Measurements of Physiological Stress Responses in C. Elegans

Summary

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

Abstract

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

Introduction

The ability of an organism to respond to changes in the intra- and extracellular environment is crucial for its survival and adaptation. This is accomplished on a cellular level through numerous protective pathways that ensure the integrity of the cell. While numerous cellular components are subject to stress-associated damage, one major involvement of cellular stress responses is to repair and protect the homeostasis of the cellular proteome. However, the compartmentalization of proteins into special structures, called organelles, poses a challenge for the cell, as it cannot rely on one centralized form of protein quality control to ensure that all the proteins within the cell are properly folded and functional. Therefore, to deal with perturbations to their proteins, organelles have evolved dedicated quality control mechanisms, which can sense misfolded proteins and activate a stress response in an attempt to alleviate the stress within that compartment. For example, the cytosol relies on the heat shock response (HSR), while the endoplasmic reticulum (ER) and mitochondria rely on their compartment-specific unfolded protein responses (UPR). The OxSR serves to alleviate the toxic effects of reactive oxygen species (ROS). Each stress response is triggered in the presence of cellular challenges and environmental insults and induces a tailored transcriptional response. The hallmarks of these responses include synthesizing molecules that re-fold misfolded proteins (such as chaperones) targeted to the proper organelle, or alternatively, remove damaged proteins by protein degradation. Failure to activate these stress responses results in accumulation of damaged proteins, cellular dysfunction propagated to systemic failure of tissues, and eventually death of the organism. The function and regulation of the different stress responses are reviewed elsewhere¹.

Many insights regarding the regulation and activity of cellular stress responses have been attributed to the nematode, Caenorhabditis elegans, a multicellular model organism in genetic research. Nematodes not only allow studying the activation of stress responses on the cellular level, but also on the organismal level; nematodes have been used to study the effects of genetic perturbations or exposure to drugs and pollutants on their growth and survival. Their quick generation time, isogeny, transparency, genetic tractability, and ease of use during experimentation make them ideal for such studies. Additionally, the relatively quick physiological response to stress (between hours and a few days) and the evolutionary conservation of cellular pathways make nematodes a prominent tool in studying stress resistance.

There are two commonly used E. coli strains used as a food source to grow C. elegans: standard OP50, a B strain in which most experimentation has been historically performed² and HT115, a K-12 strain that is used for almost all RNAi experiments^3,4. It is important to note that there are significant differences between OP50 and HT115 bacterial diets. Growth on these different bacterial sources has been shown to cause major differences in metabolic profile, mitochondrial DNA copy number, and several major phenotypes, including lifespan⁵. Some of these differences are attributed to Vitamin B12 deficiency associated with growth on OP50 bacteria, which can result in defects in mitochondrial homeostasis and increased sensitivity to pathogens and stresses. All of these phenotypes have been shown to be alleviated by growth on HT115 bacteria, which have higher levels of Vitamin B12⁶. Therefore, it is recommended that all experiments on physiological stress responses be performed on HT115 bacteria, regardless of the necessity of RNAi conditions. However, due to the ease of maintaining animals on OP50, all standard growth (i.e., maintenance and amplification of animals) can be performed on OP50, as significant differences in the experimental paradigms described here were not detected in worms maintained on OP50 as long as they were moved to HT115 post synchronization (i.e., from hatch post-bleaching with or without L1 arresting) until experimentation.

Here, the characterization of the activity of cellular stress responses using two functional methods is described. It should be noted that the protocols presented are primarily focused on cellular stress responses and their impact on protein homeostasis. First, fluorescent transcriptional reporters are utilized, which are regulated by endogenous gene promoters that are specifically activated in response to different cellular stresses. These fluorescent transcriptional reporters are based on the transcriptional induction of specific genes that are natively part of the stress response. For example, HSP-4, a heat shock protein orthologous to the human chaperone HSPA5/BiP, is activated upon ER-stress and localizes to the ER to alleviate the stress. In conditions of ER stress (e.g., exposure to tunicamycin), a green fluorescent protein (GFP), placed under the regulation of the hsp-4 promoter, is synthesized in high levels as can be assessed by fluorescent microscopy or quantitatively measured using large-particle flow cytometry of nematodes⁷. Similarly, the promoter of a mitochondrial chaperone, hsp-6 (orthologous to mammalian HSPA9), is utilized to monitor the activation of the UPR^MT8, and the promoter of the cytosolic chaperone hsp-16.2 (orthologous to the human crystallin alpha genes) is used for assessing the activity of the HSR⁹. These reporters allow a rapid characterization of the pathways activated in response to various perturbations.

Often, the reporters presented here are imaged using microscopy, which provides a qualitative output of the activation of stress responses. However, while imaging techniques provide both information on intensity and tissue location of the reporters described above, its quantification is not always accurate or robust. While it is possible to quantify fluorescent activation using imaging analysis tools, these methods are relatively low throughput and sample size is small, due to the relatively low number of animals imaged. The ease and ability to obtain large quantities of animals quickly make C. elegans an ideal model system to assay the activation of fluorescent stress reporters through the use of a large particle flow cytometer. A large-particle flow cytometer is capable of recording, analyzing, and sorting based on size and fluorescence from many live animals. Using this method, it is possible to get the fluorescent intensity, size, and also spatial (2D) information for thousands of worms. The system is controlled using FlowPilot, which allows for real-time data acquisition and analysis of the measured parameters. Here, methods for both microscopic imaging and quantitative analysis using a large-particle flow cytometer are offered as methods to measure the activation of stress responses.

