer the aqueous layer (~100 &#181;L) to a new microcentrifuge tube. 
		NOTE: Avoid the organic layer and the material in the interface.</li>
		<li>Precipitate RNA with an equal volume (i.e., 100 &#181;L) of isopropanol.</li>
		<li>Incubate at -20 &#176;C for at least 30 min, but preferably overnight. 
		NOTE: The experiment can be paused here and the RNA can be stored at -20 &#176;C.</li>
		<li>Pellet the RNA by centrifugation at &#8805;14,000 x g for &#8805;30 min at 4 &#176;C.</li>
		<li>Remove as much of the supernatant as possible without disturbing the pellet. 
		NOTE: The pellet will be small and may not be visible. The pellet may not adhere tightly to the side of the tube, so caution is necessary to avoid dislodging it.</li>
		<li>Wash the pellet with 250 &#181;L of 70% ice-cold ethanol made with RNase-free H2O.</li>
		<li>Centrifuge at &#8805;14,000 x g for &#8805;5 min at 4 &#176;C.</li>
		<li>Remove as much supernatant as possible without disturbing the pellet.</li>
		<li>Perform a quick spin at RT to remove any remaining 70% ethanol.</li>
		<li>Dry the pellet by leaving the tubes open at RT as long as needed; typically at least 20 min. Tubes can be covered with a lint-free tissue or aluminum foil to prevent contamination.</li>
		<li>Resuspend the pellet in 20 &#181;L of RNase-free H2O.</li>
		<li>Determine the RNA concentration using a small volume spectrophotometer (2 &#181;L). 
		NOTE: The experiment can be paused here and the RNA can be temporarily stored at or below -20 &#176;C.</li>
	</ol>
	</li>
	<li>Remove residual DNA by incubating with DNase I. It is recommended to use a commercially available kit (see Table of Materials) and to follow the manufacturer&#39;s instructions.
	<ol>
		<li>With this kit, prepare a 20 &#181;L reaction with 500 ng of RNA and 1 &#181;L of DNase I in a 37 &#176;C water bath for 30 min.</li>
		<li>Add 2.5 &#181;L of DNase inactivation reagent (included in the kit) to each sample and incubate at RT for 5 min with occasional flicking/vortexing.</li>
		<li>Spin down at 14,000 x g for 2 min.</li>
		<li>Without disturbing the white pellet, transfer 15 &#181;L of supernatant to a fresh microtube for cDNA synthesis.</li>
	</ol>
	</li>
	<li>Conduct cDNA synthesis. It is recommended to use a commercially available kit (see Table of Materials) and to follow the manufacturer&#39;s instructions.
	<ol>
		<li>With the kit, prepare a 20 &#181;L reaction with 15 &#181;L of DNase I-treated RNA from the previous step and 1 &#181;L of reverse transcriptase.</li>
		<li>Use the following program for cDNA synthesis: 25 &#176;C for 5 min, 46 &#176;C for 20 min, 95 &#176;C for 1 min, 4 &#176;C hold.</li>
		<li>Dilute cDNA by adding 80 &#181;L of RNase-free H2O directly to the sample.</li>
		<li>Briefly vortex, then spin down and store at -20 &#176;C until needed.</li>
	</ol>
	</li>
	<li>Perform qPCR. It is recommended to use a commercially available kit (see Table of Materials) and to follow the manufacturer&#39;s instructions.
	<ol>
		<li>With the kit, prepare a 25 &#181;L reaction containing 2 &#181;L of cDNA and 200 nM (each)&#160;of&#160;forward and reverse primers&#160;in one well of a 96-well plate.</li>
		<li>Primer sequences for measuring the heat shock genes, hsp-70 and hsp-16.2, and 18S rRNA (for a normalization control) are listed in the Table of Materials. Multiple normalization controls can be used as desired.</li>
		<li>Dilute cDNA samples 50x before measurement of 18S to ensure that the assay is in the linear range. Appropriate qPCR conditions vary with the kit and primers used (see Representative Results).</li>
		<li>Use a real-time PCR detection system (see Table of Materials) for qPCR with 40 cycles of 95 &#176;C for 5 s denaturation, 58 &#176;C for 30 s annealing, and 72 &#176;C for 30 s extension. 
		NOTE: Optimal annealing temperatures can vary by primers and conditions.</li>
		<li>Quantify using either the &#916;&#916;Ct or standard curve method12.</li>
	</ol>
	</li>
</ol>
4. Thermorecovery assay for measuring HSR at the organismal level
<ol>
	<li>Synchronize the worms (section 1.2) and maintain at 20 &#176;C until the desired developmental stage. For N2 worms, young adult worms that have not yet reached reproductive maturity are generated 60 h after the egg-laying synchronization. 
	NOTE: The developmental timing varies with each strain and the temperature at which the worms are raised. Importantly, the magnitude of HSR induction declines approximately 2-4x after the onset of reproductive maturity (see Discussion).</li>
	<li>Heat shock the worms as described in step 2.2 for 6 h.</li>
	<li>Remove the plates from the water bath, remove the paraffin film, and allow the worms to recover by incubation at 20 &#176;C for 48 h.</li>
	<li>Count the number of worms that can immediately crawl away after mechanical stimulation without jerky movement or paralysis. 
	NOTE: The 6 h incubation is optimal for examining conditions that reduce thermorecovery, but longer exposure times may be needed to look for conditions that enhance thermorecovery.</li>
</ol>

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The authors have nothing to disclose.

