Table of Contents

Table of contents DNA Methods Preparation of plasmid minipreps Zymo Plasmid Midiprep Kit Designing and storing ss-DNA oligonucleotide primers Quantifying DNA Sequencing DNA Cell Culture Methods Streaking out cultures from frozen glycerol stocks Preparation of liquid cultures of E. coli Preparation of frozen glycerol stocks of E. coli Preparation of competent cells Transformation of competent cells Transformation of Z Competent cells Growing an overnight culture Preparation of Liquid Media LB medium TB medium Ligation protocols Standard ligation protocol Quick Ligation™ protocol Polymerase Chain Reaction Techniques Basic PCR protocol Touchdown PCR protocol Site directed mutagenesis by megaprimer PCR Megaprimer PCR Cloning into expression vector SiteDirected mutagenesis by Megaprimer QuickChange 'MEGAWHOP' PCR

DNA Methods

Preparation of plasmid minipreps

Our laboratory relies upon commercial plasmid purification kits which are highly reliable and produce very homogeneous plasmid preparations. Our protocols follow, and are adapted from the manufacturer’s recommendations. We currently prefer the use of the Zymo Research Zyppy Plasmid Miniprep kits.
Zymo Research Zippy Plasmid Miniprep Kit

• Centrifuge 1.6-2.0 mL¹ of cell culture at 14 krpm for 1 min, save pellet.
• Add 600 uL of water and resuspend pellet completely by vortexing.²
• Add 100 uL of 7X lysis buffer and mix by inversion 4-6 times. The solution should turn from opaque to clear blue indicating complete lysis.
• Within 2 minutes of lysis, add 350μL of cold neutralization buffer and mix by inversion 2-3 times. The solution should turn homogeneouly yellow if properly mixed and neutralized, and a yellowish precipitate of genomic DNA should form.
• Centrifuge at 14 krpm for 4 min.
• Carefully remove supernatant (about 900 μL) to spin columns equipped with catch tubes. Do not disturb the pellet.
• Centrifuge spin columns at 14 krpm for 15 s and discard flow-through.
• Wash with 200 uL of Endo-Wash buffer, centrifuge at 14 krpm for 15 s. (It is not necessary to empty the catch tube at this point)
• Wash with 400 uL of Zyppy Wash Buffer and centrifuge for 14 krpm for 30 s.
• Centrifuge at 14 krpm for 1 min to remove last traces of PE buffer from spin column
• Place spin column in a 1.5 mL microcentrifuge tube.³
• Add 30 uL of water directly to the column matrix, let stand 1 min, then centrifuge at 14 krpm for 15 s to collect DNA.⁴
• Use DNA immediately or store at –20 °C

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Zymo Plasmid Midiprep Kit

• Add 8mL ZymoPURE P1 (Red) to the bacterial cell pellet and resuspend completely by vortexing or pipetting.
• Add 8mL ZymoPURE P2 (Green/Blue) and immediately mix by gently inverting the tube 6 times. Do not vortex! Let sit for 2-3 minutes. (A clear, purple, viscous solution indicates that the cells are completely lysed)
• Add 8mL ZymoPURE P3 (Yellow) and mix gently but thoroughly by inversion. Do not vortex! (A yellow sample with yellow precipitate indicates neutralization is complete)
• Tighten the end of the Syringe Filter and load the lysate into the filter. Wait 5-8 minutes for the precipitate to float to the top.
• Remove the lock from the bottom of the syringe and place it in a 50mL conical tube. Use the plunger and the syringe to push the solution into the conical tube. Save the lysate.
• Add 8mL ZymoPURE Binding Buffer to the lysate and mix by inverting the tube 8 times.
• Add 10mL into te 15mL Conical reservoir/Zymo-spin column assembly and centrifuge at 500xg for 2 minutes. Discard flowthrough and repeat this step until all the sample has been passed through the column.
• Add 2mL of Wash 1 to the column. Centrifuge for 1 minute at the same speed as above. Discard flowthrough.
• Add 2mL of Wash 2 to the column. Centrifuge as above. Discard flowthrough and repeat this wash step.
• Disconnect the column and discard the 15mL reservoir. Place column in a collection tube and centrifuge at 10,000xg for 1 minute.
• Transfer column to a clean 1.5mL tube and add 200uL magic water to the column matrix. Wait 2 minutes, then centrifuge at 10,000xg for 1 minute.

