In vivo Laser Axotomy in C. elegans

1. Assembling the system

The specific components of our system are outlined below. When properly assembled, the user should be able to focus the microscope with one hand, move the stage with the other (either with the mouse or the joystick), activate the laser with the foot pedal, and have a clear view of the on-screen image.

1. MicroPoint laser mounted to the epi-fluorescence port of a compound microscope. We use a Nikon 80i, but any research-grade upright or inverted scope should work. The MicroPoint system will include a custom dichroic, which should match the fluorophore that labels the cells you wish to cut. Cutting cells labeled with alternative fluorophores requires additional dichroics. Additionally, a pulse generator and foot pedal come with the MicroPoint system.
2. 100x, 1.4 NA oil objective for performing surgery and assessing regeneration, plus additional objectives as necessary for locating the sample on the slide. We use a Nikon Plan Apo VC for surgery, and find that a 4x is useful for coarse focusing and finding the sample.
3. Motorized stage with joystick. A stage with extremely high accuracy and repeatability is unnecessary, since the stage is only used for targeting. We use a Prior OptiScan II (see discussion).
4. Camera. The camera needs to provide a frame rate sufficient to focus and target the labeled neurite with the 100x objective, while visualizing the image on the computer monitor. We prefer attenuating the illumination light during the axotomy procedure to avoid potential confounding effects from photo-bleaching or damage (see 1.7, below). Thus, the camera needs to be sensitive enough to accommodate this reduced light level. We use a Hamamatsu Orca 05G, and find that imaging at 16.3 Hz is sufficient. Equivalent cameras could be substituted.
5. Computer, monitor, and imaging software that can drive the camera. Additionally, we use the imaging software to manipulate the motorized stage with a mouse (see discussion).
6. Air table. This requirement may vary depending on location, but since surgery is performed with a 100x objective, we find vibration isolation to be essential.
7. Light supply with light guide and shutter mechanism for illuminating the fluorophore. The light guide will attach to the MicroPoint. The shutter mechanism is useful to reduce the illumination intensity independently of the laser. We use a Prior Lumen 200 or a Leica EL6000, and attenuate up to 80% of total light intensity for axotomy.

2. Laser setup

A detailed protocol to initially align and focus the laser is provided with the MicroPoint laser system. We assume that procedure has been followed successfully upon initial setup. This includes aligning the laser with the crosshairs in the eyepiece. Here we provide a protocol for regular maintenance of the laser’s focus and intensity.

As noted in the MicroPoint manual, the pulsed nitrogen laser is capable of severely damaging the human eye. Care should be taken to never look directly into the laser beam. Once installed, the barrier filter should safely block radiation through the eyepieces.

Cutting neurons in an organism the size of a grain of sand can be challenging. Since the area of damage done by the laser is very small, it is essential to know the three-dimensional location of the laser’s focus in order to target the sample precisely. The laser focus is found using the mirrored slide. First, we check that the z location of the laser focus matches the focus of the imaging path. Next, the x-y location of the damage focus is marked with crosshairs in the imaging software.

Focusing in the z-plane

1. Push in the neutral density filter ND16 (Figure 1a). We use neutral density filters in the epi-illumination path to attenuate the laser while focusing. For even finer focusing, push in both the ND16 and ND4 fluorescent filters.
2. Set the MicroPoint laser to 1 pulse with the switch turned to ‘Select’ (Figure 1b), and then move the filter wheel to the appropriate long-pass barrier filter (Figure 1c).
3. Place the mirrored slide on the stage and focus on pinholes within it with the 4X objective using transmitted light.
4. Add a drop of immersion oil to the mirrored slide and refocus on pinholes with the 100X objective. Open the camera shutter with the optical path switching lever (Figure 1d) and the laser shutter (Figure 1e). If necessary, reduce transmitted light intensity so the edges of the pinholes are crisp: this will aid in focusing exactly.
5. Press the foot pedal. This should produce a small hole in the mirror if the laser is focused properly.
6. Focus the microscope slightly above the plane of the mirrored slide and shoot the laser again. This should make an even smaller hole than the first shot, or leave no mark at all. Repeat by focusing below the mirrored slide.
7. If a hole is not produced in 2.5, or if a larger hole is produced when focusing above or below the surface of the mirror, refocus the laser with the focus ring on the ablation head. Verify that the hole is still properly aligned with the crosshairs in the eyepiece. With consistent use, the z-plane focus rarely needs to be adjusted.

