Population and Single-Cell Analysis of Antibiotic Persistence in Escherichia coli

NOTE: Use sterile culture glassware, pipette tips, and growth medium. Here, E. coli cells were grown in a low-autofluorescence chemically defined medium (see Table of Materials). Inoculations were performed in the presence of a Bunsen burner to minimize the risk of contamination.

1. Cell culture and growth curve

1. Streak the strain of interest from a frozen glycerol stock on a Luria-Bertaini (LB) agar plate (supplemented with a selective antibiotic, if required), and incubate at 37 °C overnight (between 15 h and 19 h) to obtain single colonies.
   NOTE: The experiment presented here uses two strains, E. coli K-12 MG1655 corresponding to the wt strain and the isogenic MG1655 hupA-mCherry strain²². The latter strain expresses the fluorescence-tagged α-subunit of the HU nucleoid-associated protein. The hupA-mCherry reporter is integrated at the native locus of hupA. HU-mCherry serves as a proxy to follow the nucleoid dynamics in live cells as it binds to DNA in a non-specific manner.
2. Inoculate 5 mL of medium (here, 3-[N-morpholino] propanesulfonic acid-based medium [MOPS]; Table 1, Table 2, and Table 3) supplemented with glucose 0.4% and a selective antibiotic (if required) with an isolated colony in a glass tube (≥25 mL), and place the tube in a shaking incubator set to 37 °C and 180 rotations per minute (rpm) overnight (between 15 h and 19 h). Alternatively, plastic tubes, glass, or plastic flasks (≥ 25 mL) can be used instead of glass tubes.
   NOTE: MOPS medium supplemented with glycerol at a final concentration of 0.4% was used throughout the experiments described in this paper, except for the overnight cultures which were performed in MOPS glucose 0.4%. The generation time of E. coli in MOPS supplemented with glucose is shorter than that in MOPS glycerol. Using MOPS glucose 0.4% instead of MOPS glycerol 0.4% at this step ensures that the cells reach the stationary phase within 19 h. Other growth media, such as M9 or rich defined medium (RDM) supplemented with distinct carbon sources, can also be used. It should, however, be noted that the growth rate and the persistence frequency are different depending on the medium and/or the carbon used²³.
3. The following morning, centrifuge 1 mL of culture at 2,300 x g for 3 min, discard the supernatant, and gently resuspend the pellet in the same volume of phosphate-buffered saline (PBS). Measure the optical density at 600 nm (OD_{600 nm}), and calculate the volume needed for an initial OD_{600 nm} of 0.01 in a final volume of 2 mL.
4. Place 2 mL of MOPS glycerol 0.4% medium into a well of a clear bottom 24-well plate, and inoculate with the calculated overnight culture volume. Place the 24-well plate in an automated microplate reader (see Table of Materials) to monitor the OD_{600 nm} for 24 h. Set the microplate reader to measure the OD_{600 nm} every 15 min at a temperature of 37 °C and with high orbital rotation (140 rpm).
   ​NOTE: If the strain used in the experiment encodes a fluorescent reporter, make sure that its growth rate is comparable to that of the wt to avoid any artifact in the subsequent experiments, as the growth rate affects the antibiotic persistence frequency²³. Due to detection limitations at very low cell densities and the potential strain-specific effects on the OD_{600 nm}/colony forming units (CFU) relationship, it is recommended to evaluate the growth kinetics by monitoring the CFU·mL^-1 when working with uncharacterized strains.

2. Determination of the minimal inhibitory concentration of the antibiotics

NOTE: The minimum inhibitory concentration (MIC) is defined as the lowest dose of antibiotic at which no bacterial growth is observed. The determination of the MIC needs to be performed for each antibiotic and strain. In the experiments described here, the fluoroquinolone antibiotic ofloxacin (OFX) was used. The determination of the MIC allows for confirmation that the antibiotic solution has been correctly prepared, the antibiotic is active, and the strains are equally sensitive to the antibiotic. Here, the published agar dilution method was performed to determine the MIC to OFX of the different strains used²⁴. The MIC of a given antibiotic to a given bacterial strain can also be determined via the broth dilution method²⁴.

