Archives: Methods

Reference:

Lanyi, B., and T. Bergan. Methods in Microbiology, Vol 10: 93-168.

Bacterial Agglutination

Bacterial agglutination is performed using either P. aeruginosa cells autoclaved at 120^oC for 1 hour, then centrifuged and resuspended in saline or live P. aeruginosa cells grown to mid log phase, then centrifuged and resuspended in saline. (For either method resuspend the bacteria so the concentration is 5 X 10⁸ bacteria/ml.) In both methods the cell preparations are incubated with serial dilutions of antiserum for 1 hour at 37^oC. The agglutination of live cells is characterized by a coarse granular bacterial clumping, as opposed to the finer granular clumping of heat-killed P. aeruginosa with homologous LPS O-antigen-specific serum. Bacterial agglutination is scored on a scale of +1 (weak agglutination) to +4 (strong reaction). The agglutination can also be performed on glass slides using a drop each of antiserum and bacterial preparation, and the results observed after 3 to 5 minutes.

Passive Agglutination

Passive agglutination is performed using tanned sheep red blood cells and P. aeruginosa LPS (heated for 1 hour at 100^oC prior to use). Sheep erythrocytes are washed three times in saline (0.9% NaCl), adjusted to 4% erythrocytes (v/v) in PBS, pH 7.5 and 2.5mg of tannic acid in 50ml PBS is added to 50ml of the cell suspension. After a 15 minutes incubation at 37^oC with occasional mixing, the cells are centrifuged (100 x g for 20 minutes) and washed with 100ml PBS. One half of the cells are kept as a control in PBS containing 1mM NaN₃. To the other half of the cells, 20µg/ml LPS or protein is added, and the mixture is incubated for 1 hour at 37^oC with very gentle agitation at regular intervals. The antigen-coated cells are washed three times in saline and 1mM NaN₃ is added for storage at 4^oC. The cells are made up to 1% (v/v) in saline before use.

Passive agglutination is performed in 96 well conical bottom plates (Linbro, Flow Labs) using 50µl of antiserum serially diluted in saline and 50µl of 1% (v/v) antigen-coated sheep erythrocytes. Control wells contain the antiserum and tanned, non-antigen-coated sheep erythrocytes. The plates are incubated at 37^oC for 1 hour. The titre is the inverse of the highest serum dilution showing agglutination. Non-agglutinated cells give a tight button of cells at the bottom of the well, while the agglutinated cells form a mat at the well bottom.

1. Prepare before thawing:
   • Fill a test tube of 10 – 15 mls of cold media (appropriate for your cell line) with 10 – 20% fetal calf serum (use 2x the % you use for growth supplement).
   • Place in a beaker of crushed ice to CHILL. (DMSO is toxic at room temperature.)
   • Have everything in the hood that you will need: 70% alcohol, paper towels, pipets, etc.
2. Get vial from Liquid Nitrogen – use gloves and face shield. *Make sure the cover is replaced properly!!
3. “QUICK THAW” in 37ºC H₂O bath by shaking vial rapidly in water till approximately 3/4 thawed, with a small pea sized portion still frozen.
4. Remove from H₂O bath, but continue to shake vial until it has thawed completely.
5. Rinse vial with Ethanol.
6. Open vial carefully, pipet up contents and ‘layer’ it onto cold media (in the test tube you have chilling)
7. Spin at approx. 1000 rpm for 5 min.
8. Wash 1x with 10 – 15 mls of media, 1000 rpm 5 min (to wash out all DMSO). Gently resuspend the pellet.
9. Seed into a small flask T25 or T75 with appropriate volume of media containing 2x the regular amount of FCS, and the regular amount of glutamine and antibiotic.
10. NEXT DAY: (Change media if growing adherent cells to get rid of excess dead cells)
11. Check non-adherent cells to see how soon they will need fresh media and/or need to be split.
12. 1 – 4 DAYS: Passage cells when required in usual fashion.

