Quantification of Drosophila Grooming Behavior

Summary

This protocol describes a scalable individual grooming assay technique in Drosophila that yields robust, quantitative data to measure grooming behavior. The method is based on comparing the difference in dye accumulation on the bodies of un-groomed versus groomed animals over a set period of time.

Abstract

Drosophila grooming behavior is a complex multi-step locomotor program that requires coordinated movement of both forelegs and hindlegs. Here we present a grooming assay protocol and novel chamber design that is cost-efficient and scalable for either small or large-scale studies of Drosophila grooming. Flies are dusted all over their body with Brilliant Yellow dye and given time to remove the dye from their bodies within the chamber. Flies are then deposited in a set volume of ethanol to solubilize the dye. The relative spectral absorbance of dye-ethanol samples for groomed versus ungroomed animals are measured and recorded. The protocol yields quantitative data of dye accumulation for individual flies, which can be easily averaged and compared across samples. This allows experimental designs to easily evaluate grooming ability for mutant animal studies or circuit manipulations. This efficient procedure is both versatile and scalable. We show work-flow of the protocol and comparative data between WT animals and mutant animals for the Drosophila type I Dopamine Receptor (DopR).

Introduction

Grooming in Drosophila melanogaster (D. melanogaster ) is a robust innate behavior that involves the coordination of multiple independent motor programs¹. Fruit flies clean their bodies of dust, microbes, and other pathogens which could inhibit normal physiological function such as vision and flight, or lead to significant immune challenges. In sensing and responding to both mechanical² and immune activation³, flies repetitively rub their legs together or on a targeted body region until it is sufficiently clean and grooming progresses to another part of the body. Flies perform grooming movements in distinct bouts that largely occur in stereotyped patterns^1,4. A behavioral hierarchy becomes apparent as grooming signals are prioritized. Circuits and patterns of activity have been identified in support of a model that grooming programs at the top of the hierarchy occur first and suppress parallel signals from areas of the body that are groomed subsequently⁵. Highest priority is given to the head, then the abdomen, wings, and finally the thorax⁵.

The grooming program in D. melanogaster is an ideal system for studying neural circuits, modulatory molecular signals, and neurotransmitters. For instance, compromise of neurofibromin function⁶, loss of Drosophila fragile X mental retardation protein (dfmr1)⁷, and exposure to bisphenol A (BPA)⁸ all cause excessive grooming and other behaviors that are analogous to discrete human symptoms of neurofibromatosis, fragile X syndrome, and aspects of autism spectrum disorders and Attention-deficit Hyperactivity Disorder (ADHD), respectively. Grooming behavior can also be habituated differentially across mutant strains², lending this motor program to studies of behavioral plasticity. The breadth of neurological phenomena that can be modeled by Drosophila demands a novel comparative approach to measure the ability of flies to groom themselves.

The combined action of vesicular monoamine transporters and the relative abundance of dopamine and other biogenic amines in the body have been shown to mediate fruit fly grooming behavior ^9,10. Octopamine and dopamine stimulate comparable hindleg grooming activity in decapitated flies, while tyramine, the precursor of octopamine, also triggers grooming to a lesser extent⁷. Four dopamine receptors have been identified in D. melanogaster^11,12,13,14. By using the grooming assay method described in this protocol, we determined a role for the Type I family Dopamine Receptor DopR (DopR, dDA1, dumb) in hindleg grooming behavior¹⁵.

Grooming can be indirectly quantified by looking at the extent of cleanliness by which an animal can fully groom after dusting the entire body with a marker dye or fluorescent dust^5,16. The remainder of dust left on the body can be used as a relative marker for the overall behavior. Dusty flies after being given sufficient time to groom may be manifesting a specific deficit in grooming behavior. As grooming investigations have become more extensive, protocols have incorporated such practices as decapitation to add pharmacological treatments onto the neck connective nerves¹⁰, tactile stimulation of bristles to elicit the grooming response² , and video recording of behavior¹⁵. Direct observation of grooming can be easily studied by using visual observation and manually recording the frequency and duration of specific grooming events⁴.

We designed a fifteen-well grooming chamber that can be constructed with a 3D printer or laser cutter, and the blueprint designs are available for reproduction¹⁵. The design uses two joined central plates with openings matched and separated by mesh and two additional sliding top and bottom plates, from which flies and/or dye is loaded, respectively. After allowing dusted flies time to groom, we deposit them in ethanol to solubilize the dye and measure the absorbance of this solution at the wavelength of the dye. A plate reader can be used for multiple parallel samples or a single-read spectrophotometer can be used for individual samples. This method minimizes the error induced by handling and allows for grooming assays to be run on a smaller, cost-efficient scale. This method is derived and modified from the methods pioneered by Julie Simpson and Andrew Seeds, who use larger grooming chambers with heating elements for temperature sensitive circuit manipulations⁵. The following protocol showcases the quantification of grooming of the whole body as well as showing alternate methods for quantitation of dye accumulation on individual body parts. We also present sample comparison data between WT and DopR mutants, as well as methods for calculating a simple performance index for grooming behavior.

