The DAN Lab

Cell Protocols

Gram staining protocol

Note: written for potential Zymomonas isolates, but can be used for other organisms

  1. Streak strains for dense colonies from freezer stocks on ZRM agar and grow anaerobically at 30°C for >48hrs
  2. Use sterile loop to collect glob of cells and add to 100uL of ddH2O in eppy tube
  3. Mix to form a turbid solution, then use loop to spread mixture on slide
    1. Alternately, grow cells in liquid media and directly spread dense culture on slide
  4. Allow slide to completely air dry
  5. Heat-fix cells to slide by passing slide through the flame of bunsen burner so the cell side is not burnt (cell side up, away from flame), but the slide becomes almost too hot to hold (~60°C)
  6. Flood slide with crystal violet reagent, allow to sit for 1 minute
  7. Rinse slide gently but completely with R/O water
  8. Flood slide with Gram’s Iodine (mordant), allow to sit 1 minute
  9. Rinse slide gently but completely with R/O water
  10. Add decolorizing agent dropwise until liquid runs clear (~15 seconds)
    1. Take care to not over-decolorize
  11. Flood slide with safranin, allow to sit 1 minute
  12. Rinse slide gently but completely with R/O water
  13. Blot to remove most liquid, then air dry completely
  14. Observe with brightfield microscopy

Last modified: 24 January 2022; TBJ

Conjugation protocol for Z. mobilis using WM6026 donor strain

Notes

  • From Mehmet Tatli

Day 1:

   Morning

  • Inoculate 5mL ZMRG with single colony of mobilis ZM4 triple mutant and grow 24hrs anaerobically

Evening

  • Inoculate LB + 0.1mM DAP + any antibiotics needed with single colony of donor strain WM6026 containing plasmid you want to transfer

Day 2:

   Morning

  • Subculture WM6026 overnight in LB + 0.1mM DAP without antibiotic by inoculating at OD600=0.1-0.2 and grow until OD600=0.5

Midday

  • Dilute mobilis stationary phase overnight culture to OD600=0.5
  • Mix 1mL of OD600=0.5 WM6026 and 1mL OD600=0.5 mobilis in a 2mL eppie, then spin at max speed for 30 seconds (pipette gently from now on and be gentle with tube)
  • Decant supernatant from eppie, then let sit for 2-3 minutes to loosen pellet
  • Resuspend pellet in leftover media (that didn’t decant, should be ~100µL)
  • Place suspension in the middle of a mating plate as a single spot and incubate at 30C overnight

Day 3:

  • Add 1mL ZRMG to plate, pipetting up and down to ensure cells are mixed into suspension
  • Collect suspension in 2mL eppie tubes
  • Vortex vigorously for 30 seconds
  • Spin down cells and pipette off supernatant
  • Wash cells by adding 1mL ZRMG, then spinning cells down and pipetting off supernatant again
  • Carefully resuspend with 1mL ZRMG, then incubate at 30C for 4hrs in anaerobic chamber
  • After incubation, spread 100-500µL cells on ZRMG + appropriate antibiotic plate
  • Incubate plate 2-4 days for conjugant colonies to arise

Last modified: 14 July 2021; TBJ

 

E. coli Chemical Competency Cell Preparation

Notes:

  • This protocol was obtained from the Brian Pfleger lab.
  • After preparation, the transformation efficiency should be checked by transformation using plasmid DNA of known concentration.

Day 1:

  • Streak out frozen DH5α cells on an LB plate and grow overnight

Day 2:

  • Inoculate an isolated colony from LB plate into 2mL LB liquid medium. Grow overnight at 37o

Day 3:

