Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using “Cargo Mapping” Analysis

All experiments were conducted following approved protocols and according to institutional guidelines for the humane care of research animals. Neonate mice were euthanized by decapitation.

1. Preparation of Microfluidic Devices for Cell Culture

1. Prepare polydimethyl siloxane (PDMS) microfluidic devices for growth of hippocampal neurons as described by Harris and colleagues^17-19. Below are some modifications that were adapted to the cargo mapping protocol.
   NOTE: Microfluidic devices are also commercially available (Materials List), thus access to a fabrication facility is not necessary.
2. Prepare No. 1½ cover glasses (24 x 40 mm) by washing them three times with acetone, followed by three washes with 100% ethanol and three washes of water. Store coverslips in water at 4 °C until use. These treatments help ensure that coverslips are clean from debris that might interfere with plating of cells, contamination, and survival.
   NOTE: Acetone and ethanol are flammable and hazardous in case of skin contact (irritant), of eye contact (irritant), of ingestion, of inhalation. Dispose according to official regulations.
3. When ready to use, place one cover glass in a 60 mm cell culture dish per microfluidic device, and let it air dry uncovered for 45 min inside a biosafety cabinet to avoid contamination.
4. After devices are cut out from the masters and punches have been cut to create the reservoirs (Figure 1A, B), place them with the microfluidic channels side-up inside a biosafety cabinet equipped with an UV lamp for 45 min for sterilization.
   NOTE: Leaving devices under UV treatment for longer than 2 hr might affect the integrity of the PDMS.
5. Assemble the devices by placing one device onto each cover glass inside the 60 mm plastic cell culture dish so that the microfluidic channels face down. Apply gentle pressure to the top of the device (avoid touching the microchannels) to provide a non-permanent seal of the device to the cover glass. Cover each 60 mm cell culture dish with a lid.
6. Using a micropipette, hydrate the devices by adding cell culture grade water to the top two microfluidic reservoirs (1 and 2 in Figure 1A), allowing water to flow through. Remove water with a micropipette and add 1 mg/ml poly-L-lysine into microfluidic reservoir 1 (200 µl) and 2 (100 µl). The volume differential will allow for the poly-L-lysine to flow through the microchannels (Figure 1A, B).
7. To ensure that the devices inside the 60 mm cell culture dishes do not topple over inside a 37 °C incubator, place three 60 mm cell culture dishes containing the devices into a 150 mm plastic cell culture dish and cover the dish with a lid. Incubate O/N in a 37 °C incubator with 5% CO₂. Typically more than 90% of the channels are usable per device and do not contain any air bubbles or physical blockage.
8. On the next day, replace poly-L-lysine with water and leave device in incubator for at least 1 hr. Repeat this washing three times. Remove water and add Neurobasal-A growth medium (containing 2% B-27 supplement and 500 µM GlutaMAX) to each device. Leave O/N in a 37 °C incubator with 5% CO₂.
9. On the next day, plate hippocampal cells as described below.

2. Plating of Mouse Hippocampal Primary Neurons in Microfluidic Devices

NOTE: The preparation of cultured hippocampal neurons from neonate rats has been previously published in JoVE²⁵. Below are some modifications that were adapted to plate mouse hippocampal neurons in microfluidic devices, and that were optimal for transfections.

