1.3: Designing Primers for Site-Directed Mutagenesis - Biology

1.3: Designing Primers for Site-Directed Mutagenesis - Biology

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3.1 Learning Objectives

During the next two labs you will learn the basics of site-directed mutagenesis: you will design primers for the mutants you designed earlier and perform PCR amplification to make that mutant. In this handout you will review the basics of primer design while in the next handout you will learn about PCR amplification in practice.

3.2 Mini Project Flowchart

The bolded box in the flowchart below highlights the role of the current experiment in the mini project.

3.3 What is PCR? What are polymerases?

Before we begin we need to review a few definitions commonly used when we talk about site-directed mutagenesis. Site-directed mutagenesis means that we change, insert or delete a few nucleotides within the amino acid or nucleotide sequence. In other words we change relatively few, 4-5, nucleotides or amino acids in a macromolecule. Site-directed mutagenesis became significantly easier with the emergence of PCR amplification. PCR amplification means that we synthesize (make) many copies of our DNA of interest (the coding region for a protein or nucleic acid) with the help of a polymerase and a programmable machine, called the PCR machine. Polymerases are enzymes that synthesize nucleic acids using a nucleic acid template. For example a DNA polymerase is an enzyme that makes DNA using a DNA template. The sequence of the newly synthesized DNA will be complementary to that of the template. If the template sequence is AGGC the newly synthesized DNA will be TCCG. DNA polymerases are unable to initiate DNA synthesis on their own; they need a short nucleic acid, the primer. The primer is a short DNA or RNA sequence that is complementary to the template and is used to initiate DNA synthesis. The PCR machine can precisely cycle through temperature changes to accommodate the needs of DNA synthesis. For example the PCR machine can change the temperature from 95 °C to 68 °C precisely within a few seconds. You will learn more about the temperature changes necessary to accommodate PCR amplification and the mechanism of polymerization during the next lecture.

Following PCR amplification, the amplified DNA is digested using restriction endonucleases and ligated into a cloning vector. Restriction endonucleases are enzymes that cut DNA at a given sequence. For instance the restriction endonuclease EcoRI cuts the DNA strand every time the GAATCC sequence appears in the genome. Ligation means that we connect two separate nucleic acids with a covalent bond; we simply paste them together. Cloning vectors or plasmids are circular DNAs that can be replicated by the bacterial or eukaryotic host independent of replicating their own genome. This means, they allow scientists to use a bacteria or eukaryotic cell to make large amounts of the DNA that code for the protein or nucleic acid of interest. In addition, cloning vectors have features that allow easy insertion and removal of the desired DNA sequence. Bacterial cloning vectors also have a selective marker (antibiotic resistance gene). Using selective medium this marker only allow propagation of host cells that contain the cloning vector.

3.4 PCR Amplification of a Desired DNA Segment Of The Genome (Conventional Cloning)

When the project starts the first thing to do is to amplify the DNA of interest from the genome. In this section you will learn how to do that. Afterwards, you will learn how to perform site-directed mutagenesis using the Quickchange kit. Imagine you want to amplify the DNA segment below. You will need two primers: one is complementary to the beginning while the other is complementary to the end of the sequence. The primer that is complementary to the beginning of the double-stranded DNA (dsDNA) sequence is the top primer whereas the primer that is complementary to the end of the sequence is the bottom primer. Notice that the top primer anneals against the bottom DNA strand and the bottom primer anneals against the top DNA strand.

During the first cycle of PCR amplification you do NOT get the desired DNA segment. Instead, you get two DNAs: one of them starts at the beginning of our desired sequence, and the other ends with the desired DNA sequence. Both sequences extend beyond the DNA of interest (Fig. 3.1).

During the second cycle of PCR amplification you finally get the product you want. The primers are more likely to anneal to the amplified DNAs than to the original template, because you have more of the amplified DNA than of the template. As a result the desired product is synthesized.