In the literature a wide variety of temperatures, times, and equipment have been used to assay the HSR, which has introduced unnecessary caveats and led to difficulty in comparing results between laboratories. For example, temperatures ranging anywhere from 32-37 &#176;C and times from 15 min to several hours have been used to induce the HSR15. However, it is reported that lethality occurs as early as 3 h at 37 &#176;C for all stages and 1.5 h for day 1 adults15. Furthermore, we show that exposure of worms to 35 &#176;C causes lethality that is not HSF-1 dependent, making these conditions poorly suited for analysis of the HSR. In contrast, a heat shock of 33 &#176;C for 1 h is robust enough to elicit strong induction of heat shock genes, yet mild enough to not affect worm viability. Indeed, exposure to 33 &#176;C for as long as 6 h only causes 20% of worms to display abnormal movement. Therefore, we propose using a temperature of 33 &#176;C and a time of 1 h as a standardized condition for RT-qPCR and fluorescent reporter assays.
Recent experiments have revealed that developmental staging of worms for HSR experiments is particularly important. It was recently shown that in C. elegans the inducibility of the HSR declines (i.e., collapses) by &#62;50% when hermaphrodites begin egg laying5. Staging the worms correctly is critical because there are often differences in developmental timing in strains carrying mutations. If temperature-sensitive mutants are used, this will also impact results if they are not synchronized by their reproductive age. Therefore, it is recommended to carefully measure the onset of egg laying for every strain to determine when the collapse occurs. The window of time after the L4 molt and before the initiation of reproductive maturity is narrow; therefore, care must be taken so that the HSR collapse does not inadvertently cause variability in results.
In addition to developmental timing, surprisingly small changes in temperature, as little as 1 &#176;C, can have substantial effects on the HSR. For example, thermosensory neurons in C. elegans are sensitive to temperature changes as small as &#177;0.05 &#176;C16. Thus, it is imperative to use a thermometer that can accurately measure the temperature. Therefore, we propose as best practice the use of a calibrated device for temperature measurement that is precise enough to measure temperatures within &#177;0.1 &#176;C. Furthermore, a thermometer with a data-logging functionality should be used to measure temperature variations across time. Many incubators are specified to have thermal variations of more than 1 &#176;C in different parts of the incubator and across time, which can have significant effects on HSR experiments. As a best practice, we suggest using incubators that have sufficient insulation and circulation to minimize temperature fluctuations. For conducting heat shock experiments, we propose a best practice of a circulating water bath. The time it takes for an agar plate to reach a desired temperature is approximately 6-7 min in a water bath but much longer in a dry incubator15,17. However, if a circulating water bath is not available, we have shown that robust HSR induction also occurs in a dry incubator using our conditions. If a dry incubator is used, opening of the incubator for the duration of the stress should be minimized.
It is well-established that induction of heat shock genes is dependent on the master regulator of the HSR, HSF-1. Here, we present evidence that the two more indirect assays, fluorescent reporters and thermorecovery, are also dependent on HSF-1. Significantly, we found that a commonly used alternative organismal assay, thermotolerance, is not HSF-1 dependent using hsf-1 RNAi (Figure 4). Similar results have been previously reported using an hsf-1 mutant or a ttx-3 mutant, which blocks the HSR18,19,20. Together, these results indicate that the thermotolerance assay should not be used for HSR research. Furthermore, this suggests that a best practice is to test the HSF-1 dependence for any assay used to measure the HSR.
Taken together, we present a series of standardized protocols and best practices for robust and reproducible measurement of HSR induction in C. elegans. We hope that these methodologies will decrease variability in HSR experiments and increase reproducibility. Facilitating direct comparisons of HSR research between laboratories will serve to accelerate research in the HSR field. Furthermore, standardization will benefit research into aging and neurodegenerative diseases with which the HSR is intimately associated.