Designing and storing ss-DNA oligonucleotide primers

Commercially synthesized oligonucleotide primers are now quite inexpensive (typically 25¢ per nucleotide or less at scales appropriate for PCR) and can be ordered and received in a few days. We use primers from Integrated DNA Technologies, Coralville, IA. A number of design factors for successful PCR primers should be considered:

• PCR primers should be complementary to the DNA sequences flanking the 5’ and 3’ ends of the fragment to be copied; each primer should exactly match at least 18 nt of the fragment sequence
• Additional, non-complementary bases may be added to the 5’-end of either or both primers; this is commonly done to add unique restriction enzyme sequences to the PCR product to facilitate its cloning. There is generally no limit to how many “extra” bases can be added. For example, adding the sequence CCATGG to the 5’ end of a nucleotide primer would add a NcoI restriction enzyme site in the final product. If restriction enzyme sites are introduced into PCR products in this fashion it is strongly recommended that some extra bases be added to the 5’-end to facilitate the direct digestion of PCR products by restriction enzymes. The recommended additional sequence is “TGC.” So, for example, to add an NcoI site to a PCR product, the sequence TGCCCATGG should be added to the 5’-end of the complementary DNA sequence.
• PCR primers should normally have T_m values of 55-65 °C. T_m values can be estimated by the formula 4 x (G+C) + 2 x (A+T), or any number of web-based T_m calculators available on the internet can be utilized. Ideally, both PCR primers should have T_m values that are matched closely, but this is not a rigorous requirement.
• A typical PCR primer, with extra bases on the 5’-end for introducing unique restriction sites, is 24-28 nt long.
• Ideally, PCR primers should have approximately 50% GC content.
• Complementarity of the 3’-ends (especially the last three nt) for any pair of primers for PCR should be strictly avoided. Such self-complementary primers can anneal to each other and produce prodigious quantities of “primer-dimer” at the expense of the desired PCR product, especially if there is high GC content at the 3’ end of the primer. An example of PCR primers that can form “primer-dimers” is illustrated below:

• All PCR oligonucleotide pairs should be checked for possible primer-dimer formation prior to ordering or carrying out PCR.
• Reconstitute commercial primers in TE buffer at 500 pmol/uL and store at –80 deg C. Remove and dilute aliquots as required to 50 pmol/uL for routine PCR.

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Quantifying DNA

The quantity of DNA can be determined from its absorbance at 260 nm. Its quality can be estimated by measuring the A₂₆₀/A₂₈₀ ratio.

• Pipet 5 uL of plasmid miniprep DNA (approx. 0.25 ug) into 195 uL of water in a 200 uL quartz silica cuvette. Measure the absorbance at 260 nm and at 280 nm
• For ds-DNA, estimate the concentration using the formula A260 x 50 = ug/mL ds-DNA. Multiply by 40 (the dilution factor) to get the concentration of DNA in the original sample.
• The A260/A280 ratio should be greater than 1.8 for pure ds-DNA. For most purposes, however, ds-DNA less pure than this can be used successfully for most molecular biology purposes.

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Sequencing DNA

To mail out for sequencing, we utilize UC Berkeley sequencing facillities, which requires appropriate quantities of very pure vector DNA.

