1. Cell Culture

Use sterile culture tubes and pipette tips. The E. coli strain AB1157 polA-PAmCherry carries a C-terminal PAmCherry fusion of Pol1. The fusion was inserted at the native chromosomal location by replacing the wild-type gene using lambda-Red recombination as described in Datsenko et al.⁸ Functionality of the fusion protein was confirmed as judged by cellular growth rates and sensitivity to the DNA damaging agent methyl methanesulfonate (MMS). More information on construction of the cell strain can be found in Uphoff et al.⁷, Datsenko et al.⁸, and Reyes-Lamothe et al.⁹ Cell cultures are grown in M9 minimal medium to reduce autofluorescence and avoid background particles on the microscope slide. Alternatively, a nutrient-rich defined medium can be used¹⁰.

1. Streak the E. coli strain AB1157 polA-PAmCherry from a frozen glycerol stock on a Luria Broth (LB) agarose plate with selective antibiotics (here, 25 µg/ml kanamycin) and incubate at 37 °C overnight.
2. Inoculate a 5 ml LB culture from a single cell colony and grow at 37 °C shaking at 220 rpm for 3 hr.
3. Dilute the culture 1:10,000 into 5 ml minimal medium (M9 medium, MEM amino acids + proline, MEM vitamins, 0.2% glycerol) and incubate at 37 °C shaking at 220 rpm overnight.
4. The following morning, measure the optical density (OD) using a spectrophotometer and dilute the culture in 5 ml fresh minimal medium to OD 0.025. Grow for 2 hr at 37 °C shaking at 220 rpm to early exponential phase (OD 0.1).
5. Concentrate 1 ml of cells in a 1.5 ml microcentrifuge tube by centrifugation at 2,300 x g for 5 min. Remove the supernatant and resuspend the cell pellet in 20 µl residual medium and vortex.

2. Microscope Slide Preparation

1. Prepare a 1.5% low-fluorescence agarose solution in dH₂O. Use a microwave to melt the agarose until the solution is clear. Mix 500 µl of the melted agarose solution with 500 µl of 2x minimal medium by gently pipetting up and down a few times.
2. Spread the agarose solution evenly on the center of a microscope coverslip (No 1.5 thickness). This has to be done quickly before the agarose cools, avoiding bubbles.
3. Flatten the pad with a second coverslip (No 1.5 thickness). To remove background fluorescent particles, coverslips were previously burned in a furnace at 500 °C for 1 hr. Burned coverslips can be stored for weeks at room temperature covered in aluminum foil.
4. For DNA damage experiments, prepare an agarose pad containing 100 mM MMS. Follow the procedure in steps 2.1-2.3, but add 8.3 µl MMS to 500 µl of M9 medium before mixing with 500 µl of melted 1.5% agarose, for a final concentration of 100 mM MMS. (Caution! MMS is toxic and mutagenic and must be handled with gloves, mask, goggles, and lab coat).
5. Remove the top slide from the pad and add 1 µl of concentrated cell suspension onto the pad. Immobilize the cells by covering the pad with an unused burned coverslip (No 1.5 thickness, matching the microscope objective specification) and by pressing very gently on the slide. Cells should be imaged within 45 min of immobilization before the agarose pad dries. To prevent drying during longer experiments, agarose pads can be sealed using silicon gaskets.
6. For DNA-damage experiments, incubate cells immobilized on the agarose pad containing 100 mM MMS for 20 min in a humidified container at room temperature before imaging.

3. Preparing Microscopy Data Acquisition

PALM relies on the detection and precise localization of single fluorescent proteins. The sensitivity and optimal alignment of the microscope is critical for the data quality. Single-molecule fluorescence microscopes typically employ Total Internal Reflection (TIR) illumination to enhance the signal-to-noise ratio by exciting only fluorophores within a thin section above the coverslip surface. Here, imaging inside E. coli requires highly inclined illumination¹¹, which can be achieved on a TIRF microscope by slightly decreasing the angle of the excitation light. PAmCherry imaging further requires a 405 nm photoactivation laser and a 561 nm excitation laser. The fluorescence emission is recorded on an electron multiplying CCD (EMCCD) camera at a magnification resulting in a pixel length of 114.5 nm/pixel. For optimal localization precision, the pixel size should approximately match the standard deviation width of the PSF to ensure sufficient sampling without spreading the signal over too many pixels. Figure 3 shows a schematic of a minimal PALM setup. Movie 1 gives an impression of the bespoke microscope building process; see Uphoff et al.⁷ for a detailed description of the instrument.