The heat shock response (HSR) is a universal cellular stress response induced by cytosolic protein misfolding caused by temperature increases and other proteotoxic stresses. Activation of the HSR in Caenorhabditis elegans leads to transcriptional upregulation of heat shock genes such as hsp-70 and hsp-16.2. Many heat shock proteins (HSPs) function as molecular chaperones that restore protein folding homeostasis, or proteostasis, by directly interacting with misfolded or damaged proteins. The master regulator of the HSR is the transcription factor Heat Shock Factor 1 (HSF-1), whose activation is elegantly controlled via multiple mechanisms1.
The role of HSF-1 is not restricted to stress. HSF-1 is required for normal growth and development, as&#160;deletion of hsf-1 leads to larval arrest2. HSF-1 is also important during aging and age-related neurodegenerative diseases characterized by accumulation of protein aggregates and an inability to maintain proteostasis. Knockdown of hsf-1 causes accumulation of protein aggregates and a shortened lifespan, while overexpression of hsf-1 reduces protein aggregation and extends lifespan3,4. Therefore, regulation of HSF-1 at the molecular level has broad implications for organismal physiology and disease.
C. elegans is a powerful model organism for HSR research because the HSR can be measured at the molecular, cellular, and organismal levels4,5,6. Highlighting the power of this model, key advances in delineating the HSR pathway, such as tissue-specific differences in HSR regulation, have been discovered in C. elegans7,8. Furthermore, C. elegans is widely used for aging research and is an emerging system for modeling diseases linked to proteostasis disruption.
Although heat shock experiments with C. elegans can be quick and reproducible, there are several questions to consider before beginning. For example, which temperature should be used for induction of the HSR and how long should the worms be exposed? Is it better to use a dry incubator or a water bath? Which developmental stage should be used? Unfortunately, the methodologies used to investigate the HSR vary widely from laboratory to laboratory, causing confusion when selecting the best methodologies and making it difficult to compare results across the field.
We present robust and standardized protocols for using RT-qPCR, fluorescent reporters, and thermorecovery to measure the HSR. While these three approaches are complementary, they each have unique advantages and disadvantages. For example, RT-qPCR is the most direct and quantitative measurement of the HSR, and this assay can be easily expanded to include many different heat shock-inducible genes. However, RT-qPCR is the most expensive, can be technically difficult, and requires the use of specialized equipment. In contrast, fluorescent reporters have the advantage of measuring tissue-specific differences in HSR induction. However, they are difficult to quantitate accurately, can only measure induction above a certain threshold, and require the use of a fluorescence microscope. Additionally, the reporter strains described here are developmentally delayed compared to the standard N2 strain. Although newer reporter strains containing single-copy transgenes are available, they have not been tested here9. The third assay, thermorecovery, has the advantage of providing a physiologically relevant readout at the organismal level. However, this assay is arguably the least sensitive and most indirect. Finally, we discuss some common variations found in these assays and propose a set of best practices to facilitate research in this field.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>18S-forward primer</td>
			<td></td>
			<td></td>
			<td>TTGCGTCAACTGTGGTCGTG</td>
		</tr>
		<tr>
			<td>18S-reverse primer</td>
			<td></td>
			<td></td>
			<td>CCAACAAAAAGAACCGAAGT 
			CCTG</td>
		</tr>
		<tr>
			<td>AM446 rmIs223[phsp70::gfp; pRF4(rol-6(su1006))]</td>
			<td>Morimoto lab</td>
			<td><a href="http://groups.molbiosci.northwestern.edu/morimoto/" target="_blank">http://groups.molbiosci.northwestern.edu/morimoto/</a></td>
			<td></td>
		</tr>
		<tr>
			<td>C12C8.1-forward primer</td>
			<td></td>
			<td></td>
			<td>GTACTACGTACTCATGTGTCG 
			GTATTT</td>
		</tr>
		<tr>
			<td>C12C8.1-reverse primer</td>
			<td></td>
			<td></td>
			<td>ACGGGCTTTCCTTGTTTTCC</td>
		</tr>
		<tr>
			<td>CFX Connect Real-Time PCR Detection System</td>
			<td>Bio Rad</td>
			<td>1855200</td>
			<td></td>
		</tr>
		<tr>
			<td>CL2070 dvIs70 [hsp-16.2p::GFP + rol-6(su1006)]</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td><a href="https://cgc.umn.edu/" target="_blank">https://cgc.umn.edu/</a></td>
			<td></td>
		</tr>
		<tr>
			<td>EasyLog Thermistor Probe Data Logger with LCD</td>
			<td>Lascar</td>
			<td>EL-USB-TP-LCD</td>
			<td></td>
		</tr>
		<tr>
			<td>Greenough Stereo Microscope S9i Series</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hard Shell 96 Well PCR Plates</td>
			<td>Bio Rad</td>
			<td>HSS9601</td>
			<td></td>
		</tr>
		<tr>
			<td>hsp-16.2-forward primer</td>
			<td></td>
			<td></td>
			<td>ACTTTACCACTATTTCCGTCC 
			AGC</td>
		</tr>
		<tr>
			<td>hsp-16.2-reverse primer</td>
			<td></td>
			<td></td>
			<td>CCTTGAACCGCTTCTTTCTTTG</td>
		</tr>
		<tr>
			<td>iScript cDNA Synthesis Kit</td>
			<td>Bio Rad</td>
			<td>1708891</td>
			<td></td>
		</tr>
		<tr>
			<td>iTaq Universal Sybr Green Super Mix</td>
			<td>Bio Rad</td>
			<td>1725121</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser Scanning Confocal Microscope</td>
			<td>Nikon</td>
			<td>Eclipse 90i</td>
			<td></td>
		</tr>
		<tr>
			<td>MultiGene OptiMax Thermo Cycler</td>
			<td>Labnet</td>
			<td>TC9610</td>
			<td></td>
		</tr>
		<tr>
			<td>N2 (WT)</td>
			<td>Caenorhabditis Genetics Center (CGC)</td>
			<td><a href="https://cgc.umn.edu/" target="_blank">https://cgc.umn.edu/</a></td>
			<td></td>
		</tr>
		<tr>
			<td>Nanodrop Lite Spectrophotometer</td>
			<td>Thermo Scientific</td>
			<td>ND-LITE</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm M Roll</td>
			<td>Bemis</td>
			<td>5259-04LC</td>
			<td></td>
		</tr>
		<tr>
			<td>RapidOut DNA Removal Kit</td>
			<td>Thermo Scientific</td>
			<td>K2981</td>
			<td></td>
		</tr>
		<tr>
			<td>Recirculating Heated Water Bath</td>
			<td>Lauda Brinkmann</td>
			<td>RE-206</td>
			<td></td>
		</tr>
		<tr>
			<td>Traceable Platinum Ultra-Accurate Digital Thermometer</td>
			<td>Fisher Scientific</td>
			<td>15-081-103</td>
			<td></td>
		</tr>
		<tr>
			<td>TRIzol Reagent</td>
			<td>Invitrogen</td>
			<td>15596026</td>
			<td>RNA isolation reagent</td>
		</tr>
		<tr>
			<td>TurboMix Attachment</td>
			<td>Scientific Industries</td>
			<td>SI-0564</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex-Genie 2</td>
			<td>Scientific Industries</td>
			<td>SI-0236</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

