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Table of Contents
Behavior reflects nervous system activity and is dependent on multiple factors including external stimuli, past experience, neuronal structure and changes in the internal milieu of the animal. Alterations at the cellular or functional level can profoundly alter basal and evoked activity. Therefore, behavioral assays offer the researcher simple, sensitive and powerful tools to interrogate neuronal function. The sensitivity of behavioral assays can also be their greatest weakness; the behavior of animals can be dramatically affected by small changes or variations in culture or assay conditions. In this short introduction, basic considerations for behavioral assays are very briefly examined. Both general considerations and admonitions relevant to C. elegans behavioral assays are included. Specific protocols for behavioral assays contributed by individual researchers follow this more general introduction.
New behavioral assays are constantly under development and old assays are often revised. Your suggestions, comments and new assays are welcome as this chapter and the appended protocols will be updated on a regular basis.
In the following section various parameters that contribute to behavioral response are described. Not every variable will affect every behavior. But, when troubleshooting or designing an assay, these variables should at least be considered.
Controls: Clearly, controls are absolutely essential for all behavioral experiments. Because responses can vary from day to day it is important for control assays with appropriate strains or animals to be run every day in parallel with experimental animals. Both negative and positive controls should be tested daily. One way is to alternate control and experimental trials to control for extraneous variables. For example, a wild type C. elegans strain (N2) and a defective strain (glr-1) should be tested for nose touch response in addition to any experimental strains under evaluation each day. These daily controls are reported in the literature as control results along with the corresponding experimental results.
The results of an experiment can be subtly or dramatically changed by the expectations or bias of the researcher observing the behavior. Experimenter bias is usually subconscious and exists despite the best intentions of researchers; determined vigilance as to appropriate controls is required. Clearly, this entails considerable effort in both experimental design and daily effort. Ideally, all strains or animals will be scored in the behavioral assay by an observer ignorant as to the genotype, treatment and/or expected outcome. Animals or strains should be renamed and assigned code numbers prior to the experiment by another researcher to avoid bias in scoring. In some cases, only a subset of the animals is scored under these most rigorous conditions. These results of these behavioral assays are subjected to statistical analysis independent of animals tested under less rigorous conditions. If the results from the two groups are statistically equivalent, then all of the data can be pooled for reporting in the literature at the discretion of the authors and with the concordance of the reviewers. Note that these two groups of data may need to be reported independently in “Supplementary Information” for each assay.
Control strains should be raised in parallel under identical conditions this can go beyond just raising animals on the same batch of NGM plates with the same bacteria in the same box in the same incubator. Consider the controls required for behavioral assessment of animals after laser surgery. “Mock ablated” animals of the same genotype and larval stage should also be anesthetized on azide pads for the same time interval as “ablated” animals. Both “mock ablated” and “ablated” animals should be coded before behavioral testing as individual animals, not as groups of animals. This coding avoids experimenter bias toward a specific group of animals. Additional control animals (i.e. negative and positive controls) are also evaluated in parallel with the coded ablated animals to confirm that assay conditions are appropriate. The animals should also be examined to confirm that the laser microsurgery was successful prior to “decoding” the results.
Selecting the correct control animals is crucial. In the evaluation of transgenic strains, the correct control is usually not the N2 wild type strain nor is it “non-transgenic” animals from the same plate. A more appropriate control is the scoring of several independent transgenic strains generated in parallel using 1) the same transgenic marker such as GFP or phenotypic rescue, 2) the same genetic background, 3) an “empty” version of the promoter used to drive the transgene of interest and 4) that are generated by microinjection by the same experimenter. This helps decrease the corresponding problems of genetic background, marker effects on behavior, promoter copy number, and relative gene dosage. “Non-transgenic” animals from the same plate are rarely used as controls because of the mosaic nature of animals carrying extrachromosomal arrays. Unless a cell-specific marker is used to mark a specific cell of interest, “non-transgenic” animals that do not express the transgenic marker may still maintain the array and express the transgene under analysis in other behaviorally relevant cells. Strains in different labs can suffer from genetic drift resulting in altered behavior. If a transgenic strain or mutant strain is obtained from another laboratory, it is a good idea to obtain the corresponding background strain from the same laboratory.
Behavioral assessments should be made by more than one observer, should be repeated several times, and should occur on more than one day. Using more than one observer helps to avoid subconscious bias. Repeating the behavioral assay multiple times and on more than one day ensures reproducibility and consistency. Alternatively, videotaping or other optical recording of behavioral experiments permits multiple researchers to score the behavior.
Cultivation: C. elegans are generally raised on OP50 bacterial lawns on NGM agar plates. Altering the size of the lawn, the type of bacteria, the agar, and the chemicals in the NGM can affect behavioral response. Unless carefully sealed, agar plates and their bacterial lawns lose water to the atmosphere over time. Very dry culture plates should be avoided. Contamination on the culture plates can also dramatically affect behavior. Animals from contaminated plates should not be used for behavioral assays; the short cut of letting contaminated animals feed on uncontaminated plates prior to the behavioral assay is not recommended. As a general rule, mold is less deleterious than yeast or bacterial contamination, but both should be avoided depending on the assay in question. A simple bleaching protocol is sufficient to generate uncontaminated animals whose offspring can be used for behavioral assays.
Crowding and feeding status: The crowding and feeding status of animals can dramatically or subtly affect behavior. For example, animals from crowded plates perform poorly in many avoidance assays. Animals clearly behave differently depending on their feeding status and history. Animals that have been starved during development or animals that have not gone through the dauer stage can have altered behavior compared to animals that have never been crowded. Feeding status also alters behavior. Animals on the bacterial lawn respond differently in behavioral assays compared to animals off the lawn. Along the same lines, increasing time off food alters behavior.
Ambient conditions: The conditions under which assays occur can also affect the results of behavioral assays. Possible environmental factors that can be controlled include room humidity, temperature, drafts, and vibration. The dryness of assay plates can shift response. Some behavioral assays are more sensitive to this than others. Osmolarity and spontaneous reversal rate assays are quite sensitive to plate dryness. Depending on the assay, this can be controlled by pouring plates fresh daily, weighing plates poured at equal volume, or running controls on the same plates.
Ambient conditions (time of day, temperature and humidity) should always be recorded, so their effect can be considered when evaluating data or troubleshooting.
Some behavioral assays are run with plate lids on; some with lids off.
The number of animals used in population assays can affect the outcome; this should also be standardized. For example, introducing too many animals into the circular osmotic barrier used in the classical osmotic avoidance assays can result in crowding and the escape of normally defective animals.
Chemicals: The results of behavioral assays can be dramatically different depending on reagent freshness, purity and supplier. Some chemicals must be made up fresh or diluted on the day of the assay. And, impurities in chemicals (e.g., switching from a 99% pure to a 95% pure chemical) or changing suppliers can alter behavioral results. These caveats apply both to the reagents used for cultivation of the animals and the reagents used in the actual behavioral assays.
