Difference: ProtocolsRestrictionEnzymeCloning (1 vs. 5)

Revision 52018-03-22 - JeffreyBarrick

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META TOPICPARENT name="ProtocolList"

Restriction Enzyme Cloning

Restriction enzyme cloning is a bread-and-butter technique in molecular biology for modifying plasmids to contain genes or other DNA sequences of interest. While it may be more time consuming than some recently developed techniques, it is very reliable.

Changed:
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For background on restriction enzyme cloning and some pretty pictures, check out the wiki on the topic, and check out the useful diagram on this Chinese website.
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For background on restriction enzyme cloning and some pretty pictures, check out the Wikipedia page on this topic.
 
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  1. Inoculate 5 mL of liquid LB media plus the appropriate antibiotic each with a colony with insert of correct size. Grow overnight
  2. Isolate plasmid from overnight culture using a miniprep kit.
  3. Sequence the insert using the same primers you used to verify insert size. Analyze sequence to confirm correct insert.
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-- Main.MichaelHammerling - 01 Feb 2012
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Revision 42012-02-06 - JeffreyBarrick

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META TOPICPARENT name="ProtocolList"

Restriction Enzyme Cloning

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  1. Sequence the insert using the same primers you used to verify insert size. Analyze sequence to confirm correct insert.

-- Main.MichaelHammerling - 01 Feb 2012

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Revision 32012-02-01 - MichaelHammerling

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META TOPICPARENT name="ProtocolList"

Restriction Enzyme Cloning

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Materials

Deleted:
<
<
Plasmid Marker
pJEB11 →Ara
pJEB12 →Ara+
pJEB15 →Mal
 
  • Cloning plasmid (such as pUC19) and template for DNA fragment to be cloned
Deleted:
<
<
  • Electro- or chemically competent cells.
 
  • Restriction Enzymes
Added:
>
>
  • PCR Reagents
 
  • DNA Ligase
  • Electrocompetent cells or chemically competent cells of an appropriate cloning strain.
  • Antibiotic Plates to your cloning plasmid and/or the antibiotic resistance gene you are cloning
Line: 39 to 31
 
  1. Perform a restriction digest of both the cloning plasmid and the DNA fragment to be cloned to generate a linearized plasmid and DNA fragment with sticky ends. When considering how much DNA to add to the reaction, too much is preferable to too little. If performing a double digest (two restriction enzymes at the same time), be sure to use a buffer in which both enzymes have activity. Follow this protocol and the protocols that came with the restriction enzyme to plan your reaction.
  2. Run the linearized plasmid and the digested fragment to be cloned (the whole reaction for both) on an agarose gel. Be sure to include uncut plasmid and uncut DNA fragment as controls in separate lanes so you can identify the "cut" version of each. Cut out your linearized plasmid and digested DNA fragment and purify them with a gel purification kit. Quantify DNA.
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Optional Step 5: Phosphatase plasmid

>
>

Optional Step 4: Phosphatase plasmid

  If a high background of colonies of ligated vector (with no insert) is a problem (as may be the case when using only one restriction enzyme), you may use Calf Intestinal Phosphatase (CIP) to remove the phosphates from your linearized plasmid before proceeding to ligation. This will ensure that the linearized plasmid cannot ligate to itself, but must instead ligate to the insert to form a circular plasmid. You can likely ignore this step if you performed a double digest in the previous step.

Revision 22012-02-01 - MichaelHammerling

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META TOPICPARENT name="ProtocolList"
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Restriction Enzyme Cloning

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pJEB15 →Mal
Changed:
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<
  • Cloning plasmid (such as pUC19) and DNA fragment to be cloned
>
>
  • Cloning plasmid (such as pUC19) and template for DNA fragment to be cloned
  • Electro- or chemically competent cells.
 
  • Restriction Enzymes
  • DNA Ligase
  • Electrocompetent cells or chemically competent cells of an appropriate cloning strain.
Line: 28 to 28
 

Step 1: Design Primers

Deleted:
<
<
  1. Thaw one 1.7 ml microfuge tube of electrocompetent cells for each parent strain on ice.
  2. While waiting for this to thaw, place one electroporation cuvette on ice for each parent strain.
  3. Add 1 l of pACBSR and 1 l of the donor plasmid to each tube of electrocompetent cells on ice. Mix by flicking the tube with your finger. Do not pipette up and down or vortex to mix.
  4. Electroporate
  5. Pipette sample out of cuvette into original microfuge tube on ice. Add 500 l of room temperature SOC medium.
  6. Grow at 37C for 1 hr in a shaking incubator to induce antiobiotic expression.
  7. Plate 100 l and 10 l + 90 l saline on two separate LB + Cam + Kan plates.
  8. Grow plates overnight at 37C.