Using the protocols described in this manuscript, HSR induction was measured using fluorescent reporters, RT-qPCR, and thermorecovery assays. In each case, the procedure in section 1.2 was used to generate synchronized, young adult worms that had not reached reproductive maturity.
To visualize HSR induction at the cellular level, the AM446 (hsp-70p::gfp) and CL2070 (hsp-16.2p::gfp) fluorescent reporter strains were analyzed following section 2 of the protocol. In the negative control samples without heat shock, the hsp-16.2 reporter only showed normal autofluorescence, but the hsp-70 reporter had constitutive fluorescence in the anal depressor muscle as previously reported4 (Figure 1A). After 1 h of heat shock at 33 &#176;C, robust fluorescence was observed in both reporters; however, the pattern of expression was distinct depending on which reporter was used (Figure 1B). The hsp-70 reporter was brightest in the intestine and spermatheca, whereas the hsp-16.2 reporter was brightest in the pharynx. Additionally, the hsp-16.2 reporter had a high degree of worm-to-worm variability in the amount of induction as previously described, but the hsp-70 reporter did not13.
A commonly used variation of section 2 is to perform the heat shock in a dry incubator instead of a circulating water bath. Therefore, the difference between the two methodologies was also tested. It was found that both protocols resulted in robust induction of the two fluorescent reporters using our conditions, although a circulating water bath is recommended as a best practice (see Discussion) (Figure 1B).
To test the dependence of the reporters on the transcription factor HSF-1, feeding RNAi was used to knockdown hsf-1 before reporter induction was measured. It was found that fluorescence of both strains was severely reduced upon HSF-1 knockdown, indicating that these reporters are HSF-1-dependent as described in the literature4 (Figure 2). However, it was also observed that pharyngeal fluorescence persisted in both reporters upon hsf-1 knockdown, which is consistent with previous reports that the pharyngeal muscle is resistant to RNAi by feeding14.
To quantitate whole worm induction of the HSR at the molecular level, two endogenous HSPs were measured with RT-qPCR using section 3 of the protocol. Samples were measured in triplicate, a standard curve was generated for each of the primers, and a melt curve was analyzed for each sample for quality control. It was found that a 33 &#176;C heat shock for 1 h resulted in more than a 2,000x increase in relative expression for two heat shock genes, hsp-70 and hsp-16.2 (Figure 3). These results show that both endogenous genes are suitable for measuring HSR induction and that a 33 &#176;C heat shock for 1 h is sufficient to generate a substantial response. However, caution should be used in interpreting the absolute degree of heat shock induction, because the mRNA levels in the absence of heat shock are very low.
To analyze a physiological response to heat shock, an organismal thermorecovery assay was tested using section 4 of the protocol. It was found that exposure of worms to a 6 h heat shock at 33 &#176;C led to a 20% decrease in worms with normal movement after a 48 h recovery (Figure 4A). The dependence of this assay on the HSF-1 transcription factor was tested using feeding RNAi to knockdown hsf-1 before exposing worms to the stress. It was found that knockdown of hsf-1 caused a dramatic decrease in normal movement, with &#62;95% of worms showing jerky movement or paralysis after being prodded with a platinum wire pick.
We compared this thermorecovery assay to a widely used alternative organismal assay commonly referred to as thermotolerance. In the thermotolerance assay, worms are exposed to a continuous 35 &#176;C temperature using a dry incubator, and the percentage of worms alive are measured at various timepoints. Using this assay, it was found that control worms continuously exposed to 35 &#176;C died after approximately 8 h of exposure (Figure 4B). However, when the dependence of this assay on HSF-1 was tested using RNAi knockdown, it was found that inhibition of hsf-1 did not cause a decrease in thermotolerance. Similar results have been previously shown using HSF-1 mutations (see Discussion). Therefore, the use of the thermotolerance assay to measure the HSR is not recommended, and thermorecovery is the preferred method for examining the HSR at the organismal level.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61030/61030fig1.jpg" /> 
Figure 1: HSR induction measured with fluorescent reporters. (A) The basal and (B) heat-inducible expression of hsp-70p::gfp and hsp-16.2p::gfp reporter strains after 1 h of heat shock at 33 &#176;C in a water bath or incubator. Worms were raised on OP50 bacteria for 64 h, heat shocked, and then recovered at 20 &#176;C for 8 h before imaging. For reference, the no heat-shock worms in (A) were renormalized in (B) to match the range and saturation of the heat-shocked worms. Representative images of two experimental replicates are shown. Scale bar = 250 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61030/61030fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61030/61030fig2.jpg" /> 
Figure 2: HSR induction measured with fluorescent reporters is dependent on HSF-1. Strains containing the hsp-70p::gfp and hsp-16.2p::gfp reporters were raised on control (L4440 empty vector) or hsf-1 RNAi plates for 64 h, exposed to a 1 h heat shock at 33 &#176;C in a water bath, and then recovered at 20 &#176;C for 8 h before imaging. Representative images of two experimental replicates are shown. Scale bar = 250 &#181;m. <a href="https://www.jove.com/files/ftp_upload/61030/61030fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61030/61030fig3.jpg" /> 
Figure 3: HSR induction measured with RT-qPCR. N2 worms were raised on HT115 bacteria for 60 h and then heat shocked for 1 h in a 33 &#176;C water bath. The relative mRNA levels of hsp-70 (C12C8.1) and hsp-16.2 are shown normalized to the no heat-shock control. Values plotted are the mean of four biological replicates and error bars represent &#177; SEM. Statistical significance was calculated using an unpaired Student&#39;s t-test. **p &#60; 0.01. <a href="https://www.jove.com/files/ftp_upload/61030/61030fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61030/61030fig4.jpg" /> 
Figure 4: Thermorecovery, but not thermotolerance, is dependent on HSF-1. N2 worms were raised on control (L4440) or hsf-1 RNAi plates for 60 h and then shifted to either:&#160;(A) A 33 &#176;C water bath for 6 h and recovered at 20 &#176;C for 48 h before scoring for normal movement (thermorecovery), or (B) A 35 &#176;C dry incubator and removed every 2 h until dead (thermotolerance).&#160;Each assay was done with n &#8805; 30 individuals on 2 independent days. The average is shown. <a href="https://www.jove.com/files/ftp_upload/61030/61030fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Standardized Methods for Measuring Induction of the Heat Shock Response in Caenorhabditis elegans at JoVE.com