1. Preparation of the plates for MIC determination
   1. Prepare a master stock solution for the antibiotic used in the experiments by dissolving 5 mg of OFX in 1 mL of ultrapure water. Add 20 µL of 37% HCl to increase the solubility of the OFX.
   2. Melt 100 mL of sterile LB agar, and keep it at 55 °C to avoid solidification. Prepare six small glass flasks (25 mL), and pipet 5 mL of liquid LB agar medium into each flask using a sterile pipette.
   3. Dilute 10 μL of the 5 mg·mL^-1 OFX master stock solution in 90 μL of ultrapure water (1:10 dilution of the master stock).Add 0 μL, 2 μL, 4 μL, 6 μL, 8 μL, or 10 μL of the 500 μg·mL^-1 OFX solution to each of the six glass flasks containing 5 mL of LB agar medium to generate LB agar medium with final concentrations of 0 μg·mL⁻¹, 0.02 μg·mL⁻¹, 0.04 μg·mL⁻¹, 0.06 μg·mL⁻¹, 0.08 μg·mL⁻¹, and 0.1 μg·mL^-1 OFX, respectively. Mix the solution by rotating the flasks several times.
      NOTE: If the MIC of the antibiotic of interest is known, the range of concentrations should reach from below to above the actual MIC. If the MIC is unknown, a large range of concentrations with a log₂ dilution series is recommended. Make sure that the LB agar medium has cooled down before adding the antibiotic, as high temperatures may inactivate it. Nevertheless, it is important not to let the LB agar medium solidify before adding the antibiotic, as this could lead to the non-homogenous distribution of the antibiotic in the LB agar medium.
   4. Pour 5 mL of each of the six LB agar media prepared in step 2.1.3 in a dose-increasing manner into a 6-well culture plate using a sterile pipette. Let the agar cool down until solidification, and dry the plate before using.
      NOTE: Prepare the antibiotic solution, and the culture plate the day the assay is performed.
2. MIC determination assay
   1. Inoculate 5 mL of LB medium with an isolated colony in a glass tube (≥25 mL), and place the glass tube in a shaking incubator set to 37 °C and 180 rpm overnight (between 15 h and 19 h).
   2. The following morning, measure the OD_{600 nm}, and dilute the culture into a test tube to a final cell density of 1 x 10⁷ CFU·mL^-1 in PBS. For the strains tested here, 1 x 10⁷ CFU·mL⁻¹ corresponds to an OD_{600 nm} of 0.0125.
      NOTE: If the correlation between the CFU·mL^-1 and OD_{600 nm} is not known, the growth curve determined by CFU and OD_{600 nm} measurements should be established to calculate the correlation factor between the CFU·mL^-1 and OD_{600 nm}.
   3. Spot 2 μL of the previously diluted culture onto every well of the dried 6-well plate. Let the spots dry before placing the plate into an incubator at 37 °C overnight (between 15 h and 19 h).
   4. The next day, count the colonies formed in each well. The MIC corresponds to the well with the minimum concentration of antibiotic where no bacterial growth is detected.

3. Spot assay

NOTE: The spot assay method is a qualitative approach that allows the estimation of the number of viable cells (cells able to generate colonies after antibiotic stress). The spot assay is performed prior to the time-kill assay to provide insights into the viability of the strain used in the conditions tested and to inform about the dilutions needed during the time-kill assay (see section 4).

