Characterizing Microbiome Dynamics – Flow Cytometry Based Workflows from Pure Cultures to Natural Communities

1. Sampling and Fixation

1. Pure culture: deep freezing^15,17
   1. Take 2 mL of cell suspension, centrifuge for 5 min at room temperature (RT) and 5,000 x g and discard the supernatant.
      NOTE: The sampled cell suspension is described in Supplementary File 1-S1.
   2. Resuspend the cells in 1 mL of phosphate buffered saline (PBS; 6 mM Na₂HPO₄, 1.8 mM NaH₂PO₄, 145 mM NaCl with bi-distilled H₂O, pH 7.2) additionally containing 15% (v/v) glycerol as a cryoprotective agent.
   3. Incubate for 10 min on ice and shock freeze in liquid nitrogen for subsequent storage at -80 °C.
      NOTE: The protocol can be paused here. The samples are stable for at least one month.
2. Activated sludge community: formaldehyde stabilization + ethanol fixation^5,10,18
   1. Take 4 mL of the cell suspension, centrifuge for 20 min at 15 °C and 3,200 x g and discard the supernatant.
      NOTE: The sampled cell suspension is described in Supplementary File 1-S1.
   2. Add 4 mL of 2% formaldehyde in PBS and incubate for 30 min at RT.
      1. Prepare 8% formaldehyde stock solution from paraformaldehyde to avoid the conserving additive methanol added to formaldehyde shipped in liquid form. Add 4 g of paraformaldehyde to 50 mL of PBS at 70 °C. Add approximately 125 µL of 10 M NaOH solution. Stir until the paraformaldehyde has completely dissolved and let it then cool down to RT. Adjust the pH value to 7.0 with approximately 100 µL 37% HCl. Freeze the formaldehyde solution in 15 mL aliquots for storage. Do not use defrosted aliquots longer than 10 days.
         CAUTION: Formaldehyde needs to be handled and disposed with care due to its toxic and mutagenic properties. Always wear gloves while handling it. Use an exhaust hood while dissolving the paraformaldehyde to prevent exposure to toxic fumes.
   3. Centrifuge the sample for 10 min at 15 °C and 3,200 x g and discard the supernatant.
   4. Resuspend the sample in 4 mL of 70% ethanol and store at -20 °C.
      NOTE: The protocol can be paused here. The samples are stable for at least 2 months. Depending on the sample properties, the centrifugation step in 1.2.1 can also be performed for 10 min at 4 °C to save time.
3. Biogas community: drying
   1. Use a clipped 1,000 µL pipette tip to sample 200 µL of the viscous digestate into a 2 mL tube.
   2. Add 1,700 µL of PBS buffer and mix thoroughly.
   3. Place the tubes into an ultrasonic bath at 35 kHz and 80 W effective output power for 1 min to disband large cell aggregates and detach cells sticking to plant cell residues.
   4. Mix the sample thoroughly, filter the sample through a 50 µL mesh filter and divide the filtrate into four aliquots of about 400 µL.
      NOTE: Preparing aliquots at this point provides two major advantages. First, smaller volumes are easier and faster to dewater mechanically (centrifuge) and thermally (drying), but sampling of small volumes from usually thick biogas sludge can be very challenging. Second, extra samples may be used for subsequent cell sorting, backup measurements or controls.
   5. Centrifuge the aliquots twice for 10 min at 10 °C and 4,000 x g and discard the supernatant completely both times to mechanically dewater the sample as thoroughly as possible.
   6. Dry the samples in a heated vacuum centrifuge for 40 min at 35 °C, about -97 kPa and 2,500 x g to create stable pellets. Store the pellets at 4 °C in the dark.
      NOTE: The protocol can be paused here. The pellets are stable for at least 6 months.