• Go to UC Berkeley's sequencing forms page and download a current single tube sequencing form. Under PO, fill out a request for invoice https://ucberkeleydnasequencing.com/forms
• The quantity of DNA can be determined from its absorbance at 260 nm. The amount required depends on the DNA sent; for ds DNA out of high copy cell lines such as DH5a, send purified plasmid DNA at a concentration of 500 ng in less than 10µl (or dried down.)
• Custom primers (if used) should be 0.5ul of 16 pmol/uL custom primer* to each sample, or approximately 8 pmol. The total volume if sending pre-bound primers is 13 uL.
• Send each sequencing mixture (labeled clearly) per tube, packed in a secondary container, like a 50 mL Falcon tube. Send via overnight courier to

UC Berkeley DNA Sequencing Facility
310 Barker Hall
Berkeley, CA 94720-3202.

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Cell Culture Methods

Streaking out cultures from frozen glycerol stocks

Remove an LB plate with an appropriate antibiotic, and warm it to room temperature. If necessary, remove condensation by incubation in a 37 °C incubator. Label the bottom of the plate with your initials, date, and strain of E. coli used. For example the label

KMH

3-17-12

pHICA/XJB cells

indicates that individual KMH streaked out a culture of E. coli strain XJB (autolytic BL21 strain) harboring the pHICA plasmid on March 17, 2012. If this plate is discovered in the incubator or refrigerator in June 2013 (or later...sometimes a lot later) someone will realize that it needs to be discarded!
Sterilize an inoculating loop by heating in a flame until it glows cherry red over its entire length. Allow it to cool before using it to streak samples. Remove the cryovial containing the frozen glycerol stock from the –80 °C freezer, and without letting the sample thaw, take the sterile inoculating loop and scrape a little bit of ice from the surface of the frozen stock. You do not need much material: if you can see it on the loop, you have too much. Take the inoculating loop and touch it to the agar plate near one edge. In one continuous motion, drag the loop across the plate once, as shown in Figure 1:

Use one and only one of the following parameters: fileId, randomGalleryId, fgalId, attId, id, or src.
Figure 1. Streaking an agar plate to obtain single colonies (clones).

Replace the lid, invert the plate, and incubate overnight at 37 °C. If the plate has been streaked correctly, the bacterial growth will be nearly continuous and confluent near the beginning of the streak, but will thin out into single colonies somewhere farther down the plate. These single colonies represent growth from a single bacterial cell.

Streaked plates may be stored for a several weeks in the refrigerator. To keep plates from drying out, cut a strip of parafilm 1  10 cm and stretch it tightly around the edge of the plate, sealing the lid to the bottom. For longer term storage of bacterial cultures, frozen glycerol stocks should be prepared. Store plates inverted to prevent condensation from dripping on the agar surface.

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Preparation of liquid cultures of E. coli

Using sterile technique, pipet LB medium with the appropriate antibiotics, if needed, into a sterile culture tube. (For 2 mL cultures, use a 17 x 100 mm polypropylene or polycarbonate tube with a snap-top cap; for 5-10 mL culture, use a 50 mL conical polypropylene tube with a screw cap). Using a sterile pipettor tip scrape a single colony from the appropriate agar plate. Using sterile technique, tilt the culture tube until the medium is 1-2 cm from the top of the tube, and vigorously rub off the bacterial colony from the tip into the medium. For cultures in 17 x 100 mm tubes, replace the snap-top cap and adjust it so that it is in the “loose” position—this will allow in air for aerobic growth. For cultures in 50 mL conical tubes, replace the cap but leave it loosely screwed on; tape the cap to the tube to prevent it from unscrewing. Cultures are grown overnight at 37 °C with shaking on an orbital platform at approximately 250 rpm. Overnight cultures should be used immediately, or within 24 hours if refrigerated.

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Preparation of frozen glycerol stocks of E. coli

An overnight culture of E. coli should be shaken by hand to resuspend cells thoroughly. Using sterile technique, pipet 0.5 mL of bacterial culture into a sterile 1.5 mL cryovial, and dilute with an equal volume of sterile 30% glycerol. Cap the vial tightly and mix the contents of the vial completely by inverting the vial repeatedly. Glycerol stocks may be stored for a day or two at –20 °C but should normally be immediately frozen at –80 °C. Frozen stocks at this temperature, and not subjected to repeated thawing and freezing, will remain viable nearly indefinitely.