Notice that with each cycle the number of DNA sequences double. Thus after n cycles you have 2n of the desired DNA sequence. Since PCR leads to significant amplification of the desired DNA, it is often called a chain reaction.

Once the DNA of interest is amplified many times, the next step is to place the desired DNA into a cloning vector. This procedure is called cutting and pasting and includes several steps. First, the cloning vector and the amplified DNA are digested with a pair of restriction enzymes. Second, the cloning vector is purified using an agarose gel. Third, the cloning vector is treated with the enzyme phosphatase to prevent it from religating without the amplified DNA. Fourth, the cloning vector and the amplified DNA is ligated together using the enzyme DNA ligase and transformed into cells. Not surprisingly, these steps lead to significant loss of reagents and time (Fig. 3.3).

3.5 Quickchange Site-Directed Mutagenesis

In this section you will learn about Quickchange site-directed mutagenesis and how it differs from conventional PCR mutagenesis. Foremost, Quickchange site-directed mutagenesis does NOT require digestion with a conventional restriction endonuclease or ligation thereby reducing the time required for mutagenesis from a week to a few days. Quickchange has several restrictions. (1) Only a few nucleotides can be modified at a time. This indicates, it cannot be used to amplify a DNA sequence from the genome. (2) Quickchange provides less significant amplification of the target DNA sequence than conventional PCR. Therefore, extra care should to be taken to ensure that significant amount of mutated DNA is produced.

Let us walk through the steps of Quickchange mutagenesis (Fig 3.4)

Step 1: The primers in Quickchange land at the same spot in the cloning vector. One binds to the top the other binds to the bottom strand of the double-stranded DNA. Since the polymerase replicates the entire plasmid starting from the site of mutation the target DNA sequence has to be already inserted into a cloning vector (requires circular DNA). This method cannot be used to change DNA sequence on the chromosome.

Step 2: PCR amplification makes many copies of both the top and the bottom DNA strand of the cloning vector containing the mutated DNA. Quickchange mutagenesis therefore synthesizes many copies of the entire plasmid not only the DNA of interest. Note that the synthesized DNA is nicked (not a full circle), this means it cannot serve as template for further PCR cycles resulting in a less significant amplification of the target sequence. Therefore, removal of the template DNA is necessary (step 3) to ensure that significant numbers of cells that harbor the mutated DNA are produced in Step 4.

Step 3: After PCR amplification the reaction mixture is treated with a unique restriction endonuclease DpnI. DpnI digests the template plasmid (the one that does not contain mutations) leaving only cloning vectors containing your mutants. The DpnI enzyme achieves this task by digesting any nucleic acid with methylated adenosine base. Nucleic acids generated with PCR do not have methylated bases; therefore they are left intact by DpnI. No purification, phosphatase treatment or ligation is necessary after DpnI treatment, thereby reducing the time and reagent needed for mutagenesis.

Step 4: The mutated cloning vector is placed into E. coli bacteria for further studies (transformation).

An outline of each step is shown Fig 3.4.

A comparison of Quickchange and conventional PCR is shown in the table below:

PrimersComplementary to beginning and end of desired sequencePrimers land at the same spot on the cloning vector
AmplificationAmplifies DNA between primersAmplifies entire plasmid
PurposeSite-directed mutagenesis, amplification of desired DNA from genomeSite-directed mutagenesis: only changes a few nucleotides
Time4-5 days1-2 days
CaveatSlow, primer design is more complexOnly the template DNA serves as template thus it is required at a higher concentration and need to be removed prior to inserting the DNA into cells


Mutagenic primer design is illustrated below.

Primer design example:

  1. Suppose that you want to mutate the highlighted G to a C:
  2. Both primers must contain the desired mutation. The top primer anneals to the bottom DNA strand of the double stranded cloning vector. Therefore the top primer sequence will be the same, as the original sequence except it will have a C instead of a G at the appropriate spot.
  3. Primers should be between 25-45 nucleotides in length with a melting temperature of Tm=78 °C. Melting temperature should be calculated using the equation below where N is the length of the primer and values of GC content should be rounded to whole numbers.

