Nuclear Transport Assays in Permeabilized Mouse Cortical Neurons

First, the protocol describes the generation of primary neuronal cultures (step 1) and preparation of materials for the transport assay (step 2), followed by the transport assay itself (steps 3-4) and image acquisition and analysis (step 5). All methods described here were approved by the Animal Care and Use Committee (ACUC) of Johns Hopkins University.

1. Primary mouse cortical neuron cultures

1. Prepare the stock solutions.
   1. Dissection buffer: Combine 50 mL of 10x HBSS, 15 mL of 1 M glucose (in dH₂O), 5 mL of 1 M HEPES, 5 mL of 100 mM sodium pyruvate, and 5 mL of 100x penicillin-streptomycin. Bring volume to 500 mL with dH₂O. Sterile filter and store at 4 °C.
   2. Papain: Dilute papain to 30 mg/mL with dissection buffer. Store at 4 °C.
   3. DNase: Dilute DNase I to 10 mg/mL with dissection buffer. Aliquot and store at -20 °C.
   4. Plating medium: Add 5% fetal bovine serum (FBS), 2% B-27, 1% Glutamax, and 1% penicillin-streptomycin to neurobasal medium. Sterile filter and store at 4 °C.
   5. Feeding medium: Add 2% B-27, 1% Glutamax, and 1% penicillin-streptomycin to Neurobasal medium. Store at 4 °C.
2. Coat the plates.
   1. Coat the optical glass-bottom 96-well plates with poly-D-lysine (20 µg/mL) and laminin (1:100) in sterile dH₂O. Place the plate in an incubator overnight.
   2. On the day of dissection, rinse the wells three times with sterile dH₂O and replace with Neurobasal medium. Equilibrate in an incubator for at least 30 min.
3. Dissect and digest the cortices.
   NOTE: Carry out dissection on a clean bench using autoclaved tools to prevent culture contamination.
   1. Euthanize the E15-16 C57BL/6J pregnant female by cervical dislocation. Spray abdomen with 70% ethanol. Work quickly. Use scissors and toothed forceps to make a midline abdominal incision, excise the uterus, and then transfer the embryos from the amniotic sac into a Petri dish. Decapitate the embryos with scissors and collect heads in a 10 cm Petri dish containing dissection buffer.
   2. Under a dissecting microscope, use fine forceps to remove the skull and peel away the meninges. Transfer whole brains to a fresh 10 cm Petri dish containing dissection buffer. Separate the hemispheres and remove the subcortical structures.
   3. Chop the cortices with a sterile razor blade and transfer them to a conical tube containing 5 mL of dissection buffer. Add 100 µL of papain (stock 30 mg/mL, final concentration 0.6 mg/mL) and 50 µL of DNase (stock 10 mg/mL, final concentration 0.1 mg/mL). Digest in a 37 °C water bath for 10 min.
      ​NOTE: Digestion beyond 10 min is detrimental to cell survival.
   4. Remove the supernatant and wash twice with 10 mL of plating media to stop digestion. Allow the tissue to settle by gravity in between each wash.
   5. Triturate gently in 2 mL plating media with P1000 pipette until large pieces of tissue disappear (approximately 20 times). Add another 8 mL of plating media to bring the volume to approximately 10 mL and pass the cell suspension through a 40 µm cell strainer.
4. Plate and maintain neurons.
   1. Centrifuge at 200 x g for 5 min. Resuspend the cells in prewarmed plating medium. Count the cells and dilute to 500,000 cells/mL in the plating medium. Gently invert to mix and transfer to a sterile multichannel reservoir.
   2. Aspirate the neurobasal media from the wells and immediately add 100 µL of cells per well with a multichannel pipette (50,000 cells/well). Do not allow the coated wells to dry.
   3. After 24 h of dissection, change 50% of medium with the feeding medium. Continue 50% medium change every other day until day 5-7 to perform the in vitro assays.