1. Performing routine microscope alignment. Measure the 405 nm and 561 nm continuous wave laser intensities in front of the objective. Adjust the 561 nm intensity to 3.5 mW (~400 W/cm²) and 405 nm intensity to 10 µW (~1 W/cm²). Use a continuously variable neutral density filter wheel that allows gradual adjustment of 405 nm intensity from 0-10 µW. Switch off the laser illumination until the start of the experiment.
2. Place the sample on the microscope stage and bring the cells into focus in transmitted light microscopy mode (Figure 4A). The EMCCD camera gain has to be switched off to prevent damage to the camera by overexposure.
3. Define a cropped FOV to reduce data size and increase the camera read-out speed.
4. Cover the sample from ambient light and switch on the EMCCD camera gain.
5. Set the frame rate to 15.26 msec/frame (including 0.26 msec camera readout time). See “Exposure time and excitation intensities” in the Discussion section.
6. Display the camera data to check the dark background signal (Figure 4B).
7. Switch on the 561 nm laser and check the excitation background signal (Figure 4C).
8. Switch on the 405 nm laser for photoactivation of the Pol1-PAmCherry fusion proteins and increase the intensity until fluorescence PSFs appear.
9. Adjust the angle of the excitation beam to illuminate only a thin section of the sample close to the coverslip surface.
   1. To this end, the laser beam is focused into the back focal plane of a 100X NA 1.4 objective (Figure 3). Translating the focusing lens perpendicular to the beam moves the focus away from the center of the objective causing the beam to exit the objective under an angle.
   2. Aim to maximize the fluorescence intensity and minimize the background signal. Note that strict TIR excitation is optimal to image fluorophores within 100 nm of the coverslip surface, however, imaging DNA-binding proteins associated with the E. coli nucleoid requires deeper illumination up to 0.8 µm.

4. Data Acquisition

Here, we describe the general protocol for acquisition of a PALM movie. The same procedure applies for imaging Pol1-PAmCherry fusion proteins in undamaged E. coli cells and under continuous DNA damage treatment with MMS. Application of the method to fusion proteins of different molecular weight or copy number per cell will require different acquisition settings (see Discussion section).

1. Find a new field of view (FOV) of cells in transmitted light microscopy mode and focus the image. Take a camera snapshot to record the cell outlines (Figure 4A).
2. Cover the sample from ambient light and switch on the EMCCD camera gain.
3. Switch on the 561 nm laser and bleach the cellular autofluorescence and background spots on the coverslip for a few seconds before starting data acquisition. For cells grown and imaged in M9 medium and using burned coverslips there is usually very little fluorescence background; however, prebleaching could be useful for imaging cells in a rich growth medium such as LB. Note that intense illumination is toxic to cells so prebleaching should be kept to a minimum.
4. Start the acquisition of a PALM movie under continuous 561 nm excitation at 15.26 msec/frame.
5. Switch on the 405 nm laser and gradually increase the intensity over the course of the movie, reaching up to 1 W/cm². Avoid higher 405 nm intensities that cause cellular autofluorescence. Pay attention to the density of fluorescent molecules – it is important to keep activation rates low such that PSFs are clearly isolated in each frame (Figures 4D-F).
6. Record 10,000 frames/movie (depending on the number of molecules to be imaged per cell); one movie typically takes 2-3 min and requires 0.5-1 GB of hard disk space depending on the size of the FOV.
7. Repeat the acquisition procedure for multiple FOV. Note that each FOV can only be imaged once because PAmCherry fluorophores get photoactivated and bleached irreversibly.

5. Data Analysis

An automated and robust data analysis framework is essential for the performance and efficiency of the method. We use custom software written in MATLAB.