Beyond reporter analysis, the sensitivity or resistance of animals to stress can be measured using physiological stress assays. This is achieved by exposing animals to stressful environments that activate specific cellular stress pathways. Here, several methods are provided to measure sensitivity of whole animals to specific types of stressors.

ER stress is applied to C. elegans by using the chemical agent, tunicamycin, which blocks N-linked glycosylation, causing accumulation of misfolded proteins in the ER¹⁰. In C. elegans, growth upon exposure to tunicamycin results in major perturbations in ER function, and a significantly decreased lifespan¹¹. By measuring the survival of animals on tunicamycin-containing plates, ER stress sensitivity of animals can be quantified. For example, animals with ectopic UPR^ER induction and thus increased resistance to protein misfolding stress in the ER have an increased survival upon tunicamycin exposure compared to wild-type animals¹².

Oxidative and mitochondrial stress is applied to C. elegans by exposing animals to the chemical agent, paraquat. Paraquat is a commonly used herbicide, which causes superoxide formation specifically in the mitochondria¹³. Due to the specific localization of mitochondria-derived reactive oxygen species (ROS), paraquat assays are often used as a "mitochondrial" stress assay. However, superoxide is rapidly converted into hydrogen peroxide by mitochondrial superoxide dismutases (SODs)¹⁴. Hydrogen peroxide can subsequently diffuse out of the mitochondria and cause oxidative stress in other compartments of the cell. Therefore, we describe paraquat survival assays as measuring sensitivity to both mitochondrial and oxidative stress (other oxidative stress assays can be found¹⁵).

Thermotolerance assays are performed in C. elegans by placing animals in elevated temperatures. Ambient temperatures for nematodes are ~15-20 °C and thermal stress is induced at temperatures above 25 °C^16,17. Thermotolerance assays are generally performed at temperatures ranging from 30-37 °C, as animals exhibit major cellular defects at this temperature, and survival assays are completed within 24 hours^16,18. Here, two alternative methods are provided for performing thermotolerance assays: growth at 34 °C and growth at 37 °C. Together, the protocols presented here can be utilized to perform large-scale screens when combined with standard gene knock-down using RNA interference or chemical drug libraries.

The protocol can be broken into 4 broad procedures- growth of C. elegans and preparation for imaging (sections 1 and 2), imaging of transcriptional reporters using fluorescent microscopy (sections 3-5), quantitative measurements of reporters using a large-particle flow cytometer (section 6), and physiological assays to measure stress sensitivity in C. elegans (section 7).

Protocol

1. Standard growth conditions of temperatures & OP50 vs HT115

1. Standard growth and expansion
   1. Grow a culture of OP50 in LB (Table 1) or equivalent media of choice for 24-48 h at ambient temperature (~22-25 °C). Grow bacteria at room temperature as OP50 is an uracil auxotroph and there is a higher incidence of revertants (e.g., suppressor mutants) when grown at 37 °C. Long-term storage of OP50 cultures is not recommended (max 1 week at 4 °C).
   2. Seed a volume of ~100-200 µL of saturated OP50 culture onto a 60 mm NGM plate (Table 1) for maintenance of worms and 1 mL of saturated OP50 culture onto a 100 mm plate for expanding animals for experimentation.
   3. Let plates dry overnight on a benchtop.
      NOTE: 100 mm plates may need more time to dry, especially if using non-vented plates. Store all C. elegans plates in air-tight containers stored at 4 °C. Do not use plates past 6 months old as desiccation of plates will occur, which will change animal physiology on plates due to different osmotic pressure and stiffness of plates.
   4. For standard maintenance, move 10-15 young animals (eggs, L1, or L2 stages) onto a 60 mm plate, although more can be moved if dealing with mutants or transgenic animals with lower fecundity. For animals with developmental or fecundity defects, chunk a more variable pool of animals to prevent loss of the stock. If kept at 15 °C, move animals every week; for 20 °C, move animals every 3-4 days.
   5. Perform a fresh thaw of animals every 25-30 passages (~ 6 months if animals are moved once a week and kept at 15 °C).
   6. For expansion, chunk a full 60 mm plate onto 100 mm plates for expansion. As a frame of reference, animals with wild-type fecundity can have a full 60 mm plate cut into 4ths-6ths and be chunked onto a large plate at 20 °C for 2 days or 15 °C for 3 days to create a full 100 mm plate without reaching starvation.