2. Preparation of the nuclear transport assay components

1. Transport buffer (TRB, adapted from¹⁵)
   1. To prepare 5x TRB, dissolve 100 mM HEPES, 550 mM KOAc, 10 mM Mg(OAc)₂, 25 mM NaOAc, 2.5 mM EGTA, and 1.25 M sucrose in dH₂O. Adjust the pH to 7.3 and store the buffer at 4 °C.
   2. On the day of the transport experiment, prepare 1x TRB-BSA. Dilute 5x TRB in dH₂O, add EDTA-free protease inhibitor cocktail, 2 mM of DTT, and 50 mg/mL of BSA.
2. Energy regeneration mix (ERM, adapted from^9,16)
   1. To prepare 20x ERM, dissolve 2mM ATP lithium salt, 2 mM GTP lithium salt, 80 mM Creatine phosphate, and 400 U/mL Creatine kinase in 1x TRB.
   2. Freeze single-use aliquots at -20 °C.
3. Preparation of whole cell extract (WCE, source of transport receptors and Ran cycle proteins).
   NOTE: Cytoplasmic or whole cell extract (WCE) may be used, depending on the experimental goals. Here, the preparation of WCE is described as used in the initial neuron transport assays¹⁰.
   1. Grow HEK293T cells to confluence in fifteen 150 mm culture dishes. Detach with trypsin-EDTA, resuspend in media containing 10% fetal bovine serum to inhibit trypsin, and pellet cells at 200 x g for 5 min. Resuspend the cells in 10 mL of ice-cold 1x TRB with freshly added protease inhibitor cocktail.
   2. Sonicate on ice, 3 x 10 pulses, using a hand-held probe sonicator. Clarify by centrifugation at 14,000 x g for 15 min at 4 °C.
   3. Measure the protein concentration (goal 8-10 mg/mL) and snap-freeze single-use aliquots in liquid nitrogen prior to storage at -80 °C.
4. Recombinant nuclear import cargo
   NOTE: The transport protocol in steps 3-5 may be adapted for any fluorescent nuclear transport cargo, to interrogate the active or passive nuclear transport pathway of interest. Here, the protocol describes the expression and nuclear import of Rango (Ran-regulated importin β cargo), which consists of the importin β-binding domain of importin α1 flanked by the fluorescent proteins CyPet and YPet^14,17. Rango is a versatile sensor that can be used for FRET as well as nuclear import assays, where it functions as a direct importin β cargo.
   1. Transform E. coli BL21(DE3) cells with the Rango plasmid (containing an N-terminal 6-His tag) by heat-shock as follows. Thaw a 50 µL aliquot of BL21(DE3) cells on ice. Combine with 100-200 ng plasmid. Mix by tapping the tube 4-5 times and then incubate on ice for 30 min before placing on a 42 °C plate for 40 s. Transfer immediately back to ice for another 2-5 min.
   2. Add 900 µL of SOC media at room temperature and incubate on a shaker at 37 °C for 1 h before spreading the cells on a prewarmed agar plate (1.5% agar, LB media, 100 µg/mL Ampicillin). Incubate the plate overnight at 37 °C. Store the plate at 4 °C for up to 1 week or use it immediately for protein production.
   3. Start the protein production by inoculating three 25 mL of starter cultures in 50 mL tubes with increasing number of colonies (~3-10) from the BL21(DE3) plate. After incubation overnight at 37 °C on a shaker, select the cultures with OD_{600 nm} < 1.0 to inoculate a 1L culture in LB media using a 2.8 L baffle-free Fernbach flask. Grow at 37 °C until reaching OD_{600 nm} = 0.1-0.3.
      NOTE: The use of older BL21(DE3) LB agar plates or high density of starter cultures (OD_{600 nm} >1.0) could reduce the yield and purity of the protein preparation.
   4. Cool to room temperature (22-25 °C) by placing on ice. Induce protein expression with 0.3 mM IPTG (final concentration) and incubate on a shaker (80-120 rpm) at room temperature for 12-14 h.
   5. Collect the cells by centrifugation (6,000 x g, 15 min, 4 °C) and resuspend in 30 mL of ice-cold 10 mM imidazole in PBS (pH 7.4), supplemented with EDTA-free protease inhibitor cocktail. Lyse by two passes through an ice-cold French pressure cell and clarify the lysate by centrifugation (16,000 x g, 40 min, 4 °C).
   6. Incubate the lysate with high flow Nickel affinity agarose resin (30-60 min, 4 °C), using 1 mL of resin per 1 L of E. coli culture. Place the resin into the chromatography columns, wash with 10-20 volumes of ice-cold 10 mM imidazole/PBS (pH 7.4). Elute Rango with 25 mM imidazole/PBS increments (25-200 mM, pH 7.4), using a 5-fold bed volume in each step.
   7. Use SDS-PAGE to select and pool the batches with the highest purity.
   8. Prepare XB buffer by dissolving 10 mM HEPES, 100 mM KCl, 10 mM CaCl₂, 1 mM MgCl₂, and 50 mM sucrose in dH₂O. Adjust the pH to 7.7.
   9. Dialyze the protein in XB buffer and concentrate by centrifugation (7500 x g, 4 °C) on an ultrafiltration device with a 30 kD molecular weight cutoff.
   10. Measure the protein concentration by the Bradford or other preferred method, and snap-freeze single-use aliquots in liquid nitrogen before storage at -80 °C.