Assessing the effect of drugs and other pharmacologically active compounds on behavior involves several more variables. In addition to the preceding admonitions, the stability of the drug, the effect of any solvent, batch-to-batch variation in activity, and the actual concentration of the drug in the assay or assay plate should be considered. Control animals and plates should be treated exactly in parallel. For example, if the drug is dissolved in ethanol and then diluted in water, then the control animals or plates should be treated with the same diluted ethanol solution (lacking the drug). Note that increased osmolarity in the behavioral assay due to inclusion of the drug may alter behavior in some assays.
Stage of life cycle: The sex, age and life cycle stage of the animal tested in an assay is an important factor for behavior. Males are generally used only for assessment of male-specific behaviors. All animals demonstrate lethargic behavior during each molt; staging of young adults is unambiguous so adult hermaphrodites are generally used in assays. The stage of the life cycle can alter behavior in hermaphrodites although many assays can be used on larval animals. Nervous system development is largely complete by the end of the L1 stage, but some developmental changes are clearly coordinated with the L4/adult molt.
Behavioral changes occur during adult stages as well. Some assays are indiscriminate as to the age of the adult, but other behavioral assays are more sensitive. The age of animals can be standardized by “hours after molt to adulthood”, by hours since an egg was laid at a specified temperature, or by the number of eggs in the uterus. The former is preferable in most cases. Animals can also become less responsive to some sensory stimuli after several days of adulthood. Some researchers run behavioral assays only on animals from synchronized plates where adult hermaphrodites laid eggs over a two hour period (and then the parents were removed) to keep animal age constant.
The treatment of animals during the assay can also alter behavior. Most assays require that the animals be treated as gently as possible. Transfer by mouth pipette seems to be the most gentle although this in not recommended as a general lab practice. Gently moving animals with a pick is sufficient for most assays. If an individual animal is damaged in transfer and unable to move properly, then it usually is excluded from the data set. But, the rules for exclusion and the number of animals excluded should be reported.
To avoid ambiguity and inconsistency, it is critical to precisely define the criteria for scoring behavior both in the laboratory and in publications. For example, initiating backward locomotion is generally called a reversal. But, some researchers define a minimum distance the animal has to move backward to score as a bona fide reversal; some do not. Some researchers include omega turns as reversals. And, some researchers count increased backward locomotion as a reversal or response; most do not. A precise definition of the behavior to be scored is critical for analysis.
It is useful to have a “scoring worksheet” to record results for each behavioral assay. In addition to the actual scoring data, the ambient conditions, date, time, genotype, cultivation conditions, researcher name, number of animals scored, and other variables should be recorded.
The number of animals assayed is determined by the assay. As a general rule, 4 large-scale population based assays or 30 individual assays with corresponding controls should be considered a bare minimum. To achieve statistical significance, many more trials may be required.
Data from experimental strains should always be compared to data from control strains analyzed in parallel.
Statistical analysis is critical for drawing conclusions from behavioral data. At a minimum, n and p values should be presented for critical data. Additionally, standard errors of the mean or standard errors should be presented within figures and tables to allow readers to assess significance.
Reproducibility can be difficult for behavioral assays. Control strains may vary in their response from day to day. Statistically, it may be tempting to normalize the data to the control results each day to decrease variation in the reported results. Although this has been done in some cases in the literature, it is clearly more persuasive to present data that has not been normalized. If normalized data is presented, then the non-normalized data should be presented in the corresponding on-line Supplementary Information.
It is wise to confirm the phenotype of strains, sequence plasmids and carefully check other reagents upon arrival. To avoid ambiguity, order crucial strains of C. elegans from the C. elegans Genetics Center (CGC) when possible. When they arrive, freeze multiple copies for long-term storage. Most C. elegans laboratories that specialize in behavior have their own “horror stories” in which genetic drift due to continuous passaging or mislabeling have wasted months of effort. It is a wise idea to thaw an aliquot of any crucial control strains at least once a year (e.g., the wild type N2 strain). Some labs thaw bi-monthly.
Background mutations: In addition to the mutant allele, C. elegans strains often contain other changes in their DNA. Some of these are spontaneous and some are background mutations arise from the original mutagenesis. And, because backcrossing is most effective for unlinked genes, the remaining background mutations may be tightly linked to the mutant allele and gene of interest. Historically, one strategy to confirm that the gene under examination causes the phenotype of interest is to create a trans-heterozygote (examining the behavior of allele A/allele B). If the alleles do not complement in the trans-heterozygous animal, then the mutant phenotypes are likely due to alterations in the function of gene X. Multiple, independently derived alleles of the gene of interest should also be examined if they are available.
Transgenic rescue of a mutant phenotype is the best indication that the behavioral phenotype observed is due to altered function of the gene of interest. For example, cDNA rescue using a heat shock promoter and a transformation marker (described below) may restore normal function to mutant animals. This phenotypic rescue by the cDNA should be confirmed by examining the behavior of transgenic mutant animals that carry the transformation marker and the “empty” heat shock promoter construct. Alternatively, a genomic rescue construct can be used to restore normal behavior, but a mutant version of the genomic rescue construct should not restore behavior.
Integrated transgenic arrays are caused by insertion of exogenous DNA in the chromosome. Therefore the behavioral phenotype of strain carrying an integrated array can arise either from the chromosomal DNA break or from the transgenes on the integrated array. If the behavior of animals carrying the extrachromosomal array is the same as the integrated array, then the behavioral changes are due to the transgene itself.
Transgenic strains: Some markers for generating transgenic C. elegans are preferred for behavioral analysis. GFP or dsRed reporter constructs can be used as transgenesis markers although these usually require the use of a fluorescent dissection microscope for strain maintenance. Historically, transgenic rescue of an unrelated mutant phenotype has been broadly used. Commonly used markers include pha-1, lin-15, dpy-20, or unc-119. Injection of a dominant rol-6 transgene is frequently used for transgenesis C. elegans but the rolling phenotype induced by this transgene interferes with most behavioral assays.
It is important to determine if the transgenic marker used will interfere with the assessment of the behavioral phenotype. GFP or dsRed expression can have deleterious effects. pha-1, lin-15, dpy-20, or unc-119 mutant alleles or even rescued transgenic animals are defective in some behavioral assays.
RNAi knockdown of gene expression is a powerful approach to assess the role of specific genes in neuronal function and behavior. Several approaches can be used with varying efficacy to knockdown gene expression with double stranded RNA in C. elegans. The magnitude of the RNAi knockdown varies dramatically from gene to gene; this is an important consideration in interpreting experiments.
Feeding C. elegans bacteria expressing dsRNA for the gene of interest works in some cases. The nervous system is relatively refractory to RNAi compared to some other tissues, but useful results may still be obtained. Mutant strains with increased sensitivity to RNAi are often used to increase efficacy of gene knockdown; rrf-3 and eri-1 are commonly used. Of course the behavior of control animals raised on control bacterial feeding strains should be examined. For example, as control for a nose touch response RNAi experiment, a bacterial strain expressing the dsRNA corresponding to osm-10 could be fed to rrf-3 and rrf-3; glr-1 animals. osm-10 is expressed in the ASH neurons that are critical for nose touch response, but loss of osm-10 function does not perturb nose touch response. rrf-3 animals raised on osm-10 RNAi bacteria should robustly respond to nose touch; rrf-3; glr-1 animals should not as glr-1 is required for nose touch response.