2. Staging/synchronization of worms using bleaching

1. Bleaching protocol to synchronize worms
   1. Wash gravid worms (adults full of eggs) off agar plates using M9 (Table 1). Start from a non-synchronized population of worms (i.e., chunked worms from step 1.1.6) to bleach for experiments, as significant genetic drift may occur upon successive rounds of synchronization via bleaching.
   2. Move worm/M9 mixture into a 15 mL conical tube using a glass pipette; several plates of worms can be collected into a single 15 mL conical tube.
   3. Spin animals down for 30 s at 1,000 x g. Aspirate M9 supernatant. It is unnecessary to wash off residual bacteria unless there is an egregious amount of contamination or large clumps of bacteria. If this is the case, perform several washes with M9.
   4. Prepare a fresh stock of bleaching solution (Table 1). Bleaching solution can be kept for several days at 4 °C, but use freshly made bleaching solution as inefficient bleaching can result in uneven bleaching, which will cause damage to eggs before eliminating all worm carcasses.
   5. Add 2-10 mL of bleaching solution to the animals (~1 mL of bleach solution for every ~0.1 mL of animal pellet). Invert the bleach and worm mixture for ~5 min (do not exceed 10 min; note that bleaching times may vary and should be titrated specifically for each lab). Vigorously shake to help dissolve worm carcasses faster and for optimal preservation of eggs. Periodically look under a dissection microscope or put the conical tube to a light to observe when the adult worm carcasses have fully dissolved and only eggs remain in the tube.
   6. Pellet the eggs by spinning at 1,000 x g for 30 s and then aspirate the supernatant. Eggs can be centrifuged faster than worms without disrupting their integrity, so if unsure of centrifuge speed, eggs can be spun down at up to 2,500 x g without affecting their physiology.
   7. Add M9 up to 15 mL and invert the tube to clear bleach off the eggs.
   8. Repeat steps 2.1.6-2.1.7 (wash/pellet process) 2-3 more times to remove bleach.
   9. Pipet egg/M9 mix onto plates and grow at 15-20 °C for experimentation. To get a measure of how many worms to use, perform approximate egg counts by pipetting 5 µL of egg mixture onto an agar plate or slide and dividing the egg count by 5 to determine the number of eggs per 1 µL of volume. Averaging of 3 independent counts may improve accuracy. To avoid starvation, a table of recommended egg counts per plate is available in Table 2.
   10. If needed, L1 arrest animals for tighter temporal synchronization by placing egg/M9 mix in a rotator at 20 °C for up to 24 h. For wild-type animals, no defects in animal physiology were detected when L1 arresting for up to 48 h. However, mutants that are sensitive to starvation (e.g., lysosome or autophagy mutants) do very poorly with L1 arresting, and thus it is not recommended to perform this synchronization method for mutants that are known to be sensitive to starvation. If tighter synchronization is required for animals that cannot be L1 arrested, use the egg-lay method described in section 2.2.
2. Egg-lay protocol for synchronizing worms
   NOTE: As an alternative method to bleaching, an egg-lay assay can be performed. Egg-lay is used when bleaching of animals does not provide a close enough synchronization, as eggs within the egg sac of adult animals can be as different as 8-12 h apart. For experimental paradigms where it is critical for animals to be as closely staged as possible, but where L1 arresting is not possible (e.g., in starvation mutants), egg-lay assays are recommended. However, it should be noted that due to the labor involved in the egg-lay protocol, it is less feasible to perform high-scale experiments.
   1. Place 4-12 gravid adults (see Note after step 2.2.2) onto a standard OP50 or HT115-seeded NGM plate (see Section 1 above for recommendations on bacterial strains). Depending on the scale of experiments, multiple plates can be used. Be sure to carefully document how many animals are on each plate for step 2.2.
   2. Place animals at desired temperature for experiments (15-20 °C) for 4-8 h.
      NOTE: The number of hours animals are left on the plate will determine how closely synchronized the first egg laid and the last egg laid will be, so the timing can be adjusted as needed. Since shorter incubation times will mean each animal has less time to lay eggs, more animals should be placed onto plates to ensure enough eggs are laid. While the actual egg-laying rate is not fully normalized due to the tendency of animals to go through short bursts of egg-laying rather than a normalized rate of egg-lay, the average rate of eggs laid per animal can be estimated at ~5 eggs/hour for animals exhibiting wild-type fecundity¹⁹. When trying to grow animals to day 1 adult stage, time the egg-lay to have <100 eggs per plate to avoid starvation (refer to Table 2 for more details on recommended animals per plate).
   3. Remove adult animals from plates. Ensure that all adult animals are removed from plates, as animals will continue to lay eggs and result in an unsynchronized population and/or starvation of the plate.

3. Growth conditions of worms for imaging of transcriptional reporters

1. Growth of worms
   1. Inoculate bacterial culture of HT115 harboring pL4440 RNAi plasmid (EV) and/or carrying an RNAi cassette against desired target gene(s) into LB media supplemented with 100 µg/mL carbenicillin and 20 µg/mL tetracycline.
   2. Grow culture overnight (~16 hours) to saturation in a shaking 37 °C incubator.
   3. Spot 60 mm NGM RNAi plates with 200 µL of saturated bacterial culture and 100 mm NGM RNAi plates with 1,000 µL of saturated bacterial culture. Let dry at ambient temperature (~22 °C) overnight in the dark (covered loosely with aluminum foil).
   4. Place synchronized population of transgenic worms carrying fluorescent reporters (see Table 3 for complete list) onto NGM RNAi plates seeded with bacteria of choice.
   5. Grow at 15-20 °C to required stages for specific reporters as outlined below.
2. Considerations for staging of worms for experimentations
   1. Perform experiments for transcriptional reporters at day 1 of adulthood, with the exception of assays that require L4 animals. According to WormAtlas, L4 animals are obtained approximately 2.5 days (~56 h) of growth at 20 °C from the egg stage (www.wormatlas.org).
   2. For "day 1 adults," use animals at approximately 3-4 days (~65-96 h) of growth at 20 °C after plating eggs.
      NOTE: This wide range is explained as follows: ~65 h at 20 °C is when animals reach "egg-laying," which is the true "adulthood" state. 96 h is when animals enter what WormAtlas describes as "egg-laying maximal," which is when the adult is a gravid adult and has a full egg sac. This is when animals would be described as being older than day 1 and may start to display differences, and thus the protocols described here are all recommended to start in this "day 1" stage starting as early as 65 h and as late as 96 h. For the assays described here, major differences were not observed when using animals in this ~65-96 h window.
   3. For replicates of a single experiment, use a similar time point for more robust reproducibility (e.g., perform all experiments at ~65 h or ~96 h after plating eggs).
      NOTE: Some transgenic animals and mutants display slowed growth rates. For this, two recommendations exist. 1) Use staggered synchronization, where animals can be bleached at different times in order for experimentation to be performed at the same time. This is recommended when technical variability in the assay may be larger than the technical variabilities that may arise from the synchronization assay (e.g., survival assays, which run for long durations may suffer if they are not performed simultaneously). 2) Use staggered analysis, where animals can be bleached at the same time, but the assay itself is performed at different times. This is recommended for very simple assays that do not have inherent variability (e.g., RNAi-induction of stress responses).