Step 2: Perform PCR on Template to amplify desired product with restriction sites

  1. Pick three colonies from each successful transformation into separate test tubes containing 5 ml of LB medium supplemented with 0.2% L-Arabinose and 20 g/ml Chloramphenicol (Cam). The arabinose is to induce expression of the λ Red genes from the gene-gorging plasmid. Use arabinose regardless of which sugar marker you are changing.
  2. Grow cultures overnight at 37C, shaking at 120 rpm.

Day 3: Screen or Select for Desired Mutation

 
Changed:
<
<
  1. Make a 104-fold dilution of each overnight culture with two 100 l transfers through dilution tubes containing 10 ml of saline. Note that the cell density achieved after induction is considerably lower than that usually achieved during growth in LB.
  2. A. If you are gene gorging the version of the the marker you must screen for successful mutations. Plate 200 l of a 104 dilution of each culture on tetrazolium arabinose (TA), tetrazolium maltose (TM), or other suitable indicator agar containing 20 g/ml Chloramphenicol (Cam). Successful mutants will be red.
    B. If you are gene gorging the + version of the the marker you could screen for successful mutations as above, but it is easier to select for them by plating 100 l of a 102 dilution on minimal arabinose (MA) or minimal maltose (MM).
  3. Grow plates overnight at 37C.
>
>

Step 2: Perform PCR on template to amplify desired product with restriction sites

 
Changed:
<
<

Day 4: Screen for Gene-Gorging Plasmid Loss

>
>
  1. Using the primers you have designed to add restriction enzyme sites to the ends of your PCR fragment, perform PCR on your template to obtain your fragment.
  2. Analyze your PCR product via agarose gel electrophoresis to ensure that you have obtained a product of the correct size. Use a gel extraction kit to perform a gel extraction of the desired band, or if the product is very pure, use a DNA purification kit to purify the DNA. Quantify DNA concentration.
 
Changed:
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Expected results are 0-10 colonies with the marker change per 200 colonies of the other color.
>
>

Step 3: Restriction enzyme cleavage

 
Changed:
<
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  1. Pick one colony from each plate into 5 ml of LB medium in a test tube. Give each picked colony an isolate designation. Colonies from the same plate are not necessarily independent isolates, they may share undesired second-dite mutations. Picking individual isolated
  2. Grow cultures overnight at 37C, shaking at 120 rpm.
  3. Grow plates overnight at 37C.
>
>
  1. Perform a restriction digest of both the cloning plasmid and the DNA fragment to be cloned to generate a linearized plasmid and DNA fragment with sticky ends. When considering how much DNA to add to the reaction, too much is preferable to too little. If performing a double digest (two restriction enzymes at the same time), be sure to use a buffer in which both enzymes have activity. Follow this protocol and the protocols that came with the restriction enzyme to plan your reaction.
  2. Run the linearized plasmid and the digested fragment to be cloned (the whole reaction for both) on an agarose gel. Be sure to include uncut plasmid and uncut DNA fragment as controls in separate lanes so you can identify the "cut" version of each. Cut out your linearized plasmid and digested DNA fragment and purify them with a gel purification kit. Quantify DNA.
 
Changed:
<
<

Day 5: Plate to Single Colonies

>
>

Optional Step 5: Phosphatase plasmid

 
Changed:
<
<
  1. Plate 100 l of a 106 dilution of each culture on tetrazolium arabinose (TA, blue stripe) or tetrazolium maltose (TM, purple stripe). The dilution can be made with three 100 l transfers through dilution tubes containing 10 ml of saline.
  2. Grow plates overnight at 37C.
>
>
If a high background of colonies of ligated vector (with no insert) is a problem (as may be the case when using only one restriction enzyme), you may use Calf Intestinal Phosphatase (CIP) to remove the phosphates from your linearized plasmid before proceeding to ligation. This will ensure that the linearized plasmid cannot ligate to itself, but must instead ligate to the insert to form a circular plasmid. You can likely ignore this step if you performed a double digest in the previous step.
 