C. elegans promoters can also be used to express dsRNA in transgenic animals. Usually an inverted repeat of coding sequence from the targeted gene is used for expression of dsRNA, but co-expression of sense and antisense sequences from two separate plasmids can induce RNAi knockdown for some genes. Ubiquitous and inducible gene knockdown can be attempted using common heat shock vector promoters. Cell-specific knockdown is sometimes also possible as dsRNA and RNAi effects spread poorly (if at all) between neurons.
C. elegans behavioral assays encompass a wide range of behaviors and approaches. When combined with current molecular biological, electrophysiological and optical recording techniques, behavioral assays have been remarkably successful in assessing the contribution of genes and specific neurons to behavior. Generous researchers who specialize in each technique have contributed the following protocols. We hope that by providing a resource for the community, that behavioral studies in C. elegans will be made easier, more accessible and clearer for neuroscience researchers in general.
Contributed by Martin Chalfie, Columbia University, New York, NY, USA, March 2005.
C. elegans senses a variety of mechanical stimuli. These stimuli include gentle touch stimulus delivered to the body (Chalfie and Sulston, 1981; Sulston et al., 1975), harsh touch to the midbody (Way and Chalfie, 1989), harsh touch to the head or tail (Chalfie and Wolinsky, 1990), nose touch (Kaplan and Horvitz, 1993) and texture (Sawin et al., 2000). Gentle touch to the body is sensed by six sensory neurons (ALML/R, PLML/R, AVM, and PVM). An important feature of any assay for gentle touch mediated by these cells is that it not be confused with the response to other mechanical signals. Thus, care should be taken to touch the animals in a way that will not stimulate other mechanosensory neurons. In practice one should avoid too harsh a stimulus and not touch the animals too near the tip of the head or tail.
The initial and most generally used method to test for gentle touch sensitivity is to stroke the animals with an eyebrow hair that has been glued to the end of a toothpick (Chalfie and Sulston, 1981; Figure 1). Eyebrow hairs are used because they are usually not cut and are, thus, finely tapered. They also can be obtained with a minimum of discomfort. Some papers stated that the hairs are from eyelashes. I find, however, that removing eyelashes is a much more painful process. Various glues can be used to secure the hairs, but the hair should be placed so that its shaft extends straight from the end of the toothpick and does not curve away from the tip.
The hair is sterilized by dipping it into a 70% ethanol solution and drying by shaking (don't flame the hair). Sometimes bacteria accumulate on the hair; they can be removed by poking the hair into the agar on a spare plate.
Animals are touched by stroking the hair across the body just behind the pharynx (for the anterior touch response) or just before the anus (for the posterior touch response; see Figure 1). Actually touching the animals in any position along the touch receptor processes will generate a response. Touching the animals near the middle of the body (near the vulva in adult hermaphrodites), however, yields ambiguous results because both the anterior and posterior touch circuits can be activated. The animals should not be poked with the end of the hair, since this provides a stronger stimulus and can sometimes evoke a response, even in touch-insensitive animals. Similarly, animals should not be touched with the end of a platinum worm pick. Animals should not be touched at either the tip of the nose or the tip of the tail as even animals lacking the six touch sensing cells often respond.
The touch assay is really a differential assay, since animals that are dead or paralyzed will also not respond to the stroking. Routinely, animals are said to be touch-sensitive if they respond to stroking with the eyebrow hair by stopping movement toward the hair or move away from the hair (sometimes the touch stops moving animals without having them reverse their direction of movement). Touch-insensitive (Mec) animals fail to respond to the hair, but do respond to prodding with a worm pick. A partial response is one in which the animals move away from only some of the touches.
Touch mediated by the anterior touch cells can be assayed in severely uncoordinated animals by monitoring pharyngeal pumping (Chalfie et al., 1985; M. Chalfie, unpub. data). Touching animals in the anterior slow the rate of pharyngeal pumping. Another output of the touch circuitry is the defecation rhythm, which is reset by stroking the anterior of the animal with the eyebrow hair and is dependent on genes needed for the function of the touch receptor (mec-4, mec-5, and mec-9; Thomas, 1990).
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Figure 1. Using an eyebrow hair to test gentle touch sensitivity. (A) Positioning (top panel) and gluing (bottom panel) the eyebrow hair to the tooth pick. The thickened black line indicates the shaft of the hair; the gray area indicates the location of the glue. (B) Animals should be touched by stroking the hair across the body at the positions of the arrows. The six touch receptor neurons are indicated.
Wild-type animals will move (adults usually reverse direction) in response to their plate being tapped (Chalfie and Sulston, 1981). This stimulus often happens when plates are placed on the stage of a dissecting microscope. Touch-insensitive animals do not respond to this tapping. Although not as accurate a measure of touch sensitivity as touching with an eyebrow hair, this is a rapid assay that has been used to screen for touch-insensitive mutants (Chalfie and Au, 1989). Plates with F2 progeny of mutagenized parents were dropped from about 1-2 cm above the stage of the microscope and then examined for animals that did not move. Candidate touch mutants were then tested with the eyebrow hair and worm pick to determine if they were touch insensitive. Cathy Rankin and coworkers designed an electric tapper to deliver a taps at defined intervals to plates for their studies examining the touch circuitry and habituation of the touch response (Rankin, 1991; Wicks and Rankin, 1995).
The above assays treat touch sensitivity as an all-or-none phenomenon. Often, however, animals give a partial response to the touch stimulus. In addition, even touch-insensitive animals sometimes respond to the first touch. Two general methods provide a more quantitative measure of the touch response: counting the responses to multiple touches and touching with defined stimuli.
The first instance of using multiple touches with an eyebrow hair to measure the degree of insensitivity to touch was by Hobert et al. (1999). In this assay animals are touched ten times (alternating head and tail touches) and a score is giving for the number of positive responses. At least thirty animals are examined and mean percentage score is obtained. Since individual touches are not always the same, the values obtained in this assay are not truly quantitative. Nonetheless, by using multiple animals relatively subtle difference in touch sensitivity can be revealed (e.g., Hobert et al., 1999; Zhang and Chalfie, 2002). In addition, by separately scoring head and tail responses differences between the two can also be uncovered (Zhang et al., 2004). In using this method one should be careful to distinguish between animals that habituate more rapidly than wild type from those that respond less frequently to the touch stimulus. The animals with the former defect should show a response pattern in which the animals respond less frequently to successive touches; animals with the latter defect should show no pattern to the failures. The experimenter should also determine whether the animals are responding equally (in terms of number of responses) to touches in the head or tail.
von Frey hairs have been used clinically for over a century to test human touch sensitivity. They consist of a series of flexible fibers that are touched end-on to skin and bend once a specific force has been applied. The longer and thinner the fiber, the less force is required to bend it. These calibrated fibers are then used to determine the forces needed to provoke a touch sensation when placed on the skin. We have adapted this method to C. elegans by using fine monofilament nylon sutures (7-0 and smaller; Iris Chin, Miriam Goodman, and M. Chalfie, unpub. data). A short length of suture (Sharpoint, Surgical Specialties, Reading, PA) is attached perpendicularly to the end of a glass capillary (100μL volume) with epoxy and calibrated using an analytical balance. The force (in μNewtons) required for bending is calculated as the weight in (mg) found for the bent fiber X 9.8 m/s2. A limitation of this method is that fibers that deliver less than 10 μN of force are difficult to use because they bend with the air currents normally found in the lab.