    Tm = 81.5 + 0.41*(%GC) – 675/N - % of mismatch when bases are changed

    Tm = 81.5 + 0.41*(%GC) – 675/N when bases are inserted or deleted

  4. The desired mutation should be in the middle of the primer sequence with 10-15 nucleotides flanking the mutation.
  5. Primers should have a GC content of at least 40%.

Mutagenic primer sequences that fulfill the requirements above for the sample sequence are ctagggttccgcatCtcaattgacatggac (top) and gtccatgtcaattgaGatgcggaaccctag (bottom). Primer sequences are always written in the 5’ to 3’ direction this means the top and bottom primers are reverse complements of each other. In other words that they have complementary sequences and inverse chain direction to accommodate Watson-Crick pairings, but the sequence is written in the 5’ to 3’ direction.

Notes to the instructor

The experiment in Chapter 3 designs primers to alter the sequence of the Bacillus subtilis tetracycline sensor RNA ykkCD. The same protocol with minimal modifications could be used to perform site-directed mutagenesis on any nucleic acid. Two students per computer work well to maximize peer interaction while still making sure that each student has a chance to intellectually contribute to the assignments. Tablets or smart phones may also be used to complete each task thus this experiment may be used as an assignment in a lecture course. Students should be warned that primer design is an iterative process therefore several sequences have to be tried before a primer with the required GC content and TM is found. Excel may be used to calculate TM values using the equation provided. Usage of primer design programs is not recommended, because they do all the work for the students and eliminate all the educational value of this assignment.

Prelab Questions for Primer Design Lab

Define the following terms.

  1. Cloning vector or plasmid.

  2. DNA polymerase.

  3. PCR amplification.

  4. What are the pros and cons of Quickchange site-directed mutagenesis?

  5. How do you calculate primer melting temperature for Quickchange mutagenesis? Outline the equation and define each term.

Introduction to Primer Design

Lab Report Outline and Point Distribution

  1. Several sentences defining the goal/purpose of this experiment (3 pts.).
  2. Brief description of the Quickchange mutagenesis procedure. Highlight the advantages of Quickchange over “traditional” mutagenesis (6 pts.).
  3. Give your mutated primer sequence (both top and bottom) (4 pts.).
  4. Report both the percentage of GC content and the TM value (4 pts.).
  5. Explain how GC content relates to the TM of the primer. How does the TM relate to the success of your cloning experiment (8 pts.)?
  6. Explain why you chose this specific sequence for your primer. (What were you aiming for when you optimized your primer sequences?) (2 pts.).
  7. How optimal is your primer? Briefly explain (3 pts.).
  8. BLAST worksheet (30 pts.).

FAQ: How do I design primers to use with the Q5® Site-Directed Mutagenesis Kit?

For general primer design guidelines, follow the instructions below. Note that both primers do not have to be mutagenic and do not have to be phosphorylated or purified.

Substitutions are created by designing a mismatch in the center of the mutagenic primer. Include at least 10 nts that are complementary to your plasmid at the 3' end of the primer. To accommodate large mutations (from 7 to 50 per primer), changes should be incorporated at the 5&rsquoend of the mutagenic primer.
The 5' end of the second primer will begin at the base next to the 5' end of the first primer and proceed in the opposite direction on the complementary strand. This primer can be 100% complementary to the plasmid sequence or can contain mismatches, if desired. The absence of any overlap ensures that exponential (rather than linear) amplification will take place.

Deletions are created by designing primers that flank both sides of the area to be deleted. The two primers should be designed in opposite directions with their 5&rsquo ends adjacent to the area to be deleted. The primers can be 100% complementary to the plasmid sequence or can contain mismatches and/or insertions if desired.