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Preparation of competent cells

The competent cells produced according to the following procedure¹ are good for routine transformations, and typically yield approximatey 10⁶-10⁷ transformants per ug of DNA. High efficiency competent cells (>10⁸ transformants per ug DNA) are generally required for transformations of ligation mixtures, and are best purchased commercially.

• Streak out E. coli JM109 (or other strain) on an LB plate. Incubate overnight at 37 °C.
• Transfer a colony into 5 mL of LB medium in a 50 mL conical tube. Shake overnight at 37 °C.
• Inoculate 5 mL of LB medium with 50 uL of overnight culture and incubate at 37 °C with shaking to an OD600 of 0.3-0.4, about 2 hr. (The exact optical density is not that critical—even overnight cultures will produce reasonably competent cells.)
• Add an equal volume of ice-cold sterile 2x TSS buffer (20% PEG 8000-40 mM MgSO4-10% DMSO) and incubate for 5-15 minutes.
• Quickly dispense 100 uL aliquots of the suspension into chilled, sterile, microcentrifuge tubes. Immediately snap-freeze cells by immersion of the tubes into liquid nitrogen.
• Store tubes at –80 °C until needed.

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Transformation of competent cells

The following transformation protocol works well with home-made and many commercially obtained competent cells.

• Gently warm 50-100 uL of competent cells by hand until just thawed, then place in an ice bath.
• Add 1-10 uL (typically 5 uL) of ligation mixture, or 10-100 ng of DNA. Flick tube gently to mix.
• Place mixture on ice for 15-60 min. (Longer times will generate more transformants.)
• Add 500 uL of LB medium (with no antibiotic) supplemented with 5 uL of sterile 2.0 M glucose, mix gently, and incubate for 1 hr at 37 °C.
• Using a bent glass or stainless steel rod—sterilized by immersion in 95% ethanol flaming off three times— spread an aliquot of 100-400 uL (typically 400 uL) of the mixture on LB plates containing 50 ug/mL ampicillin or other appropriate antibiotic. (Note: if blue-white screening¹ is desired, add 35 uL 2% X-gal and 15 uL 100 mM IPTG to the mixture before spreading.)
• Incubate plates overnight at 37 °C, but not more than 18 hr to prevent satellite colony formation.

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Transformation of Z Competent cells

Z-competent cells (Zymo research) are high efficiency (>10⁸ cfu/ug DNA) commercial competent cells with an extremely simple transformation procedure, described below:

• Pre-warm an LB agar plate containing 50 ug/mL ampicillin or other appropriate antibiotic selection factor to 37 ºC. This step is critical to efficient transformation.
• Take a single tube (100 uL) of Z-competent cells and thaw on ice.
• Remove 25 uL of cells to a separate tube under sterile conditions.
• Add 1-5 uL of DNA to cells and mix gently.
• Incubate cells on ice for 2-60 minutes. (Longer times will generate more transformants.)
• When ready to plate, pipette the ~25 uL of transformed cells to the center of your warm, labelled agar plate. Then, pipette 50-100 uL of sterile water into the transformation tube to rinse it, and add this volume of liquid to the cells on the plate for ease of spreading.
• Using a bent glass or stainless steel rod—sterilized by immersion in 95% ethanol and flaming off three times—spread the mixture evenly over warm plates.
• Incubate plates overnight at 37 °C, but not more than 18 hr to prevent satellite colony formation.

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Growing an overnight culture

Transitioning from colonies on a plate to a liquid culture is described below:

• prepare loosely capped culture tube with 6-10* mL of LB broth containing an appropriate antibiotic.
• select a sterile p200 pipette tip and touch the end to a colony on the transformant plate
• drop the tip into the LB broth
• grow overnight, shaking, at 37°C
• spin the culture tube gently (3,000-4,000 rpm) at 4°C for 5 minutes to pellet. Resuspend by gently pipetting up and down in an appropriate volume of LB broth with antibiotic or sterile water, and use for downstream experiments.