1. Perform the localization analysis using algorithms described in Crocker et al.¹², Holden et al.¹³, HoldenI et al.¹⁴, and Wieser et al.¹⁵ PSFs are first identified in a band-pass filtered image using a Gaussian kernel with 7 pixels diameter (Figure 5A). Candidate positions correspond to PSFs with peak pixel intensities above 4.5 times the standard deviation of the background signal (Figure 5B). The locally brightest pixel per candidate PSF serves as initial guess for fitting an elliptical Gaussian function (Figure 5C). The free fit parameters are: x-position, y-position, x-width, y-width, rotation angle, amplitude, and background offset. The elliptical Gaussian mask accounts for molecule during the exposure time, which blurs and deforms the PSF.
2. Plot the resulting (x, y) localizations from all frames of the PALM movie onto the transmitted light microscopy image of the same FOV. Localizations of Pol1-PAmCherry should appear within the central area of E. coli cells (Figure 6A). If many localizations appear outside of cells, the localization threshold was set too low or the sample contained background fluorescent particles.
3. For automated tracking analysis, the MATLAB implementation of the algorithm described in Crocker et al.¹² can be used (see “Diffusion analysis” in the Discussion section). Positions that appear in subsequent frames within a user-defined tracking window are connected to form a trajectory. In the case that multiple localizations occur in the same window, tracks are uniquely assigned by minimizing the sum of step lengths. For a detailed discussion of the various considerations when calculating diffusion coefficients from single-particle tracking data, see Wieser et al.¹⁵
   1. The algorithm uses a memory parameter to account for transient blinking or missed localizations during a track. Here, we set the memory parameter to 1 frame; higher values can be used for tracking fluorophores with long-lived dark states.
   2. Choose a suitable tracking window based on the following calibration steps. For Pol1, we use 0.57 µm (5 pixels).
   3. Run the tracking algorithm for a range of tracking window parameters. Calculate the number of measured tracks per cell as a function of the tracking window to identify the smallest possible tracking window that does not split tracks (Figure 6B).
   4. Plot the resulting tracks on the transmitted light microscopy image of the same FOV to visualize the spatial distribution of molecule movement within cells. Pol1 tracks should display diffusion confined within single cells (Figures 6C-D).
   5. If a fraction of tracks appears to cross between cells this suggests that separate molecules were erroneously linked because the tracking window was chosen too large and/or the photoactivation rate was too high (Figure 6E).
   6. Plot the cumulative distribution of the step lengths between consecutive localizations (Figure 6F). The curve rises and saturates smoothly for sufficiently large tracking windows but shows a cutoff edge if the window was chosen too small.
4. To analyze the diffusion characteristics of Pol1, compute the mean-squared displacement (MSD) between consecutive localizations for each track with a total of N steps):
   MSD = 1/(N-1) ∑ _{i = 1} ^N-1 (x_i+1 – x_i)² + (y_i+1 – y_i)² .
   Include only tracks with at least 4 steps (N ≥ 5 localizations) to reduce the statistical uncertainty in the MSD values.
5. Plot a curve of MSD values over a range of lag times by calculating displacements over multiple frames (Figure 6G). The shape of the MSD curve can help to classify the observed molecular motion (Figure 6H).
6. Calculate the apparent diffusion coefficient D* per track from the MSD:
   D* = MSD/(4 Δt) – σ_loc²/Δt .
   The second term corrects for the estimated localization error (here, σ_loc = 40 nm and Δt = 15.26 msec, see Wieser et al.¹⁵).
7. Plot a histogram of the measured D* values from all tracks in the FOV (Figure 7A).
8. Identify individual Pol1 molecules that appear bound to the chromosome based on the measured D* value per track. Separate the populations of bound (sharp distribution centered at D* ~ 0 µm²/sec) and freely diffusing molecules (broad distribution centered at D* ~ 0.9 µm²/sec) by setting a threshold D* < 0.15 µm²/sec (red bars in Figures 7A and 7D).
9. Carry out the localization, tracking, and diffusion analysis for Pol1 in undamaged cells (Figures 7A-C) and in cells under DNA-damage treatment with MMS (Figures 7D-E). The fraction of bound tracks provides a direct quantitative measure of the DNA repair activity of Pol1 in vivo.