4. Induction of stress responses

1. Using hsp-4p::GFP as a readout for the activation of the UPR^ER
   1. Inducing ER stress using RNAi
      1. Prepare plates spotted with RNAi bacteria targeting the gene of interest onto NGM RNAi plates (Table 1) as in section 3. Use EV as a control for basal UPR^ER levels and RNAi knockdown of tag-335 (enzyme for N-linked glycosylation of ER resident proteins; RNAi knockdown has similar effects to tunicamycin treatment) as a positive control for activation of the UPR^ER under ER stress.
      2. Synchronize hsp-4p::GFP reporter animals using methods described in section 2.
      3. Plate eggs onto RNAi plates using criteria recommended in Table 2.
      4. Incubate eggs at 20 °C for approximately 3-4 days (~65-96 h) to perform experiments at day 1 of adulthood. For the UPR^ER experiments, there are differences in basal fluorescence of the hsp-4::GFP reporter when grown at different temperatures. Therefore, perform all experiments at 20 °C.
   2. Inducing ER stress using a chemical agent
      1. Synchronize hsp-4p::GFP reporter animals using methods described in section 2 and grow at 20 °C until animals reach the L4 stage. Transfer animals to a tube of M9. Allow worms to settle (~2-3 min by gravity or a 30 s spin at 1,000 x g), and then remove M9.
      2. Dilute tunicamycin to 25 ng/mL in M9 (a 1:40 dilution from 1 mg/mL stock). As a control, also dilute an equivalent volume of DMSO in M9.
      3. Add 25 ng/mL tunicamycin/M9 or control DMSO/M9 solution to worms (use 400-500 mL for a 1.5 mL tube or 2 mL for a 15 mL conical tube). Incubate at 20 °C on a rotating platform for 3-4 hours.
         NOTE: Tunicamycin can also be diluted directly into worm/M9 mix.
      4. Allow worms to settle, remove M9/TM solution, and wash with 1 mL of M9.
      5. Transfer animals to NGM plates or NGM RNAi plates and allow to recover overnight (or ~15-20 h) and reach Day 1 of adulthood at 20 °C prior to performing fluorescent microscopy (section 5). An overnight recovery is performed to allow for a detectable level of GFP to accumulate.
      6. Alternatively, induce hsp-4p::GFP by moving L4 animals to agar plates containing 25 ng/µL tunicamycin for 16-24 h. This has a much more robust induction of hsp-4p::GFP due to the longer duration of stress, and thus the dynamic range is much lower than the assay described above.
2. Using hsp-6p::GFP as a readout for the activation of the UPR^MT
   1. Inducing mitochondrial stress using RNAi
      1. Activate UPR^MT following the same protocol as section 4.1.1, except using RNAi knockdown of mitochondrial genes, such as cox-5b/cco-1 (cytochrome c oxidase subunit 5B; knockdown inhibits electron transport chain activity). For UPR^MT activation through perturbations in electron transport chain function, RNAi knockdown needs to be performed during early development²⁰. Therefore, perform RNAi from hatch for these experiments.
   2. Inducing mitochondrial stress using a chemical agent
      1. Prepare plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria even in experiments not involving RNAi knockdown (see Section 1). Ensure that both NGM RNAi plates (or NGM RNAi + DMSO0.2) and NGM RNAi + antimycin A plates (Table 1) are both prepared for step 4.2.2.5.
      2. Synchronize hsp-6p::GFP or hsp-60p::GFP reporter animals using methods described in section 2.
      3. Plate eggs onto seeded plates of choice using criteria recommended in Table 2. Since antimycin A is dissolved in DMSO, grow the animals on NGM RNAi + DMSO0.2 plates from hatch.
      4. Incubate eggs at 20 °C for 2 days (~56 h) to the L4 stage. Alternatively, grow animals at 15 °C for 3 days (~75 h) instead.
      5. Move worms from NGM RNAi + DMSO0.2 plates to NGM RNAi + antimycin A plates or NGM RNAi + DMSO0.2 plates as a control. Worms can be moved manually with a pick for small-scale experiments, but for large-scale experiments, we recommend washing animals with M9, settling with centrifugation, aspirating M9, and then plating to NGM RNAi + antimycin A plates.
      6. Incubate worms for an additional ~20 h and image at day 1 adult (section 5).
3. Using gst-4p::GFP as a readout for the oxidative stress response.
   1. Inducing OxSR using RNAi
      1. Alternatively, induce gst-4p::GFP using RNAi-knockdown of wdr-23 (it encodes a negative regulator of skn-1; thus, its knockdown induces OxSR) using the protocol described in section 4.1.1.
   2. Inducing the OxSR using exposure to the chemical oxidant Tert-butyl hydroperoxide (TBHP)
      1. Prepare plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria even in experiments not involving RNAi knockdown (see Section 1).
      2. Synchronize gst-4p::GFP reporter animals using methods described in section 2.
      3. Plate eggs onto seeded plates of choice using criteria recommended in Table 2. Be sure to prepare more plates than necessary as the drug treatment protocol results in loss of >10-20% of animals. For imaging, start with >100 animals, and for sorting, start with >1,000 animals.
      4. Incubate eggs in 20 °C for 2 days (~56 h) to the L4 stage. Alternatively, grow animals at 15 °C for 3 days (~75 h) instead.
      5. Wash L4 animals off the plates and split into two 15 mL conical tubes per condition. Aspirate volume down to at least 1 mL and add an equal volume of freshly made 2 mM TBHP. Use a total liquid volume to at least 2 mL, as lower volumes can cause significant death of worms when spinning. Do not wash prior to drug treatment, as residual bacteria from plates will help to ensure worms to not overstarve during incubation.
      6. Incubate worms on the rotator for 4 h at 20 °C.
      7. Wash worms by spinning at 1,000 x g, aspirating M9 + TBHP mix, and replacing with 15 mL of M9. Repeat for a second wash.
      8. Plate worms on EV RNAi to recover overnight (~16-24 h) at 20 °C. Worms can be recovered on matching RNAi of choice, but no significant differences were seen when recovered on EV RNAi, so this can be done for ease of experimental set-up. Take images of day 1 adults after recovery.
         NOTE: An overnight recovery is performed to allow for a detectable level of GFP to accumulate. Without this recovery, there is no detectable GFP signal. If a shorter-term recovery is desired, it is possible to use the assay described in section 4.3.3.
   3. Inducing the OxSR using exposure to the chemical oxidant paraquat (PQ)
      1. Repeat all steps in section 4.2.2 to get 2 batches of L4 gst-4p::GFP animals in 15 mL conical tubes per condition.
      2. Aspirate volume down to at least 1 mL and add an equal volume of freshly made 100 µM PQ. Similar to 4.3.2.5, a minimum volume of 2 mL is recommended.
      3. Incubate worms on the rotator for 2 h at 20 °C.
      4. Wash worms by spinning at 1,000 x g, aspirating M9 + PQ mix, and replacing with 15 mL of M9. Repeat for a second wash.
      5. Plate worms on EV RNAi to recover for 2 h at 20 °C, and then take images immediately after recovery.
         NOTE: 2 h of recovery was the minimal recovery required to visualize the GFP induction.
4. Using hsp-16.2p::GFP and hsp-70p::GFP as a readout for HSR activation.
   1. Inducing HSR using exposure to elevated temperatures
      1. Prepare plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria, even in experiments not involving RNAi knockdown (see Introduction).
      2. Synchronize hsp-16.2p::GFP or hsp-70p::GFP reporter animals using methods described in section 2.
      3. Plate eggs onto seeded plates of choice using criteria recommended in Table 2. Be sure to prepare 2x the number of plates necessary as half of the sample will be exposed to elevated temperatures for heat-shock induction, and the other half will serve as a non-heat-shocked control.
      4. Incubate eggs at 20 °C for approximately 3-4 days (~65-96 h) to perform experiments at day 1 of adulthood. Do not grow worms at 15 °C for heat-shock experiments as there are only minor differences between animals experiencing heat-shock out of 15 °C versus 20 °C.
      5. Move experimental groups of animals to a 34 °C incubator for 2 h. Place plates in the incubator as a single layer (i.e., no stacking of plates) to ensure the fastest and most equal distribution of heat across the plates.
      6. Move heat-shocked animals to a 20 °C incubator to recover for 2 h and then image immediately (see Section 5). Animals can be recovered longer if necessary for higher GFP induction.
         NOTE: 2 h of recovery was the minimal recovery required to visualize the GFP induction.