2. Staining

NOTE: DAPI staining has proven to deliver high resolution dot plots and is therefore particularly advantageous when analyzing communities. The DAPI concentration in the staining solution needs to be optimized to guarantee a cell gate fluorescence intensity well above the noise level but below the beads. The optimum depends on the instrument sensitivity and bead choice, can vary between sample sets due to G/C content of the contained microorganisms, and should generally be tested for newly introduced sample sets. Testing concentrations between 0.24 µM DAPI and 1 µM DAPI is recommended for the presented setup. Other dyes might be used for specific applications. The use of a SYBR Green I staining protocol is advised when absolute cell counting accuracy is prioritized over the fingerprinting analysis (Supplementary File 1-S1)^6,15. It includes less sample handling and centrifugation steps and is applicable with both fixed and vital cells. The use of vital cells further reduces sample handling steps by skipping the fixation and hence requires only one centrifugation for cell harvesting. This minimizes potential cell loss and consequently systematic measurement error. The PBS, permeabilization buffer and staining solution used during all following steps should be filtered through 0.2 µm syringe filters to reduce additional particle load in the sample. The staining of one sample containing Escherichia coli BL21 (DE3) as a biological standard with every sample pool is advised.

1. Pure culture
   1. Defrost the samples, centrifuge for 5 min at RT and 5,000 x g and discard the supernatant.
   2. Resuspend the cell pellet in ice cold PBS by repeated pipetting and adjust the OD_700nm to 0.035 (path length_cuvette = 0.5 cm).
   3. Prepare the cells for staining.
      1. Centrifuge 1 mL of adjusted cell suspension for 5 min at RT and 5,000 x g and discard the supernatant.
      2. Resuspend the cells in 1 mL of permeabilization buffer containing 0.3 M citric acid and 4.1 mM Tween20 and mix thoroughly.
      3. Incubate the sample for 10 min on ice to create permeable cell membranes for subsequent staining.
      4. Centrifuge the sample for 5 min at RT and 5,000 x g and discard the supernatant.
   4. Add 1 mL of staining solution containing 0.68 µM DAPI in 417 mM Na₂HPO₄/NaH₂PO₄ buffer (289 mM Na₂HPO₄, 128 mM NaH₂PO₄ with bi-distilled H₂O, pH 7), mix and incubate for at least 15 min at RT in the dark.
      NOTE: Precise OD adjustment is crucial to guarantee comparable measurements because the DAPI fluorescence will vary with the cell density if the DAPI concentration is kept constant. The supernatant should be removed carefully due to a generally fragile pellet to prevent cell loss.
      CAUTION: Handle and dispose DAPI with care due to its mutagenic properties.
2. Activated sludge community
   1. Mix the fixed sample thoroughly and transfer 0.6 mL into a glass tube. Add 1.4 mL of PBS and sonicate for 10 min at RT as described in step 1.3.3. Centrifuge the sample for 10 min at 4 °C and 3,200 x g and remove the supernatant. Add 2 mL of PBS and mix thoroughly. Sonicate for 5 min.
   2. Adjust the OD_700nm to 0.035 (path length_cuvette = 0.5 cm) with PBS.
   3. Prepare the cells for staining.
      1. Centrifuge the sample for 10 min at 4 °C and 3,200 x g and remove the supernatant.
      2. Resuspend the cells in 1 mL of permeabilization buffer containing 0.11 M citric acid and 4.1 mM Tween20 and mix thoroughly.
      3. Incubate for 20 min at RT to create permeable cell membranes for subsequent staining.
      4. Centrifuge the sample again for 10 min at 4 °C and 3,200 x g and remove the supernatant.
   4. Add 2 mL of staining solution containing 0.68 µM DAPI and 417 mM Na₂HPO₄/NaH₂PO₄ buffer, mix thoroughly and incubate at least 60 min at RT in the dark.
      NOTE: The use of glass tubes is advised, as certain microorganisms tend to adhere to the plastic tube walls and may be lost during the process. Incubating overnight is also possible and usually simplifies lab procedures.
      CAUTION: DAPI needs to be handled and disposed with care due to its mutagenic properties.
3. Biogas community
   1. Suspend the dried cell pellet in PBS.
      1. Add 800 µL of PBS, mix thoroughly and let the pellet soak for 15 min.
      2. Aspirate the liquid phase with a 1,000 µL pipette and move the soaked pellet to the cylindrical section of tube wall with the pipette tip. It should stick there.
      3. Move the pipette tip in a circular motion to squash the pellet along the tube walls.
      4. Wash the resulting slurry off the tube walls by dispensing the liquid phase from the pipette tip along the tube wall.
      5. Agitate the suspension by aspirating and suspending it into the pipette thrice.
   2. Wash the sample with PBS twice.
      1. Centrifuge for 5 min at 10 °C and 4,000 x g and discard the supernatant.
      2. Add 1,500 µL of PBS and mix thoroughly.
      3. Repeat the two steps above once.
   3. Place the tubes into an ultrasonic bath for 1 min as described in step 1.3.3 and, subsequently, filter each sample through a 50 µL mesh filter into an individual glass tube.
   4. Dilute 2 mL of the sample to OD_700nm 0.035 (path length_cuvette = 0.5 cm) with PBS.
   5. Prepare the cells for staining as explained in step 2.2.3 above.
   6. Add 2 mL of staining solution containing 0.24 µM DAPI and 417 mM Na₂HPO₄/NaH₂PO₄ buffer, mix thoroughly and incubate over night at RT in the dark.
      CAUTION: DAPI needs to be handled and disposed with care due to its mutagenic properties.

