4
Gene Cloning for the Analysis of Gene Expression

Huw B. Thomas and Raymond T. O'Keefe

Faculty of Biology, Medicine and Health, University of Manchester, Oxford Road, Manchester, M13 9PT, UK

4.1 Identifying Target Sequence

Identifying and defining a target sequence is an important first step in expression studies since it forms the basis for all subsequent experiments. Several well‐established and fully curated databases are available online that may be used to provide the necessary nucleotide and/or protein sequence(s). These include fully comprehensive multiorganism online resources such as the National Center for Biotechnology Information (NCBI), UCSC Genome Browser and Ensembl, or more bespoke, organism‐specific databases such as those available for yeast ( Saccharomyces Genome Database [SGD], PomBase), Zebrafish (ZFIN) or Drosophila (Flybase). Links to each of these databases can be found at the end of this chapter. Multiorganism depositories are useful for identifying orthologous genes for cross‐species expression studies, whilst single‐species databases are often updated with greater regularity.

Selection of the correct sequence, and therefore the correct database source, will depend on the specifics of the intended experiment. For example, analysis of splicing consequences of a particular exon using a mini‐gene splicing assay will require not only the sequence of the exon in question but also the upstream and downstream intron sequences and thus would be obtained from a genomic sequence database. Alternatively, cloning of a protein‐coding gene into an expression vector would require the full complementary DNA (cDNA) sequence of the open reading frame made from messenger RNA. Often both sequences are available from the same depository, but it is important to make the distinction between all available sequences for a gene/transcript/isoform of interest.

4.2 In Silico Design

For cloning experiments, it is often helpful or even necessary to visualise the vector and intended insert sequence to aid with the design of the cloning experiment. Several software packages are available that enable in silico design and annotation of a recombinant DNA construct such as Benchling, SnapGene®, pDraw and UGENE. These programs allow the identification of potential restriction sites, the design of suitable primer sequences and the confirmation of all components in a cloning vector such as antibiotic resistance or auxotrophic markers. Some of the more sophisticated cloning software provides the means to carry out a step‐wise assembly of the vector construct such that multistep cloning experiments can be modelled and, most importantly, tracked, thus providing an accurate historical record of the cloning steps carried out. Additionally, the SnapGene software suite can be used in the design of individual steps of cloning, for example calculating volumes of components for polymerase chain reaction (PCR) or predicting the results of a restriction digest experiment of a recombinant vector. All of these features simplify the experimental process and seek to reduce the likelihood of human error.

4.3 Primer Design

A useful feature of cloning software suites is the ability to design bespoke PCR primers for producing a cloned DNA fragment for a cloning experiment (Figure 4.1). Whilst this method of primer design can expedite the overall design process, the software wizards often adhere to strict default parameters and do not provide much flexibility in design options, such as primer length. In such cases, manual design may be more suitable. Many different aspects and features must be considered when designing primers; some of the more important considerations will be covered below.

Image described by caption.

Figure 4.1 A schematic representation of a primer binding. (a) Two different DNA products can be amplified from the same double‐stranded DNA template using two different pairs of primers. (b) Each primer has a complementary sequence to either the top sense strand (5′–3′) or bottom anti‐sense strand (3′–5′) and align in an anti‐parallel fashion. (c) An additional utility sequence (to aid downstream cloning techniques) may be incorporated into the final amplified product by inclusion of an extra sequence at the 5′ end of the primer sequences.

The aim of the designing process is to choose primer sequences that will accurately amplify the target sequence from the template material with high fidelity whilst having a sequence length of approximately 20 nt and an optimal annealing temperature falling between 55 and 65 °C (although examples outside this temperature range are sometimes used), with no more than a 5 °C difference between annealing temperatures for each primer [1].

4.3.1 Utility Sequences

PCR amplification of a target sequence that precedes a cloning experiment will often exploit the relative flexibility of the 5′ end of the primer sequences to introduce additional utility sequences to either end of the target sequence (Figure 4.1c). These utility sequences are designed to accommodate the downstream cloning reaction and can include adding flanking restriction enzyme sites for traditional restriction cloning, introducing an additional 15–25 nt sequence that overlaps with an acceptor vector for Gibson assembly®, providing upstream promoter sequences for phage‐derived RNA polymerases or adding homologous recombination (HR) motifs for Gateway™ cloning.

4.3.2 Annealing Temperature

The optimum annealing temperature for any pair of primers is influenced by not only the GC content of the primer/template sequence but also the polymerase used in the PCR reaction. Determining the annealing temperature to use in a PCR amplification reaction is therefore not only reliant on the chosen primer sequences. A more accurate estimation of optimal annealing temperature can be made by employing an online Tm calculator such as that provided by New England Biolabs (NEB) (https://tmcalculator.neb.com), which accounts for primer length, sequence and final concentration in addition to the polymerase and enzyme buffer used when calculating the Tm. When using primers that feature a utility sequence (as covered in Section 4.3.1), this portion of the primer sequence should be omitted from the calculations as it is not complementary to the template and will not anneal.

