An Opsono-Adherence Assay for Assessing the Opsonization of Encapsulated Bacteria by Anti-Capsule Antibodies

1. Bacterial Culture and Preparations

NOTE: The bacterial species used in this protocol is an encapsulated virulent strain, B. anthracis Ames. All manipulations with this species require appropriate biosafety and security clearances and must be performed in a Class II or Class III biological safety cabinet located in a BSL-3 laboratory. Follow institutional operating procedures for BSL-3 work and for the use of personal protective equipment when handling bacteria.

1. Prepare Brain Heart Infusion (BHI) broth by dissolving 37 g BHI powder in 1 L water and autoclaving at 121 °C for 15 min. Store broth at ambient temperature.
2. Prepare 8% sodium bicarbonate solution in water and sterilize using a 0.20 µm filter syringe. Store solution at 4 °C and use within 1 day.
3. Mix 8 g Nutrient Broth, 3 g yeast extract, and 15 g agar in 1 L water to prepare NBY agar plates. Autoclave at 121 °C for 15 min. Pour into Petri dishes and allow to solidify. Store plates at 4 °C.
   NOTE: NBY agar plates may be made with or without sodium bicarbonate. The addition of sodium bicarbonate to broth and agar plates induces the growth of mucoid colonies consisting of encapsulated bacilli. The final concentration of sodium bicarbonate in both liquid and solid media is 0.8%.
4. Prepare culture broth for B. anthracis by mixing 50% autoclaved BHI broth, 40% FBS, and 10% sodium bicarbonate solution (v/v).
   NOTE: This broth is formulated to produce short chains of encapsulated bacilli.
5. Prepare a master plate of B. anthracis by streaking it for isolation on an NBY agar plate from a spore stock one day before culturing in broth. Incubate at 37 °C.
6. Inoculate 30 mL of culture broth in a vented 250 mL flask with several colonies of B. anthracis.
   NOTE: A specific volume-to-surface ratio of broth, as specified here, is needed to produce maximally encapsulated bacilli.
7. Shake the broth culture at 125 rpm, 37 °C with 20% CO₂ and humidity for 18–24 h.
   NOTE: Growing B. anthracis at temperatures greater than 37 °C may inhibit capsule production.
8. Use negative staining with India ink to ensure sufficient encapsulation and that bacilli are in short chains (1–5 bacilli/chain) before fixation (see section 2).
9. To determine colony forming units (CFU), serially dilute 100 µL of culture 1:10 in Phosphate Buffered Saline (PBS) solution and plate culture dilutions on sheep blood agar plates. Incubate for 12–18 h at 37 °C. Count CFUs and determine the number of bacteria per mL of culture.
   NOTE: Counting can also be performed using a hemocytometer under the microscope once the bacteria have been fixed. However, this is not recommended because the bacilli are difficult to see under low magnification.
10. Add 16% paraformaldehyde (PF) to the entire volume of bacterial culture at a final concentration of 4% PF to fix the bacilli. Slowly agitate on a shaker at ambient temperature for 7 days.
    NOTE: Seven days of incubation in 4% PF is based on the United States Army Medical Research Institute of Infectious Diseases (USAMRIID) institutional standard operating procedure for fixing vegetative bacilli.
11. To remove the fixative and test for sterility, thrice wash 4 mL (10% volume) of fixed bacterial suspension in a 50 mL conical tube by centrifugation at ≥3000 x g for 45 min and using a 30 mL/wash with PBS. Store the remaining sample in 4% PF at 4 °C.
    NOTE: After pelleting, the bacterial pellet is fluffy due to the presence of a capsule. Take care not to disturb the pellet during washing. It is necessary to remove all fixative so that it does not interfere with the viability testing. If necessary, increase the number of washes.
12. For viability testing, inoculate 40 mL of a bacterial growth medium (e.g. BHI broth) with 4 mL of washed suspension (≤ 10% of broth volume) and incubate at 37 °C for 7 days. Check optical density at 600 nm before and after 7 days to measure turbidity.
13. After 7 days of incubation in broth culture, plate ≥100 µL of broth onto an agar plate and incubate for another 7 days. If no increase in turbidity and no growth on plates are seen, the original culture is considered sterile and may be transferred to a BSL-2 laboratory.
    NOTE: The Federal Select Agent Program mandates that inactivated B. anthracis can only be transferred from BSL-3 laboratories after demonstrating complete sterility of the sample. Viability testing for inactivated B. anthracis consists of culturing 10% of the sample in broth for 7 days followed by culturing on a solid medium for another 7 days. Follow institutional guidelines to receive transfer approval once sterility is established.         

2. Negative staining and light microscopy to observe encapsulation and bacilli chains

1. Mix 5 µL bacterial suspension with 2 µL of India ink on a microscope slide. Place a 1 or 1.5 µm thick cover glass on top of the suspension without creating bubbles.
   NOTE: Nail polish may be used to seal the cover glass onto the slide.
2. Observe the bacterial suspension using a 40x to 100x oil objective lens on a light microscope. Ensure that the bacilli are encapsulated by observing a zone of clearance (capsule excludes India ink) surrounding each bacterium and that the bacilli chains are short.