  • Inoculate 1mL of overnight culture into 100mL LB medium (in 500mL flask).
  • Shake at 37oC to OD600 ~ 0.25 to 0.3
    Note: This usually takes about 1.5 to 2 hours
  • Chill culture, 0.1M CaCl2, and 0.1M CaCl2 (in 15% (v/v) glycerol) on ice for 15min.
  • Centrifuge cells at 3300×g at 4oC for 10min
  • Discard medium and resuspend cell pellet in 30-40mL cold CaCl2
  • Keep cells on ice for 15min
  • Centrifuge cells at 3300×g at 4oC for 10min
  • Remove supernantant and resuspend cell pellet in 6mL 0.1M CaCl2 (in 15% (v/v) glycerol)
  • Pipette 100µL of the cell suspension into sterile 1.5mL microcentrifuge tubes
  • Freeze tubes on dry ice then transfer to -80oC freezer

Transformation of DH5α E. coli

Protocol

  • Thaw aliquots of DH5α (prepared from coli Chemical Competency Cell Preparation protocol) from -80°C on ice. Briefly spin.
  • Add DNA to DH5α tubes and flick. Briefly spin.
    • This is super variable, but > 100 ng total is ideal.
  • Let tube sit on ice for 30 minutes.
  • Heat shock cultures at 42°C for 45 seconds.
  • Leave on ice for 3-6 minutes.
  • Add 900 µL of SOC media.
  • Grow for 1 hour at 37°C.
  • Spin cells at full speed for 1 minute.
  • Remove ~800-850 µL and resuspend in remaining volume.
  • Plate on prewarmed plates and grow at 37°C overnight.

Colony PCR and colony PCR template preparation for B. subtilis

Colony PCR Template preparation

  1. Add 20µL 0.02M NaOH to PCR tube.
  2. Use sterile loop or sterile toothpick to add small amount of colony to PCR tube. Swirl to mix.
  3. Incubate at 100°C for 10 minutes.
  4. Cool down on ice immediately.
  5. Centrifuge at max speed (16000g) for 5 minutes.
  6. Pipet supernatant to new PCR tubes without disturbing cell pellets.
  7. Measure by nanodrop.
    • If >500ng/µL, dilute 50 fold with nuclease-free dH2
    • If 10-50ng/µL, use without dilution (1-2µL will be used as template).

Colony PCR using above prepared template and 2x GoTaq master mix/Add Phusion protocol as well.

  1. Check annealing temps
    • Calculate temps by going to https://tmcalculator.neb.com/#!/main
    • Set “product group” as Taq DNA polymerase
    • Set “polymerase/kit” as Taq 2X master mix
    • Set “primer concentration” as 200nM
    • Input forward and reverse primer sequences and use calculated annealing temperature for PCR
  2. Assemble the following for each colony PCR:
Component 25µL Reaction
GoTaq (in green 2mL Eppendorf tube; already has loading dye added) 12.5µL
10µM forward primer 2µL
10µM reverse primer 2µL
Colony supernatant 1-2 uL
Nuclease-free water 6.5 or 7.5 µL depending on DNA

*if assembling in bulk, combine all components except DNA, then aliquot 23-24 µL to each PCR tube*.

  1. Use the following thermocycler conditions:
Step Temp Time
Initial denaturation 95°C 5-10min
35 cycles (35x) 95°C

Annealing temp

72°C

 30s

30s

1.5min/kb
Final extension 72°C 5min
Hold 12°C Infinity (but don’t leave PCR products overnight. This can cause condensation on the thermocycler, which will damage the machine.)
  1. Perform gel electrophoresis to check PCR results
    • Prepare 1-2% agarose by dissolving Ng agarose in 100N mL 1X TBE buffer.
    • Boil/microwave the agarose solution and swirl to mix.
    • Add 2-3µL ethidium bromide.
    • Pour warm solution into mold with comb and wait for it to solidify.
    • Wait until solid, then submerge in electrophoresis chamber with 1X TBE and remove comb.
    • Load 5-10µL PCR product per well and run at 90-100V for 45-55 min.
    • Visualize on gel dock. Wang lab has a gel imager with a printer.

Notes: If no bands are seen after electrophoresis, optimize PCR conditions with an annealing temperature gradient and with and without 3% DMSO.

