DiI Dye-Filling as a Simple and Inexpensive Tool to Visualize Ciliated Sensory Neurons in C. elegans

Summary

DiI dye-filling is a method commonly used in C. elegans to visualize a subset of the ciliated sensory neurons, allowing for the identification of genetic mutations that alter sensory neuron structure or function.

Abstract

C. elegans have long been used as a simple and accessible model to study neuronal structure and the many functions of the nervous system. Of the 302 neurons within the adult hermaphrodite nervous system, 60 are classified as ciliated sensory neurons. These neurons are central to a number of C. elegans behaviors, including but not limited to chemo-, mechano-, and osmosensing, male mating, and dauer formation. For several decades now, members of the C. elegans community have used the red fluorescent lipophilic dye DiI to visualize a subset of the ciliated sensory neurons that are directly exposed to the external environment. This dye enters the ciliated ends of the neurons and distributes in a relatively uniform pattern throughout the dendrites, cell bodies, and axons. This simple and powerful method makes an excellent first-pass tool to identify genetic mutants that impart structural or functional defects in ciliated sensory neurons. Here, we present a streamlined version of this staining method to visualize the eight pairs of amphid and two pairs of phasmid neurons that are environmentally exposed in C. elegans. We discuss tips for using this inexpensive method for imaging cellular dye-filling patterns in anesthetized animals.

Introduction

Caenorhabditis elegans (C. elegans) are easy to manipulate, have fast generation times, and are low-cost to maintain. Due to these and many other advantages, C. elegans have served as a preferred model organism for studying many biological processes, especially the development and function of the nervous system. An entire issue in the Journal of Neurogenetics was recently dedicated to the historical impacts of research on this particular topic¹. They are particularly beneficial for studying the function of primary cilia, which are involved in sensing chemical and physical environmental conditions².

Adult hermaphrodite C. elegans have a total of 302 neurons, 60 of which possess primary cilia at the end of their dendritic processes³. These 60 ciliated neurons are classified as sensory neurons and are involved in many C. elegans behaviors, including but not limited to chemo-, mechano-, and osmosensing, male mating, and dauer formation^3,4. There are two subsets of ciliated sensory neurons that are exposed to the external environment, which include sixteen amphid neurons (8 pairs) in the head and four phasmid neurons (2 pairs) in the tail^3,5.

For several decades, researchers in the C. elegans community have used lipophilic dyes, such as the red fluorescent 1,1'-dioctadecyl-3,3,3'3'-tetramethylindocarbocyanine perchlorate (DiI), to visualize a number of different tissues in live animals^6,7,8,9. When animals are exposed to DiI, the dye easily and quickly intercalates into the membrane of the dendrites, axons, and cell bodies of the 20 externally exposed amphid and phasmid neurons in a relatively uniform distribution. When wild-type animals are exposed to DiI, the dye can be visualized in these neurons by fluorescent imaging during a relatively broad window of time. If there are any morphological or functional abnormalities in the primary cilia, the dye may not properly fill the neurons, and therefore, a signal may appear weaker in some or all cells or may be completely absent^6,7,10,11. Any of these outcomes can be informative of structural or functional deficits that may be present in the ciliated sensory neurons of genetic variants.

This manuscript aims to demonstrate the ease with which dye-filling can be used in C. elegans to visualize the structure of ciliated sensory neurons (Figure 1). We applied this technique in wild-type and mutant animals to demonstrate how different genetic backgrounds can show a variety of dye-filling outcomes, often related to the structural or functional integrity of their ciliated sensory neurons. We show staining at 30 min, 24 h, and 48 h post-dye-filling in a variety of different animal ages to aid in determining the optimal time course for live imaging. We also provide examples of difficulties that can arise during staining and imaging and tips to avoid these problem points. Through the use of this method, researchers at institutions of any size can begin to build upon the foundation of ciliated sensory neuron biology in C. elegans. Dye-filling is simple and cost-effective enough to be incorporated into lab activities with undergraduates to permit them the chance to work with C. elegans and fluorescence microscopy with minimal prior technical expertise. In addition, there is a significant conservation of genes involved in primary cilia biology and, more broadly, sensory neuron function between humans and C. elegans¹². Continued research on ciliary gene functions and genetic interactions in C. elegans could ultimately provide greater insight into the complexity of human ciliopathies¹³.