3. Measurement

NOTE: The exemplary sample sets were analyzed with two different flow cytometers. The PC and the ASC were analyzed with a more complex, highly adjustable and easily customizable, research centered cell sorter. For the PC analysis, this device was equipped with a laser set up as described in previous publication¹⁶, in short a blue (488 nm, 400 mW) and an ultraviolet (334–364 nm, 100 mW) argon ion laser. For the ASC analysis, newer blue (488 nm, 400 mW) and ultraviolet (355 nm, 150 mW) semiconductor lasers were installed. In both cases, the blue laser induced the FSC (bandpass filter 488 nm ± 5 nm, neutral density filter 1.9) and the SSC (bandpass filter 488 nm ± 5 nm, neutral density filter 1.9, trigger). These optical characteristics are related to cell size and cell density, respectively. The UV lasers excited the DAPI fluorescence (bandpass filter 450 nm ± 32.5 nm) for the quantification of cellular DNA content. The fluidic system was run at 56 psi (3.86 bar) with the sample overpressure at maximum 0.3 psi and a 70 μm nozzle. All parameters were recorded as peak heights instead of peak areas to improve the measurement resolution. The long-term monitoring of the biogas community was conducted with the less complex, flow cuvette based, bench top analyzer. An ultraviolet semiconductor laser (355 nm, 50 mW) was used to induce the FSC (bandpass filter 355 nm ± 5 nm) and SSC (bandpass filter 355 nm ± 5 nm, trigger) signals and excite the DAPI fluorescence (bandpass filter 455 nm C). The water used for the sheath buffer of both devices was bi-distilled and 0.1 µm filtered. The PBS used for sample dilution should be filtered through 0.2 µm syringe filters to reduce the particle noise in the measurements as much as possible.