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.

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, ...

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8.6K Views.  Florida Institute of Technology. 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.

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

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail millet. Recently, the 510 Mb genome of foxtail millet, S. italica, has been sequenced 1,2 and a 25x coverage genome sequence of the weedy relative S. viridis is in progress. S. viridis has a number of characteristics that make it a potentially excellent model genetic system including a rapid generation time, small stature, simple growth requirements, prolific seed production 3 and developed systems for both transient and stable transformation 4. However, the genetics of S. viridis is largely unexplored, in part, due to the lack of detailed methods for performing crosses. To date, no standard protocol has been adopted that will permit rapid production of seeds from controlled crosses.
The protocol presented here is optimized for performing genetic crosses in S. viridis, accession A10.1. We have employed a simple heat treatment with warm water for emasculation after pruning the panicle to retain 20-30 florets and labeling of flowers to eliminate seeds resulting from newly developed flowers after emasculation. After testing a series of heat treatments at permissive temperatures and varying the duration of dipping, we have established an optimum temperature and time range of 48 &deg;C for 3-6 min. By using this method, a minimum of 15 crosses can be performed by a single worker per day and an average of 3-5 outcross progeny per panicle can be recovered. Therefore, an average of 45-75 outcross progeny can be produced by one person in a single day. Broad implementation of this technique will facilitate the development of recombinant inbred line populations of S. viridis X S. viridis or S. viridis X S. italica, mapping mutations through bulk segregant analysis and creating higher order mutants for genetic analysis.

