A Microbiomechanical System for Studying Varicosity Formation and Recovery in Central Neuron Axons

All methods described below have been approved by the Institutional Animal Care and Use Committee (IACUC) of the Ohio State University.

1. Coverslip Preparation

1. Place one or more boxes of 12 mm or 25 mm coverslips into a glass beaker containing 70% nitric acid and incubate the coverslips at room temperature overnight.
   NOTE: Do not wash the 25 mm coverslips in the same beaker as the 12 mm coverslips. It is better to wash them separately.
2. Transfer all coverslips into a 4 L beaker filled with 2.5 L of ddH₂O and shake the beaker containing the coverslips for 1 h at 100 rpm. Use rubber bands to hold the beaker during shaking. Repeat the rinse with fresh 2.5 L ddH₂O four times.
3. Transfer cleaned coverslips to a dry flask with a metal cover. Bake the coverslips in an oven at 225 °C for 6 h and allow them to cool to room temperature.
   NOTE: The 25 mm coverslip is prone to breaking. Separate coverslips one by one and air dry, then bake them in an oven.
4. Store the dried and cleaned coverslips at room temperature in a sterilized plastic container until usage.
5. Coat the coverslips as follows:
   1. Prepare fresh coating solution in a centrifuge tube.
   2. Place each coverslip into one well of a 24-well (or 6-well) plate. Place 30 (or 100) µL of coverslip coating solution (Table 1) onto a 12 mm (or 25 mm) coverslip and swirl. Add 1 mL of sterile phosphate-buffered saline (PBS, tissue culture grade) to the well. These plates can either be used immediately or stored at 4 °C in sterile PBS for up to a week.
   3. On the day of dissection, remove the PBS. Air-dry and place the plates under UV light (30W) for 2-4 h.
      NOTE: The last step should be completed with the UV light on in a cell culture laminar flow hood. The door of the hood should be lifted slightly to allow air flow for drying the coverslips.

2. Dissection, Dissociation, and Culture of Hippocampal Neurons from Pregnant Mouse/Rat

1. Dissection and dissociation of hippocampal neurons for culture
   1. Thaw protease enzyme solution (3 mg/mL protease 23 in SLDS, Table 1) and pre-warm plating media (Table 2) at 37 °C.
   2. Dissect hippocampus and rinse with SLDS, according to the methods described in previous publication¹⁴.
      NOTE: Four hippocampi provide enough cells for one 24-well or one 6-well plate.
   3. Remove SLDS, add 2 mL of protease enzyme solution, and incubate the sample at 37 °C, 5% CO₂ for 15 min.
   4. Wash the hippocampus explants with 5-10 mL of plating medium twice. Add 5 mL of plating medium. Dissociate hippocampal explants into single cells by generating a small vortex using a pipette with a 1-mL plastic tip. Pipette up and down about 40 times, avoiding the formation of oxygen bubbles, until the solution becomes cloudy and no large pieces of explant remain.
      NOTE: Remember to leave some media during washes, the hippocampal explants should remain submerged in the medium during the whole process.
   5. Centrifuge the cells at 1,125 x g for 3 min. Remove the supernatant with the cells submerged in media. Resuspend the cells in 145 mL of plating media. Add 1 mL/well of the cell suspension to a 24-well plate (3 mL/well to a 6-well plate).
      NOTE: 145 mL of plating media is used to produce a cell suspension that has a low cell density. This volume will allow the plating of cells into six 24-well plates, eight 6-well plates, or various combinations of 24- or 6-well plates depending on the experiments to be conducted. Each well of the 24-well plate will have about 3 x 10⁴ cells⁹.
   6. Incubate the cells in the plating media at 37 °C, 5% CO₂ for 2-4 h, then remove all plating media and replace it with fresh maintenance media.
   7. After 2 days (2 days in vitro, 2DIV), replace half of the media with 2 µM Ara-C (arabinosylcytosine) dissolved in maintenance media which inhibits growth of fibroblasts, endothelial, and glial cells. After exposing cells to Ara-C for two days, replace the media with fresh maintenance media.
2. Transfection for the visualization of cell morphology
   NOTE: Transfect the cells with fluorescent constructs of green fluorescent protein (GFP), yellow fluorescent protein (YFP), mCherry, etc. to illuminate the morphology of individual neurons.
   1. Dilute the constructs from stock (1 µg/µL) into Opti-MEM media (0.8 µg of the construct in 50 µL of Opti-MEM media) and vortex. Prepare 1 construct dilution for each well.
      NOTE: A 24-well plate requires 0.8 µg of construct in 50 µL of Opti-MEM media. A 6-well plate requires 2.4 µg of construct in 150 µL Opti-MEM media.
   2. Dilute liposome-mediated transfection reagent into Opti-MEM media (1.5 µL of the transfection reagent in 50 µL of Opti-MEM media) and vortex. Prepare 1 transfection dilution for each well.
      NOTE: A 24-well plate requires 1.5 µL of transfection reagent in 50 µL of Opti-MEM media. A 6-well plate requires 4.5 µL of transfection reagent in 150 µL of Opti-MEM media.
   3. Prepare a 1:1 mixture (100 µL) of the construct in Opti-Mem media and transfection reagent in Opti-MEM media and vortex. Incubate the mixture at room temperature for 20 min.
      NOTE: Per well, there should be about 100 µL of solution containing construct, transfection reagent, and Opti-MEM media if using a 24-well plate (300 µL for 6-well plate).
   4. Remove half of the media (500 µL for 24-well plate, 1.5 mL for 6-well plate) from each well and transfer this media into a clean conical tube. Keep this tube in the incubator. Then add the 100 µL mixture (step 2.2.3) to the remaining media in the well. Allow the cells to incubate at 37 °C for 20-30 min.
   5. During the incubation (step 2.2.4), add one volume of fresh maintenance media to the media in the conical tube (step 2.2.4). Place the mixture in the same incubator (37 °C and 5% CO₂).
   6. Following the incubation, remove all media containing the transfection solution from the wells and replace it with the 1:1 mixture of old and fresh maintenance media in the conical tube (step 2.2.5).
      NOTE: Depending on the size of the constructs, cells may begin expressing within 12 h; for larger constructs, 3-4 days may be needed before the peak expression of the constructs is reached.