1. Pure culture and activated sludge community
   1. Start the instrument, laser, computer and software and prepare for sample analysis.
      1. Switch on the lasers and run for 15 min to reach the operating temperature and ensure beam stability.
      2. Fill the sheath tank with 10x sheath buffer (19 mM KH₂PO₄, 38 mM KCl, 166 mM Na₂HPO₄, 1.39 M NaCl with bi-distilled H₂O) diluted with bi-distilled H₂O to a 0.2x working solution (for cell sorting: 0.5x working solution).
      3. Prime the fluidic system with 56 psi over pressure and the waste tank with approximately 6 psi vacuum.
      4. Run the instrument for minimum 15 min in Backflush mode to rinse the sample port, tubing and nozzle.
   2. Calibrate with standard beads.
      1. Prepare the bead mixes for the calibration in the linear (1 μm blue + 2 μm yellow-green fluorescent) and logarithmic range (0.5 μm and 1 μm blue fluorescent). Dilute the bead suspension from the original stock suspensions to achieve equal concentrations.
      2. Continuously measure the linear bead mix and fit the bead peaks to a preset calibration template by manipulating the nozzle and laser optics positions to pre-calibrate the instrument in the linear range.
      3. Switch to the logarithmic range and continuously measure the logarithmic bead mix. Fit the bead peaks to their preset calibration template to fine tune the hardware of the instrument. Make final adjustments to the bead position using the gain setting of the photomultiplier tubes (PMTs).
      4. Check the position of the instrumental noise and possibly adjust it to fit the calibration template.
         NOTE: A fixed calibration template for the position of the beads needs to be prepared prior to a measurement project and cannot be changed (see Supplementary File 1-S4)! Instrumental noise can be induced by particles in the sample or the electronic noise of the PMTs and will always be recorded to some degree. It is not advisable to hide the noise completely by gain adjustment or plot offset. The abundance and position of the noise events can be a valuable source of information and should therefore be contained in the raw data. Very particle intensive samples might necessitate the subsequent removal of the noise for plotting and data analysis reasons. Control the calibration in regular intervals during a measurement day to guarantee instrument stability and therefore sample comparability.
   3. Measure a sample containing the biological standard as described in 2.1.4 (DAPI stained Escherichia coli BL21 (DE3), sampled and fixed during the stationary phase (16 h) according to the ASC protocol described in 1.2). Check the peak position in relation to their preset calibration template (see Supplementary File 1-S5).
      NOTE: The routine staining and measurement of the biological standard with every pool of samples will also serve as an internal control for the staining procedure. If the device is calibrated using beads and the biological standard does not fit its preset calibration template, the staining procedure probably needs to be repeated.
   4. Sample acquisition.
      1. Run the staining solution for 10 min to rinse the sample port and tube and ensure the stability of the DAPI fluorescence in subsequent samples.
      2. Filter the stained sample with a 50 μm mesh filter before the measurement to prevent clogging of the cytometer nozzle or flow capillary.
      3. Add the logarithmic bead mix to the sample to monitor the instrument stability and provide the possibility for a retrospective calibration check. The samples should contain between 1,000 and 2,500 events in each of the two bead gates.
      4. Create a cell gate in the instrument software during the first trial measurements of a new sample set. This gate should contain the stained cells and exclude the noise and beads.
      5. Mix the samples and measure at a maximum speed of 3,000 events s^-1 until 250,000 cells (communities) or 50,000 cells (pure cultures) are detected within the cell gate.
         NOTE: These cell counts are chosen based on the experience with the sample sets. Differing numbers may be used to shift the tradeoff between the measurement efficiency and the detection of low abundance subcommunities in either direction.
         Troubleshooting: Sudden intra-measurement shifts of the bead and cell position can be caused by air bubbles trapped in the nozzle. This necessitates the removal and cleaning of the nozzle and the rinsing of the fluidic system with sheath buffer. The instrument needs to be readjusted after remounting the nozzle (start with step 3.1.2).
      6. Backflush the instrument thoroughly with sheath buffer before shutdown and switching samples.
2. Biogas community
   1. Start up the instrument, laser, computer and software to prepare for sample analysis.
      1. Backflush at least 6 mL of sheath buffer (bi-distilled H₂O) through the tubing and sample port into a loosely attached tube using the “sheath fluid prime” function of the instrument software.
      2. Measure bi-distilled H₂O at 5 µL s^-1 to rinse the fluidic system until less than 200 events s^-1 are detected at the regular flow rate of 0.5 µL s^-1.
   2. Calibrate the system.
      1. Prepare the bead mix (0.5 μm and 1 μm blue fluorescent). Dilute the bead suspension from the original stock solutions to achieve equal concentrations.
      2. Continuously measure the bead mix in the log₄ range and fit the bead peaks to their preset calibration template by manipulating the flow cuvette and laser optic positions. Make final adjustments to the bead position with the gain setting of the PMTs.
   3. Control the calibration with the biological standard as described in step 3.1.3.
   4. Sample acquisition.
      1. Dilute 400 µL of stained sample in 1,600 µL of PBS, measure it and monitor the event count to perform a concentration test.
         NOTE: Adjusted dilution rates may be necessary for other sample sources. The event count should not exceed 1,200 events s^-1 in particle rich samples to prevent resolution loss in the forward scatter (negative example in Supplementary File 1-S2). Pure cultures can be measured with up to 2,000 events s^-1.
      2. Create a cell gate in the instrument software during these first trial measurements. This gate should contain the stained cells and exclude noise and beads.
      3. Dilute the sample according to the results of the concentration test to run at maximum 1,200 total events s^-1 and add the bead mix to the sample to monitor instrument stability and provide the possibility for a retrospective calibration check. The samples should contain between 1,000 and 2,500 events in each of the two bead gates.
      4. Mix and measure the sample until 250,000 cells (communities) or 50,000 cells (pure cultures) are detected within the cell gate.
      5. Rinse the instrument thoroughly with bi-distilled H₂O before shutdown and switching samples.