Last modified: 5 August 2021; TBJ

Last modified: 10 August 2021; LNL

B.subtilis transformation using plasmids – two steps protocol (Noah’s)

Day 0

  1. Streak out WT subtilis NCIB 3610 onto LB plate to get a single colony (harvest at the age of less than or equal to 18 hrs)

Day 1

  1. Prepare 10X MC Media (competency media):

https://docs.google.com/document/d/1USu-RptmucLvrO7NcybNfYyWxTs6g6RapJsiFK9y5iU/edit?usp=sharing

Component For 100 ml For 200 ml For 500 ml
K2HPO4 (potassium phosphate dibasic) 10.7 g 21.4 g 53.6 g
KH2PO4 (potassium phosphate monobasic) 5.2 g 10.5 g 26.2 g
Glucose (dextrose) 20 g 40 g 100 g
Na3C6H5O7 * 2H2O (Sodium Citrate Dihydrate) 0.88 g 1.8 g 4.4 g
1000x Ferric Ammonium Citrate

(2.2 g Ferric Ammonium Citrate per 100 ml ddH2O *light sensitive compound*)

 1.0 ml 2.0 ml 5.0 ml
Casein Hydrolysate (oxoid) 1.0 g 2.0 g 5.0 g
Potassium Glutamate monohydrate (L-glutamic acid) 2.2 g 4.4 g 11 g
ddH2O To 100 ml To 200 ml To 500 ml

Culture preparation

  1. Prepare one tube with 2, 3, and 4 ml of LM media (LB + 3 mM MgSO4).
  2. Inoculate a single fresh colony of WT subtilis NCIB 3610 from day 0 (age less than 18 hrs) into the 4 ml LB tube and vortex.  Then, transfer 1 ml from the inoculated tube to the 3 ml tube and vortex well. Lastly, transfer 1 ml from the original 3 ml tube into 2 ml tube.
  3. Put under constant shaking overnight (~12 to 15 hours) at room temp (~25’C)

*Alternative method: Grow the culture in LM at 37 ‘C for 3 hours instead*

Day 2 (or day 1 if using the alternative method)

  1. Dilute to 1X MC before uses: 9 mL DI + 1 mL 10X MC + 33 ul of 1 M MgSO4 (final conc o3mM)
  2. *make extra amount – 1 ml for each transformation and extra 1 ml for 2 sets of cell resuspension.
  3. Acquire first and second tube (original 3 and 4 ml tube), which is supposedly at mid-exponential phase by the time of collection, and spin-down the culture at 8000 rpm for 1-2 min.
  4. Remove supernatant and resuspend in 1 ml of 1x PBS buffer before spinning-down the resuspension at 8000 rpm for 1-2 min. Pour off supernatant again.
  5. Resuspend the cells from two tubes in 500 ul 1XMC + 3 mM MgSO4 each
  6. Check for OD600 using 50 ul of MC culture and 950 ul of DI water in semi-micro cuvettes
  7. Calculate the amount of resuspension to make your total volume of MC culture required for all transformations to be at OD600 = 0.2 and inoculate the culture
  8. For example, if you need 4 ml of MC culture, calculate the amount of dilution from original concentrated culture to make your 4 ml culture at OD600 = 0.2
  9. Incubate cultures for 5 hours rolling or constant shaking at 37 ‘C
  10. At 5 hours, add ~2 to 4 ug of plasmid DNA (or ~200-400 ng of gDNA) to 1 ml of culture for each transformation
  11. Place each tube back to rolling or shaking at 37’C for 2 hours
  12. Plate 100 ul of the culture onto selectable medium plate (LB+appropriate antibiotics)
  13. Spin-down the rest of 900 ul culture at 8000 rpm for 1-2 minutes, then remove 800 ul supernatant (leave ~100 ml). Vortex the cells and the leftover supernatant before plate the concentrated cells onto selective medium plate
  14. Place all plates at 37’C

Day 3

  1. Restreak the surviving colony on the selectable medium plate again if the single colony cannot be obtained from the original selective plate (in case when many colonies were transformed and growing on top of one another).
  2. If the single colony can be obtain, grow that colony in 3 ml liquid LB at 37’C overnight, then mix 500 ul of the overnight culture with 500 ul 50/50 glycerol in a sterile frozen culture tube.

