Fluorescence Photoactivation Imaging of Neurofilament Transport in Myelinated Mouse Axons

Abstract

Source: Boyer, N. P. et. al., Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve. J. Vis. Exp. (2020)

This video demonstrates the fluorescence photoactivation method to analyze neurofilament transport in myelinated axons of peripheral nerves. By tracking photoactivated green fluorophore-tagged neurofilaments, the technique provides insights into axonal transport in neurodegenerative diseases.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation of nerve saline solution

1. Make 100 mL of Breuer’s saline: 98 mM sodium chloride (NaCl), 1 mM potassium chloride (KCl), 2 mM potassium di-hydrogen phosphate (KH₂PO₄), 1 mM magnesium sulfate (MgSO₄), 1.5 mM calcium chloride (CaCl₂), 5.6% D-glucose, 23.8 mM sodium bi-carbonate (NaHCO₃) in double-distilled water.
2. Bubble 95% oxygen/5% carbon dioxide (carbogen) through the saline solution for at least 30 minutes prior to use. Leftover saline can be reused within one week; however, it must be reoxygenated before each use.
3. Pour oxygenated saline into a 60 mL syringe and ensure that there is minimal air remaining in the syringe.

2. Initial assembly of the nerve perfusion chamber

1. Connect the syringe and the tubing as shown in Figure 1A, placing the outflow tube into a waste flask.
2. Place the outer gasket into the perfusion chamber housing, ensuring that the flow inlet and outlet posts are aligned with the holes in the gasket.
3. Lay the inner gasket (silicone, 100 µm thick) on a #1.5 circular coverslip (40 mm diameter), carefully smoothing out any wrinkles in the gasket to ensure a tight seal. To facilitate later assembly, place the coverslip and gasket on a paper towel or task wipe with the gasket facing up.

3. Dissection and preparation of mouse tibial nerve

1. Sacrifice the animal by carbon dioxide inhalation or another institutionally approved method. Start a timer when the animal ceases movement/breathing, as experiments must only be conducted within 3 hours of sacrifice.
2. Spray the fur with 70% ethanol and remove as much as possible from the animal’s legs and back using an electric razor.
3. Using a pair of large dissection scissors, make a dorsal incision in the skin near the middle of the spine and continue the cut around the ventral aspect of the animal. Starting from this cut, slowly reflect the skin from the legs by gently pulling it away from the muscle and cutting the fascia.
4. Place the animal in a supine position on a dissection tray and pin all four paws. Optionally pin the tail to reduce movement further.
5. Using microdissection scissors, make an incision in the thigh muscles midway between the tail and knee to expose the sciatic nerve. Ensure that the nerve, which is visible through the muscle, is not cut.
6. Extend the incision dorsally and ventrally to remove the muscle. Similarly, remove the muscles of the calf, keeping cuts shallow and short to avoid damaging nerves.
7. Remove muscles until the tibial nerve is fully exposed from the point where it branches from the sciatic nerve (at the knee) to the heel (Figure 2A).
   NOTE: In all steps, including and following dissection of the tibial nerve, avoid unnecessary exposure to ambient light to minimize possible incidental activation of photoactivatable green fluorescent protein (paGFP) in the nerve.
8. Grasp the tibial nerve at the spine-proximal end with a pair of forceps and cut the nerve using a pair of microdissection scissors. Taking care not to put tension on the nerve, lift it away from the muscle, cutting any attachments.
9. Cut the spine-distal end of the tibial nerve and transfer to a small Petri dish of room-temperature oxygenated saline. From this point on in the procedure, always be certain to keep track of the proximal and distal ends of the nerve.
   NOTE: One way to do this is to mark the distal end of the nerve with an angled cut such that the taper is visible.
10. Starting from the proximal end of the nerve, gently grasp the exposed axon ends with a pair of very fine-tipped forceps.
11. With a second pair of forceps, grasp the nerve sheath proximally, and slowly pull towards the distal end of the nerve. The nerve sheath will slide along the axons with minimal resistance. Ensure that no undue tension is applied to the nerve during this process.