The sequence to be inserted should be added to the 5' end of the mutagenic primer. For insertions >6 nts, the insertion sequence can be split between the two primers. Half of the insert should be added to the 5' end of the forward primer and the other half should be added to the 5' end of the reverse primer. As described for substitutions, there should be at least 10 nts that are complementary to your plasmid on the 3' end of each primer. The maximum size of the insertion is largely dictated by oligo synthesis limitations. Insertions of up to 100 nts (50 nts at the 5&rsquo end of each primer) can routinely be accommodated using this kit.

Note: For primers greater than 60 nts long, it is recommended to have them purified by either HPLC or PAGE.

Traditional Approaches to Site-Directed Mutagenesis

Inverse PCR

For deletion or insertions of >50 bp, inverse PCR is the most popular approach. Inverse PCR uses back-to-back primers to amplify the whole plasmid, followed by ligation of the linear product forming circular DNA. This technique is also suitable for larger insertions or deletions, e.g. removing a regulatory domain from a protein.

For deletions, the selected area can be removed by designing primers that anneal at either side of the targeted deletion zone. Amplification proceeds outwards from this area, thus excluding this region from the PCR product. Once the linear product is ligated, your new construct will be lacking this deletion domain. This process is outlined schematically in Figure 1 below.

Figure 1. Protocol for inverse PCR in SDM.

Insertions can be achieved by using overlapping primers with flanking sequences that contain the appropriate extra bases. For more information on inverse PCR check out these resources:

SDM Using Modified Primers

This technique uses modified primers to incorporate small base pair changes into a plasmid, and is the method of choice for site-specific mutations. To start with, you’ll need to design some primers. But before you start:

  • Print out a copy of an amino acid codon table
  • Have your plasmid or protein sequence at the ready.
  • Make sure you know where the start codon is and in what reading frame your sequence is read.

Primer Design

Aim for SDM primers of approximately 30 bp in length with your mutated site as close to the center as possible. While it is acceptable to make primers a little longer or shorter as required, there should be a minimum of 12 bp either side of your mutated site.

If you need help with primer design, lists some really useful resources to set you on your way.

Pick the Right Reagents

Ordinary Taq polymerase just won’t cut it when it comes to SDM – you need to use a proofreading enzyme. There are a variety of commercially available polymerase kits that are up to the job, incorporating a range of features including high fidelity (proofreading capability) and hot start activation. One possibility is Takara’s In-Fusion HD Cloning Plus – an all in 1 solution that includes a high-fidelity polymerase, a PCR purification kit, cloning enzyme, and competent cells for site-directed mutagenesis.

Bear in mind that like most lab reagents, many polymerases come with their own pros and cons – generally labs will have historic reasons for picking one over the other and rarely stray from this. While sticking with what works is sensible, there is no harm in reading the literature on other available resources to make sure you have the best reagents for the job!

PCR Reaction

The PCR conditions used will vary depending on your choice of kit and polymerase, as well as the primers you have designed and the size of the product. However, the example shown below is a good starting point.

StepTemp (°C)Time (s)Cycles
Extension7220/kb plasmid18

Table 1: Example of SDM thermal cycling program

Keeping the number of cycles low will prevent unwanted mutations from occurring. Eighteen cycles should yield a reasonable amount of mutated product without incorporating unwanted mutations. Note that the number of cycles can be altered during troubleshooting if required.

Digest It!

Now that you have your PCR product ready, don’t forget the critical step – digestion! Here, you digest the parental template DNA to ensure that you only have the mutated plasmid for bacterial transformation. Standard protocols call for a 1 hour incubation at 37°C with endonuclease Dpn1 to digest all dam-methylated and hemi-methylated parental DNA, leaving you with your desired mutated plasmid.

Transform and Sequence

Transform your plasmid into competent cells just as you would with any other expression plasmid, and isolate single colonies for plasmid isolation.

Now you should have a small volume of your desired plasmid. Before you start experimenting, send a sample for sequencing to confirm the presence of your desired modification(s). It is equally as important to ensure the absence of any secondary undesired mutations.