Store the created mutants/transformed colony at -80 ‘C

Bacillus subtilis biofilms in 12-well plates

Materials:

  • subtilis freezer stock
  • LB plate (1)
  • Culture tubes (2)
  • LB medium
  • Sterile wooden dowels
  • 12-well tissue culture plates, untreated
  • 50mL falcon tube (1)
  • Metal mesh, precut to fit into the wells of the plates and autoclaved.

Note: Meshes are essential for metabolite extraction. They are not essential to the lipid extraction protocol. However, we still recommend using mesh because it facilitates biofilm growth.

  • Forceps
  • 5mL pipet & tips
  • Spectrophotometer & cuvettes
  • 1mL pipet & tips
  • 5mL Eppendorf tube (1)
  • 20µL pipet & tips
  • Tupperware with lid
  • 50mL beaker

Day 1: Streak B. subtilis plate

  1. Streak an LB agar plate for single colonies of your B. subtilis strain of interest. Incubate overnight at 37°

Day 2: Start biofilms

  1. Inoculate LB cultures
    1. Label culture tubes appropriately. We recommend starting two cultures (one is sufficient, but it’s nice to have a backup).
    2. Aliquot 3mL LB per tube. Add antibiotics if needed.
    3. Ensure that all colonies on the plate look the same; we want to use a few colonies for inoculation, so it’s important to check for any signs of variation.
    4. Collect ~3 colonies from the plate using a sterile wooden dowel. Transfer to your culture tube, swish to disperse. Blobs of colony material will be visible in the LB. Repeat for 2nd culture tube.
    5. Incubate with shaking at 200rpm, 37°C, 2-2.5hrs.
  2. Prepare 12-well plates
    1. Unpackage the sterile 12-well plates. Do not open the lids. Label the plates with strain names for each well.

Note: The outer wells are good for biofilm growth, but the inner wells don’t grow nice, reliable biofilms. We suspect a lack of aeration. You can use the center wells as negative controls for growth.

  1. Flame the forceps. Pick up a mesh by the tab, lift the plate lid and place the mesh in the well at an angle so that all meshes can fit without hitting one another. Replace lid.
  2. Grab another mesh, repeat placing until all necessary wells of the plate are filled. Include a mesh in at least 1 center well as a negative control. No need to flame your forceps between each mesh unless you violate sterility.

Note: If you drop a mesh, just move on! Do not flame the mesh. You can re-autoclave unused/dropped meshes. Always prep a few extra meshes just in case.

  1. Pour ~50mL of MSgg from your stock bottle into a sterile 50mL falcon tube. Aliquot 4.5mL of MSgg to each well of the 12well plate using a 5mL pipet. You can use the same tip for all the wells until aliquot runs out. One falcon tube aliquot does ~11 wells.
  2. Refill the falcon from the MSgg from stock bottle, change pipet tips, and continue filling wells. Repeat until all wells are filled.
  1. Check OD600 for growing LB cultures. ODs should be ~0.75. It’s important to inoculate the biofilms with B. subtilis cultures during log phase growth.
  2. Inoculate biofilms
    1. Aliquot 1mL of LB culture inoculum into an Eppendorf tube (this makes it easier to maintain sterile technique, since the culture tubes are very long and the P20 doesn’t fit nicely for sterile sampling). This is your inoculum.
    2. Add 6uL of inoculum to each of the outside wells. Leave the 2 middle wells UN-inoculated as negative controls (e. if anything grows in the middle wells, you know something is contaminated in your experiment).

Note: It’s not visually obvious when a well has been inoculated, so use consistent patterns and track your progress carefully.

  1. Incubate
    1. Transfer the 12-well plates gently to a Tupperware labeled with your name and lab info. Add a 50mL beaker of water to maintain humidity. Put on the lid.
    2. Put the tupperware in a clear spot in the 37°C room of the 5th floor core. Make sure lid is loosely set on (do not secure closures/snaps) and open the venting valve.