4.3.3 Specificity

Successful PCR relies on the accurate amplification of a single product from the template material. To this end, primers should have high specificity for a single location within the template sequence. Primers are designed to be approximately 18–25 nucleotides in length, which will usually provide sufficient specificity for the target sequence and allow easy binding to the template at the appropriate annealing temperature (Figure 4.1b). Additionally, primers should have a 40–60% GC content and have at their 3′ end a G or C to promote binding and extension of the primer. An added complexity with cloning experiments is a requirement to add a utility sequence to the amplified product by way of additional non‐homologous nucleotides to the 5′ end of the primers. These additional nucleotides reduce the percentage identity between primer and template and increase the likelihood the primers may have multiple binding locations within the template sequence. It is therefore recommended that during the primer design process, candidate primer sequences are analysed for sequence similarity to the template. This can be achieved using a program such as PrimerBlast (available through the NCBI website), which not only identifies potential additional binding sites for pairs of primers but also calculates the size of each product and the calculated Tm for determining an appropriate annealing temperature for the PCR amplification. Estimating the specificity of primers is especially important for protocols using a genomic DNA template for the PCR since the potential scope of binding sites is greatly increased compared to PCR amplification from a less complex template such as a donor vector.

4.3.4 Additional PCR Primer Design Considerations

Whilst the target template sequence will often be the most restricting factor, where possible it is advisable to adhere to some general rules when designing primers. Here are some of the more important considerations. Primers should not be based paired at their 3′ end for more than 2 base pairs with either themselves or another primer, as this base‐pairing would provide a terminus for extension by the DNA polymerase. Ensure the 3′ end of the primers contain a GC‐clamp, that is, 1–3 G or C nucleotides in the last 5 bases of the 3′ end to increase the binding potential of the primers and provide a good basis for elongation. The overall G/C content of primer sequences should not be greater than 50–60%. Avoid primer sequences that contain either multiple di‐nucleotide repeating sequences (such as ATATAT) or single nucleotides repeated more than four‐times consecutively (for example, GGGGG), as both of these features can contribute to mispriming. Choose primer sequences with a low secondary structure capacity, i.e. the likelihood of forming hairpin structures, which can lead to lower PCR efficiency and yields. A quantification of self‐complementarity is provided by the PrimerBlast software by NCBI.

4.4 Template Preparation

For most cloning experiments of model organism genes, it may be possible (or indeed preferable) to purchase a purified sample of genomic material to use as a template to amplify the target gene or sequence prior to cloning. However, in some circumstances a suitable sample may not be available or the specifics of the study design requires the template to be prepared from a particular source, for example, a sample from a patient expressing a certain genetic variant or from a specific cell/tissue type. In these instances it is important to undertake a methodology that provides the highest quality genomic DNA to ensure success in the initial amplification of a target gene and other downstream applications.

Alternatively, it is increasingly common for cloning methods to utilise de novo synthesis of a target gene/sequence rather than PCR‐based amplification from a suitable DNA template sequence [2]. Historically, this approach was costly and not a service readily available. However, an increasing number of companies now offer fully customisable synthesis of DNA fragments at a relatively low cost. There are several advantages to using de novo synthesised sequences in a cloning or expression experiment, the most significant of these being the flexibility. Since no template sequence is required, the design process is far less restrictive and sequences can be synthesised that complement the experimental design. For example, for studies involving expression of a eukaryotic protein using a bacterial system, a bespoke sequence can be designed that accurately encodes for the required protein sequence but importantly accommodates the preferred codon usage of bacteria. In instances like this, no natural template would be available for a PCR‐based approach and may hinder the success of the cloning process. Another advantage of de novo synthesis is that since sequences are highly pure and their sequence verified as standard during the manufacturing process, some of the validation steps of a cloning experiment can be avoided, which can expedite the whole cloning protocol. Indeed, it is now possible to bypass any cloning procedures and design a fully bespoke vector construct, inclusive of any target genes, sequence tags, regulatory sequences and selection markers.

4.4.1 Genomic DNA Preparation

Preparation of genomic DNA from a cell/tissue sample is a relatively straightforward process involving initial cell lysis, deproteination and finally recovery of DNA by precipitation in an alcohol‐based solution. Initial preparation of a sample will depend largely on the cell/tissue type as some cells are more resistant to chemical lysis and may need additional physical disruption, such as passage through a narrow gauge needle (≤20 G) for whole blood cells or snap‐freezing in liquid nitrogen and grinding for yeast cells, before a homogenous solution is formed. Homogenisation of the cells in the DNA lysis buffer must be achieved quickly to avoid unwanted action of cellular nucleases, which will degrade the DNA. However, the freezing of cells prior to disruption or the presence of ethylenediaminetetraacetic acid (EDTA) in the buffer will act to inhibit any DNase activity. Two important considerations of the extraction process are, firstly, during extraction, to limit the shear forces on the extracted DNA by gentle mixing and pipetting to minimise disruption of the DNA and maintain high molecular weight fragments, secondly, to remove all solvents and excess salt from the sample following extraction, which can hamper downstream processes, and, not least, to provide an accurate measurement of DNA concentration. Modern DNA extraction kits are designed to limit both the physical force and contamination levels during DNA isolation by providing optimised buffer concentrations and utilising a spin‐column component for efficient purification of DNA.