1. Established hybridomas and other cell lines can be stored in liquid N₂ using this method.
2. Have everything ready before you start working with the cells (i.e. cool media, label vials, prepare cryovials, etc.)
3. Freeze only cultures that are healthy and in the mid-log phase of growth (ie., 2-5 x 10⁵ cells/ml).
4. Have your vials already labelled with a waterproof, cryogenic pen. Include cell line name, passage number, date frozen and number of cells per vial.
5. Centrifuge the cells and resuspend in 10% DMSO/90% FCS at about 1-5 x 10⁶ cells/ml. The freezing medium should be 5ºC, not warm.
6. Put 1ml of cell suspension into cryovials and tighten the lid finger tight. Do not over tighten or the vials could explode when thawed.
7. Use the Nunc cryovials with the O-ring and inner threads, not the Nalgene ones. Put the vials in one of the cryoracks specially designed for slow, even cooling.
   1. The Nalgene cryocontainer holds 18 vials. The bottom is filled with isopropanol which must be changed after five uses. Follow the instructions on the container.
   2. The blue cryorack holds 50 vials. It is filled with water, plugged and placed plug side down. Put the vials right side up into the spaces and place the rack in this position in the -70ºC freezer. Do not place them in a “pre-frozen rack”. Do not fill 100% full or the plug will pop while freezing.
8. Record in the card file: cell line frozen; # cells; date frozen; number of vials; cane label/ #; which cane holder; your initials/ name.

Reference:

Day et.al., J. Infect. 2:39-51, 1980.

1. Preparation Of Chambers

A. Syringe Barrels:

1. remove markings with 95% ethanol.
2. use 1 ml syringes for mice & 3 ml syringes for rats.
3. cut syringe barrels into 1 cm lengths.

B. Filters:

1. cut discs from 0.22 µm pore size millipore filters.
2. use standard hole punch for mice & large 3-hole punch for rats.
3. layer discs on paper in a glass petri dish.
4. wrap well and autoclave.

2. Construction Of Chambers (in sterile hood)

1. attach filter disc to one end of chamber with a little glue applied with a sterile toothpick.
2. add 100 µl (for mice) or 500 µl (for rats) of bacterial sample (in buffer) to the chamber.
3. add second filter to open end.

3. Implantation Of Chambers

1. Anaesthetic: ketamine & xylazine mixed 1/10 in saline. Prepare 1/2 ml ketamine & 0.5 ml xylazine per mouse (amounts given per mouse listed in mouse room).
2. Insert four chambers into the peritoneal cavity of each animal via a longitudinal incision in the abdomen.
3. Close incision using surgical sutures.

4. Removal Of Chambers

1. Kill animals with CO_2.
2. Chambers are removed by making an incision into the peritoneum.

Use for producing stable exoenzyme S.

1. 1 litre
2. 30 g TSB
3. 90 ml dH₂O
4. 10 g Chelex 100-400 (other Chelex is ok)
5. Stir 6 hours at 25°C.
6. Spin 16,000 g for 30 minutes.
7. Filter through 0.45 um filter.
8. Concentrate to almost dry over a PM-30 (PM-10 is ok) Amicon membrane.
9. Discard concentrate on the filter, the rest is 10X and filtered.
10. Add nitrilotriacetic acid to 10 mM.
11. Adjust to pH 7.0.
12. Autoclave in 2 litre flask. Cool.
13. Add 20 ml sterile 50% glycerol.
14. Add 50 ml sterile 2 M MSG (or glutamate, Na salt)
15. Add sterile MgSO₄ to 1 mM

Method:

1. Grow host bacteria overnight at 37C. in Proteose peptone No 2 broth (PP2).
2. Add 0.1 ml phage stock solution and 0.1 ml of overnight bacterial culture into 4 ml of prewarmed overlay agar (0.6% agar, 1% PP2). Agar is kept at 45-49C.
3. Swirl and pour overlay agar onto PP2 plates. Allow to set and incubate at 37C. overnight.
4. Scrape off soft overlay with sterile spreader and some sterile broth. Get most of the overlay and broth into 5 mls sterile PP2.
5. Centrifuge at maximum speed in a clinical centrifuge for 15 minutes.
6. Save supernatant. Add 0.1 ml. CHCL₃ and label.
7. Do a serial dilution of this phage stock and do plate counts of 10-8, 10-10, 10-12 dilutions to obtain the titre. (eg. repeat step 2, incubate and count.)