4. Final nerve perfusion chamber assembly

1. Grasping the proximal end of the nerve, remove it from the saline and slowly lay it down onto the coverslip of the perfusion chamber within the rectangular opening of the inner gasket, maintaining gentle tension on the nerve as you lay it down so that it lies straight.
2. Place the micro-aqueduct slide over the nerve with the grooved side facing the nerve and the direction of flow parallel to the nerve. Flip the coverslip and microaqueduct assembly over, and place it within the perfusion chamber housing with the microaqueduct slide apposed to the outer gasket. The nerve and surrounding inner gasket will now be sandwiched between the coverslip and the microaqueduct slide, which are separated by the gasket, with the coverslip facing up (Figure 1B).
3. Secure the perfusion chamber by placing it in the metal housing and rotating the locking ring. Ensure that the plastic housing is fully under all metal clips and tighten well to prevent saline leakage. Overtightening may crack the microaqueduct slide or coverslip. Flip the chamber over so that the coverslip is facing down.
4. Slowly depress the saline syringe plunger to fill the perfusion chamber. Keep the inlet and outlet tubing, outlet flask, and syringe elevated above the chamber itself at all times during setup and imaging. This avoids siphoning, which can introduce bubbles or cause focus instability due to negative pressure in the chamber.
5. Transfer the perfusion assembly to an inverted microscope stage and mount the saline syringe into the syringe pump. Start the motor at an appropriate speed for a flow rate of 0.25 mL/min. Then connect and turn on the in-line solution heater set to 37 °C.
6. Connect the objective heater and set to 37 °C, apply oil to the objective, and insert the perfusion chamber into the stage mount.
7. Apply oil to the chamber heater pad and attach to the perfusion chamber. Connect and turn on the chamber heater; set to 37 °C.
   NOTE: Changes in temperature may cause bubbles to form in the perfusion chamber due to outgassing of the solution. If bubbles form, briefly increase the solution flow rate by 5-10x until bubbles clear the chamber.
8. Lock the perfusion chamber into the stage adapter and bring the objective oil into contact with the coverslip on the underside of the chamber.
   NOTE: The Bioptechs chamber with ASI stage adapter used here are designed for an inverted microscope configuration.

5. Fluorescence activation and image acquisition

1. Using brightfield illumination, focus on the layer of axons on the bottom surface of the nerve closest to the coverslip surface (Figure 2B). Myelinated axons (typically 1 - 6 µm in diameter in adult mice) can be identified by the presence of a myelin sheath, which is visible under brightfield transmitted light illumination without contrast enhancement. Schmidt-Lanterman clefts and nodes of Ranvier are also readily apparent. Unmyelinated axons are more slender (typically <1 µm diameter) and are generally present in bundles (Remak bundles), where they are generally too closely apposed to be resolved from each other.
2. If available on the microscope, activate the auto-focus system to maintain focus over the course of time-lapse imaging.
3. Acquire a brightfield reference image. Record the orientation of the nerve (spine-proximal and distal ends) with respect to images.
4. Acquire a confocal image using a 488 nm laser and an emission filter appropriate for paGFP (e.g., 525/50 nm) to record the pre-bleach autofluorescence. Keep the laser power low to minimize photobleaching, with exposure time adjusted accordingly to detect the faint signal. As an example, representative data were acquired at 5% laser power and 4 s exposures. Record the acquisition settings for use in all future experiments.
   NOTE: After photoactivation, the ideal imaging settings will produce a signal-to-noise ratio > 8 and photobleaching of less than 25% of the original signal over the course of 20 images. The axons can also be imaged by widefield epifluorescence microscopy as we did originally, but the image quality will be inferior due to lack of confocality.
5. Set the laser power to approximately 5x normal imaging power and acquire an image with an exposure time of 3-4 minutes. Though not essential, this step is recommended to bleach autofluorescence and other sources of unwanted fluorescence in order to reduce background signal and thus maximize the signal-to-noise of the photoactivated fluorescence.
6. Acquire an image with the settings used in step 5.4 to record the pre-activation autofluorescence after this bleaching step.
7. On the brightfield image, draw a line parallel to the axons with a length equal to the desired activation window size. The length of this window will vary depending on the experimental goal and parameters, but typical lengths are 5 μm for the pulse-escape paradigm and 40 μm for the pulse-spread paradigm.
8. Using this line as a guide, draw a rectangular region of interest (ROI) across the field of view perpendicular to the axons. The region must encompass all the axons to be photoactivated.
9. Determine optimal settings for photoactivation with 405 nm illumination.
   NOTE: Only perform this step and sub-steps prior to first experimental activation. Over the course of an experiment, the same photoactivation settings must be used.
   1. Activate a region of interest repeatedly using the 405 nm laser line, low laser power (e.g., 5%), and pixel dwell time (e.g., 40 µs), and one pulse, acquiring an image of the activated green fluorescent protein or GFP fluorescence after each activation. Repeat until the fluorescence no longer increases, and then quantify the fluorescence in a region of interest for each image.
   2. Plot the average fluorescence intensities versus pulse number. Select the number of pulses after which fluorescence no longer increases as the optimal number of pulses for activation.
10. Activate the paGFP fluorescence in the region drawn in step 5.8 by patterned excitation with 405 nm light. Ensure that an image is acquired just prior to and just following activation.
    NOTE: The ideal paGFP activation will produce a clearly defined region of fluorescence with sharp boundaries contained within the region of interest or ROI.
11. Start a 1 minute timer as the activation finishes. At the end of 1 minute, start acquisition of a timelapse series.
    NOTE: The 1-minute delay is necessary to allow for the increase in fluorescence that is observed following the photoactivation of paGFP. For the pulse-spread method, a 5-10 minute acquisition period with 30-second timelapse intervals is sufficient for the measurement of the initial slopes in the central and flanking windows to measure velocity and directionality. For the pulse-escape method, an acquisition period of 30-150 minutes with 5 or 10-minute timelapse intervals permits analysis of the long-term pausing behavior of the filaments.
12. Save all images acquired, as well as the ROI used for fluorescence activation.
13. Move to a new region of the nerve and repeat steps 5.1-5.11. If the new region is along the same axon, it must be at least 500 µm from the previously activated region to avoid detection of fluorescent neurofilaments that moved out of the other activated region. The acquisition of the final timelapse must finish before the end of the 3-hour window.
    NOTE: It is possible that the preparation may be viable for longer than 3 hours, but we have not confirmed that. With proficient dissection and preparation, between five and eight 10-minute timelapse image sets may be acquired within this 3-hour window.
14. After the final timelapse image series is acquired, stop the flow of saline, disconnect the solution and chamber heaters, and remove the perfusion apparatus from the microscope stage.