Changed:
<
<

Day 6: Patch for Plasmid Loss

>
>

Step 5: Ligate

 
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  1. Patch 6-12 colonies from each plate on three LB (or TA/TM) plates with 20 g/ml Chloramphenicol, with Kanamycin, and - last - with no antibiotic.
  2. Grow plates overnight at 37C.
>
>
Following the protocol for NEB T4 DNA Ligase, ligate your product into your linearize plasmid.
 
Changed:
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Day 7: Save a Freezer Stock of the New Strain

>
>

Step 6: Transform ligation reaction

 
Changed:
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<
  1. Pick from a patch that grows without antibiotic and not with either antibiotic present. Usually a majority of patched colonies have successfully lost both plasmids.
  2. Grow cultures overnight at 37C and archive 2 × 1 ml frozen copies in 10% glycerol.
>
>
  1. Depending on the concentration of DNA used in the ligation reaction, transform 1-5 microliters of your ligation reaction into electrocompetent cells or chemically competent cells.
  2. If transforming a selectable marker such as antibiotic resistance, plate the appropriate amount of transformed cells on plates containing the selective agent, grow overnight at 37 degrees (or the appropriate temperature for your plasmid, if other) and proceed to Step 7. If performing a blue/white screen or other kind of screen, proceed to Step 6A.
 
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Test for Marker Neutrality

>
>

Step 6A: Blue/White Screening

If cloning a nonselectable piece of DNA, you will have to perform a screen of some kind to verify its presence. A common screen of this type is the blue/white screen. This is done with plasmids such as pUC19, where the multiple cloning site is located in the middle of the LacZ gene. As a result, a successful clone will disrupt the LacZ gene. Be sure to use an appropriate LacZ knockout strain when doing a blue/white screen.
  1. Make plates to perform the blue/white screen by spreading X-Gal and IPTG on plates with the appropriate antibiotic for your plasmid. Stocks of X-Gal and IPTG should be available in the -20. If not, make your own!
  2. Plate the appropriate amount of transformation on the plates and grow at 37 degrees overnight.
  3. Plates the next day should have some blue, but mostly white colonies. Blue colonies indicate functional LacZ, which means that your insert was NOT cloned successfully. Therefore, you want to test white colonies, which indicate that LacZ has been disrupted. Pick several of these colonies and proceed to step 7
 
Changed:
<
<
Competitive fitness assays should be used to test for non-neutral mutations that sometimes occur during strain construction by this method. Testing three independent isolates usually assures one success.

References

  1. Herring, C.D., Glasner, J.D., and Blattner, F.R. (2003) Gene replacement without selection: regulated suppression of amber mutations in Escherichia coli. Gene 311, 153-163.
  2. Sean Sleight's Detailed Procedure.
>
>

Step 7: Verify insert via PCR and sequencing

 
Added:
>
>
  1. Perform colony PCR using a fraction of each colony on 5-10 colonies using primers outside the multiple cloning site of your vector. Use Taq polymerase for this. Run products on an agarose gel and check for product of expected size if insert is present.
  2. Inoculate 5 mL of liquid LB media plus the appropriate antibiotic each with a colony with insert of correct size. Grow overnight
  3. Isolate plasmid from overnight culture using a miniprep kit.
  4. Sequence the insert using the same primers you used to verify insert size. Analyze sequence to confirm correct insert.
  -- Main.MichaelHammerling - 01 Feb 2012

Revision 12012-02-01 - MichaelHammerling

Line: 1 to 1
Added:
>
>
META TOPICPARENT name="ProtocolList"

Restriction Enzyme Cloning

Restriction enzyme cloning is a bread-and-butter technique in molecular biology for modifying plasmids to contain genes or other DNA sequences of interest. While it may be more time consuming than some recently developed techniques, it is very reliable.

For background on restriction enzyme cloning and some pretty pictures, check out the wiki on the topic, and check out the useful diagram on this Chinese website.