This tests for the presence of electron transport system in Pseudomonas species.

Materials:

• Filter paper
• 1% N,N,N’N’ tetramethyl-p-phenylenediamine

Methods:

From Colonies:

1. Pre-wet filter paper with oxidase reagent (N,N,N’,N’-p-phenylenediamine) and allow to dry.
2. Pick a bacterial colony with a sterile toothpick.
3. Gently scratch the colony onto the filter paper. A blue color is produced by Pseudomonas species.

From Liquid Culture:

1. Spot the filter paper with 10 to 20 µl of culture.
2. Add 10 µl of freshly made oxidase reagent and look for production of a blue color.

NOTE: If the filter paper is pre-wetted with oxidase reagent, false positive can occur.

1. Grow an overnight culture of H103 (wildtype) in 10 ml of minimal medium containing 1/20 normal amount of sulfate.
2. Subculture the overnight 1/40 into fresh medium containing no sulfate.
3. Grow to OD₆₀₀ = 0.1-0.2.
4. Add unlabelled methionine to 1 mM (0.1 ml of 0.1 M stock of autoclaved unlabelled methionine into 10 ml of cells) and ³⁵S-methionine to a final concentration of 50 µCi/ml. Return cells to incubator.
5. To monitor uptake of label, remove 1 ml samples at 30 minute intervals, filter cells through 0.45 micron filters, wash twice with 0.1 M LiCl, dry and count filters.

1. Grow bacteria to an OD₆₂₀ of 0.8.
2. Wash 3x in PBS containing 2% FCS.
3. Resuspend to the original volume with 1mM MgCl₂. Dispense 0.8 ml into Eppendorf tubes.
4. Add monoclonal or polyclonal serum and vortex (3 on Vortex Genie Fisher)
5. Incubate for 45 min. at 37ºC with agitation.
6. Wash 3x with PBS containing 1% FCS, 1 mM MgCl₂. (Microfuge for 1 min)
7. Incubate at 37ºC for 45 min in PBS containing 2% FCS and a 100-1000 fold dilution of rabbit anti-mouse IgG.
8. Repeat step 6 and then step 7 this time with goat anti-rabbit Ab coupled to FITC.
9. Wash with PBS.
10. Add 10 µl to a slide. Add a cover slip and with a fluorescence microscope measure the fluorescence emitted at 525 nm.

Another method uses bacteria fixed on slides. For each incubation step place the slides in a humid chamber for 10 min. at 37C. The slides are washed by dipping in PBS.

1. Grow the strain of bacteria to be frozen, in 5 to 10 mls of appropriate media, until an OD₆₀₀ = 0.6-0.8 is reached OR aseptically (using a sterile swab) scrape bacteria from a freshly grown (24 – 48 hr) plate and resuspend in 2 to 5 mls of a rich broth such as Mueller Hinton or LBNS.
2. Using a sterile pipette, add dimethyl sulfoxide (DMSO) so the final concentration is 7% (i.e. 70ul DMSO per 1 ml of cell suspension). Note: DMSO is considered a sterile solution but can be filter sterilized using a 0.22 um nylon filter.
3. Vortex.
4. Aseptically pipet 1.0 to 1.5 mls of bacterial suspension (with DMSO) into each cryovial. Make sure the cryovial is labelled with a waterproof, cryogenic pen (black color lasts better in the freezer). Freeze at -70^oC.