Haynes:TransformationPlasmids

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(Quick Method: Chemically competent cells + plasmid mini prep, Quick Method)
(Traditional Method: Chemically competent cells + ligation)
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<- [[Haynes:Protocols | Back to Protocols]]
<- [[Haynes:Protocols | Back to Protocols]]
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=Traditional Method: Chemically competent cells + plasmid mini prep=
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=Traditional Method: Chemically competent cells + ligation=
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# Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
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<div style="width: 800px">
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# Incubate DH5α Turbo competent cells on ice just until thawed. You will need 30 μL cells per ligation.
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This method is useful for increasing the efficiency of plasmid up-take.  
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Thaw TSS cells on ice.
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Add DNA, pipette gently to mix (1μl of prepped plasmid is more than enough).
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# Incubate chemically competent cells on ice just until thawed. You will need 30 μL cells per ligation.
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Note: If you are adding small volumes (~1μl), be careful to mix the culture well. Diluting the plasmid back into a larger volume can also help.
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# Add 30 μL thawed cells to to 1 sterile 2.0 mL tube (per ligation).
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Let sit for 30 minutes on ice.
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# Add the total ligation reaction to the cells in each 2.0 mL tube. Pipette up and down '''gently''' to mix the cells and DNA.
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Note: If you are in a rush, you can shorten this incubation time to 5-10 min.
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# Incubate on ice for 10 min.
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Incubate cells for 30 seconds at 42oC.
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# Heat shock: Transfer the tubes to 42°C for exactly 45 seconds (heat shock) on a heat block or water bath, then immediately place the tubes on ice for 1 minute.
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Note: According to the original TSS paper and qualitative experience (JM), this step is completely optional and may actually reduce transformation efficiency.
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# Add 750 μL sterile SOC medium to each sample.
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I tested this with DH5a Z1 and pUC19 and found that heat shock at 42C for 30 sec improved transformation efficiency 10-fold (Paul Jaschke)
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# Recovery: Close the caps tightly. Place the tubes in the shaking incubator, secured in a sideways position with tape. Incubate the tubes, with shaking, at 37°C for 45 minutes.
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Incubate cells on ice for 2 min.
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# Pre-warm the agar plates: Incuabte the selection agar plates (one per sample) at 37°C during the 45 minute recovery period.
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Add 1 mL SOC (2XYT and LB are also suitable, original paper suggests LB + 20mM glucose) at room temp.
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# Pellet the cells by centrifugation at top speed for 3 minutes.
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Incubate for 1 hour at 37oC on shaker.
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# Discard the SOC supernatant.
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Note: Can also save some time here by reducing incubation to ~45 min.
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# Resuspend the pellet in 100 μL LB medium (plus proper antibiotic).
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Spread 100-300 μl onto a plate made with appropriate antibiotic.
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# Pipette the total volume of cells onto the agar; spread using sterile glass beads.
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Grow overnight at 37 °C.
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# Incubate the inverted plate(s) overnight at 37°C to get colonies.  
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Save the rest of the transformants in liquid culture at 4 °C. If nothing appears on your plate, you can spin this down, resuspend in enough medium to spread on one plate and plate it all. This way you will find even small numbers of transformants.
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Note: To store the colonies long term, seal the plate with parafilm and keep the plate at 4°C (inverted).
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hugh kingston 09:46, 4 February 2008 (CST):I tried a few variations on this protocol, and found
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</div>
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using SOC instead of LB + 20mM glucose increased efficiency 3 fold
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heat shock of 42°C 45s increased efficiency 15-20 fold compared to no heat shock
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<br><br>
=Quick Method: Chemically competent cells + plasmid mini prep=
=Quick Method: Chemically competent cells + plasmid mini prep=
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# Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
# Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
# Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).
# Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).
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Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.
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Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.<br>
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Note 2: This method can be used for ligations, where construction is very straight-forward and your DNA material is highly concentrated. It works well for "forced" ligations (sticky overhangs, no unwanted annealing)
</div>
</div>
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=Chemically competent cells + iGEM Registry sample, Quick Method=
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<br><br>
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=Quick Method: Chemically competent cells + iGEM Registry sample=
<div style="width: 800px">
<div style="width: 800px">
# Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
# Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
-
# Incubate DH5α Turbo competent cells on ice just until thawed. You will need 30 μL cells per ligation.
+
# Incubate DH5α Turbo competent cells on ice just until thawed. You will need 30 μL cells per plasmid sample.
# <font color="blue">Locate the desired well in the Registry plate. Inject 10 μL sterile dH<sub>2</sub>O into the well and pipette up and down to resuspend the dried DNA and indicator dye. Transfer each DNA solution to a  sterile 0.5 mL tube.</font>
# <font color="blue">Locate the desired well in the Registry plate. Inject 10 μL sterile dH<sub>2</sub>O into the well and pipette up and down to resuspend the dried DNA and indicator dye. Transfer each DNA solution to a  sterile 0.5 mL tube.</font>
# Make a "negative control" sample by simply putting 10 μL sterile dH<sub>2</sub>O in a tube without DNA.
# Make a "negative control" sample by simply putting 10 μL sterile dH<sub>2</sub>O in a tube without DNA.
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# Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
# Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
# Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).
# Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).
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Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.
Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.
</div>
</div>