3. Determination of the optimal plasma membrane permeabilization conditions

NOTE: Due to variations between each batch of neurons, optimize the permeabilization conditions for each batch of neurons. Perform this on the same day before performing nuclear transport assays.

1. In a plastic 96-well staging plate, prepare a serial dilution of Tris-HCl pH 7.5 (to cause osmotic swelling), at 0 µM, 10 µM, 20 µM, and 40 µM in each of three BSA concentrations: 50 mg/mL, 100 mg/mL, and 150 mg/mL (for molecular crowding/mechanical support). Prewarm the solutions to 37 °C.
2. Rinse the neurons once in prewarmed PBS. Remove the PBS and transfer Tris/BSA from the staging plate. Return to the incubator for 4 min.
3. Remove from the incubator and place the culture plate on ice. Rinse 2 x 5 min in 1x TRB-BSA (as prepared in step 2.1.2).
4. After the final rinse, add 70 kD of Texas Red-labeled dextran (0.6 mg/mL) and Hoechst (1:10,000) in 1x TRB-BSA. Leave the plate on the bench at room temperature for 15 min, protected from light.
5. Image using a confocal microscope to identify the buffer and the BSA concentration in which the plasma membrane permeabilization percentage is the highest, but the nuclear membrane still restricts entry of the 70 kD dextran (goal ≥ 80%). Use these conditions for subsequent experiments.

4. Nuclear import assay

1. Prepare the transport reactions
   NOTE: To enable all reactions in the plate to be started simultaneously, prepare reaction mix in a plastic 96-well staging plate on ice, prior to neuron permeabilization. Carry out a minimum of two technical replicates per condition. Include controls on each plate.
   1. Prepare the base reaction mix consisting of 2.5 mg/mL of WCE, 1x ERM, fluorescent cargo (200 nM Rango), and Hoechst 33342 (10 mg/mL, 1:10,000) in 1x TRB-BSA, sufficient for 50 µL per well. OPTION: Include 70 kD Texas red-labeled dextran to permit monitoring of nuclear pore integrity throughout the transport reaction, keeping in mind that confocal microscopy is needed to image nuclear dextran exclusion.
      NOTE: The WCE concentration provided is a reasonable starting point for assay optimization. Perform empiric testing of each batch of WCE to determine the optimal concentration required for efficient nuclear import of the desired cargo. Avoid combining data from replicates prepared with different batches of WCE since import efficiency may vary.
   2. Prepare the controls including (1) Cargo alone: Rango, but no WCE or ERM and (2) Inhibitor: fluorescent cargo, ERM, WCE, and 100 µM importazole (IPZ, a small molecule inhibitor of importin β¹⁸). As an alternate inhibitor, use 0.8 mg/mL of wheat germ agglutinin (WGA)¹⁹.
      NOTE: Equilibrate the inhibitors in the assay mix containing WCE for at least 30 min before initiation of transport. For maximal inhibition, post-permeabilization rinse steps in corresponding wells should also include the inhibitor.
2. Permeabilize and rinse the neurons
   1. Permeabilize the neurons according to the optimal, batch-specific protocol identified in step 3.
   2. Remove the culture plate from the incubator and immediately place it on ice. Rinse 2x for 5 min in 1x TRB-BSA. Include an inhibitor in the rinses for designated wells.
3. Run the transport reaction.
   1. Following the final rinse, remove 1x TRB-BSA. Working quickly, use a multichannel pipette (50 µL/well) to transfer the premixed transport reactions onto the permeabilized cells.
   2. Immediately load the plate in a high content microscope at room temperature to initiate time-lapse imaging (see step 4.4) or place the plate on the bench, protected from light, and allow the reaction to proceed for the desired time period (e.g., 30-120 min) prior to fixation and imaging.
   3. If desired, fix in 4% paraformaldehyde/PBS for 15 min, rinse 2x for 5 min in PBS, and transfer to 50% glycerol/PBS for later imaging. Keep the fixed plates protected from light at 4 °C (stable for up to 1 week or more).
4. Live imaging
   NOTE: Apply any preferred image acquisition and analysis method at this stage.
   1. Load the plate in a high content microscope. Run an initial test plate to optimize and save imaging parameters, including magnification, focus, exposure times (Hoechst and FITC), and intervals prior to running experimental samples. Save the parameters to reload with each subsequent plate.
      NOTE: For quantitative analysis, set the exposure time for the fluorescent import cargo (FITC) well below saturation at steady state or the latest timepoint to be used. Establish this using the test plate, targeting half-maximal saturation to permit variations in intensity across experimental conditions. Collect the Hoechst images for each frame and timepoint to permit nuclear identification during subsequent automated analysis.
   2. Adjust the focus offset, if needed, using Hoechst as the focus wavelength and run the imaging protocol. Collect the images every 5 min for 30 min. Adjust the interval and reaction time as needed based on the imaging speed and kinetics of the import reaction of interest.