Focusing in the x-y plane

8. Follow steps 2.1 through 2.3 above and press the foot pedal to make a small hole in the mirror slide.
9. Initiate ‘Live’ capture mode from the Elements ‘Acquire’ menu. In the ‘Measure’ toolbar select ‘ROI editor’, then select the ‘double cross’ tool and drag the crosshairs to align them with the center of the new hole. Click ‘Save ROI’ and then ‘Exit editor’ and resume Live capture mode.
10. Activate the ‘mouse XY’ setting in Elements to manipulate the stage with the mouse.
11. Pull the neutral density filter(s) back to the original position(s), set the laser pulses to 10, and remove the mirrored slide.

Adjusting the power of the laser

12. The power of the laser should be adjusted to the minimum power necessary to cut the neuron of your choice. Adjust the power by moving the attenuator plate (Figure 1f) while cutting living worms (described below). If the laser power is too high, it will cause peripheral damage or blast a hole in the worm. If the laser power is too low, it will not sever the neuron. Once the system has been set up, the attenuator position rarely needs to be changed. If the laser becomes weak, and the attenuator plate needs to be moved to continue cutting, it could be a sign that the dye in the dye cell needs to be replaced (see discussion).

3. Immobilize the worms

1. Prepare a solution of 3% molten agarose in M9 (22mM KH2PO4, 42mM Na2HPO4, 85mM NaCl, 1mM MgSO4). Dispense 5 mL of molten agarose into a 15 mL falcon tube. Fill another tube with water. Place tubes in a modular heating element set to 55°C. Fill the heating element with water to create mini water baths for each tube. Note: agarose can be prepared in large quantities in advance and re-melted for subsequent experiments.
2. Place two layers of labeling tape on each of two slides and a clean slide between these slides. Dispense a drop of agarose using a plastic bulb pipette to the middle of the clean slide and place another slide perpendicular to the first so that the resulting agarose pad is the thickness of two pieces of tape (Figure 2). Remove one of the two slides from the pad after 30 seconds. Rinse and store the plastic pipette in the water-filled falcon tube for use with subsequent slides.
3. Next, place 3-5 μL of 0.10 μm or 0.05 μm polystyrene beads in the center of the pad. Add 5-10 worms expressing fluorescent protein in the neurons of interest. If the neurons of interest are asymmetrically distributed on the right or left side of the worm, use a platinum pick to flip worms so that the correct side is facing up. Worms must be placed on slides within 10 minutes of preparing the agarose pads.
4. Carefully place a cover slip on the worms. After placing the cover slip, do not move it relative to the agarose pad as this will disrupt the cuticle and destroy the worms. Place the prepared slide on the microscope stage.

4. Cut neurons

1. Direct light into the eyepieces with the optical path switching lever. Focus on the worm to be cut with the 4X objective, place a drop of oil on the cover slip, and switch to the 100X objective. Return the optical path to the camera.
2. Use the mouse to move the neuron of your choice under the center of the crosshairs. Press the foot pedal to fire the laser and sever the neuronal process. If needed, two sets of pulses can be used to sever the neuron. At first, it is helpful to track precisely which neurons have been severed in individual worms in order to assess subsequent regeneration. When performed correctly, laser axotomy produces a small break in the neuron without causing significant damage to the surrounding skin tissue or other neurons (see Figure 3). In certain cases, a small scar may form around the site of injury.
3. Recover the worms by removing the cover slip in a direct upward movement, being careful that the worms remain on the agarose pad. Do NOT slide the coverslip off. Then, cut the pad around the worms with needle-nose forceps and place the section of agarose on a freshly seeded NGM plate containing OP50⁸. Place 10 μL of sterile M9 on the pad to free the worms from the beads and remove agarose.