Typically, we mount the capillary on a manual micromanipulator with the fiber perpendicular to the agar surface, position a worm underneath the fiber, and make contact by moving the z-axis of the micromanipulator. Animals are touched in the same locations as described above for testing body touch with eyebrow hairs, and the response is noted. Ninety percent of wild-type hermaphrodites respond to a 10 μN of force; virtually all animals respond to forces >100 μN. In contrast, hermaphrodites homozygous for mec-3 and mec-6 null mutations fail to respond to a 100 μN force. Approximately 10% of mec-4 (u253) animals respond to a 100 μN force (the difference may reflect some activity from the PVD cells, which are defective in mec-3 animals).
Extracted from the literature. Comments from researchers who use this assay would be appreciated.
Harsh touch to the body is measured by prodding animals with a platinum wire in the midsection of the body (Chalfie and Sulston, 1981; Way and Chalfie, 1989). Nonmoving gravid adults are prodded at or just posterior to the vulva. Animals respond by initiating locomotion, usually by backing up. Animals to be assayed should be grown in the continuous presence of food. Animals who are starved and animals that have passed through the dauer stage often fail to respond to harsh touch regardless of the functional status of the PVD sensory neurons.
Harsh touch is assessed in animals in which the function of the ALM and PLM neurons have been perturbed as these neurons mediate response to both gentle and harsh touch. Response to harsh touch is mediated by the PVD sensory neurons and perhaps the FLP sensory neurons. The phenotype is called “touch aberrant” (abbreviated Tab) to distinguish it from “Mechanosensory defective” (abbreviated Mec) animals. mec-3, unc-86 and lin-32 animals are Mec and Tab; mec-4, mec-2 and mec-6 animals are nonTab and Mec (respond to harsh, but not gentle touch).
Although the animals are reported to avoid the edge of an agar chunk, little is known about this behavior.
C. elegans dauers can occasionally be found flailing at the top of mold filament independent of plate orientation. (Croll and Matthews, 1977) An overtly similar behavior called nictation is reported to play a role in the dispersal of parasitic nematodes. An assay for this behavior has not been developed.
Contributed by Anne Hart, Massachusetts General Hospital and Harvard Medical School, Boston, MA USA, March 2005.
The nose of C. elegans moves in a rhythmic dorsal/ventral pattern as animals feed on a standard bacterial lawn. (C. elegans lay on their side on agar plates.) This rhythmic nose motion is called foraging in this context. Aberrant foraging rates and patterns of movement can be observed in some strains/animals. The rate of basal foraging is dependent on the RMD motorneurons and on glr-1 function (Hart et al., 1995).
Animals respond to touch to the side of the nose by rapidly moving their nose away from the stimulus; this is called head withdrawal. Response is again mediated by the RMD motorneurons and requires glr-1 function. (Hart et al., 1995) A hair-similar in size to the hair used in the nose touch assay, is placed on the agar plate so that the side of the animal's nose will just barely touch the hair at the maximal extent of the foraging motion. Animals respond by rapidly moving the nose away from the hair. Significant practice is required learn where to lay down the hair and to learn to discriminate between the rapid withdrawal motion versus normal foraging motion. Foraging and head withdrawal assays should be undertaken by observers blind as to genotype or treatment.
Thin bacterial lawns for these assays resemble those used in nose touch assays. They can be prepared by releasing a single drop of an overnight OOP50 culture over a standard NGM plate. Allow the liquid to soak into the plate. To prevent bacterial growth and too much thickening of the bacterial lawn, the plate should be used within a few hours or should be sealed with Parafilm and stored at 4 degrees for weeks. Allow stored plates to return to room temperature before use.
Contributed by Catharine Rankin, University of British Columbia, Vancouver, BC, Canada, March 2005.
The tap-withdrawal protocol measures the responses of a worm to a single tap or a series of successive taps (trains of taps), given to the side of a 4 cm Petri plate filled with 10 ml of NGM agar. The taps/trains cause the worm's environment to vibrate briefly, and this causes the adult worm to swim backwards for some distance (termed a reversal). This technique produces a quantifiable measure of the magnitude of this reversal response. For this assay worms are placed on the center of a Petri plate and are videotaped through the lens of a dissecting microscope. The mechanical tapper is arranged to hit the center of one side of the plate. The tapper is composed of an electromagnetic relay run by a stimulator. (Rankin et al., 1990) The tapper delvers a force of approximately 1 to 2 Newtons per tap which are transmitted through the dish and the agar to stimulate the worm. These experiments are usually run with the lids off of the plates while taps are being delivered. The effects of drugs on the tap response can be assessed by adding the drug plus vehicle to the agar and comparing the results to a group that was tested with just the vehicle for the drug. The length of the reversal to tap can be affected by several variables: 1) The intensity of the tap; Chiba and Rankin (Chiba and Rankin, 1990) showed that the tap response is graded and that stronger stimuli produce larger reversals. 2) The age of the worm; Chiba and Rankin (Chiba and Rankin, 1990), showed that as worms got larger so too did reversals. 3) The agar in the plates; as agar plates age they dry out and lead to differences in the ease with which worms can move. This can be controlled for by only using plates within a certain age range, or by weighing plates prior to use. 4) The reversal response can also be altered by the ambient temperature and humidity in the room where the testing is done. This is probably the result of changes in the agar and ease of movement; it is an important variable to control. At the very least, experimental and control animals must always be run together, on the same day, at about the same time under all of the same conditions. By taking note of temperature and humidity conditions for each animal it will be possible to determine whether variations in either variable affected the outcome of the experiment.
Prior to undertaking tap reflex experiments, it would be advisable to also read the protocols in the Learning, Adaptation and Habituation section on the tap reflex.