If the sequencing returns with a positive result – congratulations, you are an SDM pro! But if the sequencing returns with a negative result – don’t fret! You can always try it again – our expert tips for troubleshooting SDM should get you back on track!

Nowadays, decreasing costs of oligonucleotide synthesis and advances in synthetic biology means synthetic approaches are gaining traction over site-directed mutagenesis. Furthermore, the emergence of CRISPR/Cas9 technology has also simplified gene editing such that mutagenesis can now be performed in vitro and in vivo in a few simple steps. Nevertheless, SDM continues to be a mainstay in the molecular biology toolbox, and is not going anywhere anytime soon.

Primer Design

As a rule of thumb, 11 bp of complementary sequence on either side of the desired mutation (usually 1-3 mismatched bases) is sufficient for your primers to successfully anneal to the plasmid of interest during the PCR reaction. Ideally, your primers should be free of palindromic and repetitive sequences, but if present, a minor extension can usually ensure that the 3’-base(s) do not form secondary structures. The introduction (or ablation) of a restriction site through mutagenesis vastly facilitates the subsequent process of screening for succesfully mutated clones. Forward and reverse primers are designed to be complementary, but each primer may extend beyond the complementary region as long as an overlap with a minimum 6 bp is maintained. This overlap ensures that the PCR generates a nicked circle rather than a linear product (see figure).


A high purity plasmid prep significantly increases the success rate of site directed mutagenesis. In addition, you may want to try different concentrations of template (e.g. 0.1-1.0 ng/μl). Smaller plasmids (

3 kb) are generally more efficiently amplified than larger constructs, but plasmids as large as

6 kb can be amplified fairly easily by simply following the polymerase manufacturers’ PCR protocol. Be sure to adjust the extension time to match the size of your template. Amplification of GC-rich plasmids is facilitated by the addition of DMSO to the PCR reaction (usually around 3% final concentration). The DMSO reduces secondary structures of the DNA template, and may also decrease the annealing temp of the primers. Because you will be using a methylation-dependent enzyme (DpnI) to eliminate the parent plasmid from the PCR products, the plasmid template should be isolated from a methylation competent bacterial strain (e.g. DH5α which is dam+). Methylation deficient bacterial strains can be identified by the dam13(-) mutation – you’ll want to steer clear of these strains when preparing the plasmid template for site directed mutagenesis.


To ensure that you don’t introduce undesired mutations through the PCR process, you need a high-fidelity polymerase. There are many high fidelty polymerases on the market you need one with 5’->3’ polymerase activity (for amplification), 3’->5’ exonuclease activity (increases amplification fidelity), and no 5’->3’ exonuclease activity (which could potentially truncate the template). Importantly, you also need a DNA polymerase that produces blunt-ended PCR products (e.g. Phusion, Pfu, and Vent polymerases). Note that certain polymerases such as Taq generate A-overhangs (a feature used in TA-cloning). Such non-complementary bases at the 3’-end interfere with plasmid reconstitution, and consequently Taq polymerases cannot be used for site directed mutagenesis.


To remove the template DNA (unmodified plasmid) a restriction digest with DpnI is used. DpnI is unique in that it cleaves only DNA that is methylated at the adenosine of the GATC recognition site.

Transformation: After the PCR reaction, no ligation is required since the E. coli you transform your PCR products into will efficiently patch up the DNA. The resistance marker from the parental plasmid provides a mean for selecting for transformants which have taken up the mutagenized plasmid. Note that any residual parental plasmid (usually from incomplete DpnI digest) can also form colonies under these conditions.