3. Setting Up the Puffing Pipette Apparatus

NOTE: There are five basic components to this set-up: the glass pipette, the tubing, the syringe, the micromanipulator, and the buffer (Hank's). Any buffer that has physiologically relevant salt concentrations could be used in this set-up.

1. Pull a borosilicate pipette to have around 45-µm-diameter opening tip using a micropipette puller and the specifications in Table 3.
   NOTE: It was found that 45 µm is a suitable size for the diameter of the pipette opening under experimental conditions. Slight variations in the opening size should not significantly affect results. It is important to note that larger departures from this number should still work for varicosity induction by adjusting the height of the syringe that is linked to the pipette via tubing, but this will require recalibration of the pressure value.
2. Insert the un-pulled end of the pipette into thin rubber tubing.
   NOTE: The tubing wall should be thin and rigid to minimize the fluid resistance, ensuring that the height of the syringe is directly proportional to the pressure of fluid flowing through the pipette with negligible resistance. The tubing should be no longer than about 0.6 m in length to further minimize the resistance.
3. Connect the other end of the tubing to a 10-mL plastic syringe through a connector with a turn-able valve.
   NOTE: The plastic syringe needs to be held at a constant, measurable height to ensure an accurate measure for the assumed pressure flowing from the pipette. A height of 190 mm was utilized, since this provides minimal pressure but reliably induces varicosities in axons. Other height values may also be used if the pressure cause the cells to detach from the coverslip surface. It is recommended to use the same setting throughout the experiment.
4. Insert the pipette into the screw top of the micromanipulator to hold the pipette in place. Screw the metal, flat topped screw attached to the micromanipulator arm so that it holds the un-pulled end of the pipette just above where the rubber tubing is attached. Angle the pipette at a 45° angle with the surface of the neuron-containing coverslips.
   NOTE: The micromanipulator should be constructed so that the pipette can be held without constricting the rubber tubing. The micromanipulator must allow motion of the pipette in the x, y, and z directions to accurately position it within range of the cells. A graphic as well as a photograph of the apparatus is provided below (Figure 1).
5. Turn on a fluorescence filter, open the aperture, and ensure a fluorescent spot can be seen coming through the objective. Using the micromanipulator, move the pipette into a position that lines up the tip with the fluorescent spot and lower the pipette into a position just above the height of the cell culture dish (35 mm x 10 mm). Once it is in position, turn on the transmitted light and turn off fluorescence.
6. Using tweezers, add one coverslip (with the cells facing up toward the pipette) from the cell culture plate to the cell culture dish containing about 2 mL of Hank's buffer at room temperature.
7. Place the culture dish containing the coverslip onto the scope.
8. Using the fine focus knob and 20X objective, focus on a plane about four full turns above the cells (about 0.4 mm). Use the micromanipulator to position the pipette tip so that it is focused and in the center, left-hand side of the plane as seen through the eye-pieces.
   NOTE: Dendrites are projections that should be in the same frame as the cell body they originate from. To induce axonal varicosities, one should follow longer projections from the cell body to medial and distal axons.

4. Calibrating the Pressure of the Pipette Utilizing a Stretchable-membrane System

1. Setup the puffing pipette apparatus with the exact same parameters as described above over a system containing a microfabricated silicone membrane (500 µm in diameter and 50 µm thick) and focus the microscope onto the surface of the membrane.
   NOTE: This system was designed to measure small pressure values and an explanation of the system has been published before¹⁰. This measurement only needs to be done once for puffing experiments if the exact same setting of puffing is used.
2. Focus the microscope on the arrays of microdots (4 µm in diameter and 10 µm apart, as measured between centers). Turn the stop valve and allow fluid to flow out of the pipette. Examine the deflection of the membrane in response to the fluid pressure with a confocal microscope.
3. Utilize a linear model to determine the corresponding pressure associated with deflection of the membrane.
   NOTE: While a recent study found that a deflection of w= -3.143 ± 0.69 µm was obtained for 190 mm syringe height, this produced a pressure of 0.25 ± 0.06 nN/µm² from the current pipette setup, which is within the physiological and pathological range, though the syringe height is not the sole determinant. Other factors influence the exact pressure value on the neurons as well, including pipette opening, positioning (distance and angle) of the pipette tip relative to the cells, and even the flexibility of tubing^4,10.