5. Imaging using a stereo microscope or low-magnification wide-field/compound microscope

1. Preparation of worms for fluorescent microscopy
   1. Pipette 5-10 µL of 100 mM sodium azide on top of a standard NGM plate (i.e., no bacteria).
      NOTE: Sodium azide concentration can be brought down as low as 10 mM, although the most robust immobilization was observed at 100 mM sodium azide with no detectable effect on fluorescent signal.
   2. Under a dissecting microscope, pick 10-20 animals from experimental plates, and transfer into the spot of sodium azide. Animals should cease movement shortly after landing in sodium azide, and sodium azide itself will evaporate within seconds.
   3. Once sodium azide has evaporated, line up animals to desired imaging setup. Move the animals side by side with anterior and posterior sides in the same orientation for all animals. Image animals immediately.
      NOTE: No changes in reporter signal were observed for any of the transcriptional reporters in Table 3 for up to 15 min after paralyzing in sodium azide.
2. Image acquisition using a stereomicroscope
   NOTE: For this protocol, a Leica M205FA microscope equipped with a Leica DFC3000G monochromatic CCD camera, standard Leica GFP filter (ex 395-455, EM 480 LP), and LAS X software was used. Recommended settings for exposure times can be found in Table 4.
   1. Launch LAS X program.
   2. Start a new project: Open the acquisition tab, click on Open Projects and click Folder to open a new project. This project can be renamed by right-clicking the folder and scrolling down to Rename or clicking F2. In the acquisition tab, there are also options to adjust exposure time and zoom to desired settings.
   3. Position the worm sample under the microscope objective and locate the correct focal point of worms using the bright-field setting to minimize fluorescent bleaching. The center of the sample is where the line of eggs is clearly visible and not fuzzy. Set the exposure time, zoom, focus, and bright-field condensers to desired settings.
   4. Acquire an image using the Capture Image button.
   5. Save the image in .lif (Leica Image File) format as this saves all raw images and metadata. A TIFF can also be exported by right clicking the image (or project) and under the Save as option, click TIFF. This will store all channels (e.g., bright-field and GFP) with any modifications (e.g., if contrast was adjusted, this will be saved into the TIFF).
3. Quantitative analysis of fluorescent images
   1. If quantitative analysis will be performed, take 3-D images. This is performed by clicking the z-section option labelled with "z" at the top right. Z-sections will be active if this box is red.
   2. Optimize z-sections by selecting the range and slice thickness in the bottom left of adjustable options. Whenever possible, use the System Optimized button for optimal settings.
   3. Capture the image and store the image as described above in section 5.2. Line up worms with spaces between them for easier measurements.
   4. Import TIFF images into the imaging software of choice (e.g., ImageJ).
   5. For ImageJ, from the Analyze menu, choose Set Measurements. Check the following: area, mean gray value, integrated density, display label.
   6. Using the ROI (region of interest) tool, draw a ROI. Measure each worm individually. From the Analyze menu, choose Measure or press M. Draw a ROI in the background where there are no worms, and then measure the background in the same manner. Copy or save measurements that appear in the Results window.
   7. Subtract the background integrated density from the integrated density of each measured ROI. The background intensity for a ROI is defined as the product of the background mean gray value and the area of the ROI drawn.
4. Image acquisition using a compound/wide-field microscope
   NOTE: For imaging of transcriptional reporters using a compound/wide-field microscope, this protocol uses a Revolve ECHO R4 microscope equipped with an Olympus 4x Plan Fluorite NA 0.13 objective lens, a standard Olympus FITC filter (ex 470/40; em 525/50; DM 560), and an iPad Pro for the camera and to drive the ECHO software. Recommended settings for exposure times can be found in Table 4.
   1. Use the touchpad to launch the control program. Create a new album and file name.
   2. Position plate under the objective lens. Set the exposure time and fluorescence intensity by using the baseline (EV/control treatment) and positive control, so that the signal is visible but not saturated.
   3. Save a bright-field image and GFP/FITC image.

6. Quantitative measurements of reporters using a large-particle flow cytometer

NOTE: Growth and preparation of worms for large-particle flow cytometer analysis can follow the same paradigms as sections 1-5 for preparation of worms for fluorescent imaging, with the exception that a larger number of animals are required. Use >500 animals per condition, as some animals are lost during manipulation, not all animals pass the filtering criteria during quantification, and some animals are not properly read by the flow cytometer. Wash animals ready for sorting off plates in 5-10 mL of M9 solution into 15 mL conical tubes for subsequent sorting on the flow cytometer.

1. Sorting setup using a large particle flow cytometer
   1. Before turning on the flow cytometer