Importantly, the variety of DNA extraction kits available is extensive. This variety reflects not only the range of cell types and DNA sources routinely utilised within a research context but also the scale and format of the intended DNA preparation. For example, bespoke kits are available for use on blood, urine, bacteria, bone or cultured cells. Similarly, several of these kits are available in either a single‐tube (1.5 ml) scale or in a multiwell plate scale, allowing up to 96 samples to be processed concurrently. Thus it is important to choose the best available protocol/kit for the sample in question to maximise the integrity and purity of the final DNA extraction.

4.4.2 RNA to Complementary DNA (cDNA) Conversion

Following isolation of an RNA sample, it is necessary to convert the RNA to complementary DNA (cDNA) before it can be used in a PCR reaction or downstream cloning experiments. Conversion to DNA also serves to stabilise the sample, as DNA is far less susceptible to degradation than RNA. Conversion is achieved using a retroviral‐derived reverse transcriptase, which transcribes the RNA to generate a complementary strand of DNA. A typical RNA to cDNA conversion reaction requires several key components. The first is a suitable RNA template, which could be a total RNA preparation or a specific RNA transcript. Other essential components include: a reverse transcriptase enzyme, dNTPs for cDNA synthesis, primers for the initiation of transcription, a suitable reaction buffer, an RNase inhibitor and finally a reducing agent such as dithiothreitol (DTT) to prevent oxidative damage to the protein components of the reaction.

The primers used are often one of three types: gene‐specific for conversion of a specific gene or locus within an RNA sample; oligo‐d(T), which primes at the poly‐adenosine tail of a mature mRNA transcript (thereby preferentially enriching full‐length mRNA within a total RNA sample); or random hexamers, which bind indiscriminately across the length of RNA template and act to prime all RNA fragments equally irrespective of whether they are full‐length or not. Optional alternatives to oligo‐d(T) primers are anchored oligo‐d(T) primers, which are modified to include 2 random residues at their 3′ end; this increases their likelihood of annealing to the 5′ end of the poly(A)‐tail and prevents priming from within the poly(A)‐tail, ultimately leading to increased cDNA yields. However, oligo‐d(T) priming is less suitable in circumstances where the RNA template may be degraded since any degradation of the 3′ poly(A) tail will reduce the efficiency of the priming and therefore the reverse transcription.

Several reverse transcription kits are available commercially, which include all the necessary components for a simple conversion reaction. Typically the template RNA is incubated with dNTPs and primers and heated to ensure all components are disassociated from each other before being rapidly cooled (usually by placing on ice) to aid the annealing of the primers to the template RNA. The remaining reaction components are added and the reaction proceeds at a temperature optimal for the enzyme activity. Recent bioengineering of reverse transcriptase enzymes has meant that modern kits can convert total RNA to cDNA in as little as 15 minutes. Once converted the cDNA sample may be used in a PCR reaction to amplify a gene of interest (GoI) for cloning.

4.4.3 Plasmid DNA Preparation

Plasmids are circular DNA molecules that are used as a vehicle for other DNA sequences. Plasmids can be easily and specifically maintained and propagated in bacteria as they contain sequences that allow both their replication in bacteria as well as resistance to specific antibiotics that can be used to select and grow only cells containing a desired plasmid. Many different types of plasmids have been developed and are either available commercially, directly from researchers or from a plasmid repository like Addgene (https://www.addgene.org). An important step in a cloning experiment is the isolation of plasmid DNA. This step may involve the initial isolation of a plasmid to be used in a cloning experiment or alternatively the isolation of a newly cloned recombinant construct. In either situation, careful extraction is required to ensure the sample is of high integrity but importantly is made up of plasmid DNA alone and is not contaminated by genomic DNA. Proprietary plasmid purification kit protocols are optimised to minimise the risk of contamination by genomic DNA.

4.4.4 RNA Preparation

Since RNA is much more susceptible to both physical and enzymatic degradation than DNA and that common ribonucleases are highly stable and resistant to heat denaturation, extra precautions must be taken when isolating RNA. These precautions include cleaning the work area and technical equipment (such as pipettes) with an RNase inhibitor such as RNaseZap™ (Thermo Fisher) and ensuring all steps of the RNA isolation protocol are carried out on ice (unless specified otherwise).

