PROCEDURE
Analysis and selection of optimal probes ● TIMING 70 min
1ǀ Stain bacterial samples according to the protocol referred to in Step 3 of ‘Preparation of probed samples’. Apply 2.5 ml of each membrane permeable probe (SYTO 9, SYTO 13, SYTO 17, and SYTO 40) at received concentrations to the diluted pure cultures separately.
2ǀ Analyze samples stained with SYTO 9, SYTO 13, SYTO 17, and SYTO 40 by FCM. Use the emission detectors for cells stained by SYTO 9, SYTO 13, SYTO 17, and SYTO 40 as mentioned above.
▲ CRITICAL STEP The FCM parameters that should be assessed in order to determine optimal probes for each strain are: the degree of staining, good emission signals, low false signals (from debris), and optimum number of events of cells displayed in FCM histograms.
3ǀ To confirm that PI is the best choice for detecting dead cells (those with damaged membranes), compare the effectiveness of PI, SYTOX Blue, and 7-AAD. All of these stains are believed to detect cells with damaged membranes. With the same heat-treated samples (discussed below), different numbers of stained cells were observed microscopically and with the FCM (data not shown) with the four stains.
▲ CRITICAL STEP PI was ultimately chosen to identify and enumerate the dead cells because it consistently gave the highest cell counts (data not shown).
4ǀ Plate an aliquot of cells in late log phase after exposure to membrane impermeable probes and also cells in late log phase that have not been exposed to any of the membrane impermeable probes on respective standard nutrient agar.
▲ CRITICAL STEP This step ensures that exposure to PI, SYTOX Blue, and 7-AAD have not affected culturability.
5ǀ Assess the optimal incubation time of these three fluorescent probes through the aide of an epifluorescence microscope.
? TROUBLESHOOTING
▲ CRITICAL STEP The performance of a probe depends largely on the incubation time. Except P. syringae, other strains used in this study could be stained successfully within 20 min. We found that 60 min of incubation in the dark was optimal for these bacterial strains.
Analysis using Liquid Counting Beads
1ǀ Run liquid counting beads with bacterial cells and compare the number of events detected by FCM. Detailed procedures are discussed below.
2ǀ Determine the optimal voltage settings of the PMT of emission spectra for the liquid counting beads so that bead region is separated from the cell region in the side-scatter (SS) vs. respective emission spectra of each probe.
? TROUBLESHOOTING
▲ CRITICAL STEP The bead concentration may vary from lot-to-lot, and should be verified using the information provided by the manufacturer.
Confirmation to avoid cell clumping during staining ● TIMING 20 min
1ǀ Cells at concentrations greater than ~104/ml caused high abort rates in the FCM and concentrations less than ~ 104/ml resulted in interference with the electronic signal of the sheath fluid, which was subject to other false-positive signals.
2ǀ To ascertain that clumping did not occur, pure culture samples were vortexed for 2 min and diluted with the appropriate broth to obtain a cell number close to ~104/ml.
? TROUBLESHOOTING
▲ CRITICAL STEP Typically 108/ml of cells in the pure culture gave 0.8 OD600 which varies depending on the type of spectrophotometer. This OD600 value can be used in the dilution process to achieve ~104/ml of cells.
3ǀ A total of 10 to 15 ml of diluted stained cells were passed through a sterilized 0.20 mM pore-size and 25-mm-diameter hydrophilic membrane.
4ǀ The captured cells on the membrane were observed under the epifluorescence microscope, and cells were not clumped. These initial experiments confirmed that ~104 cells/ml resulted in an abort rate close to zero, which suggests good detection and minimal clumping of cells.
5ǀ A specific amount of liquid counting beads in appropriate broth dilution media and purified NP water were run and assessed separately to identify the effect of noise, which typically arises from the presence of debris in the growth media.
▲ CRITICAL STEP These control experiments were conducted to determine the voltage and threshold settings, which indicate the signal level required to distinguish cells from debris; these settings are important control parameters required to establish an accurate protocol.
Preparation of stained samples ● TIMING 65 min
1ǀ Vortex samples of pure cultures of different cells for 2 min.
2ǀ Dilute samples of pure cultures with appropriate broth to obtain a cell number close to 104/ml. Observe cells under the epifluorescence microscope to ensure that clumping has not occurred.
▲ CRITICAL STEP Determine the number of CFU/ml after dilution by plating 50µl in triplicate of serial dilutions on respective agar, followed by incubation at 37 °C for 20 h.
3ǀ Add 2.5µl of each fluorescent probe (SYTO 9, SYTO 13, SYTO 17, and SYTO 40) separately to 1 ml of diluted individual samples. Vortex samples for 30 sec and then incubate in the dark for 60 min.