Protocol

1. Preparation

1. Prepare an aspirator for moving live Drosophila from a culture vial to the grooming chamber. Aspirators allow transfer of conscious animals to the behavioral chambers to ensure that anesthesia does not affect subsequent behavioral observation.
   1. Using scissors cut 1.5 feet of tygon tubing ID ⅛", OD ¼".Cut at least 1 inch off of the tip of a 1 mL disposable micropipette tip. Hold a 1 cm square piece of mesh (opening 0.196 inch) over the cut tip.
      NOTE: Most 1mL tips have a gradation line where the tip sits in the package box, typically cut to that line.
   2. Snugly place a fresh 1 mL micropipette tip over the mesh/cut tip. Observe the mesh form a tight barrier between the two tips. The inside of the new tip creates a holding chamber for flies.
   3. Cut the extreme tip of the new micropipette tip to widen the opening enough to allow passage of a single fly. The hole will roughly match the opening of the grooming chamber (approximately 1.5-2 mm in diameter).
   4. Snugly fit the nested tips onto the end of the tygon tubing. Push the tips into the tube to ensure a tight fit to allow for vacuum pressure.
   5. On the other end of the tubing, cut the tip of a 200 μL micropipette tip and fit the narrow cut end into the tubing. This is the "mouth" side of the aspirator.
2. Prepare dust aliquot tubes. Weigh approximately 5 mg of Brilliant Yellow dye on tared weighing paper. Pour the dust into a 0.6 mL microcentrifuge tube and tightly close the cap.
   NOTE: We advise the use of nitrile gloves during aliquot preparation, as some latex gloves are permeable to the dye.
3. Repeat step 1.2) for all samples in the experiment (one for every fly in each chamber).
4. Prepare ethanol (EtOH) tubes: Pipet 1 mL of 100% EtOH into 1.5 mL microcentrifuge tubes. Label tubes for genotypes or conditions.
5. Repeat step 1.4) for all samples in the experiment (one for every fly in each chamber). Cap and save the EtOH tubes for completion of the grooming assay. Also include at least one "Blank" sample which will only be 1 mL EtOH without a fly for negative controls.
6. Assemble each grooming chamber by screwing the sliding top plate (with the smaller holes) but leaving the bottom plate (with the larger holes) removed for addition of dust.
7. Place each necessary grooming chamber flat on a table with the top face down. Fly entry side is face-down.
8. Load 5 mg Brilliant Yellow dye aliquot into each chamber by tapping the tube against the surface of the chamber above the desired well. Tap the chamber against the table to ensure that all of the dust falls through the mesh to the top plate.
9. Screw the bottom plate onto each chamber such that the wider ends of the conical openings face outward and the holes do not rest over the wells. Secure the bottom plate tightly so that it does not slide and release dye.
10. Flip the chamber over and knock it flat against a table a few times so that the dust falls through the mesh to rest on the bottom plate.
11. Slide the top plate of the chamber into the "open position" so that the small holes align with the fifteen wells, allowing the introduction of flies into individual wells.

2. Fly-dusting and Grooming

1. Load one fly into the mouth aspirator by gently sucking through one end of the aspirator as if using a straw. Deposit the flies by lightly blowing and directing the opening toward the target.
2. Aspirate one fly into each well used for the experiment. After loading a well, slide the top plate so that the fly is trapped in the chamber and place tape over the opening of that well during loading of the whole chamber to prevent the fly's escape while loading other wells. If multiple flies are aspirated into a single well, leave that opening uncovered until only one fly remains.
3. After all of the necessary wells are filled with flies, flip the chamber such that the bottom plate is upward so that the dust has the potential to coat the flies by falling through the mesh.
4. Tightly secure the top and bottom plates by tightening the screws, and vortex the chamber to dust the flies for 4 s.
5. Knock the chamber (top plate facedown) against a table twice so that dye falls through to the other side. Then, flip the chamber over (top plate face-up) and knock the chamber twice to ensure dye settles on the floor of the bottom chamber away from the fly held in the top chamber.
6. For control flies which are not allowed to groom, immediately proceed to Step 3 of the protocol. For flies that will complete the assay, proceed to Step 2.7.
7. Rest the chamber flat on a table or countertop with the top plate upward for thirty minutes to allow the flies to groom. Place the chamber in a noise and vibration-free space to keep your environment constant for all grooming experiments (lighting levels, humidity, and temperature). A tabletop incubator at RT or 25 °C with humidity controls is ideal.