Once the experiment has been run and videotaped, then the behavior must be scored. Using stop-frame video analysis, each worm's behavior is scored by tracing its body from the videosimage onto a transparent acetate sheet on the video monitor, going frame by frame through the tape. The adult worm will usually (90% to 95% of the time) respond to the tap with a reversal response, with the size and number of the responses decreasing as stimuli are repeated through the habituation run. In a typical reversal response, a stopped or forward-moving worm moves backwards for a distance (usually less than 1 or 2 worm lengths) and then either remains still or re-initiates forward movement in a new direction. To score the response, a person notes the pre-tap position of the worm, and then traces the total distance the worm reversed (i.e. track length) onto the acetate sheet. Because the head of the worm often makes small side-to12 side foraging movements, reversals are most accurately scored by tracing the path of the tail. In order to have consensus a scoring manual should be developed with rules for scoring variations from this behavior (i.e. the reversal must occur within 1 second of the tap, a pause is scored as 0 backward movement, if the worm is already swimming backwards when the tap is delivered this is a “missing data point” etc). In some cases, a worm will respond with acceleration forward rather than a reversal. In a typical acceleration response, a stopped or forward-moving worm either initiates forward movement or increases its speed, respectively. Generally, in experiments with adult wild-type worms, these acceleration responses are simply scored as missing data points (an adult worm will accelerate to tap only about 5% of the time). Under other circumstances or protocols, such as with larval, mutant or laser-ablated animals, the acceleration response occurs more often and is therefore scored. If a worm accelerates after the tap, the acceleration is scored by measuring the distance the worm moves (i.e. track length) during the 1-sec interval before the tap and subtracting it from the distance the worm moves during the 1-sec interval after the tap. If the measurement for the second after the tap is ≥1.75 X the measurement for the second before the tap then the response is considered an acceleration. It is important to note that the reversal response and the acceleration response are two qualitatively different outcomes, which cannot easily be compared. These “scored” tracings are then scanned into a computer and the length of each is measured using NIH image. The values from this program, representing the reversal/acceleration magnitude, can then be directly transported into a statistical package for data analysis. The frequency of reversals and accelerations is also an important measure of the behavior of the worm, and are scored simply by counting the number of instances of either response.
Contributed by Anne Hart, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA, March 2005.
C. elegans responds to gentle touch to the nose by initiating backward locomotion. (Kaplan and Horvitz, 1993) This response is the “nose touch response” or Not assay. Response to nose touch is mediated by the ASH, OLQ and FLP sensory neurons. The two polymodal ASH sensory neurons mediated 60% of the wild type response to nose touch. Their ciliated, sensory endings are located at the tip of the nose where they detect light touch to the nose, high osmolarity (Hart et al., 1999) and volatile repellent chemicals (Troemel, 1999).
Nose touch is assayed by laying a hair on the surface of the plate in front of the animal. As an animal moves forward, it contacts the hair with the tip of the nose perpendicular to the direction of movement. Some practice is required to anticipate where the hair must be laid down for an animal to run into at 90 degrees. Normal animals immediately initiate backward locomotion. Defective animals either hurdle over the hair or slide their nose along the hair. An individual animal is tested no more than 10 times in a row to avoid inducing habituation. You can drop the assay plate to induce lethargic animals to move or reverse direction, but you cannot touch the animal to facilitate the assay.
Nose touch can be assayed either on very thin bacterial lawns or off food. Response on food is more robust. Animals that are moved to thin lawns recover within 10 minutes and respond in roughly 90% of trials. Response by animals who have wandered off the bacterial lawn is slightly diminished; animals respond in roughly 80% of trials. Response is dramatically reduced in animals who are physically moved by the investigator to a plate lacking food; animals respond in roughly 30% of trials (Chao et al., 2004).
The hair should be wiped off every few trials to eliminate bacteria and water that may adhere. Normal animals will ignore a wet or slimy hair. A folded “Kimwipe” can be used or clean bare fingers. In the latter case, avoid using lotions, creams or other surfactants. Don't flame the hair. Individual hairs can be reused used for weeks.
The hair used in the Not assay must be of an appropriate size. Too thick will result in all animals responding; too thin will cause even animals of mutant genotype to respond. Hair thickness has definite variation between individuals; arm hair from a female researcher or an eye lash from a male researcher would be a good place to start. The root of the hair is taped to the small end of a glass Pasteur pipette or to a thin wooden stick. Many hospitals stock cotton swabs on thin wooden sticks that are ideal. Slightly thicker hairs are required for nose touch assays on food. Under the microscope it is clear that each hair tapers to a very fine tip. Generally the very end of a hair is too thin and the animal must impact a thicker (and less distal) part of the hair in the nose touch trial.
Thin bacterial lawns for nose touch assays can be prepared by releasing a single drop of an overnight OOP50 culture over a standard NGM plate. Allow the liquid to soak into the plate. To prevent bacterial growth and too much thickening of the bacterial lawn, the plate should be used within a few hours or should be sealed with Parafilm and stored at 4 degrees for weeks. Allow stored plates to return to room temperature before use. If the bacterial lawn is too thick, then the animals may crawl through the food over or under the hair and avoid the nose touch stimulus.
Each hair is tested by comparing the response of N2 and glr-1 animals. (Hart et al., 1995). N2 animals should respond robustly; glr-1 animals should fail in 90% of trials. (Note that eat-4 animals are so defective in their response to nose touch that they are not useful as a control. Lee et al., 1999; Raizen and Avery, 1994) At a bare minimum at least 5 animals should be tested at 10 trials each. Response is expressed as % response.
C. elegans adapt to nose touch after a rapid series of repeated trials. The nose touch trials must be given in fairly rapid succession to adapt the animals' response-without accidentally hitting the animal over the head with the hair. To measure habituation, compare the % response in the first 10 trials to the % response in the trials 40 through 50.
See also: “Chemotaxis and chemoaversion, quadrant assay, version 2” and “Drop assay”.
Contributed by Anne Hart, Massachusetts General Hospital and Harvard Medical School, Boston, MA, USA, March 2005.
Avoidance of high osmolarity (concentrated solutions) by C. elegans was first described by Culotti and Russell (1978). and revisited by various researchers over subsequent years. The original assay, described in an updated format below, tests the rate at which C. elegans will cross an osmotic barrier on the surface of the agar. The barrier is usually circular in shape-referred to as a ring. Animals are placed in the center of the ring (an area of low osmolarity) so they are surrounded by the high osmolarity ring which will they avoid as a noxious stimulus. Animals escape the ring by crossing the high osmolarity barrier. Outside the ring is normal NGM agar, a region of low osmolarity. In practice, the rate at which animals escape is dependent on several parameters. Particularly critical are plate dryness, the number of animals in the ring versus the size of the ring, and the rate of diffusion of the osmotic barrier. The assay is dependent on the number of times that animals encounter the osmotic barrier; special consideration must be taken when assessing osmotic avoidance for uncoordinated or lethargic animals. Hermaphrodites are assayed.
Control strains must be used each day. N2 animals are generally used as positive controls as they robustly avoid the ring barrier. Generally, che-3 or osm-10 animals are used for negative controls. Uncontaminated, uncrowded cultures must be used. Animals that have been starved during their life, those close to running out of food, animals that have passed through dauer or animals from culture plates that are very dry should not be assayed.
The number of animals to be tested determines the size of the ring. A 5/8th inch diameter ring holds 30 to 100 animals for population assays; a 3/8thinch diameter ring holds 1 to 5 animals. Too many animals for a specific ring size leads to aberrantly high escape rates; too large a ring for the number of animals leads to aberrantly low escape rates.