1. To prepare the LB agar plates for spot assays, pour 50 mL of LB agar into a square Petri dish (144 cm²). Prepare one square Petri dish per time point. Let the LB agar solidify, and dry the plates before use.
2. Inoculate 5 mL of medium (MOPS glucose 0.4%) with an isolated colony in a glass tube (≥25 mL), and place the tube in a shaking incubator set to 37 °C and 180 rpm overnight (between 15 h and 19 h).
3. The next day, measure the OD_{600 nm}, and dilute the culture into fresh temperature-adjusted medium (37 °C, MOPS glycerol 0.4%) in a glass tube (≥25 mL) to a final OD_{600 nm} of ~0.001. Let the culture grow overnight in an incubator at 37 °C at 180 rpm (between 15 h and 19 h).
4. On the following day, measure the OD_{600 nm}, and incubate the culture to a final OD_{600 nm} of 0.3.
5. While incubating, prepare 96-well plates by placing 90 μL of 0.01 M MgSO₄ solution in every well except the wells of the first row (row A). The 96-well plates will be used for 10-fold serial dilution in step 3.7.
   NOTE: In this experiment, two strains (wt and hupA-mCherry strain) were tested in triplicate at seven different time points. As one plate comprises 12 columns, one plate can be used for two time points, and, thus, a total of four plates were prepared.
6. At an OD_{600 nm} of 0.3, withdraw 200 μL of each bacterial culture. These samples correspond to the t0 (untreated) timepoint prior to the antibiotic treatment and allow the determination of the CFU·mL^-1 before the antibiotic treatment.
7. Centrifuge the samples at 2,300 x g for 3 min. During the centrifugation time, add the desired concentration of OFX to the liquid culture, and continue to incubate at 37 °C while shaking.
   NOTE: Here, OFX was used at a concentration of 5 μg·mL^-1 (corresponding to the MIC multiplied 83-fold). In a previous study, this concentration was used to characterize the persistence phenomenon under OFX exposure²⁵. The antibiotic concentration used for the time/kill assays can vary depending on the antibiotic, the culture medium, and the state of bacterial growth.
8. After centrifugation, resuspend the cell pellet in 200 μL of 0.01 M MgSO₄ solution. Place 100 μL in the empty well of the 96-well plate prepared in step 3.5. Perform a dilution by transferring 10 μL of the well in row A into the well in row B containing 90 μL of 0.01 M MgSO₄. Continue the serial dilutions by transferring 10 μL of the well in row B into the well in row C. Repeat until reaching a dilution of 10⁻⁷ (with every transfer being a 10-fold dilution).
   NOTE: In this experiment, six cultures (three for the wt strain and three for the hupA-mCherry strain) were tested for every timepoint. According to the protocol, a muti-channel pipette is used to perform the serial dilutions for the six strains at the same time. To minimize the technical error, the pipette tips are changed between every transfer.
9. Spot 10 μL of each dilution onto the LB agar plates prepared in step 3.1.
   NOTE: A multi-channel pipette is used to simultaneously spot the same dilution (e.g., a 10⁻⁴ dilution) for the six cultures. During the spotting, the first stop of the multi-channel pipette should be used without pushing the second stop, as this can result in microdroplets being dispensed onto the plate.
10. At relevant time points after the antibiotic addition, withdraw 200 μL of the culture, and perform serial dilutions as described in step 3.8. Spot 10 μL of each dilution as described in step 3.9.
    NOTE: For the experiment described here, seven time points were collected (t0, t1h, t2h, t3h, t4h, t5h, t6h). For strains exhibiting increased antibiotic susceptibility as compared to the wt, multiple washes in MgSO₄ 0.01 M can be performed to remove residual antibiotics.
11. Incubate the plates at 37 °C overnight (between 15 h and 19 h). The next day, count the number of colonies at the two highest dilutions for which colonies can be detected. Ideally, the spots on the agar plates for these dilutions contain between 3 to 30 colonies allowing the accurate determination of the CFU·mL^-1 for each sample.
12. Calculate the survival ratio by dividing the calculated CFU·mL^-1 of each timepoint by the CFU·mL^-1 of the initial population at t0.

4. Time-kill assays

NOTE: While spot assays are an easy-to-use method to estimate the survival rate of a given strain for a given antibiotic, time-kill assays give a higher-resolution survival rate and are performed to accurately quantify the bacterial viability. The profile of the killing curve can be used to determine whether a given bacterial strain is sensitive, tolerant, or resistant to the antibiotic in a given condition. Moreover, time-kill assays allow the determination of the time of antibiotic exposure needed to detect the persistence phenomenon (beginning of the second slope of the biphasic killing curve) as well as the persistence frequency.