If a restriction site was introduced (or ablated), bacterial colonies can be screened by identifying the presence (or absence) of that particular site with fragment length polymorphism (RFLP) analysis. In this process, if your mutation introduces an additional restriction site into your plasmid, then, when the plasmid is digested with the appropriate restriction enzyme and run on a gel, one of the bands present in the digest products of the unmodified plasmid will be split into two smaller bands (Figure B, digest A). In contrast, if your mutation ablates a restriction site, digestion with the appropriate enzyme will merge two smaller bands visible in a digest of the unmodified plasmid into a larger band (not shown). As a footnote a similar approach can be used to identify Cas9/CRISPR-induced genome modifications.

Occasionally, multimerization of the PCR primers can cause duplication of the primer sequence in the resulting plasmid. An additional restriction digest, which excises a short region (<400 bp) proximal to the target site, can identify these duplications (slightly larger bands sizes relative to the original template). Due to the small difference in size, the fragments should be separated on a high percentage agarose gel (

3%). Note that primer duplication will evade detection in the initial RFLP analysis simply because the introduced restriction site is also duplicated.


The E. coli gene sequence and mutation site were copied to the corresponding txt files as described previously. The sequences flanking mutagenic codon in the primer were set 11–21 bps in length thus there were 11 possible lengths for both sides, respectively. Totally, the script searched 121 primers, of which the Tm values were calculated for each flanking sequence. The GC content of each qualified primer was then calculated to see if in the range of 40–60%. The primers agreeing with both conditions were written in the output file. In case that no primers satisfy these two conditions simultaneously, the script also classifies those only fulfilled the first requirement in mutation_output_primer_suboptional file as a backup.

Screenshot of output file R600D_output_primer_optional.txt, in which the primers generated by this script for site-directed mutagenesis are listed, is shown in Figure 2. The primers are listed from low to high Tm to facilitate reading and comparing. In this file, the first pair of primers, R600D_forward_0_5 and R600D_reverse_1_3 look good enough in terms of Tm and GC content. Generally, the flanking sequences should not be too short so as to assure binding to templates at the step of annealing in PCR reaction. If the primers are too long, the Tm would be too high. In many cases, there may be multiple pairs of primers suitable for site-directed mutagenesis. For instance, the second or third pairs of primers also are good enough. With the primer list, it would be simple to find the mutagenic primer. The Tm value is calculated with the method based on nearest-neighbor thermodynamics using biopython API. However, users can switch to other methods by modifying the script, that is, Tm_Wallace: “rule of thumb” or Tm_GC: empirical formulas on the basis of GC content. 7


In this study, we have developed a PCR-based method for site-directed mutagenesis of large plasmids (SMLP). In the SMLP method, we take advantage of several advanced techniques, including a high-efficiency DNA polymerase for large DNA fragment amplification, and two separate PCR reactions, and the recombinational ligation with a 5′–3′ exonuclease. These techniques were combined to form a high-efficiency protocol for the mutagenesis of large plasmids. This method exhibits excellent reproducibility when it is utilized to generate mutants for the plasmids up to 17.3 kb. We show that the SMLP method can be applied to the generation of point substitution, deletion, and insertion mutations for both small and large plasmids, and it has been confirmed successful for site-directed mutagenesis through three-fragment assembly. This method is simple, low-cost, and especially suitable for the laboratories that require the mutagenesis of a gene cloned in a large plasmid.

Role of site-directed mutagenesis in the CRISPR-CAS9:

In modern times, various methods and ready to use kits for site-directed mutagenesis are available. However, one of the most important merits of the site-directed mutagenesis is in the gene editing, especially in the CRISPR-CAS9.

Any point mutation can be introduced in vivo with the help of the CRISPR-CAS9 system into the genome of a model organism.

Here, in the CRISPR-CAS9, the CAS9 is the nuclease which is used to cleave the DNA. Once it induces a double-stranded break, the mutation is inserted through the homologous-direct repair.

Read more on the homologous-direct repair (HDR): What is gene editing and CRISPR-CAS9?