5. Score neurons for regeneration

1. Prepare agarose pad as outlined in step 2.2.
2. Place worms in 3-5 μL of polystyrene beads. Apply a coverslip. Slides can be prepared sequentially, or alternatively all the worms can be prepared in advance and the slides kept in 10 cm culture dishes, each containing a damp Kimwipe to keep the agarose moist.
3. Use the 100X objective to visualize the severed neurons. In many neurons, a neuronal stump distal to the cut site will remain. We use the presence of this remnant as an indication that the neuron was indeed severed (Figure 4). The presence of this stump makes it possible to distinguish a neuron that has been cut and regenerated from one that was not cut. Note: both the distal and proximal stumps initially retract after being cut.
4. Assess regeneration. A variety of parameters can be determined. One simple assay is whether the injured neuron has formed a growth cone (Figure 4a) and regenerated, or not (Figure 4b). Alternatively, neurite length can be traced and compared.

6. Representative results

As an example, we describe characterization of regeneration of the γ-Aminobutyric acid (GABA) motor neurons. These neurons, which reside in the ventral nerve cord and extend processes circumferentially to the dorsal nerve cord, are essential for proper locomotion⁹. GABA neurons can be visualized by expressing a genetically encoded fluorophore, such as green fluorescent protein (GFP), under the control of the unc-47 or unc-25 promoters (strains EG1285 and CZ1200, respectively, available from the C. elegans Genetic Center).

Mount L4-stage worms as described, flip the worms so the GABA neurons on their right sides are facing up, and place the cover slip. Cut 1-3 of the posterior commissures in each worm, midway between the dorsal and ventral cords. Avoid cutting commissures that cross or fasciculate with other commissures, as these are difficult to score later. Recover worms as described.

18 to 24 hours after a successful axotomy, the worms will have recovered from surgery and exhibit normal locomotory and egg-laying behaviors. Discard worms that are dead or sick. Remount the remainder as described, and assess regeneration. We usually aim to assess at least 30 cut neurons per experimental condition. In oxIs12 animals, typically 60-70% of severed L4 GABA neurons will have initiated a growth cone structure, evidenced by a broadening of membrane at the tip and the extension of several neurite branches, and migrated from the cut site (Figure 4a). In many cases, the growth cones will have migrated to reconnect with the dorsal nerve cord, an adjacent neuronal commissure, or the distal stump. The remaining 30% of neurons will have either initiated no growth, forming a stump proximal to the site of axotomy, or will have extended only small filopodia (Figure 4b).

Figure 1. The UV pulsed laser axotomy system. Specific components are: (a) ND filters (b) pulse selector (c) filter wheel lever (d) optical path switching lever (e) laser shutter and (f) attenuator plate.

Figure 2. Preparing an agarose pad. To prepare an agarose pad of the desired thickness, two layers of tape (green) are placed on two slides. A slide is placed between the first two, and a drop of agarose is then added to the clean slide. Finally, a fourth slide is placed perpendicularly to the first three, resulting in a pad that is the thickness of two pieces of tape.

Figure 3. Representative GABA neuron axotomies. In a typical experiment, GABA neuron commissures are severed at the middle of the lateral aspect of the worm. A severed commissure is shown immediately before (left panel) and after (right panel) axotomy. All images were taken with a 100X oil objective.

Figure 4. Representative results of GABA neuron axotomies. 24 hr post-axotomy severed axons are scored for regeneration. A regenerating axon with a growth cone (arrow) is shown in (a) while a non-regenerating axon is seen as a proximal stump (arrow) in (b). The distal stump of each cut axon is shown in each panel (arrowhead) and is an indication that the axon has been severed.