Protocol

1. Preparation of solutions

1. Prepare the bleach solution. Combine 2 mL of bleach, 500 µL of 10 M NaOH, and 7.5 mL of dH₂O in a 15 mL centrifuge tube.
   NOTE: Bleach solution remains stable for up to 1 week when stored at room temperature (RT).
2. Prepare the M9 buffer. Combine 3 g of KH₂PO₄, 6 g of Na₂HPO₄, and 5 g of NaCl in 1 L of sterile water and autoclave to sterilize. Once cooled completely to RT, add 1 mL of sterile 1 M MgSO₄.
3. Prepare the DiI stock solution. Add 10 mg of DiI to 5 mL of dimethylformamide (DMF) for a final concentration of 2 mg/mL DiI in DMF. DiI is light-sensitive, so always cover the solutions with aluminum foil. DiI in DMF can be stored at -20 °C for many years and does not require thawing between uses.
   CAUTION: DMF is both flammable and harmful when in contact with skin or inhaled. Work with this solution under a chemical hood, away from open flames, and wearing proper personal protective equipment (PPE).

2. Isolation of synchronized populations by bleach preparation

NOTE: Gather all of the equipment and solutions necessary for all steps of the process prior to beginning. Bleach preparations are very time-sensitive, so having the necessary materials before starting the process ensures protocol success.

1. Wash unstarved, gravid adult animals off plates using 1 mL of M9 buffer. Allow the animals to collect at one edge of the plate by tilting the plate slightly.
   NOTE: Typically, aim to collect ~50-100 unstarved, gravid adult animals for a reliable yield of eggs. However, bleach preparation can be performed on nearly any number of gravid adult animals. The number of gravid adult animals available for collection can vary widely depending on the genetic background of the animals. A greater number of animals collected usually yields a greater number of eggs.
2. Using a glass Pasteur pipette, collect and transfer the animals in the M9 buffer to a labeled 1.5 mL microcentrifuge tube.
3. Centrifuge the animals at 350 g for 30 s. This speed is slow enough to collect the live animals at the bottom of the tube without causing any harm.
4. Decant most of the excess M9 buffer, leaving the collected worms at the bottom of the tube undisturbed.
5. Add 1 mL of the prepared bleach solution and allow the animals to float in the solution until they begin to disintegrate, inverting the tube regularly to encourage tissue breakdown. View the tissue breakdown and release of eggs by periodically checking the tube under a dissecting microscope. This process usually takes between 5 min and 10 min.
6. Once the tube contains mostly released eggs with only a few visible animal tissues remaining, centrifuge at maximum speed for a standard tabletop microcentrifuge (typically between 15,000-21,000 g) for 30 s to collect the eggs at the bottom.
7. Decant as much of the bleach solution as possible using a glass pipette while careful not to disturb the pelleted eggs. Add 1 mL of M9 buffer to halt the degradation of the eggs by the remaining bleach solution.
   NOTE: Steps 2.6 and 2.7 should be carried out quickly. Delays in centrifugation, removal of bleach solution, and washing with M9 buffer can result in low egg yields due to degradation.
8. Centrifuge the tube again at 21,000 g for 30 s. Decant the supernatant without disturbing the pelleted eggs.
9. Wash twice more with 1 mL of M9 buffer, decanting liquid between each wash without disturbing the pelleted eggs.
10. After the final wash, decant most of the M9 buffer, then resuspend the eggs in the remaining liquid by flicking the tube or shaking vigorously.
11. Using a sterile glass pipette, transfer a drop of the solution containing the eggs to the surface of a fresh NGM plate with E. coli. Aim to place the drop next to the E. coli and add around 100 eggs per plate. These eggs are approximately synchronized in age.
    NOTE: In cases where the egg yield is high enough that obtaining 100 eggs per drop is difficult, additional M9 buffer can be added to the tube containing eggs to dilute their concentration.

