Bimolecular Fluorescence Complementation-Coupled Photoactivated Localization Microscopy

1. Cloning

1. Determine the configurations to clone and choose a linker. Tag proteins with the fragments on the N- or C-terminus as described below so they do not disrupt their proper localization. Use a flexible linker such as (GGGGS)x2.
2. Genetically tag the proteins of interest to the PAmCherry1 fragments. As one option, use the cloning plasmids listed in the Materials List containing the fragments RN (PAmCherry1 residues 1-159) and RC (Met plus PAmCherry1 residues 160-236) with flanking MCS sequences.
   1. Linearize the plasmids containing the fragments RN and RC with inverse PCR (or restriction digest) at the point of insertion. Primers for inverse PCR are listed in Table 1. Below is an example PCR protocol used in the author's lab (see Materials List for the exact materials used in this protocol).
      1. Mix the PCR reaction together, bringing up to a final volume of 50 µl with water: to a thin-walled PCR tube, add the water, 25 µl of 2x master mix, 0.5 µl each of the forward and reverse primers (20 µM diluted stock, 0.2 µM final concentration), and 1 ng of template. Use the following cycling conditions: 98 °C for 30 sec, 30 cycles of 98 °C for 7 sec and 68 °C for 2 min, and a final extension at 72 °C for 10 min.
   2. PCR the genes of interest for insertion to the linearized plasmids containing RN and RC fragments. Perform the PCR as in step 1.2.1 with the extension time at 68 °C. Use primers with overhangs that contain a flexible linker sequence, such as GGAGGTGGAGGTAGTGGTGGAGGTGGAAGT for (GGGGS)x2, and 15 base pair homology to the ends of the linearized RN and RC plasmids for recombination.
      Note: The primer overhangs for the proteins of interest are presented in Table 2 if using the primers in Table 1 to linearize the plasmid backbone.
   3. Digest the PCR reaction with Dpn1 to eliminate the PCR template. Add 0.5 µl of Dpn1 (20 U/µl) directly to the 50 µl of the PCR reaction. Incubate at 37 °C for 1 hr and heat-inactivate at 80 °C for 20 min. If the background template persists in later steps, increase the enzyme amount or lengthen the digestion time.
   4. Purify the PCR products via a spin column as described by the manufacturer.
   5. Perform a ligation-independent reaction to combine a linearized plasmid containing an RN or RC fragment with the gene of interest. Measure the concentration of the purified PCR products: combine 50 ng of linearized plasmid and 50 ng of the gene of interest with 1 µl of enzyme premix and bring the total volume to 5 µl with deionized water. Incubate at 50 °C for 15 min, place on ice, and add 20 µl of TE buffer.
   6. Transform recombinant plasmids into competent bacteria.
   7. Select 2-4+ (as desired) colonies for O/N culture. Add 5 ml of LB broth and 5 µl of kanamycin (50 mg/ml stock) into a 14 ml polypropylene round-bottom tube, and add the selected bacteria colony with a sterilized inoculating loop. Incubate at 37 °C O/N while shaking at 225 rpm.
   8. Freeze 1 ml of O/N culture for glycerol stock and miniprep the remainder as described by the manufacturer.
   9. Perform sequencing to determine a good clone. If using the cloning plasmids in the Materials List, use M13 forward and reverse primers (in separate reactions) as necessary to obtain the complete sequence of the fusion construct. Compare with the expected sequence using an alignment tool such as BLAST from the National Center for Biotechnology Information (NCBI) – http:// blast.ncbi.nlm.nih.gov/BlastAlign.cgi.
   10. Transfer the sequence-verified constructs to an expression plasmid via a recombinase reaction.
       1. Add 75 ng of the destination plasmid—such as pcDNA for transient transfections or pLenti for lentiviral packaging—and 50 ng of the recombinant cloning plasmid to 1 µl of enzyme premix and bring up the total volume to 5 µl with deionized water. Incubate at 25 °C for 1 hr, then add 5 µl of TE buffer.
          Note: For lentiviral packaging and stable cell line generation, use a different destination plasmid for each construct, for example, one with puromycin resistance and the other with neomycin. This allows for a double selection of transduced cells. Also consider the promoter for each, such as the constitutive cytomegalovirus (CMV) promoter for high expression levels, or CMV with the TetO operator for tunable doxycycline-regulated expression.
       2. Transform 5 µl into competent bacteria and miniprep the plasmids as in steps 1.2.6 to 1.2.8, but using the appropriate antibiotic (typically ampicillin).
3. Determine the optimal BiFC configuration.
   1. Co-transfect expression plasmids into U2OS cells (or cell of choice).
      1. Add 350 µl of phenol red-free and antibiotic-free DMEM supplemented with 10% FBS into each well of an 8-well #1.5 glass bottom chamber slide. Plate about 7.5 x 10⁴ cells per well so cells are about 70%-90% confluent the next day.
      2. The day after plating, co-transfect expression plasmids with different configurations in each well using the preferred reagent and as described by the manufacturer.