RNA isolation protocols include examples where cell disruption and RNA purification is achieved through a combination of a homogenising spin‐column and a cleanup spin‐column. More commonly, cells/tissues are disrupted using a phenol‐based solution such as TRIzol® (Sigma) to solubilise biological material and simultaneously denature proteins. Subsequent addition of chloroform results in phase separation of the sample, where proteins are retained in the organic phase, DNA resolves to the interface and the RNA remains in an aqueous phase. An added advantage of a TRIzol isolation is that each phase constituent (RNA/DNA/protein) can be retained and individually isolated using specific reagents and methodology. However, more optimised methods are available for extraction of protein or DNA from cell/tissue samples and, as such, RNA extraction alone is the predominant application of the TRIzol reagent. The highest integrity of RNA can be achieved using a combination of phenol/chloroform extraction with TRIzol followed by a column cleanup using a proprietary kit that includes a DNase digest step to eliminate any DNA carried over.

4.5 Cloning Methods

4.5.1 Gateway Cloning

Gateway cloning is a well‐established easily reversible method for cloning that relies on the rapid shuttling of a sequence from a donor vector to a destination vector utilising homologous recombination sequences present in both vectors [3]. The use of universal recombination sequence pairs (either L‐R or B‐P) has meant that many genes can be commercially purchased already cloned into a donor vector harbouring the necessary homologous recombination motifs. Furthermore, several destination vectors have been designed to reflect the downstream application of the cloned gene, for example vectors optimised for expression in yeast, bacteria or mammalian cells, in addition to vectors that have been designed to be used in specific circumstances such as antibody production, protein–protein interactions or RNAi assays. Cloning is carried out by incubating both vector populations in the presence of a proprietary recombinase enzyme (Clonase™) specific for either L‐R of B‐P homologous recombination pairs. An important benefit of this method of cloning is that recombination is irreversible but that successful recombination alters the recombination sites to that of the other possible motif pair. For example, recombination of sequences by using the L and R motifs results in the recombined sequences becoming B and P, respectively. This property provides the ability to reverse the reaction or subclone by utilising a different Clonase enzyme.

4.5.2 Gibson Cloning

Gibson assembly is a cloning technique developed in the Craig Venter Institute by Dr Daniel Gibson and colleagues [4]. Gibson cloning can be used to assemble up to five fragments in a single reaction regardless of fragment length or end compatibility. Gibson assembly has become a popular choice for molecular cloning since it requires fewer technical steps and reagents and thus is often faster and cheaper than other cloning methods. Additionally, Gibson assembly is sequence independent and can be carried out without the need for any restriction enzymes.

Gibson assembly relies on two important characteristics: firstly, all fragments feature an overlapping portion to their assigned adjacent fragment and, secondly, the simultaneous activity of three enzymes within the reaction mix.

Fragments are amplified with bespoke primers to incorporate flanking utility sequences. Here the additional sequence is a 15–25 nt overlapping sequence specific to either another fragment (where multiple fragments are to be joined together) or with the vector backbone sequence. An important advantage of Gibson cloning is that through careful design of the primer sequences, fragment sequences may be altered during the cloning protocol. For example, cloning and mutagenesis can be achieved by introducing a short indel or single nucleotide polymorphism (SNP) into the fragments. This mechanism can be useful to model the effects of small mutations within the target sequence where only a wild type template sequence is available.

During a Gibson assembly reaction, fragments are incubated with the Gibson assembly mix that contains three enzymatic components: a 5′ exonuclease, a high‐fidelity DNA polymerase and a DNA ligase. The action of the exonuclease degrades a short portion of the 5′ strands on each fragment, thus revealing complementary 3′ sequences between adjoining fragments. The DNA polymerase repairs any gaps in sequence between fragments and finally each fragment is ligated together by the actions of the DNA ligase, producing a fully assembled construct. The reaction mix has been engineered such that all three enzymatic reactions can occur during a single incubation step at 50 °C.

4.5.3 Restriction Enzyme Digestion and Ligation Cloning

Restriction digest cloning takes advantage of the availability of many different restriction endonucleases that target a wide variety of short DNA sequences to cut a DNA strand. New England Biolabs (www.neb.com) is just one company that sells an extensive portfolio of restriction enzymes that can be used for cloning. Restriction enzymes can digest DNA to produce either blunt ends, 5′ overhanging ends or 3′ overhanging ends. The 5′ or 3′ overhangs are usually preferred as they add specificity and directionality in the cloning process. In the design stage it is preferable to incorporate two different restriction enzyme sites into the 5′ end of the PCR primers that will leave overhangs on each end of the DNA after digestion to allow directional cloning of the DNA into the vector. Many cloning vectors are designed with multiple restriction enzyme sites that allow the insertion of DNA sequences that have been digested with compatible restriction enzymes. A vector is first cut with two restriction enzymes and is then either gel purified or column purified to remove the unwanted piece of DNA. It is also useful to dephosphorylate the cut vector with either calf intestinal phosphatase or Antarctic phosphatase to reduce re‐ligation of the vector later on. Antarctic phosphatase has the advantage of being able to be heat inactivated [5]. The DNA to be inserted into the vector is also cut with the same two restriction enzymes, or other compatible restriction enzymes, and purified to remove the unwanted pieces of DNA from the ends. Mixing the digested vector and DNA together in the presence of T4 DNA ligase inserts the DNA into the vector.