Preparation of Liquid Media

LB medium

This is a standard liquid medium used to grow cultures of E. coli. The recipe can be found in Recipes. All solid reagents should be added to the appropriate size Wheaton bottle (or other autoclavable container), and water added. When most of the solids have been dissolved (it is not necessary to completely dissolve all the solids) the solution should be neutralized by the addition of the indicated amount of NaOH. The bottle should be loosely capped—which should be secured from untwisting with tape—and autoclaved for 20 minutes at 250 °F. The solution should cool to 50 °C before any antibiotics, if needed, are added. (If you are able to hold the bottle in an ungloved hand without discomfort, it is cool enough for the addition of antibiotics.) After the addition of antibiotics, the solution should be swirled vigorously to mix thoroughly.
Media prepared without antibiotics can be stored at room temperature until opened and used the first time. Thereafter media should be stored in the refrigerator. Media prepared with antibiotics, especially ampicillin, should be stored in the refrigerator immediately.

TB medium

This is a standard “rich” medium used to grow E. coli cultures to very high cell density, and is thus useful for protein overexpression. The recipe can be found in Recipes. It is important to note that the nutrient media and the phosphate buffer solutions must be prepared and autoclaved separately. Once the solutions have cooled, they are combined using sterile technique, and antibiotics and any other additives added at that time. This medium should be swirled vigorously to mix and stored in the refrigerator if not used immediately.

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Ligation protocols

Standard ligation protocol

Ligation is a critical step in recombinant DNA methodology which accomplishes the “stitching together” of two disparate fragments of DNA. This is a common point of failure in cloning attempts. A robust protocol for ligating a PCR product to a linearized vector follows.

• Mix the following reagents in a 1.5 mL microcentrifuge tube, adding the enzyme(s) last:
  • 2-3 uL of linearized vector (approx. 50 ng)
  • 4-5 uL of PCR product
  • 1 uL 10x T4 ligase buffer with 10 mM ATP
  • 1-2 uL of T4 ligase (3-6 Weiss units per uL)
  • water to 10 uL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Incubate in the refrigerator (4 °C) overnight. This solution may be used directly to transform cells. Store unused portion at –20 °C.

Quick Ligation™ protocol

Our laboratory has experienced excellent ligation results using Quick T4 Ligase (New England Biolabs). In addition, this protocol takes substantially less time than traditional ligation, and is the preferred method whenever practical. The basic procedure follows.

• Mix the following reagents in a 1.5 mL microcentrifuge tube:
  • 1-3 uL of linearized vector (approx. 50 ng)
  • 4-5 uL of PCR product
  • water to 10 uL
• Add 10 uL of 2x Quick Ligation Buffer and mix gently
• Add 1 uL of Quick T4 Ligase and mix thoroughly
• Centrifuge briefly to collect the solution in the bottom of the microcentrifuge tube
• Incubate at room temperature for 5 minutes
• Chill on ice; use immediately for transformation or store at –20 ºC

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Polymerase Chain Reaction Techniques

Basic PCR protocol

PCR is used to amplify a particular DNA fragment which is flanked by sequences complementary to two flanking ss-DNA oligonucleotide primers. The following protocol is appropriate for Pfu Ultra, which does not need pre-annealing prior to adding polymerase:

• Mix the following reagents in a 0.5 mL PCR tube:
  • 1 uL of plasmid template (approx. 50 ng)
  • 0.5 uL of oligonucleotide primer #1 (50 pmol/ uL)
  • 0.5 uL of oligonucleotide primer #2 (50 pmol/ uL)
  • 2 uL dNTP mix (2.5 mM each)
  • 2.5 uL 10x polymerase buffer
  • water to 24.5 uL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Add 0.5 uL of Pfu Ultra(2.5 units/ uL, Stratagene) (See note)
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Perform PCR using the following segments:
  • Segment 1: 95°C for 2 min
  • Segment 2: 25-30 cycles of 95°C (30 s) → 62°C (30 s) → 72 °C (1 min or 1 min/kb of target). The annealing temperature (62°C) may be adjusted from 50-72°C as required to get product.
  • Segment 3: 72 °C (10 min) for extension, and then hold at 4 °C or place on ice.
• Dilute 20 uL of the PCR mix with 4 uL of 6x agarose gel loading buffer and run out on a 1% low-melt agarose gel
• Visualize bands on a UV transilluminator, cut out and purify desired PCR product (see Electrophoresis Protocols?)

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Touchdown PCR protocol

• Mix the following reagents in a 0.5 mL PCR tube:
  • 1 uL of plasmid template (approx. 50 ng)
  • 0.5 uL of oligonucleotide primer #1 (50 pmol/uL)
  • 0.5 uL of oligonucleotide primer #2 (50 pmol/uL)
  • 2 uL dNTP mix (2.5 mM each)
  • 2.5 uL 10x polymerase buffer
  • water to 24.5 uL
• Flick the tube gently to mix, spin briefly in a microcentrifuge to collect the contents of the tube in the bottom
• Add 0.5 uL of Pfu Ultra(2.5 units/ uL, Stratagene) polymerase
• Perform 20 cycles of 95°C (30 s) → 65 °C (30 s) → 72 °C (1 min or 1min/kb of target); but instead of the middle temperature staying constant, it is lowered 0.5-1°C per cycle, with the final temp being 45-55°C.
• Perform an additional 20 cycles of 95°C (30 s) → 50 °C (30 s) → 72 °C (1 min or 1min/kb of target)
• At the end of cycling incubate for 10 min at 72 °C, and then hold at 4 °C or place on ice.
• Dilute 25 uL of the PCR mix with 5 uL of 6x agarose gel loading buffer and run out on a 1.5% low-melt agarose gel in two wells of 15uL
• Visualize bands on a UV transilluminator, cut out and purify desired PCR product (we use a Zymo gel extraction kit)

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Site directed mutagenesis by megaprimer PCR

Megaprimer PCR

This two-step PCR-based method is certainly one of the simplest and most efficient methods of introducing specific point mutations into a DNA fragment coding for a protein. The method is summarized in Figure 2 below:

In the first round of PCR one flanking oligonucleotide primer (primer A) is paired with an oligonucleotide primer (mutant primer M1) which is designed to introduce a point mutation in the desired location within the gene. The PCR product for this reaction (A-M1) is gel purified and used as a primer in a second round of PCR with the other flanking primer (primer b) to produce a DNA product corresponding to the entire gene, with the desired mutation included. The mutant primer should be designed in such a way as to have at least 9 exactly complementary nucleotides flanking the base mismatches required to introduce the desired mutation. Normally, the mutant codon should be checked against a usage table of codons for E. coli to ensure that the codon is not rarely used for protein translation.

A typical protocol for site-directed mutagenesis follows:

• Mix the following PCR mix in a 0.5 mL PCR tube:²
  • 1 uL of plasmid template (approx. 50 ng)
  • 0.5 uL of oligonucleotide primer #1 (50 pmol/uL)³
  • 0.5 uL of mutant primer (50 pmol/uL)
  • 2 uL dNTP mix (2.5 mM each)
  • 2.5 uL 10x polymerase buffer
  • water to 24.5 uL
• Perform PCR according the protocol previously described
• Dilute 25 uL of the PCR mixture with 5 μL of 6x agarose gel loading buffer.
• Load 15 μL of the sample into each of two lanes of a 1½% low-melt agarose gel and separate by electrophoresis; for PCR products, φX174/Hae III markers or a PCR ladder are necessary to locate the DNA fragment of the correct length, and should be loaded in a nearby lane
• Cut out the PCR product from the agarose gel, extract and purify using Zymo gel extraction kit.⁵
• For the second PCR reaction, mix in a 0.5 mL PCR tube:
  • 1 uL of plasmid template (approx. 50 ng)
  • 0.5 uL of oligonucleotide primer #2 (50 pmol/uL)
  • 10 uL of PCR product from first PCR⁴
  • 2 uL dNTP mix (2.5 mM each)
  • 2.5 uL 10x polymerase buffer
  • water to 24.5 uL
• Perform PCR according the protocol previously described
• Dilute 15 uL of the PCR mixture with 3 uL of 6x agarose gel loading buffer, and run out on a 1½% agarose gel and separate by electrophoresis, using a PCR ladder in an adjacent lane to help identify the DNA fragment of the correct length.
• Cut out the PCR product from the agarose gel, extract and purify using Zymo gel extraction kit.⁵