Materials

Plasmid Marker
pJEB11 →Ara
pJEB12 →Ara+
pJEB15 →Mal

  • Cloning plasmid (such as pUC19) and DNA fragment to be cloned
  • Restriction Enzymes
  • DNA Ligase
  • Electrocompetent cells or chemically competent cells of an appropriate cloning strain.
  • Antibiotic Plates to your cloning plasmid and/or the antibiotic resistance gene you are cloning
  • XGal and IPTG (for blue/white screens)

Step 1: Design Primers

  1. Thaw one 1.7 ml microfuge tube of electrocompetent cells for each parent strain on ice.
  2. While waiting for this to thaw, place one electroporation cuvette on ice for each parent strain.
  3. Add 1 l of pACBSR and 1 l of the donor plasmid to each tube of electrocompetent cells on ice. Mix by flicking the tube with your finger. Do not pipette up and down or vortex to mix.
  4. Electroporate
  5. Pipette sample out of cuvette into original microfuge tube on ice. Add 500 l of room temperature SOC medium.
  6. Grow at 37C for 1 hr in a shaking incubator to induce antiobiotic expression.
  7. Plate 100 l and 10 l + 90 l saline on two separate LB + Cam + Kan plates.
  8. Grow plates overnight at 37C.

Step 2: Perform PCR on Template to amplify desired product with restriction sites

  1. Pick three colonies from each successful transformation into separate test tubes containing 5 ml of LB medium supplemented with 0.2% L-Arabinose and 20 g/ml Chloramphenicol (Cam). The arabinose is to induce expression of the λ Red genes from the gene-gorging plasmid. Use arabinose regardless of which sugar marker you are changing.
  2. Grow cultures overnight at 37C, shaking at 120 rpm.

Day 3: Screen or Select for Desired Mutation

  1. Make a 104-fold dilution of each overnight culture with two 100 l transfers through dilution tubes containing 10 ml of saline. Note that the cell density achieved after induction is considerably lower than that usually achieved during growth in LB.
  2. A. If you are gene gorging the version of the the marker you must screen for successful mutations. Plate 200 l of a 104 dilution of each culture on tetrazolium arabinose (TA), tetrazolium maltose (TM), or other suitable indicator agar containing 20 g/ml Chloramphenicol (Cam). Successful mutants will be red.
    B. If you are gene gorging the + version of the the marker you could screen for successful mutations as above, but it is easier to select for them by plating 100 l of a 102 dilution on minimal arabinose (MA) or minimal maltose (MM).
  3. Grow plates overnight at 37C.

Day 4: Screen for Gene-Gorging Plasmid Loss

Expected results are 0-10 colonies with the marker change per 200 colonies of the other color.

  1. Pick one colony from each plate into 5 ml of LB medium in a test tube. Give each picked colony an isolate designation. Colonies from the same plate are not necessarily independent isolates, they may share undesired second-dite mutations. Picking individual isolated
  2. Grow cultures overnight at 37C, shaking at 120 rpm.
  3. Grow plates overnight at 37C.

Day 5: Plate to Single Colonies

  1. Plate 100 l of a 106 dilution of each culture on tetrazolium arabinose (TA, blue stripe) or tetrazolium maltose (TM, purple stripe). The dilution can be made with three 100 l transfers through dilution tubes containing 10 ml of saline.
  2. Grow plates overnight at 37C.

Day 6: Patch for Plasmid Loss

  1. Patch 6-12 colonies from each plate on three LB (or TA/TM) plates with 20 g/ml Chloramphenicol, with Kanamycin, and - last - with no antibiotic.
  2. Grow plates overnight at 37C.

Day 7: Save a Freezer Stock of the New Strain

  1. Pick from a patch that grows without antibiotic and not with either antibiotic present. Usually a majority of patched colonies have successfully lost both plasmids.
  2. Grow cultures overnight at 37C and archive 2 × 1 ml frozen copies in 10% glycerol.

Test for Marker Neutrality

Competitive fitness assays should be used to test for non-neutral mutations that sometimes occur during strain construction by this method. Testing three independent isolates usually assures one success.

References

  1. Herring, C.D., Glasner, J.D., and Blattner, F.R. (2003) Gene replacement without selection: regulated suppression of amber mutations in Escherichia coli. Gene 311, 153-163.
  2. Sean Sleight's Detailed Procedure.

-- Main.MichaelHammerling - 01 Feb 2012

 
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