Fundamentals: Ligands, Complexes, Synthesis, Purification, and Structure Identification of Protein Ligands in the Absence of Structure Using trans-Substitution

As described above, site-directed mutagenesis can be used to help determine the identity of ligands to metal sites in proteins for which high-resolution structural information is lacking. A potential ligand residue can be replaced by a residue that is either unable to coordinate metal, or has altered coordination properties, and the spectroscopic, biochemical, and/or functional properties of the protein are examined. If these properties are altered (the variant protein may fail to interact with the metal, have very different spectroscopic features, or may lack enzymatic activities attributed to the metal site), it is reasonably likely that the substituted residue may indeed act as a metal ligand. However, such defects may also result from substitution of a residue that plays a critical role in stabilizing the overall structure of the metalloprotein, without directly bonding to the metal.

The trans-substitution method outlined in Figure 3 provides a stringent test of ligand identity that avoids misassigning structurally critical non-ligating residues as ligands. In cases where substitutions of non-ligating residues alter the properties of a metalloprotein by overall disruption of structure, it is unlikely that structure can be restored by addition of an exogenous surrogate ligand. In the absence of metal–ligand interactions, the affinity of small molecules for preformed cavities in proteins is fairly low. 54–57 If deletion of a side chain disrupts structure, then the cavity to which the side-chain surrogate might bind would not be preformed, and the energy penalty to restructure the cavity (and the surrounding protein) would further decrease affinity. Thus, recovery of metal binding, wild-type spectroscopic features, and/or reactivity towards substrates as a result of adding an exogenous mimic of the deleted side chain is a strong indication that the side chain is in fact a metal ligand.

The chemical flexibility of the trans-substitution method can also enhance ligand identification. The addition of exogenous ligands that have predictable effects on spectroscopic features associated with a metal center can provide direct evidence of coordination. For instance, addition of exogenous ligands with different coordination properties may produce characteristic changes in electronic absorbance spectra. 58 Alternatively, addition of deuterated exogenous ligand to a paramagnetic metalloprotein may result in the loss of one or more hyperfine-shifted resonances in an 1 H-NMR spectrum. 59–61 Likewise, the addition of conservatively modified or isotopically substituted exogenous ligands may alter ligand–metal vibrational features in resonance Raman spectra through a mass effect, demonstrating direct bonding between the exogenous ligands and metal.

One of the first examples of the use of trans-substitution to determine protein ligand identity is the trans-substitution of an axial heme ligand in heme oxygenase (HO). Heme oxygenase is an enzyme that catalyzes the conversion of its own heme to biliverdin. As described above, site-directed mutagenesis had implicated His25 as an axial ligand to the heme substrate, as replacement with alanine produced a hemoprotein with very different spectroscopic features, and no catalytic activity. 26 However, addition of imidazole (imd) to H25A HO restored both the spectroscopic and catalytic properties of the unsubstituted enzyme. 62 Notably, the band associated with the heme iron–histidine nitrogen vibration [ν(Fe–Nim)] that was missing from the resonance Raman spectrum of H25A HO was restored upon addition of imd. Furthermore, by addition of N-methylimidazole (N-meimd), the frequency of the ν(Fe–NN−meim) band was shifted in a way that is consistent, both in direction and in magnitude, with the expected mass effect on a diatomic oscillator. 62 These trans-substitution results strongly support the assignment of His25 as the axial ligand to the heme in this enzyme.

Trans-substitution has also been used to identify the metal ligands in the β1-subunit of soluble guanylate cyclase (sGC). This enzyme catalyzes the nitric oxide-dependent conversion of GTP to cGMP. 63 Soluble guanylate cyclase had previously been shown to bind a heme 64 through a single axial histidine 65 further studies had localized the heme-binding region to an N-terminal 385 residue region of the β1-subunit. 66 A sequence alignment of β1 subunits shows four conserved histidines in the heme binding region of sGC. Substitution of three of the four conserved histidines has no effect on the heme content of this fragment, whereas a fourth histidine substitution, H105A, prevented heme from combining with protein expressed in E. coli. 67 Although these results are consistent with histidine 105 serving as axial ligand, it is also possible that the loss of heme binding in the H105A variant results from misfolding. However, by adding imidazole to the growth media, the H105A polypeptide was recovered with a stoichiometric equivalent of heme. 67 The trans-substituted complex could be prepared in states with similar electronic absorbance spectra to the wild-type complex. 67 Again, resonance Raman spectroscopy was used to confirm that exogenous imidazole was acting as an axial ligand. 67