Cloning into expression vector

• Prepare a restriction digestion using 5 uL of the concentrated, purified PCR product.
• Re-purify the digested DNA using Zymo gel extraction kit.
• Perform a ligation of 4-5 uL of a solution of this DNA fragment to the desired vector digested with the same enzymes (and perhaps dephosphorylated as well)
• Transform competent E. coli cells with this ligation mixture, spread on LB plates with the appropriate antibiotic, and grow up overnight at 37 °C
• Pick 2-3 colonies from this plate, grow up overnight in 5 mL of LB medium with the appropriate antibiotic, and prepare plasmid minipreps for each selected clone.
• Perform restriction digestions of miniprep plasmid DNA with the appropriate restriction endonucleases, and run out the products on a 1-1½% agarose gel to verify the plasmids contain the desired insert DNA.
• For clones showing evidence of the appropriate DNA insert in the vector, prepare frozen glycerol stocks from some of the overnight liquid culture. These cultures can be used to prepare samples of plasmid DNA for sequencing, or for conduction pilot overexpression trials (described later).

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SiteDirected mutagenesis by Megaprimer QuickChange 'MEGAWHOP' PCR

The advent of extremely high-fidelity DNA polymerases has made whole-plasmid PCR a practical method of introducing mutations into cloned gene products. The following method is a modification of the Stratagene QuikChange protocol that uses an easily synthesized megprimer PCR product as the mutagenic primer instead of the somewhat more problematic long oligonucleotides used in the standard QuikChange method, and is based on the method of Chen et al.¹. This is by far our preferred method for constructing site-directed variant plasmids.

• Construct the desired mutated gene or gene fragment using a one- or two-step megaprimer PCR method as described in the previous section.
• Set up a whole-plasmid PCR using the following reagents:
  • 1 uL of plasmid template miniprep (approx. 50 ng)²
  • 5 uL of mutated gel-purified PCR product (approximately 100 ng)
  • 2 uL dNTP mix (2.5 mM each)
  • 2.5 uL 10x Pfu Ultra polymerase buffer
  • water to 24.5 uL
• Mix thoroughly and denature at 95°C for 2 min
• add 0.5 uL (1.25 U) Pfu Ultra polymerase
• Mix thoroughly and perform PCR as follows:
  • Perform 25-30 cycles of PCR:
    • 95°C for 1 min
    • 60°C for 1 min
    • 72°C for 1 min per kb of DNA product (e.g., 5 min for a 4000 bp plasmid + 1000 bp gene product)
  • Product extension at 72°C for 7 min
• Cool PCR mixture to 4°C
• Transfer PCR mixture to a 1.5 mL microcentrifuge tube and add 1 uL (10U) of DpnI to digest the original template plasmid DNA.
• Mix thoroughly and incubate at 37°C for 2-3 hours.
• Use 1-5 uL of this mixture to transform Z-competent cells as described in molecular biology. Spread 10-100 uL of transformed cells on pre-warmed LB-ampicillin plates.
• Clones that grow on LB-ampicillin plates must be screened by DNA sequencing to identify successfully mutated transformants.