4. Cell Sorting

NOTE: The cell sorting procedure requires additional instrument set up and is performed according to published protocols¹².

1. Start up and calibrate the instrument as described in steps 3.1.1-3.1.3, but use a 0.5x working solution of the sheath buffer.
2. Set the DD frequency and DD amplitude to find the droplet break-off.
3. Mount the deflection plates, charge them and decide for 2- or 4-Way Sort Mode.
4. Perform the internal drop delay calibration with 2 μm yellow-green beads to define the time span that takes a particle or cell to travel from the interrogation point (laser hits cell) to the last drop attached to the stream.
5. Sort 6 times 20 beads on a glass slide and count them with a microscope to verify the drop delay calibration.
6. Prepare the sorting gate template.
7. Use the “single and one-drop mode: highest purity 99%” at a rate not higher than 2,500 total events s^-1 to sort 500,000 cells in maximum 4 gates at a time (4-Way Sort Mode).
8. Harvest the cells by centrifugation (20,000 x g, 6 °C, 25 min) and freeze the pellet at -20 °C for later DNA isolation and sequencing.
   NOTE: Avoid sorting cells in the gates with less than 5% relative abundance, as the sorting time is inversely proportionate to the relative abundance. The individual steps are described in detail in the cell sorter manual. The required cell numbers are heavily dependent on the respective use cases and extraction protocols. Numerous methods have been applied: ddPCR (1,000 cells^17,19), 16S rDNA or mcrA amplicon sequencing (500,000 cells^6,10,15) and proteomics (up to 1.1 x 10⁷ cells^16,17). The cell sorting will be particularly advantageous when combined with a metagenomics approach because of the lower diversity in the sorted subcommunities.
   Caution: Do not touch the deflector plates during the sorting procedure due to high voltages.

5. Data Analysis

NOTE: The cytometric measurement yields “flow cytometry standard” .fcs data files with a generally uniform data structure. However, different cytometer manufacturers use slightly adapted versions and older instruments might predate the 2010 introduction of the latest FCS standard 3.1. This can lead to dot plot scaling issues, which prevent the direct comparison of the results gathered with different instruments. However, the individual data points are always stored in the lines of a matrix, with the respective scatter and fluorescence intensity values in the different columns. The presented microbiome analysis pipelines use the distinctive cell size and DNA content to describe microbial subcommunities. These parameters are correlated to the FSC and DAPI fluorescence channel intensities (cell sorter: FL-4, analyzer: FL-1) and result in subcommunity clusters when visualized in two-dimensional FSC vs. DAPI fluorescence plots. These dot plots allow the interpretation and analysis of flow cytometry data and are usually logarithmically scaled. They can be displayed from .fcs files using proprietary cytometer software or third party solutions and are the basis for the automated (Cytometric Histogram Image Comparison, CHIC) and semi-automated (Cytometric Barcoding, CyBar) community analysis.