Revision as of 19:48, 4 March 2013

<- Back to Protocols

Traditional Method: Chemically competent cells + ligation

This method is useful for increasing the efficiency of plasmid up-take.

  1. Incubate chemically competent cells on ice just until thawed. You will need 30 μL cells per ligation.
  2. Add 30 μL thawed cells to to 1 sterile 2.0 mL tube (per ligation).
  3. Add the total ligation reaction to the cells in each 2.0 mL tube. Pipette up and down gently to mix the cells and DNA.
  4. Incubate on ice for 10 min.
  5. Heat shock: Transfer the tubes to 42°C for exactly 45 seconds (heat shock) on a heat block or water bath, then immediately place the tubes on ice for 1 minute.
  6. Add 750 μL sterile SOC medium to each sample.
  7. Recovery: Close the caps tightly. Place the tubes in the shaking incubator, secured in a sideways position with tape. Incubate the tubes, with shaking, at 37°C for 45 minutes.
  8. Pre-warm the agar plates: Incuabte the selection agar plates (one per sample) at 37°C during the 45 minute recovery period.
  9. Pellet the cells by centrifugation at top speed for 3 minutes.
  10. Discard the SOC supernatant.
  11. Resuspend the pellet in 100 μL LB medium (plus proper antibiotic).
  12. Pipette the total volume of cells onto the agar; spread using sterile glass beads.
  13. Incubate the inverted plate(s) overnight at 37°C to get colonies.

Note: To store the colonies long term, seal the plate with parafilm and keep the plate at 4°C (inverted).



Quick Method: Chemically competent cells + plasmid mini prep

  1. Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
  2. Incubate DH5α Turbo competent cells on ice just until thawed. You will need 30 μL cells per plasmid sample.
  3. Dilute 0.5 μL plasmid DNA (concentration not important) in 10 μL sterile dH2O in sterile 0.5 mL tubes.
  4. Make a "negative control" sample by simply putting 10 μL sterile dH2O in a tube without DNA.
  5. Add 30 μL thawed cells each tube of diluted DNA (and the negative control). Immediately place on ice and incubate for 10 min. (Do not heat shock; No 30 min. recovery is required for Amp or Kan resistance)
  6. Label the pre-warmed plates with the antibiotic name, strain name, DNA name, your initials, and the date.
  7. Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
  8. Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).

Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.
Note 2: This method can be used for ligations, where construction is very straight-forward and your DNA material is highly concentrated. It works well for "forced" ligations (sticky overhangs, no unwanted annealing)



Quick Method: Chemically competent cells + iGEM Registry sample

  1. Warm selection agar plates at 37°C (one for each plasmid, plus one for a zero plasmid control) for at least 15 min.
  2. Incubate DH5α Turbo competent cells on ice just until thawed. You will need 30 μL cells per plasmid sample.
  3. Locate the desired well in the Registry plate. Inject 10 μL sterile dH2O into the well and pipette up and down to resuspend the dried DNA and indicator dye. Transfer each DNA solution to a sterile 0.5 mL tube.
  4. Make a "negative control" sample by simply putting 10 μL sterile dH2O in a tube without DNA.
  5. Add 30 μL thawed cells each tube of diluted DNA (and the negative control). Immediately place on ice and incubate for 10 min. (Do not heat shock; No 30 min. recovery is required for Amp or Kan resistance)
  6. Label the pre-warmed plates with the antibiotic name, strain name, DNA name, your initials, and the date.
  7. Pipette the total volume of cells + DNA onto the agar; spread using sterile glass beads.
  8. Incubate the inverted plate(s) overnight at 37°C to get colonies. Seal the plate with parafilm and store the plate at 4°C (inverted).

Note: The negative control will show you the number of “background” colonies so that you can determine whether your transformation worked, or is just the result of selection failure.

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