4.5.4 Ligation Independent Cloning

In ligation independent cloning, the 3′–5′ exonuclease and 5′–3′ polymerase activities of T4 DNA polymerase are utilised to produce long complementary single‐stranded overhangs (10–15 nucleotides) in both the PCR product and the plasmid, which can be annealed together. PCR primers are designed to contain sequences complementary to the ends of a plasmid cut with a restriction enzyme. Specific conditions where only one nucleotide is used with the T4 DNA polymerase exonuclease activity removes nucleotides from the 3′ end of the DNA to a certain position to prepare the plasmid and PCR product with complimentary single‐stranded overhangs. The plasmid and PCR product are then annealed and this mixture is transformed into bacteria, where the nick between the PCR product and the plasmid is repaired during the replication process, completing the insertion of the PCR product into the plasmid [6].

4.5.5 TA Cloning

TA cloning takes advantage of Taq DNA polymerase, which has a non‐template dependent terminal transferase activity that incorporates a single deoxyadenosine (A) at the 3′ end of PCR products [7]. These PCR products with 3′A overhangs can then be incubated with a plasmid vector that has a 3′T overhang and then ligated together with DNA ligase. T overhang plasmids can be obtained commercially or can be produced by restriction enzyme digestion or by addition of T overhangs to blunt ended plasmids [8]. The disadvantage of TA cloning is that Taq DNA polymerase does not have 3′–5′ exonuclease proofreading activity so there is the possibility of introducing random mutations. If introduction of random mutation is a concern then the DNA can first be amplified with a proofreading polymerase and then 3′A's added with Taq polymerase. Also insertion of a DNA fragment by TA cloning is not directional, so potentially 50% of the resulting clones will not be in the correct orientation. A variation on TA cloning is TOPO TA cloning, where plasmids are available with the 3′T end of the plasmid fused with topoisomerase I, which catalyses the ligation of the introduced A overhang PCR product [9].

4.6 Uses for Cloned DNA Sequences

The ability to clone either the gene sequence or cDNA into a plasmid vector is essential for the analysis of gene function. A wide variety of plasmid vectors that contain certain features that facilitate gene functional analysis are either commercially available or available through Addgene (www.addgene.org). For example, to determine the subcellular localisation of a protein in living or fixed cells, cDNA sequences can be cloned in frame with sequences that code for a number of fluorescent proteins like the green fluorescent protein (GFP), enhanced GFP or mCherry. The addition of small epitope tag sequences, like the V5, FLAG, MYC or HA tag, by cloning a cDNA in frame with one of these short (8–10 amino acid) peptide epitope tags, can allow for recognition of the protein fused to the tag with a specific antibody for the tag. Tagging then allows applications like western blotting, immunoprecipitation and immunofluorescence to be carried out without the need for a specific antibody to your protein of interest. The small size of these epitope tags means they usually do not influence the function of the tagged protein. Affinity tags are another type of tag that can be fused to a protein through cloning, which allows the affinity tagged protein to be purified with a specific affinity resin. Popular affinity tags are a 6–8 Histidine tag, glutathione‐S‐transferase (GST), maltose binding protein (MBP) or calmodulin binding peptide (CBP).

Cloned genes can also be placed under the control of different promoter sequences that allow one to tune the expression of the cloned gene. For example, in yeast there is a selection of promoter sequences that have been engineered into plasmid vectors that provide a range of expression levels from low to high. Additionally, there are promoters that can be turned on or off by the addition of chemical inducers or inhibitors. In yeast there is the popular GAL1 promoter that is active when cells are grown in the presence of galactose and repressed when grown in the presence of glucose. With these different promoters the effects of either under‐ or overexpressing a certain gene product can be determined. Another well‐used system for regulating expression in mammalian cells is based on the tetracycline resistance operon in bacteria. The Tet‐On and Tet‐Off systems have been developed to either turn on or off the expression of a gene in the presence of tetracycline or doxycycline. In summary, these are the most common uses of a cloned gene or cDNA, but once you have a gene cloned into a plasmid vector it can then be used for many different applications.

4.7 Verifying Cloned Sequences

Following successful cloning of a gene into a vector (using any of the methods described above), constructs will need to be validated to ensure the recombinant sequence is accurate and devoid of any errors. Nucleotide level verification is usually achieved by targeted Sanger sequencing of the plasmid, but often a carefully planned restriction digest of the plasmid can be used to screen several candidate colonies to eliminate any unsuccessful clones prior to undertaking the more expensive protocol of verification by sequencing.