The dryness of the assay plates is a critical consideration and can be the critical variable to control. The objective is to obtain a large number of plates of uniform dryness. Plate dryness is assessed by weighing the plates. Once you establish the correct plate weight in your lab, then (if the plate volume poured remains unchanged) finding good plates for the assay is relatively straight-forward.
NGM agar plates are allowed to sit out on the bench overnight after a large batch is poured. It is important that an equal volume of media is poured into each plate; an automated media dispenser is ideal. The next day the plates are inverted and then placed in a plastic box. The length of time required for plates to dry depends on atmospheric humidity in the laboratory and how the box is covered. In winter in Boston it can take only a few days for plates to dry in a box with a closed (but not tightly sealed) lid. In summer, it can take weeks. To identify plates of uniform dryness, plates are removed from the box and individually weighed. If a constant amount of plate media is dispensed during plate pouring, then the weight of each plate is directly related to how dry the plate is (how much evaporation has taken place). If the amount of media is a constant over weeks or months, then plates can be often selected based on weight alone-once the correct weight is determined in behavioral assays. Be sure to remove any condensation from the plate lid before weighing.
The only definitive test to determine the correct weight (and dryness) is to run the osmotic avoidance assay with the control strains. A good rule of thumb to start with here is that animals crawling on the agar surface of an appropriately dried plate should have a visible, but small meniscus.
Alternative approaches: The drying process can be hastened by covering the box with a large Kimwipe (as opposed to a lid) although uniformity of plate dryness may decrease. Or, plates can be dried in a sterile hood for a few hours to hasten the drying process-but the plates must be allowed to equilibrate with the lids closed for several hours afterward.
Prepare 8M glycerol in water. This corresponds to a 60% solution of pure glycerol in water; “pure glycerol” from Sigma is a 13M solution. Add a tiny bit of bromophenol blue to color the 8M glycerol and the resulting rings blue. 8M glycerol aliquots can be stored in the −20°C freezer for years; aliquots are stable at room temperature for days.
Depending on the number of animals to be assayed, either ultra fine point Sharpie lids (∼3/8th inch) or metal test tube caps (∼5/8th inch) are obtained. C. elegans S Basal is required for population assays.
Osmotic barrier rings must be printed onto the agar of the assay plate so they will be ready as soon as the animals finish rinsing in the S Basal (see next paragraph). Usually four assay plates can be run in one batch. Obtain 4 additional 6cm Petri dish lids and four metal test tube caps. Pipette about 25 to 30μl of 8M glycerol into the surface of each inverted lid. Place the open end of the cap down onto the inverted lid bisecting the drop of glycerol. Spin the cap against the surface of the lid to spread the glycerol around the circumference of the cap. Lift the cap. If there is a bubble of glycerol closing the cap, then pop it. Place the lid with glycerol very gently onto the agar of an assay plate. Try to avoid marring the surface of the agar. Let the glycerol soak into the agar; this usually takes between 2 and 5 minutes. Lift the cap off. Check to make sure the annular osmotic barrier is uniform. Always run the control animals in duplicate (e.g., N2 and osm-10) first to confirm that the assay plates are working. It is a good idea to run control interspersed with experimental animals during the assays.
Rinse animals off culture plates and into a 1.5ml microfuge tube with 1ml of S Basal using a Pasteur pipette. No need to close the tube lids. Let animals settle by gravity for 5 minutes, remove excess S Basal and add another 1ml. After 5 to 10 minutes, move animals in a minimal amount of liquid to the center of the osmotic ring. Remove excess liquid by very gently blotting with a rolled up Kimwipe. Once the excess liquid is removed and animals start crawling, the timing begins for that assay plate. Record the time the assay starts. Replace plate lid and avoid disturbing animals by dropping or tapping the plate during the assay. Repeat till all four assays are underway. When the assays are done, wipe off the caps and lids so they can be reused.
Animals should move actively within the ring during the assay period. N2 animals should avoid entering the osmotic barrier for at least 10 minutes; 50 to 80% of osm-10 or other control animals should either die in the osmotic barrier or cross it and escape the ring. Those who die in the ring are counted as escapers. If too many N2 animals escape, then the plates are likely too wet. If too few osm-10 animals escape, then the plates are likely too dry.
The number of animals that have escaped at 10 minutes is recorded along with the number of animals that remain in the ring. The % escaping animals is calculated. At least 5 assays per strain or genotype are suggested; these should be determined on at least two different days.
The rate at which animals encounter the osmotic barrier is important in determining their response. If limited animals are available-or individual animals need to be tested, then smaller rings should be used.
Rings can be generated using ultra fine point Sharpie lids and 20 μl of 8M glycerol as described above. Use the open end of the Sharpie lid. Animals to be assayed are gently (and rapidly) picked from the bacterial lawn to an empty plate (to eliminate food) to the assay plate. Up to 5 animals can be used in a ring of this size. At least 7 rings with at least 4 animals for each genotype is a good staring point. As described above, control strains should be run before experiments begin.
If the osmotic response of individual animals must be assessed (e.g., laser ablation experiments), then animals should be assayed at least 4 times. Once an animal crosses the osmotic barrier, then the animal is immediately returned to the bacterial lawn for at least 15 minutes.
If small numbers of animals are used, then it is convenient, and perhaps more accurate, to record the time the animals cross the barrier, not the % escaping. While the assay is running, plates are checked every minute and the number of animals who have escaped is recorded. The average time to escape is then calculated.
Contributed by Gert Jansen, Erasmus Medical Center, Rotterdam, The Netherlands, March, 2005.
We discriminate three different responses using the quadrant plate chemotaxis assay for salts as developed by Steven Wicks (Wicks et al., 2000):
Chemo-attraction to 0.1–100 mM NaCl or other salts.
Chemo-aversion of 1 M NaCl.
Gustatory plasticity, i.e. the response of C. elegans to 25 mM NaCl after 15 min pre-exposure to 100 mM NaCl (Jansen et al., 2002).
Briefly, pairs of opposite quadrants of four-quadrant Petri plates (Falcon X plate, Becton Dickinson Labware, USA) are filled with 16 ml buffered agar (2% agar, 5 mM K2HPO4/KH2PO4 MgSO4) either containing a dissolved attractant or no attractant. The plates are left open to solidify and dry for 60 min. Then the plates are closed and stored for use on the same day. Just before the assays are performed, i.e. just before washing off worms, adjacent quadrants of the assay plates are connected with a thin layer of molten agar (without attractant). Assays are always performed in duplicate, try to control for environmental influences by placing the plates on the bench in different orientations. In addition, assays should always be repeated on separate days.