         1. For each well, add 125 ng of each expression plasmid into 50 µl of serum-free reduced media in a 0.6 ml tube and pipette up and down to mix. Add 1 µl of the transfection reagent down the center of the tube and directly into the media. Agitate gently to mix and let the reaction sit for 30 min.
         2. Reduce the media in each well of the chamber slide by about half. Add the transfection mixture dropwise to the wells and shake gently to mix. Incubate cells at 37 °C and 5% CO₂ for 24-48 hr. Change the media the day after transfection.
   2. Image cells on a fluorescence microscope as in Protocol 2 beginning with step 2.1.4.
      Note: The sample may be imaged on a standard fluorescence microscope if the expression levels are high and the camera is sensitive enough for the relatively low photon output of PAmCherry1. Reconstituted PAmCherry1 molecules can be activated with an ultraviolet filter set (405 nm) prior to imaging with a green excitation (561 nm) filter set.
4. Where applicable, create a negative control by, for example, introducing a point mutation into one of the proteins of interest that disrupt the interaction. Perform site-directed mutagenesis on the cloning plasmid of the protein to be mutated and of the chosen configuration.
5. Generate a stable cell line by packaging the constructs into viral particles and infecting U2OS cells (or cell line of choice). Consider an inducible (tetracycline) expression system so the expression can be induced prior to imaging if prolonged irreversibility of BiFC is an issue, or if tunable expression is desired.
   CAUTION: Working with viruses is classified as Biosafety Level 2. While recent generations of viral packaging systems have greatly reduced the likelihood of producing replication-competent viruses, some risks are still present. References can be found at http://www.cdc.gov/ biosafety/publications/.
   1. Repeat step 1.2.10 using a lenti- or retroviral destination vector. Increase the O/N culture to 100 ml for a midiprep and greater purity of the plasmids.
   2. On Day 1: Plate about 2 x 10⁶ 293T/17 cells into a 10 cm dish for each construct to be packaged.
   3. On Day 2: Prepare a transfection reaction using a transfection reagent of choice and as described by the manufacturer.
      1. Prepare a premix of packaging plasmids with final concentrations of 250 ng/µl for packaging plasmids 1 and 2, and 100 ng/µl for packaging plasmid 3 in sterile water. Add 10 µl of the packing plasmid premix and 2.5 µg of the target plasmid to 1 ml of serum-free reduced media in a 5 ml polypropylene round-bottom tube and pipette up and down to mix. Add 20 µl of transfection reagent down the center of the tube and directly into the media.
         1. Agitate gently to mix and incubate at RT for 30 min. Remove about half the media from the cells and add the transfection mixture in a dropwise manner. Shake gently to mix and incubate at 37 °C and 5% CO₂. Incubate 10 ml of media per plate in a tube with the cap loosened for temperature and CO₂ equilibration.
   4. On Day 3: Gently change the media with the pre-incubated media and return the cells to the incubator. Incubate another tube of media with the cap loosened. Note: Syncytia, or the formation of large multi-nucleated cells, can be a sign that the packaging is going well. However, cells are in a fragile state and can be easily detached. When changing media, tilt the pipettor so that it is horizontal and add the media dropwise into one edge of the plate.
   5. On Day 4: Harvest the virus by removing the media with a syringe and filtering through a 0.45 µm pore size filter into a clean 50 ml tube. Gently replace the media with the pre-incubated media.
   6. On Day 5: Harvest again as done previously in step 1.5.3.3 and combine common filtrate into the same 50 ml tube. Add 6 ml of virus concentrator to each tube and mix. Store at 4 °C for at least 24 hr until the virus precipitates.
      Note: Depending on the health of the cells, the virus can be harvested a third time.
   7. Concentrate the virus by centrifuging at 1,500 x g for 15 min at 4 °C. Aspirate the supernatant and resuspend the pellet in 250 µl of media. Viruses can be used immediately for infection or stored in 50 µl aliquots at -80 °C for up to one year.
   8. Plate about 3 x 10⁵ U2OS (or target) cells per well into a 6-well plate (2 ml of DMEM with 10% FBS per well) for infection, so cells are about 50% confluent at the time of infection.
   9. The following day, remove about half the media so about 1 ml remains. Coinfect with 50 µl of concentrated virus for each construct. Add 1 µl of polybrene (8 mg/ml) to each well. Incubate at 37 °C and 5% CO₂. Change media the next day and incubate for one more day before antibiotic selection.
      Note: The amount of virus to add can vary depending on the desired rate of infection. Viruses can be titrated using qRT-PCR.
   10. Add antibiotics to select infected cells. Separate wells with uninfected cells that have not seen the virus are required as canaries to test the efficacy of the selection. Incubate for 2-7 days, or until selection is complete.
       Note: Effective concentrations should be titrated initially, but they are typically 1.0 µg/ml for puromycin and 1.0 mg/ml for neomycin.
   11. Expand the cells into a larger 10 cm dish and proceed to Protocol 2.