3. Dye-filling procedure

1. Label as many 1.5 mL microcentrifuge tubes as needed for the number of strains being dye-filled. Create foil covers for each tube to protect light sensitive DiI solution, once added.
2. Wash animals of a desired developmental stage from the age-synchronized NGM plates using approximately 1 mL of M9 buffer. Ensure that each plate contains approximately 100 animals hatched from the 100 eggs placed on the plate during bleach synchronization. Collect the animals into the properly labeled 1.5 mL microcentrifuge tubes.
   NOTE: Dye-filling works well on a wide variety of developmental stages. This study demonstrates dye-filling in animals ranging from larval stage 3 (L3) to adults (48 h post-L3).
3. Centrifuge animals at 350 g for 30 s to collect them at the bottom of the tube.
4. Decant the M9 buffer without disturbing the animals collected at the bottom of the tube.
5. Wash twice more with 1 mL of M9 buffer, decanting liquid between each wash without disturbing the collected animals. On the last round of washing, decant to leave 200 µL of M9 buffer remaining in the tube.
6. Cover all tubes with aluminum foil to protect from light exposure and add 1 µL of DiI in DMF directly to the M9 buffer in each tube.
   NOTE: If changing the amounts of M9 buffer and DiI, make sure to maintain a 1:200 ratio of DiI in the M9 buffer.
7. Rock or shake all tubes at RT for 2 h.
8. Centrifuge tubes at 350 g for 30 s to collect animals at the bottom of the tubes.
9. Decant the liquid containing the DiI from the microcentrifuge tubes. Wash with M9 buffer three times, decanting liquid between each wash.
10. After decanting from the final wash, transfer animals in a small (~50 µL) volume of M9 buffer to fresh NGM plates with E. coli.
11. Allow animals to recover after staining on fresh NGM plates for at least 30 min. Cover the plates with aluminum foil until observation to preserve the DiI stain. Observe animals up to 24 h after dye-filling with no significant loss in fluorescent signal. At or after 48 h, the signal can begin to fade (see Figure 2).

4. Imaging dye-filling

1. Prepare a 2% agarose in a sterile water solution. Fully dissolve the agarose using a microwave.
   NOTE: Agarose (2%) in water solution can be reused multiple times until repeated water loss increases agarose concentration to a point where the solution becomes too viscous.
2. Obtain glass slides and 22 mm x 22 mm coverslips to prepare agarose pads. Using a glass pipette, place a small drop of the molten 2% agarose in water on one glass slide and quickly place a cover slip on top of the agarose drop. Once the agarose pad is fully solidified, remove the coverslip, leaving the agarose pad on the glass slide.
   NOTE: Avoid the formation of bubbles in agarose pads, as they can obstruct the imaging of animals.
3. Add 5-10 µL of anesthetic (10 mM levamisole) to the surface of the agarose pad. Transfer 5-20 animals to the anesthetic, then place the cover slip back on top of the agarose pad.
4. Observe animals using a stereo, compound, and/or confocal fluorescence microscope, depending on available equipment and screening and imaging needs. DiI has an excitation peak at 550 nm and an emission peak at 564 nm; thus, view it using a tetramethylrhodamine/cyanine dye 3/yellow fluorescent protein (TRITC/Cy3/YFP)-compatible filter cube.
   NOTE: Appropriate microscope settings, such as exposure time, gain, and laser power, needed to obtain clear images of dye-filling can vary between different types of microscopes and should be independently determined by users for their unique needs.

Results

Adult N2 worms imaged 24 h after dye-filling demonstrate clear fluorescent signal spread relatively evenly throughout the amphid neurons (Figure 1A,A') and phasmid neurons (Figure 1D,D'). In these animals, the dendritic projections and cell bodies of the amphid neurons can be easily distinguished. There are no clumps of dye in the dendritic projections, nor are there any interruptions in...

Discussion

xSuccessful dye-filling relies on careful consideration of the developmental stage and genetic background of the animals, as well as the elapsed time until imaging. Some genetic mutations disrupt the structure and/or function of the externally exposed ciliated sensory neurons, resulting in animals that are unable to dye fill properly. Therefore, dye-filling of novel C. elegans mutants can be used as a simple first-pass identifier for defects in sensory neuron structure or function. However, one limitation of thi...

Materials

Name	Company	Catalog Number	Comments
DiI	Biotium	60010	Not water soluble, make 2 mg/mL solution in DMF. Solution is light sensitive, cover with foil. Store at -20 °C. Solution good for many years.
Levamisole hydrochloride	Fisher	AC187870100	10 mM solution in M9 Buffer. Store at -20 °C. Solution good for many years.
M9 Buffer	IPM Scientific	11006-517	Available for purchase, but also easy to make in house following recipe in protocol.
N2 (C. elegans strain)	CGC	N2	C. elegans wild isolate
YH2125 (C. elegans strain)	n/a	n/a	Strain generated in Yoder Laboratory (Bentley-Ford et al, 2021)