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

We have developed a methodology for performing crosses in Setaria viridis (S. viridis). The method involves pruning the panicle prior to a hot water treatment to kill viable pollen. Crosses are performed following a well-controlled growth regime and typically result in the recovery of 1 to 7 cross-pollinated seed/s per panicle.

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail ...

A New Application of the Electrical Penetration Graph (EPG) for Acquiring and Measuring Electrical Signals in Phloem Sieve Elements

Electrophysiological properties of cells are often studied in vitro, after dissociating them from their native environments. However, the study of electrical transmission between distant cells in an organism requires in vivo, artifact-free recordings of cells embedded within their native environment. The transmission of electrical signals from wounded to unwounded areas in a plant has since long piqued the interest of botanists. The phloem, the living part of the plant vasculature that is spread throughout the plant, has been postulated as a major tissue in electrical transmission in plants. The lack of suitable electrophysiological methods poses many challenges for the study of the electrical properties of the phloem cells in vivo. Here we present a novel approach for intracellular electrophysiology of sieve elements (SEs) that uses living aphids, or other phloem-feeding hemipteran insects, integrated in the electrical penetration graph (EPG) circuit. The versatility, robustness, and accuracy of this method made it possible to record and study in detail the wound-induced electrical signals in SEs of central veins of the model plant Arabidopsis thaliana1. Here we show that EPG-electrodes can be easily implemented for intracellular electrophysiological recordings of SEs in marginal veins, as well as to study the capacity of SEs to respond with electrical signals to several external stimuli. The EPG approach applied to intracellular electrophysiology of SEs can be implemented to a wide variety of plant species, in a large number of plant/insect combinations, and for many research aims.

A New Application of the Electrical Penetration Graph EPG for Acquiring and Measuring Electrical Signals in Phloem Sieve Elements

Electrical Penetration Graph (EPG) is a well-established technique for studying the feeding behavior of stylet-bearing insects. Here we show a new application of EPG as a non-invasive tool for the acquisition of intracellular electrophysiology recordings of sieve elements (SEs), the cells that form the phloem vasculature in plants.

Electrophysiological properties of cells are often studied in vitro, after dissociating them from their native environments. However, the study of ...

Methods of Soil Resampling to Monitor Changes in the Chemical Concentrations of Forest Soils

Recent soils research has shown that important chemical soil characteristics can change in less than a decade, often the result of broad environmental changes. Repeated sampling to monitor these changes in forest soils is a relatively new practice that is not well documented in the literature and has only recently been broadly embraced by the scientific community. The objective of this protocol is therefore to synthesize the latest information on methods of soil resampling in a format that can be used to design and implement a soil monitoring program. Successful monitoring of forest soils requires that a study unit be defined within an area of forested land that can be characterized with replicate sampling locations. A resampling interval of 5 years is recommended, but if monitoring is done to evaluate a specific environmental driver, the rate of change expected in that driver should be taken into consideration. Here, we show that the sampling of the profile can be done by horizon where boundaries can be clearly identified and horizons are sufficiently thick to remove soil without contamination from horizons above or below. Otherwise, sampling can be done by depth interval. Archiving of sample for future reanalysis is a key step in avoiding analytical bias and providing the opportunity for additional analyses as new questions arise.

Repeated soil sampling has recently been shown to be an effective way to monitor forest soil change over years and decades. To support its use, a protocol is presented that synthesizes the latest information on soil resampling methods to aid in the design and implementation of successful soil monitoring programs.

Recent soils research has shown that important chemical soil characteristics can change in less than a decade, often the result of broad environmental ...

Development of New Methods for Quantifying Fish Density Using Underwater Stereo-video Tools

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and estimating fish abundance. We developed and implemented a rotating stereo-video camera tool that covers a full 360 degrees of sampling, which maximizes sampling effort compared to stationary camera tools. A variety of studies have detailed the ability of static, stereo-camera systems to obtain highly accurate and precise measurements of fish; the focus here was on the development of methodological approaches to quantify fish density using rotating camera systems. The first approach was to develop a modification of the metric MaxN, which typically is a conservative count of the minimum number of fish observed on a given camera survey. We redefine MaxN to be the maximum number of fish observed in any given rotation of the camera system. When precautions are taken to avoid double counting, this method for MaxN may more accurately reflect true abundance than that obtained from a fixed camera. Secondly, because stereo-video allows fish to be mapped in three-dimensional space, precise estimates of the distance-from-camera can be obtained for each fish. By using the 95% percentile of the observed distance from camera to establish species-specific areas surveyed, we account for differences in detectability among species while avoiding diluting density estimates by using the maximum distance a species was observed. Accounting for this range of detectability is critical to accurately estimate fish abundances. This methodology will facilitate the integration of rotating stereo-video tools in both applied science and management contexts.

We describe a new method for counting fishes, and estimating relative abundance (MaxN) and fish density using rotating stereo-video camera systems. We also demonstrate how to use distance from camera (Z distance) to estimate species-specific detectability.

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and ...