1. Prepare LB agar plates for the time-kill plating assay. Pour 25 mL of LB agar into a Petri dish (±57 cm²). Prepare at least two Petri dishes per timepoint (two dilutions per timepoint per strain are plated).
2. Let the LB agar solidify, and dry the plates before adding five to eight sterile glass beads to each plate. Invert and label the plates according to the strain/condition/timepoint.
   NOTE: Glass beads allow the spreading of bacteria on the agar plates during step 4.7 and step 4.9. Alternatively, the cells can be dispersed on the agar plate using a spreader.
3. For each sample, prepare 10-fold serial dilution glass tubes containing 900 µL of 0.01 M MgSO₄ solution. The number of dilution glass tubes per sample that needs to be prepared corresponds to the dilutions needed to detect colonies in the spot assay.
4. Inoculate 5 mL of medium (MOPS glucose 0.4%) with an isolated colony in a glass tube (≥25 mL), and place the tube in a shaking incubator set to 37 °C and 180 rpm overnight (between 15 h and 19 h).
5. After 16 h of growth, measure the OD_{600 nm}, and dilute the culture into fresh temperature-adjusted medium (37 °C, MOPS glycerol 0.4%) in a glass tube to a final OD_{600 nm} of ~0.001. Let the culture grow overnight in an incubator at 37 °C at 180 rpm (between 15 h and 19 h).
6. On the following day, measure the OD_{600 nm}, and incubate the culture to a final OD_{600 nm} of 0.3.
7. At an OD_{600 nm} of 0.3, withdraw 100 μL of the culture, dilute according to the data obtained in the spot assay (at an OD_{600 nm} of 0.3 and without antibiotic, a 10⁻⁵ dilution usually gives 200-300 colonies), and plate 100 µL on the LB agar plates prepared in steps 4.1-4.2. Gently shake the plates to avoid the beads contacting the dish edges, as this can lead to a non-homogenous spreading of the bacterial cells on the LB agar medium. This first sample prior to the antibiotic treatment corresponds to the t0 time point (CFU·mL^-1 before antibiotic treatment).
8. Add the desired concentration of OFX, and continue to incubate at 37 °C while shaking.
   NOTE: Here, OFX was used at a concentration of 5 μg·mL^-1 (corresponding to the MIC multiplied 83-fold).
9. At relevant time points after the antibiotic addition, withdraw 100 μL of the culture, dilute according to the data obtained in the spot assay, and plate 100 µL on the LB agar plates prepared in steps 4.1-4.2. Plate the cells as described in step 4.7.
   NOTE: For the experiment described here, seven time points were collected (t0, t1h, t2h, t3h, t4h, t5h, t6h). For strains exhibiting an increased antibiotic susceptibility as compared to the wt, multiple washes in MgSO₄ 0.01 M can be performed to remove residual antibiotics.
10. Incubate the plates at 37 °C overnight (between 15 h and 19 h). The next day, count the number of colonies at the two highest dilutions for which colonies can be detected. Ideally, the plates should contain 30-300 colonies to allow an accurate determination of the CFU·mL^-1 in each sample.
11. Calculate the survival ratio by normalizing the CFU·mL^-1 at each time-point by the CFU·mL^-1 at t0. Plot the log₁₀ normalized CFU·mL^-1 as a function of time.

5. Microfluidic time-lapse microscopy imaging

NOTE: The following section describes the preparation of the microfluidic plate as well as the time-lapse image acquisition and image analysis procedure. The aim of this experiment is to observe and analyze the persistence phenotype upon antibiotic treatment at the single-cell level. The data collected during this experiment can be used to generate a wide range of results depending on the question addressed and/or the fluorescent reporters used during the experiment. In the experiment described here, quantitative analysis of the cell length and HU-mCherry fluorescence²², reflecting the nucleoid organization in persister and non-persister cells, was carried out.