5. Image analysis

1. In the microscope Review Plate dialog, open Hoechst and FITC images from a representative frame. Under Run Analysis tab, select Translocation-Enhanced module, and configure the settings. Set Hoechst as the compartment image and FITC as the translocation probe image.
2. Under compartments, adjust approximate nuclear width, intensity above local background, and minimum/maximum area. Activate Auto Separate Touching Compartments as the neurons tend to cluster. View results by selecting Test Run. Adjust the settings to maximize the accuracy of neuronal nuclei identification while limiting erroneous inclusion of non-neuronal nuclei or debris.
   NOTE: Glial contamination, if present, can typically be excluded at this step based on nuclear size. Determination of optimal compartment identification settings is a trade-off between accuracy and cell number and must be set empirically depending on the assay, cell type, and culture density. Stringent parameters will omit numerous cells but include fewer errors, while liberal parameters will be more inclusive but contain more errors. If different cell populations (i.e. disease mutants) or treatments are included that may affect nuclear size, we recommend setting size/intensity parameters that are broad enough to accommodate all conditions to avoid the need for multiple parameter sets within an experiment.
3. To avoid artifacts from the nuclear edge, define regions of measurement several pixels inside and outside the compartment edge defined by Hoechst staining. Depending on magnification and binning, set 2-4 pixels for inner region distance, and 1-2 pixels for outer region distance. Adjust the outer region width (1-2 pixels).
4. Select the desired background estimation method (default of Auto Constant is recommended). In Configure data log, select the desired output parameters, including mean inner intensity, mean outer intensity, and inner/outer mean intensity (the nuclear to cytoplasmic (N/C) ratio). Save the analysis parameters.
   NOTE: The inclusion of internal positive and negative controls within each experiment is strongly encouraged to ensure that parameters maximize sensitivity to detect the biological event of interest.
5. Use the Plate Utilities dialog to run analysis on the desired plate(s) using the saved parameters and export the measurements.
6. Review and summarize the data by the treatment condition. OPTION: Apply filters to remove non-physiologic data, such as probe intensity = 0 and extremes of N/C ratio (i.e., <0.1 and >100). Equally apply any filters across all conditions and experiments.
7. As an additional correction for baseline fluorescence, normalize the data to the control wells in which lysate and ER were omitted. To facilitate comparisons across biological replicates (i.e., independent neuronal culture preparations), express the final nuclear import data as a percent of time 0.