Methods for Image-based Surveys of Benthic Macroinvertebrates and Their Habitat Exemplified by the Drop Camera Survey for the Atlantic Sea Scallop

Underwater imaging has long been used in the field of marine ecology but decreasing costs of high-resolution cameras and data storage have made the approach more practical than in the past. Image-based surveys allow for initial samples to be revisited and are non-invasive compared to traditional survey methods that typically involve nets or dredges. Protocols for image-based surveys can vary greatly but should be driven by target species behavior and survey objectives. To demonstrate this, we describe our most recent methods for an Atlantic sea scallop (Placopecten magellanicus) drop camera survey to provide a procedural example and representative results. The procedure is divided into three critical steps that include survey design, data collection, and data products. The influence of scallop behavior and the survey goal of providing an independent assessment of the U.S. sea scallop resource on the survey procedure are then discussed in the context of generalizing the method. Overall, the broad applicability and flexibility of the University of Massachusetts Dartmouth School for Marine Science and Technology (SMAST) drop camera survey demonstrates the method could be generalized and applied to a variety of sessile invertebrates or habitat focused research.

Image based surveying is an increasingly practical, non-invasive method to sample the marine environment. We present the protocol of a drop camera survey that estimates the abundance and distribution of the Atlantic sea scallop (Placopecten magellanicus). We discuss how this protocol can be generalized for application to other benthic macroinvertebrates.

Underwater imaging has long been used in the field of marine ecology but decreasing costs of high-resolution cameras and data storage have made the ...

Experimental Protocol for Examining Behavioral Response Profiles in Larval Fish: Application to the Neuro-stimulant Caffeine

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and toxicological studies. Using automated digital tracking platforms, recent efforts in neuropharmacology are leveraging larval fish locomotor behaviors to identify potential therapeutic targets for novel small molecules. Similar to these efforts, research in the environmental sciences and comparative pharmacology and toxicology is examining various behaviors of fish models as diagnostic tools in tiered evaluation of contaminants and real-time monitoring of surface waters for contaminant threats. Whereas the zebrafish is a popular larval fish model in the biomedical sciences, the fathead minnow is a common larval fish model in ecotoxicology. Unfortunately, fathead minnow larvae have received considerably less attention in behavioral studies. Here, we develop and demonstrate a behavioral profile protocol using caffeine as a model neurostimulant. Though photomotor responses of fathead minnows were occasionally affected by caffeine, zebrafish&#160;were markedly more sensitive for photomotor and locomotor endpoints, which responded at environmentally relevant levels. Future studies are needed to understand comparative behavioral sensitivity differences among fish with age and time of day, and to determine whether similar behavioral effects would occur in nature and be indicative of adverse outcomes at the individual or population levels of biological organization.

Here, we present a protocol to examine larval zebrafish and fathead minnow locomotor activities and photomotor responses (PMR) using an automated tracking software. When incorporated in common toxicology bioassays, analyses of these behaviors provide a diagnostic tool to examine chemical bioactivity. This protocol is described using caffeine, a model neurostimulant.

Fish models and behaviors are increasingly used in the biomedical sciences; however, fish have long been the subject of ecological, physiological and ...

An Induction System for Clustered Stomata by Sugar Solution Immersion Treatment in Arabidopsis thaliana Seedlings

Stomatal movement mediates plant gas exchange, which is essential for photosynthesis and transpiration. Stomatal opening and closing are accomplished by a significant increase and decrease in guard cell volume, respectively. Because shuttle transport of ions and water occurs between guard cells and larger neighboring epidermal cells during stomatal movement, the spaced distribution of plant stomata is considered an optimal distribution for stomatal movement. Experimental systems for perturbing the spaced pattern of stomata are useful to examine the spacing pattern's significance. Several key genes associated with the spaced stomatal distribution have been identified, and clustered stomata can be experimentally induced by altering these genes. Alternatively, clustered stomata can be also induced by exogenous treatments without genetic modification. In this article, we describe a simple induction system for clustered stomata in Arabidopsis thaliana seedlings by immersion treatment with a sucrose-containing medium solution. Our method is easy and directly applicable to transgenic or mutant lines. Larger chloroplasts are presented as a cell biological hallmark of sucrose-induced clustered guard cells. In addition, a representative confocal microscopic image of cortical microtubules is shown as an example of intracellular observation of clustered guard cells. The radial orientation of cortical microtubules is maintained in clustered guard cells as in spaced guard cells in control conditions.

An Induction System for Clustered Stomata by Sugar Solution Immersion Treatment in Arabidopsis thaliana Seedlings

The goal of this protocol is to demonstrate how to induce clustered stomata in cotyledons of Arabidopsis thaliana seedlings by immersion treatment with a sugar-containing medium solution and how to observe intracellular structures such as chloroplasts and microtubules in the clustered guard cells using confocal laser microscopy.

Stomatal movement mediates plant gas exchange, which is essential for photosynthesis and transpiration. Stomatal opening and closing are accomplished by a ...