1. Bacterial cell culture for microfluidic time-lapse microscopy
   1. Inoculate 5 mL of medium (MOPS glycerol 0.4%, supplemented with a selective antibiotic if required) with an isolated colony in a glass tube (≥25 mL), and place the tube in a shaking incubator set to 37 °C and 180 rpm overnight (between 15 h and 19 h).
   2. The next day, measure the OD_{600 nm}, and dilute the culture in fresh temperature-adjusted medium (37 °C, MOPS glycerol 0.4%) in a glass tube to a final OD_{600 nm} of ~0.001. Let the culture grow overnight (between 15 h and 19 h) in a shaking incubator at 37 °C and 180 rpm to obtain an early exponential-phase culture on the next day.
2. Preparation of the microfluidic plate and time-lapse microscopy imaging
   NOTE: Microfluidic experiments can be performed in commercially available microfluidic devices (as described here) or in in-house produced microfluidic systems.
   1. Remove the conservation solution (if present) from every well of the microfluidic plate, and replace it with a fresh culture medium.
      NOTE: If the microfluidic plate contains a waste outlet well, the conservation solution of the outlet well should be removed but not replaced by the medium.
   2. Seal the microfluidic plate with the manifold system by clicking on the Seal button or through the microfluidic software (first select Tool, followed by Seal Plate).
      NOTE: To seal the plate, a uniform pressure should be applied to the plate and the manifold by manually squeezing the plate against the manifold. If performed correctly, the note "sealed" should appear on the ONIX2 interface. It is important to not apply any pressure onto the glass slide to avoid any potential risk of breaking the manifold.
   3. Once sealed, perform a first priming sequence (click on Run Liquid Priming Sequence on the microfluidic software interface).
      NOTE: Run Liquid Priming Sequence corresponds to 5 min of perfusion at 6.9 kPa for wells 1-5, followed by 5 min of perfusion at 6.9 kPa for well 8, and a final perfusion round for well 6 for 5 min at 6.9 kPa. The Run Liquid Priming Sequence allows the removal of the conservation solution that may still be present in the channels connecting the different wells.
   4. Incubate the plate in a thermostatically controlled cabinet of the microscope at the desired temperature (here, 37 °C) for a minimum of 2 h before the start of the microscopy imaging.
   5. Start a second Run Liquid Priming Sequence before beginning the experiment.
   6. Seal off the microfluidic plate by clicking on Seal off on the microfluidic software interface. Replace the medium in well 1 and well 2 with 200 μL of fresh medium, in well 3 with 200 μL of fresh medium containing the antibiotic (here, OFX at 5 μg·mL⁻¹), in well 4 and well 5 with 200 μL of fresh medium, in well 6 with 200 µL of fresh medium, and in well 8 with 200 μL of the culture sample (from step 5.1.2) diluted to an OD_{600 nm} of 0.01 in the fresh medium.
   7. Seal the microfluidic plate as described in step 5.2.2, and place the plate onto the microscope objective inside the microscope cabinet.
      NOTE: Make sure to place a drop of immersion oil onto the microscope objective before placing the microfluidic plate.
   8. In the microfluidic software, click on Cell Loading to allow the cell loading into the microfluidic plate.
      NOTE: The Cell Loading step comprises 15 s of perfusion at 13.8 kPa for well 8, followed by 15 s of perfusion at 27.6 kPa for well 6 and well 8, and a final perfusion round for well 6 for 30 s at 6.9 kPa. The density of the cells in the microfluidic plate is critical for the experiment. The first part of this microfluidic protocol consists of growing bacteria for 6 h in a fresh medium before the antibiotic treatment. After 6 h of growth, the cell density has to be sufficient to detect the rare persister cells (in the conditions used in this study, the persister cells generate at a frequency of 10⁻⁴). If the cell density is too high, it is difficult to distinguish the individual cells, which prevents precise single-cell analysis. As the growth rate is directly dependent on the medium, the density of cells in the microscopy fields should be assessed before launching the experiment.
   9. Set an optimal focus using transmitted light mode, and select several regions of interest (ROIs) where an appropriate cell number is observed (up to 300 cells per field).
      NOTE: Select at least 40 ROIs to make sure that the rare persister cells are imaged.
   10. On the microfluidic software, click on Create a Protocol. Program the injection of fresh medium at 6.9 kPa for 6 h (well 1-2), followed by the injection of the medium containing the antibiotic at 6.9 kPa for 6 h (well 3), and, finally, the injection of fresh medium at 6.9 kPa for 20 h (wells 4-5).
       NOTE: A bacterial culture diluted to an OD_{600 nm} of 0.01 allows bacteria to grow in the microfluidic chamber for 6 h, ensuring that the cells are in the exponential growth phase. Depending on the number of cells introduced into the microfluidic device during the loading step (see step 5.2.8), the duration of the growth phase can be adapted to obtain up to 300 cells per ROI. As persistence is a rare phenomenon, increasing the number of cells per ROI improves the possibility of observing persister cells. The cell number should, however, not exceed 300 cells per ROI, as this makes single-cell analysis tedious.
   11. Perform microscopy imaging in time-lapse mode with one frame every 15 min using transmitted light and the excitation light source for the fluorescent reporter. Here, a 560 nm excitation light source for the mCherry signal was used (580 nm LED at 10% power with filter 00 [530-585 ex, 615LP em, Zeiss] and 100 ms exposure for mCherry). The Zeiss-compatible Zen3.2 software was used for cell imaging.
3. Image analysis
   NOTE: The opening and visualization of the microscopy images are performed with the open-source ImageJ/Fiji software (https://fiji.sc/)²⁶. The quantitative image analysis is performed using the open-source ImageJ/Fiji software and the free MicrobeJ plugin (https://microbej.com)²⁷. In this protocol, the MicrobeJ 5.13I(14) version was used.
   1. Open the ImageJ/Fiji software on the computer, and drag the hyperstack time-lapse microscopy images into the Fiji loading bar. Use Image > Color > Make Composite to fuse the different channels of the hyperstack. If the channels of the time-lapse experiment do not correspond to the desired color (e.g., the phase contrast is shown in red instead of gray), use Image > Color > Arrange Channels to apply the appropriate color to the channels.
   2. Open the MicrobeJ plugin, and detect the bacterial cells using the manual editing interface. Delete the automatically detected cells, and manually outline the persister cells of interest frame by frame.
      NOTE: Different settings can be used to automatically detect individual cells. Manual detection was used here as the analyzed persister cells form long filaments, which are rarely detected correctly using automatic detection.
   3. After detection, use the Result icon in the MicrobeJ manual editing interface to generate a ResultJ table. Save the ResultJ file, and use the ResultJ table to gain insights into different parameters of interest of the single-cell analysis. In the case of this protocol, the mean fluorescence of the HU-mCherry intensity, the cell length, and the cell area of individual cells were exported.