      1. Make sure the sheath liquid bottle is not empty. Prepare sheath liquid from the 250x stock at least a few hours prior to using the sorter, as there is a small amount of detergent in the sheath fluid, which can cause bubbles that can cause artifacts during acquisition.
      2. Ensure that all waste containers are not full.
   2. Turning on the flow cytometer
      1. Turn on the air compressor. Turn it to Auto. Check the pressure gauge - it should be around 30 psi.
      2. Turn on the instrument. Use the power switch that is next to the power cord on the left of the instrument.
      3. Turn on lasers. A 488 nm light source is usually sufficient for most experiments, although a 561 nm light source needs to be used if higher excitation is required for red fluorescence.
      4. Open the FlowPilot software; the instrument should make a series of clicks switching on the different valves.
      5. Turn on the lasers in the software window by clicking Start. Initiate laser in the Argon laser control popup window by clicking Run. This should cause the laser to turn on and reach around 12 mW. The 488 light source level will go up to around 12.
      6. Click Done to close the window.
   3. Checks on software before progressing
      1. Check pressure gauges -Look at the 4 pressure values displayed in the bottom of the window. The values should be around the original setup (Sheath 5.5-5.7; Sample 5.7-6.0; Sorter 3.1-3.3; Clean 8.5-8.7). If it looks similar, check the box next to Pressure OK.
      2. Check fluidics -To make sure there are no air bubbles and debris blocking the flow of sheath/sample through the flow cell, click Clean several times.
      3. Check sheath flow rate -For this, collect sheath for 60 s. Switch Off Sort, then switch On Sheath in the manual controls to start the flow of sheath. Collect in a 15 mL tube for 60 s; the flow rate should be ~9-10 mL/min.
   4. Cleaning before use of the flow cytometer
      1. Put ~3-5 mL of 10% bleach solution into the collection 'cup' and click Acquire. Let run for ~30 s, click Abort, and remove excess with vacuum.
      2. Rinse the collection 'cup' with deionized water and remove with vacuum. Repeat 2x.
      3. Put ~3-5 mL of COPAS cleaning solution into the collection 'cup' and click Acquire. Let run for ~30 s, click Abort, and remove excess cleaning solution with vacuum.
      4. Rinse the collection 'cup' with deionized water and remove with vacuum. Repeat 2x.
      5. Put ~3-5 mL of M9 solution into the collection 'cup' and click Acquire. Let run for ~30 s, click Stop, and remove excess M9 solution with vacuum.
2. Running samples on sorter
   1. Adjust laser PMT power and size gating based on the condition that causes the brightest activation of the transcriptional reporter of interest. Recommended settings can be found in Table 4.
   2. Add prepared worms to 'cup'. Click Acquire. Watch to make sure that all the liquid is not taken up into the machine; this will cause the flow cytometer to take in air and create bubbles in the detector.
   3. Click Abort when the sample is low and/or enough animals have been collected.
   4. Click Setup | Data Storage | Gated Only. This will save the data only based on the size constraints. Click Store Gated and save gated data. Click Erase to erase data.
   5. Rinse the collection 'cup' with deionized water and remove with vacuum. Repeat 2x.
   6. Repeat steps 6.2.1-6.2.5 with the rest of the samples.
3. Calibration/ Quality Control - if necessary
   NOTE: This requires running control 42 µM GYR fluorescent particles provided, to calibrate the 488 laser.
   1. Press the metal lip on the top of the sample cup to remove the air tube. Unscrew the cap and use a syringe to remove liquid from the sample cup.
   2. Mix the bottle of control particles well before use and add few milliliters into the sample cup. Close the cap and put the air back on by pressing it on till it clicks in place.
   3. In the software, go to the Tools option and click Run Control Particles.
   4. For control particles, reset the PMT values to: GREEN - 325; YELLOW - 365; RED - 575.
   5. Click Acquire. The sheath should turn on followed by sample. Once the beads start to go through the flow cell, the flow rate will be seen at the bottom of the screen. Optimally, the flow rate should be between 5/s and 15/s. If the flow rate is too low or zero, turn the sample valve physically clockwise to increase flow rate. If the flow rate is too high, turn the sample valve physically anti-clockwise to decrease flow rate.
      NOTE: Normally, under bead saver mode, 500 beads are read before switching off. The data can be erased and beads re-read.
   6. Once the reading is completed, check for clean single peaks for the 5 parameters as well as the CV values. The Coefficient of Variance (CV) should be <15%. Also, make sure the CV values for the three different fluorescent channels are close to each other.
   7. Make a record of the QC check: under the File tab, click Save as screen image.
4. Cleaning and shutdown
   1. Use a vacuum to remove the sample from the sample cup and repeat section 4.1.4.
   2. Put ~3-5 mL of deionized water into collection 'cup' and click Acquire, let run for ~ 30 s, and then click Abort. Leave some distilled water behind in the sample cup.
   3. Empty the sample recovery cup and the waste bottle.
   4. Turn off software. Under the File tab, click Exit. In the pop-up menu, click Turn Off Without Purging.
   5. Turn off the laser, turn off the instrument, and then turn off the air compressor. Close the hatch to cover the instrument.