2. Imaging Fixed Cells

1. Prepare a sample for imaging
   1. Clean an 8-well #1.5 glass-bottom chamber slide by adding 250 µl of 1 M NaOH for at least 1 hr and wash thoroughly with PBS.
      Note: Untreated chamber slides typically result in high background signal. Additionally, cells may be sensitive to residual NaOH and failure to thoroughly clean the glass surface (as many as 10x washes) may make cell adhesion difficult. If necessary, incubate the cleaned chamber slide with PBS O/N after washing. For sensitive cell lines, plating at a higher density (e.g., at 50%-60% initial confluency) may help.
   2. Plate about 5.5 x 10⁴ of the U2OS stable expression cells per well in 350 µl of phenol red-free DMEM with 10% FBS so that cells are healthy and not over-confluent when imaging.
   3. Perform necessary treatments for the experiment, such as adding tetracycline to induce protein expression.
   4. Fix the cells immediately before imaging.
      1. Prior to fixation, prepare fresh paraformaldehyde (PFA) solution — 3.7% PFA in 1x PHEM with 0.1% glutaraldehyde (GA). For 10 ml PFA solution, weigh 0.37 g of PFA and transfer to a 1.5 ml microcentrifuge tube. Add 1 ml of distilled water and 30 µl of 1 M NaOH and vortex. Heat at 70 °C and vortex every 1-2 min until PFA is completely dissolved.
      2. Transfer dissolved PFA to a 15 ml conical tube and add 3.8 ml distilled water, 5 ml 2x PHEM buffer, 20 µl of 25% glutaraldehyde, and vortex to mix. Stock, 2x PHEM buffer is 120 mM PIPES, 50 mM HEPES, 20 mM EGTA, and 16 mM MgSO₄, with pH adjusted to 7.0 with 10 M KOH. This fixative solution can be stored at 4 °C for several days.
         CAUTION: PFA and GA are both hazardous. Prepare the solution in a fume hood and wear proper protective equipment when handling both chemicals. Avoid inhalation or direct skin contact.
      3. Remove the growth medium. Wash quickly with 500 µl of PBS and add 250 µl of the PFA fixative per well. Incubate the cells for 20 min at RT.
   5. Remove the PFA fixative from the sample and add 350 µl of PBS or imaging buffer (100 mM Tris with 30 mM NaCl and 20 mM MgCl₂, pH 8.5) per well.
   6. Vortex 100 nm gold particles to break up aggregates and add 35 µl per well (10x final dilution) for tracking stage drift during imaging.
2. Acquire images
   1. Turn on the microscope. Power on the 405 and 561 nm lasers, but keep the shutters (internal or external) closed at this point. Turn on the EMCCD camera and allow it to cool down. Ensure the 561 nm filters are in place.
   2. Open the acquisition software and set the exposure time to 100 msec and the EMCCD gain to 300 (range 1 – 1,000).
   3. Add immersion oil to the objective and secure the sample to the microscope stage.
   4. With either a bright field or the 561 nm laser on (~1 kW/cm² ), bring the sample into focus.
   5. For imaging Ras and other membrane proteins, use a 60X apochromat TIRF objective with a 1.49 numerical aperture and bring the microscope into TIRF configuration. Adjust the excitation laser so that it is off-centered when hitting the back aperture of the TIRF objective; this causes the laser to deflect upon reaching the sample. Keep adjusting the laser until the critical angle is reached and the laser is being reflected back. Search for a cell to image with several gold particles in view. Set a region of interest that encloses the cell (or a region within the cell) and the gold particles.
      ​Note: TIRF decreases the penetration depth of the excitation laser and reduces the out-of-focus background signal. Additionally, drift correction is likely to be more accurate when more gold particles are in view.
   6. If available, engage the autofocus system to correct for sample drift in the z-direction. Otherwise, adjust the focus manually throughout the acquisition if the image goes out of focus.
   7. Begin acquisition with the 405 nm laser off and the 561 nm laser on (~1 kW/cm²) in case sufficient activation is occurring already (typically if expression levels are high). Otherwise, turn on the 405 nm laser at the lowest factory power setting (0.02 mW or 0.01 W/cm²) and increase gradually (0.1 mW at a time) as necessary until there are several tens of molecules per frame or so that single molecules are well separated.
   8. As data acquisition continues, gradually increase 405 nm laser power to keep the spot density roughly constant. Note: At lower 405 nm excitation power, some PAmCherry1 can already be activated. As these molecules photobleach, the population of remaining photoactivatable PAmCherry1 molecules decreases, requiring a higher photon flux to maintain the same density of molecules emitting fluorescence in each frame.
   9. Continue image acquisition until high 405 nm power (2.5 – 10 W/cm²) does not activate more activation events.
      Note: The total acquisition time depends on the expression level and efficiency of the complementation.

Table 1. Primers for linearizing cloning plasmids containing PAmCherry1 fragments. The cloning plasmids containing RN and RC can be linearized with inverse PCR at the point of insertion at either the N- or C-terminus of the fragments. Sequences are listed 5' to 3'.

Table 2. Overhangs for gene of interest primers that match Table 1 primers.

The 15 base pairs of homology for the ligation-independent reaction is generated from primer overhangs. Additionally, the overhangs include the sequence for a (GGGGS)x2 flexible linker. The flexible linker sequence can be added to the primers in Table 1 instead. Sequences are listed 5' to 3'.