1. Before mouse brain dissection, pre-warm Dulbecco’s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum (FBS; without antibiotics), Mixture A (125 mg DL-cysteine, 125 mg bovine serum albumin (BSA) and 3.125 g of D-glucose in 500 ml of phosphate buffered saline (PBS); filter sterilize through a 0.22 µm filter), and deoxyribonuclease I solution (DNase I: 50 mg of DNase I and 0.5 ml of a 1.2 M MgSO₄ solution in 100 ml Hank’s Balanced Salt Solution (HBSS) filter sterilize through a 0.22 µm filter) in a 37 °C water bath.
   1. Pre-warm Neurobasal-A growth medium inside a 37 °C incubator with 5% CO₂ to equilibrate the pH.
2. Dissect hippocampi from 1-3 day old neonate mice according to established protocols²⁶. Keep hippocampi in a 15 ml conical tube containing 10 ml dissection buffer (HBSS containing 10 mM HEPES pH 7.4, 50 mM glucose, 100 U/ml penicillin and 100 µg/ml streptomycin), on ice. Put up to 4 hippocampi per 15 ml conical tube.
3. Perform the remaining protocol is under sterile conditions inside a biosafety cabinet.
4. Dilute 45 units of papain in 5 ml of Mixture A, vortex, and incubate for 5 min at 37 °C until papain powder is dissolved. Add 1 ml of DNase I solution. Filter the mix through a 0.22 µm syringe filter unit.
5. Carefully aspirate HBSS’ from 15 ml conical tube containing hippocampi. Add 10 ml cold (4 °C) HBSS to hippocampi and let them settle to the bottom of the tube. Carefully aspirate HBSS from 15 ml conical tube.
6. Add 1 ml papain/DNase I mix to up to 4 hippocampi and incubate for 20-30 min at 37 °C. Tilt conical tube every 5 min to bathe hippocampi with mix. Alternatively place hippocampi in a 37 °C water bath shaker with gentle shaking (~100 rpm).
7. Aspirate papain/DNase I mix and wash hippocampi twice with pre-warmed DMEM (37 °C) containing 10% FBS.
8. After second wash, add 2 ml of pre-warmed DMEM containing 10% FBS to hippocampi and manually triturate hippocampi by pipetting up and down ~10-12 times using a 1 ml pipette tip to dissociate neurons.
9. Let triturate settle for ~1 min as this will allow non-dissociated neurons to settle to the bottom of the conical tube. Transfer supernatant containing dissociated neurons to a new 15 ml conical tube avoiding transfer of larger tissue that has settled at the bottom of the tube.
10. Spin cells at 1,000 x g for 2 min and carefully remove supernatant without disturbing the cell pellet. Resuspend cells in 80 μl Neurobasal-A growth medium by pipetting gently.
11. Remove medium from microfluidic reservoir 1 and 1’. Apply 20 µl of cells (~275,000) to microfluidic reservoir 1, and allow them to flow through. Place device in 37 °C incubator with 5% CO₂ for 20 min.
12. Look under microscope to ensure that cells have adhered to the cover glass. Add Neurobasal-A growth medium to top off all reservoirs. Return device with cells to a 37 °C incubator with 5% CO₂.
13. Check devices every ~2-3 days and top off reservoirs with Neurobasal-A containing 2% B-27 supplement (without GlutaMAX) to avoid evaporation. Neurons start to extend their axons through microfluidic channels ~2-3 days after plating. Subject neurons to transfection typically 7-10 days after plating.

3. Transfection of Hippocampal Neurons Grown in Microfluidic Device

1. Per device, prepare a mix of 1.2 µl Lipofectamine 2000 in 30 µl Neurobasal-A, and a mix of 0.5 µg fluorescent cargo fusion plasmid in 30 µl Neurobasal-A and incubate 5 min at RT. Add Lipofectamine mix to DNA mix and incubate for 20 min at RT. Add 60 µl of Neurobasal-A containing 4% B-27 supplement to the DNA/Lipofectamine mix and mix by flicking the tube.
2. Remove all media from microfluidic reservoir 2 and 2’ and carefully fill up wells with Neurobasal-A containing 2% B-27 supplement without spilling over the medium into the other compartments.
3. Remove media from microfluidic reservoir 1 and 1’. Add 120 µl of DNA/Lipofectamine mix to microfluidic reservoir 1 and let it flow. Return device to 37 °C incubator with 5% CO₂ for 3-4 hr.
4. Remove all media from microfluidic reservoir 1 and 1’ and replace with Neurobasal-A containing 2% B-27 supplement for 24 hr (or until ready to image). Typically, 5-7 axons that grow through microchannels are transfected under those conditions, which represent approximately 1-3% transfection efficiency consistent with efficiencies published for hippocampal cultured cells ²⁶.