4.7.1 Bacterial Transformation

Transformation of the vector into a bacterial strain enables propagation of the plasmid and subsequent isolation and purification ready for verification by Sanger sequencing. In addition, some forms of cloning provide a means to positively select for successfully cloned constructs during the transformation step. For instance, homologous recombination cloning methods (Gateway) often employ a negative selection (or suicide) gene such as ccdB incorporated into the portion of the plasmid to be replaced during successful cloning. Providing the construct is transformed into a bacterial strain that is sensitive to the expression of the ccdB gene, then only constructs that have successfully replaced the ccdB gene during the cloning reaction will grow. Examples of Escherichia coli strains sensitive to ccdB expression include Top10 and EC100D. More commonly, transformed cells are cultured on solid media containing an antibiotic selection in which genetic resistance is provided by expression of the transformation plasmid. This resistance ensures only transformed colonies will be cultured. Antibiotics such as ampicillin, kanamycin and tetracycline are some of the more common selection markers used.

4.7.2 Mini/Midi/Maxi DNA Prep

The volume of cell culture and required mass of purified plasmid will determine what scale of preparation to use for plasmid purification. A standard mini‐prep protocol will require 2–5 ml of overnight culture and yield up to 10 µg of pure plasmid DNA. Often, this amount is sufficient for both verification protocols and any downstream applications. However, larger yields may be achieved by scaling the volume of overnight culture up to 100 ml and using a midi‐prep kit for up to 100 µg of plasmid or in rare circumstances a maxi‐prep kit may be used to purify up to 0.5 mg of plasmid from approximately 500 ml of bacterial culture. The principles of the purification process are analogous regardless of the kit scale used but mini‐prep kits offer the highest extraction efficiency whilst also having the easiest and shortest protocol.

4.7.3 Sequence Confirmation

Confirmation of plasmid constructs by Sanger sequencing is the gold‐standard technique for ensuring that the cloned plasmid is accurate to the nucleotide level. Most commonly provided as an external third‐party service, ‘plasmid to sequence’ can be achieved in as little as 24 hours. Although most service providers offer low cost sequencing and require only a small volume of purified plasmid (5–15 µl) as a starting point, the cost of the service can be reduced by additional preparation steps being carried out in‐house prior to samples being sent to the sequencing provider; these steps include florescent labelling and terminator reactions.

Each sequencing reaction requires a specific sequencing primer to determine the location and direction of the sequencing reaction. Sequencing service providers offer a range of ‘universal primer’ sequences, such as T7, T3, M13‐for and M13‐rev that match sequences found in the most commonly used plasmid constructs. However, if no universal primer sequence is available or the novel fragment of DNA to be sequenced is larger than the 800–1000 bp length of a single sequencing reaction, then a single (or multiple) bespoke sequencing primer must be used. These primers can be included with the sample when sending to the sequencing provider or, alternatively, the sequencing provider may manufacture the primer prior to sequencing the sample. This primer can then be used in any subsequent sequencing reactions.

4.8 Applications of Gene Constructs

The above passages describe the necessary steps, protocol variations and common considerations when designing or implementing a study involving the cloning of a target gene into an expression vector. Once such a vector has been produced (and adequately verified), the range of applications for which a recombinant vector may be used is extremely large.

4.8.1 Recombinant Expression

A construct may be used simply to generate recombinant RNA, DNA or protein, in which case the choice of expression organism (or in some cases a particular strain) is crucial.

4.8.1.1 Bacterial System

Bacterial expression offers the fastest and easiest way to produce excess amounts of a cloned target gene. However, whilst several competent E. coli strains are commercially available, subtle differences in their features lend themselves to particular applications. Some of these features are designed to aid the cloning or transformation process itself, such as different antibiotic resistance or optimised plasmid replication for a high copy expression, whilst other strains have been manipulated to aid downstream expression steps. For example, the Rosetta™ and Origami™ strains have been specifically engineered for use in protein expression studies. Rosetta strains are designed to be used when expressing eukaryotic proteins requiring a ‘universal’ codon translation from mRNA to protein and thus are not restricted to the native codon usage of E. coli [10]. Likewise, Origami strains are commonly used in protein expression studies due to the mutation of their thioredoxin reductase (trxB) and glutathione reductase (gor) genes [11]. These mutations greatly enhance disulphide bond formation in the cytoplasm and leads to a much higher yield of active protein compared to comparable expressions in other bacterial host strains. Enhancement of disulphide bonds is mostly important for expression of extracellular proteins that normally exist in a more oxidising environment. Intracellular proteins (cytoplasmic/nuclear) function in a more reducing environment and do not generally have disulphide bonds.