Animals are prepared by bleaching gravid adults and washing the eggs with M9 and growing the animals for approximately 72 hrs at 25°C. During a 15 min wash, the well-fed, young adult nematodes are washed three times with CTX buffer (5 mM KH2PO4/K2HPO4 pH 6, 1 mM CaCl2 and 1 mM MgSO4) and 100–200 worms are placed at the intersection of the four quadrants in 5 ul CTX buffer. Most of the buffer can be removed with a pipette, or a tissue. The distribution of the worms over the four quadrants is determined after 10 min. A chemotaxis index (CI=(A-C)/A+C, where A is the number of worms over quadrants 1 and 3, C is the number of worms over quadrants 2 and 4) was calculated at each time point. As attractants we use 100 microM, 1, 10, and 100 mM NaAc, NaCl, NH4 Ac and NH4Cl. As repellent we use 1 M NaCl.
To test the plasticity of the response of C. elegans to salts, animals are pre-exposed to the compound tested during the washing steps (Jansen et al., 2002). We found that the optimal concentration for pre-exposure was 100 mM NaCl. The 100 mM salt was simply added to the CTX washing buffer. Optimal pre-exposure time was 15 min, longer pre-exposure was toxic, especially for some mutants, while shorter pre-exposure had less effect. Subsequently, the animals are tested in a standard water-soluble compound chemotaxis assay for their attraction to the salt used during pre-exposure. We found that chemotaxis to 25 mM NaCl worked best.
Contributed by Stephen Wicks, Boston College, Chestnut Hill, MA, USA, March, 2005.
This assay was developed to assay attraction from or avoidance of soluble compounds, or to compare concentrations of a single compound (Wicks et al., 2000). The assay is robust enough to characterize known mutants, or to screen for new mutants which fail to approach or avoid soluble compounds such as salts, sugars, amino acids and other potentially biologically significant compounds. This assay is effective (chemotaxis index of 0.99 from some compounds), probably because the gradients that develop between adjacent quadrants are very high, and thus demonstrating a preference for one compound over another, or a discrimination of two concentrations, is an easy task for the worm. The assay relies on the use of segmented plastic Petri plates. A variety of manufacturers sell appropriate plates. Ideally, the plates should have no (or at least only a very small) raised arm - a recent innovation that facilitates robotic handling of plates-restricted to the periphery. A plate will have 4 equally sized quadrants. Each quadrant should have a volume capacity of 10-12 mls. and is filled with agar containing the dissolved compound of interest solublized in molten agar. In practice we generally fill opposite pairs of quadrants with the same compound-containing agar. Thus, each quadrant is adjacent to two quadrants containing the other compound or concentration of interest. Each quadrant is overfilled such that surface tension provided by the sterilized plastic is just sufficient to keep the agar from spilling, and the agar is allowed to dry. It is important to keep the volume in each quadrant the same as in all other quadrants. Once the agar has solidified, the tops of each plastic wall that separates the four quadrants is coated with agar containing no attractant to provide a smooth, continuous surface over which animals move. It is also allowed to dry and cool. A population of animals is then spotted at the intersection of the quadrants in a small (4-10 microliter) volume of low salt buffer. The population will segregate over the quadrants that contain the preferred taste. Animals present in each quadrant are counted at several time points to assess adaptation and changing taste preferences.
Procedure 1. Quadrant plate preparation
Plates are available from Falcon. Cases of 500 plates (Falcon #351009 X PLATE Petri dish with lid) each of which is 100 mm (diameter) by 15 mm (deep) are typically ordered.
Taxis Agar is composed of 2% agar (MOPS buffered). We have found that for most salts, 10% of 0.1 M MOPS (we pH the MOPS to 7.2 with 25% ammonium) is sufficient to control the pH, although this does not appear to be true for amino acids or other free bases. The desired attractants are dissolved in solution prior to autoclaving. After autoclaving, the solutions are brought to 1 mM CaCl2 and 1 mM MgSO4. Don't add TWEEN or any other detergents; these disrupt the surface tension of the agar too much. As an alterative we also have been using phosphate buffered agar.
After the agar cools to about the temperature that you could comfortably hand pour (55-60 degrees C), an autopipetter is used to distribute 12.6 ml of agar (the exact volume will depend on the plate used) into each quadrant of the plate. Typically, I place similar attractants into opposite quadrants of the plate. The surface on which the agar is distributed should be level, as this volume will bring the agar level above the plastic spacers between quadrants; surface tension keeps the agar from spilling from one quadrant into adjacent ones. (Also, don't shake or bump the bench or table!).
Let the agar cool and harden. Then, a few minutes prior to testing a population, draw about 0.5 ml of buffered agar (with CaCl2 and MgSO4) into a glass pipette and gently but briskly fill the “valleys” between adjacent agar-filled quadrants on each plate. I have tried using a lower agar concentration for this step to avoid building any significant “topography” for the worms and that works well also, but, with care this is not necessary.
A population of animals is washed off of plates and rinsed once with a low-salt CTX-buffer solution (1mM CaCl2, 1mM MgSO4, 5mM potassium phosphate, pH=6.6), placed over the centre of the plate in about 10ul of CTX Buffer and allowed to partition over time. The worms will often show a preference almost immediately (within 30 s) even prior to the evaporation of the buffer.
The preference for one kind of moiety (or concentration of moiety) over the other is generally expressed as fraction of worms over that moiety divided by total number of worms per plate (although, obviously it could also be expressed as [# worms over attractant - # worms over control] / Total # worms, as with the odorant assays from the Bargmann lab). This is calculated at various time points (generally, I use 10 min 20 min, 30 min, 45 min, and 60 min-sometimes also 5 min if the buffer has evaporated).
Genetic screens for animals defective for avoidance of aversive compounds were carried out on assay plates with 103-104 worms per plate. The assay plates were prepared by pouring 5 ml of chemotaxis agar (1.5% agar, 75 mM NH4Cl, 10 mM MOPS, pH 7.2 with NH4OH, and 0.25% Tween 20) into a 9 cm Petri dish. A thin line (25 μl) of an aversive compound (e.g., copper as 150 mM CuSO4, 2 % SDS, or high osmotic pressure as 4 M fructose; Ward, 1973) was poured across the midline of the Petri dish and another 25 μl traced in a semicircle around one half of the dish (Figure 2). Young adult worms were washed twice with S-basal buffer to remove bacteria. Immediately after the compound was absorbed into the chemotaxis agar, worms were placed on one side of the assay plate in a small volume of buffer. Excess buffer was carefully removed with tissue paper. A volatile attractant (2 μl of isoamyl alcohol 1:10 dilution in ethanol) was placed across the assay plate 1 cm from the opposite edge with the aversive semicircle. Normal, healthy worms approached the attractant, but stopped abruptly at the boundary formed by the repulsive compound. Individuals that passed through the aversive barrier were isolated and allowed to produce self progeny. To quantify the behavior of a given strain, an aversion index (A.I.) was calculated by dividing the number of worms that, one hour after placing the worms on the plate, crawled through the aversive compound by the total number of worms on the plate (approximately 250–500). This index includes all animals that attempt to crawl up the side of the plate.
Contributed by Paolo Bazzicalupo and Massimo Hilliard, Istituto Internazionale di Genetica e Biofisica, Naples, Italy, March, 2005.