4. “Cargo Mapping” Analyses

1. Live Imaging of cargo movement
   1. Perform the imaging on an inverted epifluorescence light microscope equipped with a 37 °C incubator and a CO₂ control chamber.
   2. Mount the coverslip with hippocampal neurons grown in a microfluidic device onto the microscope stage. To avoid neuronal death, work expeditiously to maintain the samples at the microscope for no longer than 1 hr.
   3. Using a high-resolution objective (100X), find axons of transfected cells that extend through channels of microfluidic chambers and record the number of the channels in which the transfected axon was found. Count the number of channels using a hand tally counter.
      1. For optimal recording of live movement and subsequent colocalization analysis, choose axons that grow relatively flat through the channels so that imaging of most vesicles can be performed in focus.
   4. Remove most of the medium from reservoirs 2 and 2’ with a 1 ml plastic transfer pipette. To facilitate subsequent alignment of fixed images with movement trajectories on kymographs, align the right edge of the microchannels with the field of view during imaging (Figure 1B, C).
   5. To facilitate fixing of the samples during imaging, perform live imaging with two people. Alternatively, if live imaging is performed by a single person, use a microscope system equipped with drift correction.
   6. Start imaging live cargo movement with time-lapse specifications specific to the transport dynamics of the cargo being analyzed (microscope specifications and settings for imaging YFP-PrP^C are specified in the legend of Figure 2 and Materials List).
   7. After sufficient live movement data have been collected, have one person fill up reservoirs 2 and 2’ with pre-warmed 4% paraformaldehyde in PBS containing 0.04 g/ml sucrose to fix the cells. Avoid touching the microfluidic device, as this will disrupt the focus of the live imaging. Have the other person adjust the focus while the fixative is added. Fixation is successful when immediate stop of movement is observed. Add fixative to reservoir 1 and 1' afterwards.
      NOTE: Temporary photobleaching of the cargo fluorescence might also be observed, but the fluorescence persists after IF staining for motor proteins is completed (next step).
      NOTE: Paraformaldehyde is harmful by inhalation, in contact with skin, and if swallowed. Irritating to eyes, respiratory system and skin. Dispose according to official regulations.
2. IF staining of motor proteins for colocalization analysis
   NOTE: For quantitation of relative levels of motor proteins (as determined by antibody staining), associated to individual moving cargoes, validate and titrate antibodies to ensure that the antibody-antigen binding is saturated. This is done by determining the specificity of the antibody using neurons in which the protein of interest has been either knocked-out or knocked-down, and by performing IF analysis with increasing antibody concentrations. For negative controls, neurons are stained with secondary antibodies in the absence of primary antibodies. More detailed description of antibody validation can be found in Encalada et al.¹⁶, and Szpankowski et al.²⁰.
   1. Remove the coverslip with hippocampal neurons grown in microfluidic device from the microscope stage. Do not detach the microfluidic device from the coverslip, as the microchannels need to remain intact in order to superimpose the live and fixed images. Perform the following steps in a humidity chamber.
   2. To ensure the flow of solutions into the microchannels, maintain a buffer volume differential between reservoirs 1, 1’, and 2, 2’ of at least 30 µl. in all subsequent steps. For example, add 100 µl media volume to microfluidic reservoir 2, 2’, and 70 µl media volume to microfluidic reservoir 1, 1’.
   3. Remove fixative from chambers of the microfluidic device and wash cells three times with PBS waiting 10 min between washes.
   4. Permeabilize cells by adding 0.1% Triton X-100 diluted in PBS for 5 min to all reservoirs.
   5. Remove solution from all reservoirs and wash with PBS. Repeat washing three times waiting 10 min between washes.