4.8.1.2 Yeast System

For studies of eukaryotic processes, transformation of a recombinant gene vector into a yeast host enables the expression of that gene in a simple eukaryotic system. Yeast transformations are more straightforward, requiring fewer reagents and less time than transfections into higher eukaryotic cells such as human cell lines. Additionally, certain types of plasmid vectors, such as the pRS family vectors [12] are cross‐compatible between bacterial and yeast systems, allowing convenient propagation in E. coli and maintenance in Saccharomyces cerevisiae.

Yeast transformation efficiency and accuracy can be aided by incorporating the genes for a step in the biosynthetic pathways for uridine (URA3), leucine (LEU2) or histidine (HIS3) in a plasmid, providing successfully transformed colonies the ability to grow in specific nutrient‐deficient media in strains where these genes have been deleted or mutated. A huge range of plasmid constructs offering different combinations of antibiotic resistance and biosynthetic markers make for greater flexibility when designing an experiment involving transformation and exogenous expression of a recombinant gene.

4.8.1.3 Human Cell Transfection

For studies involving human or mammalian genes, transfection of a recombinant gene construct into a suitable cell line is a prerequisite for many analytical methodologies. For example, a recombinant protein may be expressed in a mammalian cell line to investigate its localisation, interacting partners or maturation process (such as transcript splicing or protein folding).

Cells used for transfection are most commonly well‐established cell lines such as the human cell lines HeLa, HEK‐293 or K562, or mammalian cell lines CHO and COS cells. Whilst these cell lines are relatively easy to maintain and transfect, the multiple genomic alterations they have undergone may make them poor candidates for in vivo studies. Fortunately, several different primary cell lines are commercially available allowing greater specificity in the choice of host cell. Importantly, commercially available transfection regents are routinely optimised to many of the most commonly used cell types, ensuring that differences between cell lines are accounted for during the transfection protocol, often meaning bespoke methodologies for different cell lines. However, transfection efficiency and complexity of culture methods can vary greatly from cell line to cell line. As such, experimental design often results in a compromise between experimental success and suitability of design.

4.8.2 In Vitro Production of RNA

Many plasmid cloning vectors contain, adjacent to their multicloning sites, recognition sequences for phage RNA polymerases (T7 and/or T3) that can be utilised to produce RNA by any gene cloned into the vector. The plasmid vector is usually digested with a restriction enzyme downstream of the sequence from which RNA will be produced and mixed with a phage RNA polymerase, optimised transcription buffer and the four ribonucleotides to produce an RNA by run‐off transcription. The ability of phage RNA polymerase to incorporate radioactively labelled or modified nucleotides facilitates downstream detection. The RNA produced can be used for in vitro translation to produce proteins, for probes to use during in situ hybridisation and northern blotting, for structural studies or as templates for RNA processing reactions.

4.9 Case Study: Cloning of a Human Missense Variant Exon into a Minigene Splicing Vector

With the rise in population, Genome Wide Association Studies (GWAS) – which seek to genetically characterise populations by capturing and analysing many thousand individual genomes – several rare genetic diseases are increasingly being associated and linked to low frequency allele mutations in a number of human genes. In many instances these genes are poorly characterised and the functional outcomes of the single nucleotide polymorphisms (SNPs) described are as yet undefined. Whilst missense mutations in highly conserved residues often hint at a structural change to the protein leading to a change in its functional capacity (or often loss of function), examples are emerging where single missense mutations are altering a conserved splice site, a splicing factor binding motif or activating a cryptic splice site [13]. Understanding how these nucleotide variations may be affecting splicing of the pre‐mRNA prior to translation can give us a greater understanding of the functional consequences of such rare genetic mutations in the pathology of their associated disease/conditions.

The following case study will describe in full the methods that may be used to identify, isolate and generate a variant‐containing exon into a mini‐gene splicing vector. This vector can then be used to investigate the splicing consequences of the variant exon compared to the wild type exon sequence.

4.9.1 Methods

4.9.1.1 Prerequisites

  • A human missense mutation to interrogate for splicing effects.
  • Affected gene name and exon number.
  • Precise mutation coordinate (genomic or cDNA) and polymorphism details (e.g. c.1305A > C).
  • pSpliceExpress Vector (available from Addgene – Plasmid #32485) [14].

4.9.1.2 Background

The pSpliceExpress vector construct is an example of a mini‐gene vector that contains two exon sequences (in this instance derived from the rat insulin gene INS2) flanking a pair of homologous recombination (HR) motifs. The portion of the vector between the HR motifs contains the ccdB gene that allows for positive selection of vectors deficient of this portion when the construct is transformed into TOP10 competent cells. The vector is designed so that during the cloning procedure, a single fragment (usually containing a single exon with flanking intronic sequence) is homologously recombined, thus replacing the ccdB portion with an exonic sequence and producing a mini‐gene construct containing three exons. This construct can then be transfected into a human or mammalian cell line, where the expressed transcripts will be subject to splicing by the endogenous splicing machinery. The effect of a single nucleotide variation between the WT mini‐gene and variant mini‐gene on its splicing pattern can then be compared by extracting the RNA population from the cells and carrying out reverse transcription PCR.