Trans-substitution has also been used to identify ligands and gain structural and functional insight in a transmembrane hemoprotein, the cytochrome b subunit of Azobacter vinelandii hydrogenase. 68 Eight histidines were replaced by various residues in this study, including four conserved histidines in the putative transmembrane helices. Substitution of the four transmembrane histidines with alanine resulted in proteins that were unable to catalyze O2-dependent H2 oxidation, suggesting that these histidines act as ligands to two transmembrane hemes. When imidazole was added to H194A, one of the transmembrane variants, O2-dependent H2 oxidation activity was restored, supporting the assignment of histidine 194 as a heme ligand. 68 In a second assay of activity, imidazole restored activity of both the H194A and the H208 variants. Interestingly, imidazole also restored catalytic activity in proteins with tyrosines at positions 194 and 208. The recovery of activity with addition of imidazole to the H194 and H208 variants supports the idea that these histidines act as heme ligands. The success of the trans-substitution method in heme oxygenase suggests that this method may have general applicability for ligand identification and characterization in integral membrane metalloproteins such as those involved in electron transport, where structural data is difficult to obtain.

High mutagenesis efficiency

High mutagenesis efficiency. Multiple base mutagenesis is common, and we tested a 12 base substitution, insertion and deletion using a pUC19 plasmid. A random 12 base substitution was carried out within a single mutated primer. Alternatively, a random 12 base oligonucleotide containing a stop codon was inserted into wild type pUC19 plasmid, followed by deletion of the exact 12 bases to restore the wild type plasmid.

High mutagenesis efficiency. The mutagenesis efficiency for a 12 base substitution, insertion or deletion was above 90%. The performance of GeneArt Site-Directed Mutagenesis kit was comparable to the latest generation of kits from competitor 'Q'.

Site-Directed Mutagenesis of Plasmids

We frequently use both the Phusion system from New England Biolabs and the Site-Directed Mutagenesis Kits from Stratagene for site-directed mutagenesis. Below are examples of each procedure.

Using Site-Directed Mutagenesis to Install BsaI sites into a plasmid for subsequent use in Golden Gate Cloning
Authors: Eldon Chou, Caroline Ajo-Franklin, contributions from: Guillaume Cambray
Overview:This procedure describes the first part of making a promoter library in a given plasmid. Specifically this protocol describes using the Phusion site-directed mutagenesis kit to install BsaI sites flanking the promoter in the plasmid. To complete the library, Golden Gate cloning is used with dsDNA oligos to generate plasmids with different promoters, which is described in a separate protocol.

Materials & Equipment needed:

  • parent plasmid from which to create variants, i.e. pEC86
  • 10x T4 Ligase buffer, NEB # B0202S
  • T4 Polynucleotide kinase, 10 U/uL NEB # M0201S
  • Phusion Site-Directed High G-C Mutagenesis kit, # M0531S (or 5x High G-C buffer, dNTPs, DMSO, & Hot Start Phusion DNA Polymerase)
  • DpnI, NEB # R0176S
  • NEB Buffer 4 (comes with DpnI)
  • BsaI HF, NEB # R3535S
  • PCR machine

Critical Steps

Important Safety Concerns:
Waste Disposal & Clean-up:
Day 0: Design mutagenesis primers

Day 1: Phosphorylate mutagenesis primers and do pcr mutagenesis of plasmid
Note: The first step (phosphorylation) is unnecessary if you ordered phosphorylated primers.

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