1. Cytometric Histogram Image Comparison
   Note: The code, detailed documentation and a manual for this package are available at http://www.bioconductor.org/packages/release/bioc/html/flowCHIC.html. They have also been published²⁰.
   1. Install and load the R package, set the working directory containing the .fcs files and set up a file list by executing:
      > source("https://bioconductor.org/biocLite.R")
      > biocLite("flowCHIC")
      > library(flowCHIC)
      > # Set the working directory to the path where your FCS files are located
      > # Type the path into the quotes
      > setwd("")
      > # Get a list of the filenames of the FCS files located in your working directory
      > files <- list.files(getwd(),full=TRUE,pattern="*.fcs")
      > # Get a list of the filenames of the FCS files included to the package
      > files <- list.files(system.file("extdata",package="flowCHIC"),
      + full=TRUE,pattern="*.fcs")
      > # Read the first FCS file as flowFrame
      > frame <- read.FCS(files[1],alter.names=TRUE,transformation=FALSE)
      > # Get a list of the parameter/channel names
      > unname(frame@parameters@data$name)
   2. Create images for the cytometric raw data by executing the "fcs_to_img" package function:
      > # Create the histogram images using default parameters
      > fcs_to_img(files)
   3. Define subsets of the histogram images by executing the "img_sub" package function:
      > # Create the histogram image subsets using default parameters
      >  img_sub(files,ch1="FS.Log",ch2="FL.4.Log")
   4. Calculate the Overlap and XOR values to conduct the image analysis by executing the "calculate_overlaps_xor" package function:
      > # Save the names of all subset images as a list
      > subsets <- list.files(path=paste(getwd(),"chic_subset",sep="/"),full=TRUE,pattern="*.png")
      > # Calculate overlap and XOR images and write values to two new files
      > results<-calculate_overlaps_xor(subsets)
   5. Create non-metric multidimensional scaling (NMDS) plots for similarity analysis by executing the "plot_nmds" package function:
      > # Show a NMDS plot of the samples
      > plot_nmds(results$overlap,results$xor)
2. Cytometric Barcoding
   NOTE: The code, detailed documentation and a manual for this package are available at http://www.bioconductor.org/packages/devel/bioc/html/flowCyBar.html. They have also been published^11,12.
   1. Develop a master gate template for use on all samples.
      1. Load the .fcs files into the analysis software using the drag and drop functionality.
      2. Double click a measurement and choose the x- and y-axis parameters from the respective dropdown lists to open a FSC vs. DAPI fluorescence plot.
      3. Use the polygon drawing tool to reproduce the cell gate previously used to control the number of analyzed cells in steps 3.1.4.5 and 3.2.4.4 and name it accordingly. Drag the cell gate entry in the sample list on the all samples group to pool it.
      4. Double click the cell gate and correct the axis assignment to visualize the cell gate events.
      5. Define the subcommunities prevalent in the sample set with the elliptical gates tool following the published guidelines¹² and use the scanning of the SSC or another third parameter²¹ to ensure the integrity of the gate template. Pool, control and adjust the subcommunity allocation on the other samples.
   2. Export the relative subcommunity abundances into a .txt file for use in the R script.
      1. Open the table editor with the edit tab.
      2. Add columns for every subcommunity that was defined in step 5.2.1.5 and set the output statistic to “frequency of parent” and name them.
      3. Create the table in the “table editor” after choosing saving format and destination.
         NOTE: The analysis software directly provides relative subcommunity abundances. They can also be obtained through division of the event count in the individual subcommunity gate by the event count in the cell gate. The values have to be saved in a tab delimited matrix in a .txt file that can not contain any n.a. fields and needs to meet some additional requirements to be read by the R code (see manual and FAQ for specific instructions).
   3. Install and load the R package and load the data by executing:
      > source("https://bioconductor.org/biocLite.R")
      > biocLite("flowCyBar")
      > # Type the path into the quotes
      > setwd("")
      > # Load dataset
      > data(Cell_number_sample)
      > # Show data
      > Cell_number_sample[,-1]
   4. Normalize the relative abundances by executing the "normalize" package function:
      # Normalize data, print 2 digits
      normalize(Cell_number_sample[,-1],digits=2)
   5. Create a bar code of the normalized and a boxplot of the original relative abundances by executing the "cybar_plot" package function:
      Normalized_mean<-normalize(Cell_number_sample[,-1],digits=2)
      Normalized_mean<-data.frame(data.matrix(Normalized_mean))
      # Plot with default parameters
      cybar_plot(Normalized_mean,Cell_number_sample[,-1])
   6. Create an NMDS plot of the original relative abundances.
      > Normalized_mean<-normalize(Cell_number_sample[,-1],digits=2)
      > Normalized_mean<-data.frame(data.matrix(Normalized_mean))
      > # NMDS plot of normalized cel numbers
      > nmds(Normalized_mean)
   7. Create the correlation analysis of the normalized relative abundances and other parameters by executing the "correlation" package function:
      > # Load dataset
      > data(Corr_data_sample)
      > # Show correlation values
      > Corr_data_sample[,-1]
      > # Run correlation analysis
      > correlation(Corr_data_sample[,-1])
      NOTE: CHIC and CyBar rely on relative abundances. However, flow cytometry can also determine absolute cell numbers, which might be of great interest for biotechnological applications. To determine the absolute cell number, the number of cells in the gate of interest is divided by the analyzed sample volume. The analyzed sample volume can be determined by adding bead suspension with a known concentration into the sample (formula in Supplementary File 1-S7). The bench top analyzer is additionally equipped with a true volumetric counting feature that directly quantifies the volume in the sample tube during the measurement. It is advised to use a SYBR Green I staining protocol for cell counting.