Results

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
14 x 22 Rectangle gasket 0.1mm	Bioptechs	1907-1422-100	Inner gasket
2-deoxy-D-glucose	Sigma	D6134
30mm Round gasket w/ Holes	Bioptechs	1907-08-750	Outer gasket
35 x 10mm dish	Thermo Fisher	153066	Dissection dishes
40mm round coverslips	Bioptechs	40-1313-0319
60mL syringe - Luer-lock tip	BD	309653
Andor Revolution WD spinning-disk confocal system	Andor		Outfitted with Perfect Focus and FRAPPA systems
Calcium chloride	Fisher	C79
Coverslips	Fisher	12-541-B	For fluorescein slide
D-(+)-glucose solution	Sigma	G8769
Dissecting pins	Fine Science Tools	26001-70
Dissection forceps	Fine Science Tools	11251-30	Fine tipped forceps
Dissection microscope	Zeiss	47 50 03
Dissection pan with wax	Ginsberg Scientific	568859
Dissection scissors	Fine Science Tools	14061-09	Initial dissection scissors
FCS2 perfusion chamber	Bioptechs	60319-02-03
Fluorescein sodium	Fluka	46960
Inline solution heater	Warner Instruments	SH27-B
Laminectomy forceps	Fine Science Tools	11223-20	Initial dissection forceps
Magnesium sulfate	Sigma-Aldrich	M7506
Microaqueduct slide	Bioptechs	130119-5
Microscope slides	Fisher	12-544-3	For fluorescein slide
Microscope stage insert	Applied Scientific Instrumentation	I-3017
Objective heater system	Okolab		Oko Touch with objective collar
Objective oil - type A	Nikon	discontinued
Plan Apo VC 100x 1.40 NA objective	Nikon	MRD01901
Potassium chloride	Fisher	P217
Potassium phosphate	Sigma-Aldrich	P0662
Sodium bicarbonate	Sigma-Aldrich	S6297
Sodium chloride	Sigma-Aldrich	S7653
Syringe pump	Sage Instruments	Model 355
Tubing adapter - female	Small Parts Inc.	1005109
Tubing adapter - male	Small Parts Inc.	1005012
Tygon tubing	Bioptechs		1/16" ID, 1/32" wall thickness
Vannas spring scissors	Fine Science Tools	15018-10	fine scissors