A High-throughput Assay for the Prediction of Chemical Toxicity by Automated Phenotypic Profiling of Caenorhabditis elegans

Applying toxicity testing of chemicals in higher order organisms, such as mice or rats, is time-consuming and expensive, due to their long lifespan and maintenance issues. On the contrary, the nematode Caenorhabditis elegans (C. elegans) has advantages to make it an ideal choice for toxicity testing: a short lifespan, easy cultivation, and efficient reproduction. Here, we describe a protocol for the automatic phenotypic profiling of C. elegans in a 384-well plate. The nematode worms are cultured in a 384-well plate with liquid medium and chemical treatment, and videos are taken of each well to quantify the chemical influence on 33 worm features. Experimental results demonstrate that the quantified phenotype features can classify and predict the acute toxicity for different chemical compounds and establish a priority list for further traditional chemical toxicity assessment tests in a rodent model.

A High-throughput Assay for the Prediction of Chemical Toxicity by Automated Phenotypic Profiling of Caenorhabditis elegans

A quantitative method has been developed to identify and predict the acute toxicity of chemicals by automatically analyzing the phenotypic profiling of Caenorhabditis elegans. This protocol describes how to treat worms with chemicals in a 384-well plate, capture videos, and quantify toxicological related phenotypes.

Applying toxicity testing of chemicals in higher order organisms, such as mice or rats, is time-consuming and expensive, due to their long lifespan and ...

Using Caenorhabditis elegans for Studying Trans- and Multi-Generational Effects of Toxicants

Information about toxicities of chemicals are essential in their application and waste management. For chemicals at low concentrations, the long-term effects are very important in judging their consequences in the environment and on human health. In demonstrating long-term influences, effects of chemicals over generations in recent studies provide new insight. Here, we describe protocols for studying effects of chemicals over multiple generations using free-living nematode Caenorhabditis elegans. Two aspects are presented: (1) trans-generational (TG) and (2) multi-generational effect studies, the latter of which is separated to multi-generational exposure (MGE) and multi-generational residual (MGR) effect studies. The TG effect study is robust with a simple purpose to determine whether chemical exposure to parents can result in any residual consequences on offspring. After the effects are measured on parents, sodium hypochlorite solutions are used to kill the parents and keep the offspring so as to facilitate effect measurement on the offspring. The TG effect study is used to determine whether the offspring are affected when their parent is exposed to the pollutants. The MGE and MGR effect study is systematical used to determine whether continuous generational exposure can result in adaptive responses in offspring over generations. Careful pick-up and transfer are used to distinguish generations to facilitate effect measurement on each generation. We also combined protocols to measure locomotion behavior, reproduction, lifespan, biochemical and gene expression changes. Some example experiments are also presented to illustrate the trans- and multi-generational effect studies.

Using Caenorhabditis elegans for Studying Trans- and Multi-Generational Effects of Toxicants

Trans- and multi-generational effects of persistent chemicals are essential in judging their long-term consequences in the environment and on the human health. We provide novel detailed methods for studying trans- and multi-generational effects using free-living nematode Caenorhabditis elegans.

Information about toxicities of chemicals are essential in their application and waste management. For chemicals at low concentrations, the long-term ...

Leaf Area Index Estimation Using Three Distinct Methods in Pure Deciduous Stands

Accurate estimations of leaf area index (LAI), defined as half of the total leaf surface area per unit of horizontal ground surface area, are crucial for describing the vegetation structure in the fields of ecology, forestry, and agriculture. Therefore, procedures of three commercially used methods (litter traps, needle technique, and a plant canopy analyzer) for performing LAI estimation were presented step-by-step. Specific methodological approaches were compared, and their current advantages, controversies, challenges, and future perspectives were discussed in this protocol. Litter traps are usually deemed as the reference level. Both the needle technique and the plant canopy analyzer (e.g., LAI-2000) frequently underestimate LAI values in comparison with the reference. The needle technique is easy to use in deciduous stands where the litter completely decomposes each year (e.g., oak and beech stands). However, calibration based on litter traps or direct destructive methods is necessary. The plant canopy analyzer is a commonly used device for performing LAI estimation in ecology, forestry, and agriculture, but is subject to potential error due to foliage clumping and the contribution of woody elements in the field of view (FOV) of the sensor. Eliminating these potential error sources was discussed. The plant canopy analyzer is a very suitable device for performing LAI estimations at the high spatial level, observing a seasonal LAI dynamic, and for long-term monitoring of LAI.

An accurate estimation of leaf area index (LAI) is crucial for many models of material and energy fluxes within plant ecosystems and between an ecosystem and the atmospheric boundary layer. Therefore, three methods (litter traps, needle technique, and PCA) for taking precise LAI measurements were in the presented protocol.

Accurate estimations of leaf area index (LAI), defined as half of the total leaf surface area per unit of horizontal ground surface area, are crucial for ...

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

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Leaf Area Index Estimation Using Three Distinct Methods in Pure Deciduous Stands