7. Physiological assays to measure stress sensitivity in C. elegans

1. Measurement of ER stress sensitivity using tunicamycin exposure
   1. Prepare NGM RNAi DMSO plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria even in experiments not involving RNAi knockdown (see Section 1). Remember to also seed NGM RNAi TM plates (see Table 1). Seed a sufficient amount of plates: plan for ~5-7 sets of NGM RNAi DMSO plates and ~2-3 sets of NGM RNAi TM plates.
   2. Synchronize animals of choice using methods described in section 2.
   3. Plate eggs onto NGM RNAi DMSO seeded plates of choice using criteria recommended in Table 2. Be sure to prepare 2x the number of plates necessary as half of the sample will be transferred to NGM RNAi TM plates.
   4. Incubate eggs at 20 °C for approximately 3-4 days (~65-96 h) to day 1 of adulthood.
   5. At day 1, prepare lifespans by transferring animals onto separate plates. To conserve plates (as TM costs are high), use 8 plates of 15 animals per condition, for a total of 120 animals per condition. This allows a manageable number of animals per plate for scoring and allows a sufficient amount of animals for statistical analyses, even with some censorship.
   6. For the first 5-7 days, move adult animals away from progeny every day onto a new plate until progeny are no longer visible. During this stage, censor animals that are bagged, exhibit vulval protrusions/explosions, or crawling up the sides of the plates, as these are not deaths associated with ER stress sensitivity. Note that TM treatment causes arrest in animals, and thus only 1-2 moves of these animals every 2-3 days is sufficient to minimize the costs associated with producing TM-containing plates. Wild-type animals have an average survival of ~15-17 days on DMSO and 12-14 days on tunicamcyin.
   7. After animals have stopped producing progeny, score lifespans every 1-2 days until all animals are scored as dead or censored. TM-treat animals every day during day 6-14 of adulthood for higher resolution.
2. Measurement of mitochondrial and oxidative stress sensitivity using exposure to paraquat
   1. Prepare plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria even in experiments not involving RNAi knockdown (see Introduction).
   2. Synchronize animals of choice using methods described in section 2.
   3. Plate eggs onto NGM RNAi seeded plates of choice using criteria recommended in Table 1. The assay calls for ~60-100 animals per condition, so prepare accordingly.
   4. Incubate eggs at 20 °C for approximately 3-4 days (~65-96 h) to day 1 of adulthood.
   5. Prepare a fresh vial of 100 mM paraquat in M9 solution.
   6. Pipette 50-75 µL of M9+paraquat into as many wells of a flat-bottom 96-well plate as desired. It is generally recommended to have ~8-10 wells per condition containing ~8-10 animals per well. This allows for an easily visible number of animals per well with ~80 animals per strain.
   7. Pick 8-10 animals per condition and transfer them into each well containing M9+paraquat. Use a pick to transfer animals into the wells rather than pipetting to avoid differences in volume and unintended changes in paraquat concentrations.
   8. Every 2 h, score for death of animals in each well. Tap the plates gently, which will cause live animals to thrash or bend. Note that it is possible that live animals are sometimes paralyzed long enough to be scored as dead. Therefore, if the number of live animals exceeds the number of live animals from a previous timepoint, it is likely that the animal was alive and should be unscored (e.g., if at hour 4, 2/10 animals are scored as dead, and at hour 6, only 1/10 animals are dead, hour 4 should be rescored as 1/10 animals dead).
3. Measurement of heat sensitivity using exposure to elevated temperatures
   1. Prepare plates spotted with RNAi bacteria targeting the gene of interest as in section 3. Use HT115 bacteria even in experiments not involving RNAi knockdown (see Section 1).
   2. Synchronize animals of choice using methods described in section 2.
   3. Plate eggs onto seeded plates of choice using criteria recommended in Table 1. Have 60-100 animals per condition for thermotolerance assays. For thermotolerance assays, use L1 arrest or egg lay assays for the best synchronization as there is major variability based on age of animals in the assay.
   4. Incubate animals at 20 °C for approximately 3-4 days (~65-96 h) to perform experiments at day 1 of adulthood. Do not grow worms at 15 °C for heat-shock experiments as there is a minor difference between animals experiencing heat-shock out of 15 °C versus 20 °C.
   5. At day 1, prepare animals by transferring them onto separate plates. It is generally recommended to have ~10-15 animals per plate with 4-6 plates, for a total of 60 animals. This allows a manageable number of animals per plate for scoring and allows for minimal time for animals to be pulled out of elevated temperatures.
   6. Place animals into a 37 °C incubator and score every 2 h. Start scoring for thermotolerance at 37 °C at hour 5, as little to no death occurs prior to 5 hours. Median thermotolerance is accomplished at ~9 hours, so hour 7, 9, and 11 are critical timepoints, although due to incubator and lab-to-lab variability, this may need to be titrated per lab. Moreover, any methods to decrease variability will help (e.g., not stacking plates, placing plates in the same area within a single incubator, minimizing time the incubator is opened or closed, taking as few plates out of the incubator at a time to minimize time animals spend outside of 37 °C, etc. For a full guide, see²¹).
   7. As an alternative to step 7.3.6, place animals at 34 °C instead of 37 °C. Median thermotolerance at 34 °C is at a little over 14 h, so time points 12, 14, and 16 are critical for 34 °C thermotolerance assays.