▲ CRITICAL STEP Samples must be incubated for 60 min in the dark to ensure that they have been stained thoroughly. Prepare the probes immediately after thawing and avoid light exposure during thawing and staining processes.
Flow Cytometric Analysis of SYTO probed samples ● TIMING 10 min
1ǀ Complete area scaling and quality control adjustments (as listed in Equipment setup) prior to running samples. It is one of the most important steps for the accurate and unbiased measurements and detection of cells.
▲ CRITICAL STEP Quality control of the FCM must be done using the CST beads prior to starting the FCM measurement. Refer to Table S1 in the supplemental material for voltage settings, threshold parameters, and flow rates pertaining to each probe/cell combination. Fluorescence was used for thresholding. The FSC and SSC parameters were varied to set the populations inside the histogram. These settings are only for the strains and probes used in this study. These settings will be different and must be optimized when different cells and probes are analyzed. The FCM allows saving each setting and operating conditions for specific cells and probes.
? TROUBLESHOOTING
▲ CRITICAL STEP This study validated that cell numbers lower than 104/ml results in zero abort rate. Use this cell concentration and the settings in Table S1 as the starting parameters for different cells and probes which were not used in this study.
2ǀ Inject 1 ml of stained cells into the FCM without liquid counting beads
▲ CRITICAL STEP This step is used to optimize the settings and operating conditions each time before the stained cells are injected into the FCM.
3ǀ Transfer a known volume of stained cells into a sterilized tube, and then add the appropriate amount of liquid counting beads. Vortex this sample for 10 sec. Inject sample into the FCM without changing the settings and the operating conditions as indicated in Table S1.
4ǀ Analyze the number of events/sec in the acquired data set. If <500 events/ sec, add 25µl of beads; if >1000 events/sec add 50µl of beads.
▲ CRITICAL STEP If the number of events/sec goes above 1000, dilute stained cells 1 to 2-fold with the appropriate broth to keep the abort rate close to zero. Note the dilution factor which will be used to calculate the cell density at different physiological conditions.
5ǀ Gate (polynomial gate) the cells in forward (FW) vs. side scatter (SS) plots. To obtain the total number of events in the cell region creates a one-dimensional histogram gate in the histogram for cells stained with a specific probe and create another one-dimensional histogram gate for the liquid counting bead region and display in a histogram. The cells and the bead regions should be separated and will not overlap if the setting in Table S1 is followed and optimized.
▲ CRITICAL STEP Record the same number of total events for all conditions. We recorded 5000 events for each case and each run in the FCM. The recorded total events should be consistent from lot-to-lot injection of stained cells into the FCM. Record the numbers in the cell and bead regions using each gate that was created and as mentioned above in Step 5.
6ǀ Calculate the number of cells per unit volume by FCM using the following equation:
[# of events in cell region / # of events bead region] * [# of beads per test / test volume] * dilution factor .........................………….. (1)
▲ CRITICAL STEP Acquire the data from three or four independent experiments for each condition. Use the same accurate pipette filler for the same experiment for adding probes and beads into the diluted cultures. The accuracy of the FCM methods depends not only on the FCM parameters, but also on the pipette fillers and pipetting skills.
7ǀ Total cells (live and dead) were counted using the SYTO probes, and the dead cells were enumerated using PI. Live cells (culturable and nonculturable cells) were calculated by subtracting the number of PI-stained cells from that of SYTO-stained cells. The nonculturable cell numbers were estimated by subtracting the number of plate count (culturable) cells from that of live cells.
Heat Inactivation ● TIMING 6 h 20 min
1ǀ Dilute previously grown cultures in LB or King’s broth to a final concentration of 104 cell/ml.
2ǀ Transfer 1-ml of these diluted cells into separate 2-ml sterilized culturing tubes and place in a 72°C heat block containing nanopure water in each well.
▲ CRITICAL STEP Prepare the heat block at least 30 min prior to inserting the tubes containing the cells to ensure that the final temperature has been reached.
3ǀ Prepare multiple tubes for each diluted cell and for each heating condition. Submerge tubes in hot water for 5, 10, and then 15 min. Following heat exposure, bring cells to room temperature (in 5 to 6 min).
4ǀ Transfer two culture tubes of each treatment of each heat-exposed cells to an incubated shaker (same conditions used during pure culture of each type of cell) for 6 h. Cells that were or were not incubated after heat treatments and brought to room temperature were diluted and plated on the respective standard nutrient agars for culturable cell counts (CFU/ml) after performing appropriate dilutions using the respective broth.
Flow Cytometric Analysis of Heat Shocked Cells ● TIMING 70 min
1ǀ Add 2.5µl of propidium iodide (PI) at received concentration to 1-ml of heat exposed cells. As necessary, the PI-stained cells are diluted with the appropriate broth when the abort rates are greater than zero.