5. Puffing to Induce Varicosities

1. Once the pipette is in position, focus the microscope on a plane that contains a region of interest. Position the cells and processes in the left half of the imaging field (seen by live capture of the camera) within the puffing area, while the right half is outside of puffing area.
   NOTE: Due to extensive outgrowth of axons and dendrites of neurons, it is unlikely that all the processes of a neuron are within the same puffing area. This is actually one advantage of this system, which allows the examination of the local effect of puffing. There are two types of puffing that can be utilized to induce varicosities: long duration puffing and pulse puffing.
2. Puffing
   1. Long duration puffing
      1. Position the syringe at the height, 190 mm above the coverslip containing cultured neurons⁴. Begin time-lapse imaging of area of interest with an optimized exposure time revealing clear neuronal morphology. Capture at least 15 frames at a 2 s interval (30 s total) as the baseline. The interval can be adjusted based on the experiment. At frame 16, open the turn-able stop valve of the syringe and allow fluid to flow for 75 frames (150 s). At frame 75, turn the valve off so that fluid flow stops. Stop video capture.
         Note: This should induce axonal varicosities in neurons 7-9 DIV within 5-10 s, whereas older neurons take longer to form varicosities.
   2. Pulse (2 s duration) puffing
      1. Position the syringe at the proper height (190 mm). Begin time-lapse imaging and capture at least 15 frames (30 s) at a 2 s interval. At frame 16, open the valve of the syringe and allow fluid to flow, then close the valve after 2 s. Wait 5-20 s (depending on the frequency to be tested) before opening the valve again. Repeat opening and closing of the valve to create fluid pulses, and continue for a total period of 150-300 s. Continue time-lapse imaging for 20 min to capture the recovery stage.
         ​NOTE: When the interval is longer than 10 s, the pulses induce drastically less varicosities. At a 20 s interval, no varicosities can be induced by the pulses⁴. To obtain reliable, consistent varicosity formation, it is advised the syringe height should be around 190 mm. Syringe height and fluid pressure should not be raised to a point where cells begin detaching from the coverslip surface.
   3. Before a day of use, clean the set-up (pipette, tubing, and syringe) by flowing about 10 mL 70% Ethanol through the set-up. Following the ethanol wash, flow 10 mL of Hank's buffer (or desired buffer) through the set-up to prime the system for that day's use. After finishing experiments for the day, clean the set-up as previously described with ethanol to prevent growth of contaminants overnight.

6. Complimenting Methods

NOTE: The puffing assay can be combined with a variety of different techniques that are discussed in the Introduction section. Here, the focus is on the core techniques that are most frequently used together with the puffing assay, including high-resolution fluorescence timelapse imaging, calcium imaging, and recovery assays. These are discussed below.

1. Conduct high-resolution fluorescent time-lapse imaging by following the protocol below
   1. Track the movement of fluorescently-tagged proteins within neurons with a 100X oil lens. Neurons are transfected with the proper cDNA construct. Capture for 150 frames (total 300 s) with a 2 s interval.
      NOTE: Sometimes, multiple-color time-lapse imaging is performed to image the movement of more than one fluorescently-tagged protein simultaneously.
   2. Generate a kymograph either through the video capturing software or through ImageJ (NIH).
      NOTE: The speed and directionality of travel can be determined using the kymograph. *Be sure to note the direction the soma lies from the region captured. Travel towards the soma is retrograde, while travel away from the soma to the axonal tip is anterograde.
   3. Repeat this process following puffing, or a kymograph can be created using the image capture during the puffing assay.
      NOTE: The effects of puffing on axonal transport of mitochondria and synaptic proteins were shown in a recent paper⁴.
2. Calcium imaging
   1. Add 1 µL (for a 24-well plate) or 3 µL (for a 6-well plate) of 5 mM Fluo-4 AM to the culture media in one well and incubate at 37 °C for 30 min.
   2. Wash the cells twice with 1 mL Hank's buffer.
      NOTE: Loaded neurons emit green fluorescence through a FITC filter set. Loci denote regions of appreciable calcium within the cell. When combined with puffing, an increase in intensity (increase in internal calcium level) can be seen.
3. Varicosity recovery
   NOTE: Immediately following a puffing experiment, a 20 min recovery experiment can be carried out.
   1. Capture images for 300 frames (1,200 s) with a 4 s interval.
      NOTE: A cell transfected with soluble YFP/mCherry/GFP should show a moderate level (less than 50%) of recovery by the end of the imaging session. Axonal recovery depends on a number of factors, such as the developmental stage of cultured neurons⁴. This can also be combined with high-resolution fluorescent imaging and calcium imaging described above.