3. Preparation of Samples and Absorbance Analysis

1. Keeping the chamber level with the top plate upward, gently unscrew the bottom plate and remove it from the chamber, taking care not to lose dye.
2. Place the chamber on a Fly CO₂ pad with low airflow until the flies are anaesthetized.
3. Unscrew the top plate from the chamber. Carefully use forceps to grab a leg and move one dusted fly from each chamber and deposit it in a 1.5 mL microcentrifuge tube containing EtOH. The transfer time for 15 chambers each containing one fly is approximately 3-5 min. Experimenters should factor in transfer times if/when staggering experiments with multiple chambers to ensure efficient staging of experiments.
4. Once each microcentrifuge tube contains 1 fly, close the cap and invert the tube three times to agitate and mix the solution of dye and ethanol.
5. Incubate tubes containing ethanol and flies for 5 h at RT to ensure full removal of the dye from the fly into solution. After the 5 h, briefly vortex each tube (2 s) to ensure clearance of the dye from the flies.
6. Aliquot 50 µL of the experimental 1.5 mL tube into a well of a 96-well plate if available, taking note of the location of each sample on the plate. Add 200 µL of EtOH to dilute the sample 5x. The dilution avoids ceiling effects of heavily dusted flies or large grooming defects.
7. Use a plate reader to analyze each sample at 397 nm. Alternatively, measure absorbance for individual samples and blanks on a standard spectrophotometer in the visible spectrum if a plate reader is unavailable.
8. Read and record the sample through the plate reader and save the spreadsheet with the recorded samples on the plate.

4. Quantification of Results

1. Compile results for all samples and measure the variance for each genotype or condition.
   NOTE: Dye accumulation at time 0 min and at time 30 min are regarded as separate conditions using statistical analysis by one-way ANOVA and Bonferroni correction or other corrections for multiple parallel comparisons.
2. Calculate grooming percentage difference for each genotype/condition using the following equation: [(dye accumulation mean value at time 0' - dye accumulation mean value at time 30') / dye accumulation mean value at time 0] x 100.
3. Express the percentage for each genotype and condition in bar plots and compare across genotypes and conditions.

Results

The grooming assay yields quantitative data to assess behavioral performance based on the relative remainder of accumulated dye left on the bodies of flies after a set time of measurement for grooming (30 min). Sample images of the sliding grooming chamber design and major steps of the assay are highlighted in Figure 1. Flies aggregate a significant amount of dye from immediate dusting by vortexing in the presence of dye (Figure 2d, 2e). Dusted flies can ...

Discussion

The grooming assay is relatively straightforward, but we would caution experimenters to pay special attention to the following issues. Maintaining a tight seal by tightening the screws on the top and bottom plates after introducing flies and dye is essential for reproducible results. The Brilliant Yellow dye is very fine and loose joints will allow losses of dye from the edges of the chamber. The irregularity in dye content for each well could easily throw off grooming quantification as non-uniform dusting will increase ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
High-Flex Tygon PVC Clear Tubing	McMaster-Carr	5229K54	ID 1/8", OD 1/4", used with micropipettor tips and mesh to construct mouth aspirators
Micropipette tips (1 mL and 200 μL)	Genesee Scientific	24-165, 24-150R
Nylon Mesh Screen, 2 x 2.6"	McMaster-Carr	9318T44	Used to construct grooming chamber and mouth aspirators
Dumont #5 Forceps	Roboz Surgical Instrument	RS-5050
Brilliant Yellow Dye	Sigma-Aldrich	201375-25G	we recommend use of nitrile gloves while handling this product
Vortexer	Fisher Scientific	12-812	set to "touch"
Ethanol	Carolina Biological Supply	86-1282
1.5 mL microcentrifuge tubes	VWR International	10025-726
0.65 mL microcentrifuge tubes	VWR International	20170-293	tubes can be reused with successive assays
UV 96-well plate	Corning	26014017
BioTek Synergy HTX Platereader	BioTek	need to download catalog to access product number	http://www.biotek.com/products/microplate_detection/synergy_htx_multimode_microplate_reader.html?tab=overview
Gen5 Microplate Reader and Imager Software	BioTek
Microsoft Excel	Microsoft		https://www.microsoftstore.com/store/msusa/en_US/pdp/Excel-2016/productID.323021400?tduid=(65d098c0e83b86c952bdff5b0719c83f)(256380)(2459594)(SRi0yYDlqd0-LI..ql4M2LoZBEhcBljvIA)()
Drosophila Incubator	Tritech	DT2-CIRC-TK
1/4" acrylic plastic	McMaster-Carr	8473K341
8 - 32 nuts	McMaster-Carr	90257A009
8 - 32 x 1" hex cap screws	McMaster-Carr	92185A199	the bottom plate needs to be tapped for this size screw
8 - 32 x 1/2" hex cap screws	McMaster-Carr	92185A194	the second plate from the top needs to be tapped
2 - 56 3/8" flat head phillips machine screws	McMaster-Carr	91500A088	these hold the two middle plates together
0.175" ID, 1/4" OD, 0.34" aluminum pipe	McMaster-Carr	92510A044	Manufactured in-house; product listed is approximately the same dimensions and should work for size 8 screws.  These act as sheaths for the 1" screws and set the hex cap up slightly from the surface of the top plate