3. FITC-labeling of bacteria

1. Dissolve fluorescein isothiocyanate (FITC) in dimethyl sulfoxide at a concentration of 2 mg/mL.
2. Measure the volume of inactivated bacterial stock.
3. Wash the stock 3x in PBS to remove the fixative.
4. Add FITC solution at a final concentration of 75 µg/mL. Slowly agitate the mixture for 18–24 h at 4 °C.
5. Thrice wash the bacteria by pelleting at ≥3000 x g, 45 min, and using 30 mL PBS/wash to remove excess FITC. Ensure that the fluffy pellet is not disturbed during washing. Resuspend the bacilli in PBS at the same start volume.
6. Aliquot and store the bacilli in microtubes at -20 °C until use. Do not freeze/thaw bacilli multiple times as the bacilli may lose their capsule.

4. Bacterial Opsonization

1. Heat inactivate a small volume of test sera (from non-human primates, NHPs) at 56 °C for 30 min in a heat block.
2. Thaw and wash FITC-labelled bacteria with PBS 2x by pelleting at 15000 x g for 3–5 min. Resuspend in PBS at the same start volume.
3. In a 96-well round bottom plate (plate #2), serially dilute test sera in cell medium. In another 96-well round bottom plate (plate #3), add 10 µL of diluted test sera into each well.
   NOTE: In every plate, PBS and normal monkey sera were included in place of diluted test sera as negative controls. We also chose one test serum as a positive control to generate a standard curve to normalize all assays done on different days. The positive control test serum was arbitrarily chosen because it was plentiful.
4. To plate #3, add 10 µL freshly reconstituted baby rabbit complement.
   NOTE: Once reconstituted, discard the remaining baby rabbit complement; do not refreeze and thaw.
5. To plate #3, add the appropriate volume of FITC-labeled bacilli for a multiplicity of infection of 1:20. For example, add the volume that contains 5 x 10⁴ x 20 bacilli for 5 x 10⁴ cells. Top off each well in plate #3 with the appropriate volume of cell medium to total 105 µL.
   NOTE: Plate #1, which contains cells, receives 100 µL of opsonized bacteria with the remaining 5 µL as the void volume. We tested each dilution of the test sera in duplicate.
6. Incubate plate #3 at 37 °C for 30 min in the presence of 5% CO₂ with humidity to opsonize bacilli.

5.Adherence Assay and Fluorescent Labeling of Mammalian cells

1. Maintain all reagents at 37 °C before use.
2. Place plate #1, which contains adherent cells, on a 37 °C heat block.
3. Use a multichannel pipettor to remove the spent medium from wells and quickly replace it with 100 µL of opsonized bacteria from plate #3. Ensure no bubbles have been generated in the wells during pipetting. Incubate plate #1 at 37 °C with 5% CO₂ and humidity for 30 min for adherence.
4. Wash each well 5x with 150 µL PBS on top of a 37 °C heat block to remove unattached bacilli.
   NOTE: Care must be taken to ensure that the cells are not accidentally dispersed during washing.
5. Dilute 16% PF with PBS 1:4 to make 4% PF. Fix cells with 4% PF for 1 h by adding 150 µL to each well. Wash cells 2x with PBS.
6. Mix 2 µL HCS (high-content screening) Orange Cell Stain in 10 mL PBS. Add 150 µL of this staining solution to fixed cells and incubate for 30 min at ambient temperature.
7. Wash wells 2x with PBS.
8. Store at 4 °C until microscopy.

6. Microscopy

NOTE: In general, a high-throughput automated fluorescence microscope will facilitate imaging for the OAA. If an automated system is not available, fluorescent images of adherent bacilli can be taken manually. In this study, adherent bacilli and cell monolayers were imaged using the Zeiss 700 laser scanning microscopy system with specialized equipment; our system was composed of the Zeiss Axio Observer Z1 inverted microscope equipped with an automated stage, the Definite Focus module, 40 x NA 0.6 objective, Axio Cam HRc camera and Zen 2012 (Blue Edition) software. The images acquired are widefield images, not confocal images. The following protocol is intended as a basic microscope setup that is applicable to a variety of automated microscopy systems.

1. Incubate plate #1 at ambient temperature for 30 min. Turn on the microscope and related equipment.
   NOTE: Acclimation at ambient temperature prevents condensation from forming on the glass plate during imaging. In addition, it prevents defocusing due to temperature shifts.
2. Place the plate on the stage and focus on the cells using a bright field or phase contrast.
3. Select 96 well format as the plate setting. Select channel settings.
   NOTE: FITC's peak excitation and emission wavelength are 495/519 nm; the HCS orange cell stains are 556 and 572 nm. Other fluorophores may be used depending on the filters that are available on the microscope.
4. Select setting to Tile mode. Indicate the number of images to be acquired.
   NOTE: The tiling function allows the capture of multiple locations in a sample well and can be set to acquire an image in a tiling or a random format. We imaged 10 random areas per well.
5. Change acquisition mode to "black & white" or "monochrome" and binning ≥ 2×2.
   NOTE: Setting the binning from 1×1 to 2×2 or 4×4 would increase microscopy speed but would also result in a lower resolution of the image. For OAA, it is sufficient to use these settings as the assay enumerates both the macrophages and the adherent bacteria.
6. Turn on the autofocus or focusing module. Indicate how often the autofocusing function is to be performed. Turn on the Z stack function, if available. Indicate the number of Z stacks that will capture the thickness of the cell monolayer and adherent bacilli.
   NOTE: The number of Z stacks chosen will increase microscopy time but will ensure an image with the correct focus is taken at each Z stack.
7. Designate a file folder to save images to. Run the automated imaging program.