2ǀ Incubate the cells in the dark for 60 min.
▲ CRITICAL STEP P. syringae samples must be incubated for 60 min in the dark to ensure that they have been stained thoroughly. Other cells can be stained within 20 min.
3ǀ Inject the sample into the FCM
4ǀ Follow the same protocols for FCM analysis methods as used in the section entitled Flow cytometric analysis of SYTO probed samples to create two one-dimensional gates in the cell and bead regions in the respective histograms for each cell (either control or heat inactivated for different periods). Calculate the number of cells in each heat inactivation condition using equation 1.
Confirmation of gating protocols using cell sorting ● TIMING 20 min
1ǀ Prior to cell sorting, follow the initialization procedures as discussed in the section entitled Equipment Setup.
2ǀ The histograms produced by the optimized methods typically were symmetric with small shoulders. To evaluate the minor populations represented in the shoulders of histograms, sort them separately from the main population.
▲ CRITICAL STEP This step is used to differentiate between cells and debris
3ǀ After optimizing voltage setups and threshold values create two one-dimensional gates for the stained cells in the shouldering of the histograms and also for the major population in the symmetrical or asymmetrical (skewed) distribution of histograms ▲ CRITICAL STEP Be sure that optimal settings and parameters have been established for different SYTO probes and their combinations with different cells (see Table S1).
4ǀ Stain 8ml of diluted pure culture cells (~104 cell/ml) with respective SYTO probes.
5ǀ Inject samples into FCM and sort. It takes ~10 min to sort the cells in an 8 ml sample in high-throughput mode.
▲ CRITICAL STEP Analyze populations in the two gates by sorting in high-throughput mode and collect in 5-ml sterilized culture tubes in the sorting chamber.
6ǀ The material in the 5-ml tubes is vortexed and filtered separately through two sterilized 0.20µm pore size GE Osmonics, PCTE 25 mm diameter hydrophilic membranes. To recover the remaining cells from the tubes, rinse the tubes with filtered fluorescence-activated cell sorter (FACS) flow sheath fluid several times, vortex, and filter through the respective membrane. Two tubes will contain populations from two one-dimensional gates as mentioned above.
▲ CRITICAL STEP Be careful to not lose materials in the tubes. Mark the gate name on the respective tube to make sure the sorted materials in each tube corresponds to the appropriate gate. To confirm cell sorting efficiency, the sorted cells in each tube can be injected separately into the FCM. Sorted cells in each tube should only be visible in the respective gate.
7ǀ Place membranes under the calibrated epifluorescence microscope and observe to determine the accuracy of sorting. ▲ CRITICAL STEP Perform the microscopic observation of cells on the membrane surfaces as soon as possible before the fluorescent intensities of probes fades.
8ǀ The sorted samples can be used for subsequent RNA-based assays or qPCR or for further downstream molecular assays for healthy (viable) and unhealthy (dead or damaged) populations which not performed in this study. However, in another study11 we utilized this protocol to perform RNA-based and qPCR assays for the mammalian cells.
Confirmation of gating protocols using DNA extraction and agarose gels. ● TIMING 70 min
1ǀ Repeat steps 1-6 in the section entitled Confirmation of gating protocols using cell sorting.
▲ CRITICAL STEP When sorting, prepare the DNA isolation kit.
2ǀ Place resulting membranes in PowerBead Tubes from the PowerSoilTM DNA isolation kit.
3ǀ Extract DNA according to the manufacturer’s protocols within 30 min after sorting.
4ǀ Run samples on an agarose gel for 30 minutes at 94 volts.
▲ CRITICAL STEP The agarose gel solution with ethidium bromide can be prepared in advance. Prior to pouring it in the gel box with comb, warm the gel for 2-4 min to liquify. ! CAUTION Wear the laboratory grade mask while warming and pouring the gel into the gel box.
5ǀ After running the gel, take images using a FluroChemTM IS-8800. The amount of DNA extracted from the cells on the membrane is reflected in the thickness of the bands on the agarose gel and gives a relative estimation of cell density in each sorted tube.
▲ CRITICAL STEP Wash the gel with the 1x TBE buffer for 2-5 min prior to imaging to remove excess ethidium bromide staining. Pour 100 ml of 1x TBE buffer into a square shape sterilized plastic container and place the container on a lab shaker. The size of the container should be twice as big as the gel is. The shaker speed should not be more than 20 rpm.
? TROUBLESHOOTING
▲ CRITICAL STEP It is highly recommended that prior to attempting this step, the users have significant experience in the DNA extraction, running agarose gels and imaging gels.