These two assays were developed to study the avoidance response of individual worms to water soluble repellents. They can efficiently be used to score populations as well. Reference to and practical use of these tests can be found in Hilliard et al. (2004) and in Hilliard et al. (2002). These papers also report some controls strains (especially mutants tested) that can help researchers understand and define the usefulness of these assays. Main use of the assays: determine the phenotype of single animals to characterize existing mutants, isolate new mutants, characterize laser-operated animals, study spatial resolution of sensory organ, study water-soluble repellent adaptation. Possible other applications include response to volatile repellent.
In this assay, a small drop of repellent is delivered near the tail of an animal while it moves forward. Once in contact with the tail, the drop surrounds the entire animal by capillary action reaching the anterior sensory organs. If the substance is sensed as repellent the animal stops moving forward and starts moving backward. If the substance is not sensed as a repellent the animal will continue its forward movement.
Animals: Well fed young adult hermaphrodites. With some experience and appropriate controls, younger larvae and males can also be tested. Animals are washed off the plates were they have grown, put on unseeded NGM plates and allowed to rest at room temp for 10-15 minutes before testing. We always challenge the worms to be tested with a few drops of buffer alone (and they should not respond). Occasionally populations of worms appear hyper-reactive. In these cases the test should be postponed to avoid false positives.
Plates: The tests are conducted on unseeded NGM plates. In some cases a more controlled composition of the medium may be desirable (e.g., without Ca and/or Mg ions, lower salt concentration etc.). NGM can certainly be substituted by other media but appropriate controls are necessary. It is possible to use M13 plates (plates in which the agar is dissolved in M13 buffer, see below, in which repellents are usually dissolved) to reduce possible interference due to the change in composition between buffer and medium.
Best results are obtained with fresh, non wet plates such that the tracks of the moving worms can be detected but will disappear in few minutes. Assays are done at room temperature as long as it does not exceed 24-25 °C. Plates should be allowed to equilibrate to the room temp.
Repellents: All repellents are dissolved in: 30 mM Tris-HCl pH 7.0, 100 mM NaCl, 10 mM KCl (M13 buffer). We have used quinine, other alkaloids, Denatonium etc. at 10mM final concentration. Cu++ and Zn++ were also used at 10mM. For Low pH avoidance we used 0.1 or 0.2 M Acetic acid. For osmotic avoidance we used Glycerol at 1M. It is advisable to determine dose response curves if new repellents are tested. With appropriate controls buffer composition, repellent concentration and the composition of the medium can be modified.
Delivery of the stimulus and scoring: Drops are delivered using 10μl glass capillaries (Blaubrand intraMARK or equivalent) pulled by hand on a flame to reduce the diameter of the tip. The capillary is mounted in a holder with rubber tubing and operated by mouth pipetting. In case of toxic compounds, all the mouth operations can be performed using a 1-5 ml syringe. We calculated the average drop size to be about 5 nanoliters. The size of the drop has to be large enough such that the liquid surrounds the animal and can reach the anterior sensory organs but not so large that the animal looses adherence to the substrate and swims in it. The animal usually starts a backward motion within 1 second of the delivery of the drop. The response to each drop is scored as positive if the animal reacts within 4 seconds. The avoidance index (a.i.) is the number of positive responses divided by the total number of trials. The a.i. will range between value of 1 and 0 where higher values indicate stronger repellent response, lower values indicate weak response. As described below the a.i. can be used to define the response of single animals as well as of populations.
In the single animal assay (for example to determine the phenotype and thus the genotype of one animal derived from a cross-or to determine the effect of cell ablation on one animal) repeated drops are delivered to the same animal. The single animal is transferred from the growing plate to the test plate with a platinum wire or an eyelash and allowed to rest and adapt to the new plate for 5-10 minutes before starting the assay. An ISI (Inter Stimuli Interval) of at least two minutes is used between successive drops to the same animal. Each animal is tested with no more than 20 successive drops; the animal is then transferred onto a new seeded plate and allowed to recover for one or two hours before starting a new set of experiments. No more than 3 sets of 20 drops each were conducted per day on each animal. In most cases the phenotype is accurately determined with just 10 drops. The avoidance index (a.i.) will be the number of positive responses divided by the total number of trials.
Two methods can be used to define the responses of populations of worms:
10 to 50 (usually 30) well-fed adult animals of the same population are placed each on a small NGM unseeded agar plate and allowed to rest for 5-10 minutes. Each animal is then tested with several successive drops (5-30) as described in the single animal assay. The results of all animals belonging to the same population are then combined. This is our preferred population method since it provides data on the individual variation within the population and the results are amenable to more accurate statistic analysis.
A population of well-fed adult animals is washed with buffer and placed on a large unseeded plate. The animals are allowed to rest for 15 minutes and then at least two series of 50 animals of the same population are challenged each with a single drop of the chosen substance. Each response is recorded as either positive or negative. The a.i. is the number of positive avoidance responses divided by the total of trials (drops delivered). This is obviously faster than the other method and often it is more than satisfactory.
The dry drop test resembles the drop test except that the animal encounters the repellent substance after the drop has been absorbed into the agar. In this case the drop is delivered about 0.5-1 mm anterior to the animal (head stimulus). This assay allows spatial resolution between anterior and posterior sensory organs since it prevents the capillary action (present in the drop test) that brings the substance and thus the stimulus on the entire body surface. In the drop test both amphid and phasmid neurons are stimulated, in the dry drop test only amphid neurons are stimulated. Thus a difference in response between drop test and dry drop test may indicate a role of the phasmid neurons in determining the response.
It is also possible to separate stimulation of anterior and posterior sensory neurons by confronting the animals with two dry drops. The first just anterior to the animal nose; the second just posterior to the animal tail (0.3-0.5 mm). With this strategy, the animal is forced to encounter the repellent first anteriorly with the head (e.g., amphids) and then, during the backward movement, with the tail (e.g., phasmids). In this case, the duration of the backward response is the measure of the effect of the posterior sensory organ. For instance with SDS as a repellent, the backward response is reduced from an average of 3 seconds if only head stimulus is applied, to 2 seconds if the repellent is applied also on the tail as described in Hilliard et al. (2002).
Other variations: Another interesting use of the dry drop test was done in the Maricq lab (Mellem et al., 2002). In their analysis, instead of simply scoring the response as either negative or positive, they measured the delay of the response from the time when the animal encountered the repellent on the agar to the time when the backward response begins.
The drop test can be used to investigate the adaptation of animals to water soluble repellent substances. Two adaptation paradigms have been used:
Worms are exposed to successive stimuli (drops) and the response to each drop is recorded as either positive or negative. If the ISI (inter-stimulus interval) is short enough (10 to 30 seconds) the avoidance index (determined by testing 10 or more worms) undergoes a decrease on the second stimulus and will undergo further reductions on successive stimuli. The adaptation is reversible and after an adequate period of rest the a.i. will go back to normal. The main parameter that can be varied in the assay is the length of the ISI. If the ISI is long enough no adaptation will be detected.
Worms are exposed to the repellent continuously for a relatively long period of time and then the a