4.9.1.3 Identification and Amplification of Target Exon

Using the official gene name or an appropriate accession number locate the database entry for the GoI on the NCBI or Ensembl website. Save the target gene's genomic sequence (including intron portions) in FASTA format. Load the genomic sequence into SnapGene (or an equivalent sequence viewer) and using the variant coordinates, annotate the location of the polymorphism seen between WT and disease associated alleles. Save the sequence of the mini‐gene vector (pSpliceExpress) into a separate file accessible by SnapGene.

4.9.1.4 In Silico Design

Highlight the portion of the GoI defined as 100 nt upstream (5′) of the start of exon to the site of mutation and save as a new sequence named Fragment 1. Likewise, highlight and save the portion of the GoI defined as the first nucleotide downstream of the mutation site to 100 nt downstream (3′) of the end of the exon, as Fragment 2. Use the ‘Gibson assembly’ software wizard available in SnapGene to design overlapping primers for the assembly of Fragment 1 and Fragment 2 into the pSpliceExpress digested vector (choose NheI and XbaI as restriction sites used to linearise the vector and 15–25 nt of overlap between fragments). See Figure 4.2. SnapGene will design two pairs of primers, one for each fragment. Rename these primers F1‐for, F1‐rev, F2‐for and F2‐rev. Primers F1‐for and F2‐rev will have an additional utility sequence at their 5′ ends that matches the sequence 5′ and 3′ (respectively) to the insertion site on the pSpliceExpress vector. Primers F1‐rev and F2‐for will overlap each other, covering the area including the mutation site. Before exporting primer sequences from SnapGene software, manually alter the sequence of the F1‐rev and F2‐for primers at the location that relates to the variant/mutation site. For example, if missense mutation is G > A, alter the G > A in the F2‐for primer and the corresponding C > T in the F1‐rev.

Image described by caption.

Figure 4.2 Primer design for a Gibson assembly of a mini‐gene construct. (a) Mini‐gene construct consisting of an exon of interest flanked by at least a 100 bp adjacent intronic sequence. Exon may contain necessary variant SNP or may need introducing during PCR using mutagenic primers. (b) Primer design for simultaneous Gibson assembly and site‐specific mutagenesis. Primers F1‐for and F2‐rev include a utility sequence consisting of a 15–25 nt overlap with the pSpliceExpress vector. Primers F1‐rev and F2‐for are mutagenic and introduce a complementary SNP during amplification of Fragment 1 and Fragment 2 (see inset box). Using only primer F1‐for and F2‐rev primers will produce a WT Fragment 3 deficient of SNP but including overlapping sequence ends. (c) Overview of a Gibson assembly cloning method. (i) Overlapping double‐stranded DNA fragments are digested at their 5′ ends, revealing complementary 3′ sequences between fragments. (ii) Fragments anneal due to complementarity. (iii) 3′ ends are extended by DNA polymerase and DNA nicks are sealed by ligase activity, producing a seamless assembled construct.

4.9.1.5 Fragment Amplification

PCR amplify each fragment with newly designed primers using a high fidelity DNA polymerase such as Phusion. Use F1‐for and F1‐rev for ‘Fragment‐1’, F2‐for and F2‐rev for ‘Fragment‐2’ and finally use F1‐for and F2‐rev for a single ‘WT‐Fragment’ absent of a variant missense mutation. Use 50–250 ng of human genomic DNA as a template in a 50 µl reaction containing 1 µl dNTPs (10 µM), 1 µl forward primer (10 µM), 1 µl reverse primer (10 µM), 10 µl 5× HF buffer, 0.5 µl Phusion polymerase, made up to 50 µl with nuclease free water. Use the following thermocycler settings. Initial denaturation at 98 °C for 1 minute, followed by 35 cycles of 98 °C for 30 seconds, calculated annealing temperature (55–72 °C) for 20 seconds and extension at 72 °C for 30 s/kb of amplicon, followed finally by a further extension at 72 °C for 10 minutes.

Run a small volume (5–10 µl) of each reaction on a suitable percentage agarose gel to ensure that only a single product has been amplified in each reaction. Use Primer‐BLAST or SnapGene to calculate the product size for each pair of primers and check band heights against expected product size. Clean up the remaining PCR reactions using an QIAquick PCR purification kit (Qiagen) and measure the concentration of each PCR product.

Incubate 1 µg of purified pSpliceExpress vector with 1 µl each of restriction enzymes NheI and XbaI (NEB) for one hour at 37 °C in the presence of a suitable buffer, with a total reaction volume of 50 µl. Halt enzymatic activity by an incubating reaction at 65 °C for 20 minutes. Run the digested vector at 70 V for 90 minutes on a 0.8% agarose gel and excise band relating to the fragment at 4.2 kb (Figure 4.3). