Electrotransformation

Competent Cell Preparation

Choice of Culturing Conditions

Two techniques have been 

For n transformations of v volume:

Materials

1. SOB (+antibiotic if necessary)

   Tryptone	20 g/L	–	Dissolve with stir bar. Check pH.
   Yeast extract	5 g/L	–
   NaCl	0.5 g/L	8.56 mM
   KCl	0.746 g/L	2.5 mM
   Milli-Q water	Reserve Mg soln volumes
   NaOH, 1 M	→ pH 7.0
   Autoclave 15 min ≤ 1 L, 20 min 1–1.4 L. Or filter sterilize* after adding components below.
   MgCl2 sterile	10 mL/L 1 M	10 mM	autoclaved or filtered*
   MgSO4 sterile	10 mL/L 1 M		autoclaved or filtered*
   Glucose sterile	20 mL/L 1 M 7.2 mL/L 50% 18 mL/L 20%	20 mM 3.6 g/L	For SOC only. Can be spiked-in.
   Store at 4° for best long-term preservation of nutrients. Aliquots of SOC can be frozen to inhibit contaminant growth. * Detergent-free membranes (PES, NYL, CN) are preferred for filter-sterilization.

2. Culture tubes for precultures (detergent-free).
3. Culture flasks large enough to hold growth medium volume (detergent-free).
4. Refrigerated shaking incubator space for the culture flasks.
   • Only a refrigerated shaker can sustain 18–23°C temperature.
5. Ice bucket/tray large enough to swirl flasks in. Access to ice.
6. Chilled centrifuge bottles/lids or tubes appropriate for holding culture volumes and balancing. (detergent-free)
   • Bottles must be able to collectively hold culture(s) without any exceeding 80% capacity. Filling bottles higher will result in leakage into rotor.
7. Chilled sterile water to balance multiple centrifuge bottles, if needed.
8. 4°C centrifuge and rotor compatible with centrifuge bottles.
9. (opt.) A serological pipette at -20°C for adding n v volume CCMB80 before going to cold room.
10. Labeled cryoboxes for comp cell aliquots, prechilled at -80°C.
11. (opt.) Liquid nitrogen, dewar, and slotted ladle, if flash freezing.
12. Materials on a mobile lab bench cart for the cold room:
    1. n tubes (e.g. 200 µL tubes) for cell aliquots, labeled/marked and arranged in clean tube racks or tip rack+boxes. Blade, if planning to cut apart tube strips.
    2. Extra tubes or bag of tubes, for aliquotting any excess comp cells, if desired.
    3. Serological pipettor and few pipettes appropriate for resuspending and transferring cells.
    4. A 5 or 50 mL tube per strain to hold resuspension.
    5.  (opt.) Electronic repeater pipette and a 5 mL combitip per strain and some extras. Reservoirs, if opting to us a multichannel pipette.
    6. Micropipette and tips appropriate for aliquot volumes ≥v. (Sometimes useful to aliquot remaining volume even when using electronic repeater pipette.)
    7. Ethanol spray
    8. Paper towels
    9. Extra gloves
    10. Trash bin/bag
    11. (opt.) If flash freezing, box/bag to collect all tubes in before flash-freezing.
        

Protocol

1. Grow preculture to saturation:
   Inoculate ≥1/400 culture volume, 0.05nv  , SOB (+selection, if necessary) with colonies (best) or a scrape of frozen stock, and incubate shaking 18–22°C 12–18 hr until saturated.
   • Use a fresh, trusted source of the strain. For most reliable results, inoculate a seed culture from colonies/patch from <2 week-old plates streaked from a frozen stock. Plasmid-bearing strains ought to be used from even fresher plates to minimize plasmid mutations.
   • Incubation of preculture at 30° or 37°C seems to be fine for 10⁹/µg efficiency , though 20–23°C is recommended in the original protocol.
   • NEB Turbo grown in 20°C SOB  saturates at OD≈3.5.
2. Inoculate 25nv mL SOB with ≈0.2% saturated seed culture, to a final OD₆₀₀≈0.01.
   Incubate 18–22°C, shaking 250 rpm for flasks.
   Grow culture to OD₆₀₀=0.2–0.26 (early exponential phase), 9–15 hr. Periodically monitor OD starting at 8 hr.
   • Transformation efficiency from LB cultures decreases linearly between OD=0.3–0.6. ⁽¹⁾  The same might apply here.  Do not allow the culture to grow to an OD ₆₀₀  > 0.26 for maximum efficiency.
   • Several tenfold lower-efficiency at stationary phase, but supposedly still fine for transforming pure plasmid or the simplest cloning.
   • ≈12–15 hr required for 18° culturing, less for higher temperatures.
3. Prepare the mobile lab bench cart for the cold room as described. Prechill both the mobile lab bench cart (with its contents) and the centrifuge to 4°C ≥45 min before cell harvest (next step)  ^{[measured with infrared thermometer]} .
4. When culture reaches OD₆₀₀≈0.2–0.26, immediately harvest cells: decant into prechilled centrifuge bottles and balance them. Use chilled sterile water if necessary.
   Centrifuge 1000×g, 10–15 min (depending on volume) in a prechilled 4°C centrifuge.
5. Check for a small pellet. Gently decant medium away from pellet, shaking the bottle to drain. Absorb the last of the medium on the lip with a paper towel. Return bottles to ice.
6. Gently resuspend cells in ⅓ culture volume (8nv mL) of chilled CCMB80 buffer, by swirling. Combine volumes to reduce vessels.
7. Incubate suspension on ice 20 min.
8. Harvest cells as before (step 4–5).
9. Gently resuspend in 1/25 volume (nv mL) chilled CCMB80 buffer by swirling and striking.
   
   • Flat-bottom centrifuge bottles allow easier gentle resuspension of pellets.
   • Original Hanahan / CSH protocol instructs 12-fold concentration in CCMB80 buffer. The OpenWetware version instructs 25-fold concentration.
   
   Perform remaining steps with pre-chilled materials in the cold room for maximum efficiency.
   Decant or pipette cells into 5 or 50 mL conical tubes on ice for easier access with pipette for aliquotting. Finish gently resuspending any visible remaining cell clumps using P1000 tip if necessary.
10. Aliquot into tubes in the cold room (for top efficiency) or on cold block or ice. You can use a repeater pipette or multichannel pipette from a chilled reservoir.
11. Cap the tubes while they are racked, and slice apart if using tube strips. Minimize touching the tubes to keep them cool. Check that all caps are fully in the tube.
    Dislodge tubes from racks into a bag/box without touching the bottom (your hands are warm). This is easiest by pushing them out from the bottom using a spare rack or tip box.
12. (opt.) Flash freeze in liquid nitrogen for supposedly higher efficiency.
    • Flash freezing improves competence, but simply freezing is not estimated to noticeably reduce competence, maybe within twofold ^[SPB].
    • Residual ethanol from a dry-ice-ethanol bath fails to dry from tubes/boxes at -80°C. Freeze tubes in a bag if using this method, and pour tubes into box.
13. Quickly move tubes to prechilled, labeled -80°C freezer boxes, either directly from 4°C, or if flash freezing, directly ladled out of liquid nitrogen dewar or dry-ice/ethanol bag (so as not to heat tubes).
    If not freezing, proceed to transformation right away.

Detergent Residue

According to Tom Knight ⁽⁴⁾ : Detergent is a major inhibitor of competent cell growth and transformation. Glass and plastic must be detergent-free for these protocols. The easiest way to do this is to avoid washing glassware and simply rinse it out. Autoclaving glassware filled ¾ with deionized water is an effective way to remove most detergent residue. Media and buffers should be prepared in detergent-free glassware and cultures should be grown in detergent-free glassware.

According to the Cold Spring Harbor protocol ⁽⁷⁾ : "Detergents and organic contaminants are strong inhibitors of transformation reactions. To avoid problems caused by residual detergent in glassware, use disposable plastic tubes and flasks wherever possible for preparation and storage of all solutions and media used in transformation. Organic contaminants present in the H₂ O used to prepare transformation buffers can reduce the efficiency of transformation of competent bacteria. H ₂ O obtained directly from a well-serviced Milli-Q filtration system (Millipore) usually gives good results. If problems should arise, treat the deionized H ₂ O with activated charcoal before use."

OM_Eporator_4309_900_010-05_052016_en.fm (eppendorf.com)

Transformation

Summary for frozen lab aliquots

1. Thaw comp cell aliquots (50–100 µL) slowly on ice or above cold block for a few min.
   Use one aliquot per assembly reaction. 
   Prechill electroporation cuvette and DNA on ice. Label recovery tubes or block seal.
   Prewarm recovery medium to 37° (opt.)
2. Add pure DNA ≤10 µL to cells and mix twice by pipetting.
3. Incubate cold on 1°C block / ice for 5 min (opt.)
4. Load cell+DNA suspension to cuvette.
5. Electroporate: tap cuvette on table, wipe electrodes, and electroporate at 1.7 kV for 1 mm cuvette.
6. Recovery: within a few seconds, resuspend cells in ½–1 mL SOC and move to recovery vessel.
   Incubate at growth temp (e.g. 37°C), 1 hr shaking, 2–3 hr for nonselective integration.
7. Plate on selective agar and incubate.

Detailed

To wash and reuse electroporation cuvettes:

• Fill cuvette with 10% bleach to brim, no more than 20 min.
  Pour out bleach and flick out remnants.
• Fill with tap water; flick out water × 5
• Fill with MilliQ water; flick out water × 5
• Dry in a 37° rack