   6. Remove solution from all reservoirs and apply blocking buffer containing 10% Normal Donkey Serum and 3% protease free and immunoglobulin G (IgG)-free BSA in PBS and incubate for at least 30 min at RT.
   7. Spin primary antibody stock solution at 20,000 x g for 5 min to remove precipitates. Dilute antibody to an appropriate concentration in blocking buffer. Replace the blocking buffer from microfluidic device with antibody solution. Incubate samples for 2 hr at RT or O/N at 4 °C.
   8. Remove solution from all reservoirs and wash three times with PBS, waiting 10 min between washes.
   9. Spin secondary antibody stock solution at 20,000 x g for 5 min to remove precipitates. Dilute antibody to an appropriate concentration in blocking buffer. Replace PBS with secondary antibody solution. Incubate samples for 1 hr at RT.
   10. Remove solution from all reservoirs and wash three times with PBS, waiting 10 min between washes.
   11. Replace PBS with ~10-20 µl mounting medium by directly pipetting mounting medium into the opening of the channels connecting the reservoirs. Let mounting media dry for ~20 min – 1 hr at RT.
   12. Store devices at 4 °C protected from light to avoid photobleaching, and image axons within two days (maximum one week) of staining.
   13. After staining is completed, mount coverslip with hippocampal neurons grown in microfluidic device onto the microscope stage. Locate the region of the transfected axon imaged in step 4.1.1-4.1.7.
   14. Acquire Z-stack images (300 nm steps) for each of the three channels: cargo, motor 1, motor 2, using a high-resolution objective (e.g., a high numerical aperture oil or water objective).
       NOTE: Acquisition of Z-stacks is important as axons often do not grow perfectly flat in microchannels and therefore not all fluorescent puncta are in the same focal plane.
3. Cargo Mapping
   1. Generate a kymograph of cargo movement using image analysis software. The ImageJ plugin developed by Rietdorf and Seitz (EMBL) is recommended for generating kymographs (for download see Materials List).
      NOTE: ImageJ is a public domain, Java-based image processing program developed at the National Institutes of Health^27,28 (for download see Materials List).
   2. Align the fixed images of the fluorescent cargo (generated in step 4.2.14) manually by superimposing them onto the position of the trajectories in the kymograph at the point of fixation (Figure 2A, B), using a commercially available graphing software program (Materials List). Manually align by first superimposing the fixed cargo image onto the kymograph and manually choosing the cargo puncta that corresponds to a specific trajectory on the kymograph. For best alignment results, use the Z slice that is in best focus for the cargo mapped. Several Z slices may be used.
   3. For each image in the Z-stack (cargo, motor 1 and motor 2; generated in step 4.2.14) determine the X-Y coordinates and intensity amplitudes for each fluorescent puncta using an established algorithm to fit 2D Gaussians to the point spread function that represents each point source in each of the three channels^16,20,21 (Figure 2D).
      NOTE: To obtain the X-Y coordinates, a custom built MATLAB software package called “Motor Colocalization” was developed^16,20, which incorporates a Gaussian fitting algorithm developed by Jaqaman, Danuser and colleagues^21-23. The software package and detailed instructions for its use can be obtained upon request.
   4. For each of the individually cargo puncta that were mapped onto the mobile or stationary trajectories on the kymograph, select the X-Y coordinates and intensity amplitudes from the results of the “Motor Colocalization” program (these cargoes are individually labeled in Figure 2B). Use several Z-stack images acquired in step 4.2.14) to map more cargoes that are found on different focal planes.
   5. Compare the individual X-Y coordinates for the mapped cargo to those obtained for each of the motor channels in the corresponding Z slice (Figure 2D). Determine and select the X-Y motor coordinates that are within a 300 nm radius of the cargo. For cargoes and motors that have signal within the same focal plane, use the “Motor Colocalization” software package to determine the puncta that are located within a 300 nm radius.