Results

Using transcriptional reporters to measure activation of stress responses
Here, fluorescent transcriptional reporters are used, which serve as robust tools to measure activation of most stress responses in C. elegans. GFP expression is driven under the promoter of canonical targets of master transcriptional regulators involved in responding to compartment-specific stresses. A comprehensive list of commonly used transcriptional reporters is available in

Discussion

Here, methods to interrogate cellular stress responses in C. elegans, using fluorescent transcriptional reporters and physiological stress survival assays are described. The reporters all utilize GFP expression driven under the promoter of a downstream transcriptional target of the transcription factors involved in mounting cellular stress responses. The use of hsp-4p::GFP modulated by XBP-1s-mediated UPR^ER, hsp-6p::GFP controlled by ATFS-1-mediated UPR^MT, gst-4p::GFP<...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Antimycin A	Sigma-Aldrich	A8674	for mitochondrial stress
Bacto Peptone	Fisher Scientific	DF0118072	for NGM plates
BD Difco granulated agar	VWR	90000-782	for NGM plates
Calcium chloride dihydrate	VWR	97061-904	for NGM plates
Carbenicillin	BioPioneer	C0051-25	for RNAi
Cholesterol	Sigma-Aldrich	57-88-5	for NGM plates
COPAS Biosorter	Union Biometrica	350-5000-000	equipped with a 488 nm light source.
COPAS Cleaning Solution	Union Biometrica	300-5072-000	to use with COPAS
COPAS Sheath Solution	Union Biometrica	300-5070-100	to use with COPAS
DMSO	Sigma-Aldrich	472301	solvent for drugs
IPTG dioxane free	Denville Scientific	CI8280-4	for RNAi
LB Broth Miller	Fisher Scientific	BP1426500	for LB
M205FA stereoscope	Leica	10450040	equipped with a Leica DFC3000G monochromatic CCD camera, standard Leica GFP filter (ex 395-455, EM 480 LP), and LAS X software
Magnesium sulfate heptahydrate	VWR	EM-MX0070-3	for NGM plates, M9
Paraquat	Sigma-Aldrich	36541	for oxidative/mitochondrial stress
Potassium Chloride	Fisher	P217-500	for bleach soluton
Potassium phosphate dibasic	VWR	EM-PX1570-2	for NGM plates
Potassium phosphate monobasic	VWR	EM-PX1565-5	for M9
Revolve	ECHO	75990-514	equipped with an Olympus 4x Plan Fluorite NA 0.13 objective lens, standard Olympus FITC filter (ex 470/40; em 525/50; DM 560), and an iPad Pro for camera and to drive ECHO software
Sodium Azide	Sigma-Aldrich	71289-50G	for imaging
Sodium Chloride	EMD Millipore	SX0420-5	for NGM plates, M9
Sodium phosphate dibasic	VWR	71003-472	for M9
Tert-butyl hydroperoxide	Sigma-Aldrich	458139	for oxidative stress
Tetracycline hydrochloride	Sigma-Aldrich	T7660-5G	for RNAi
Tunicamycin	Sigma-Aldrich	T7765-50MG	for ER stress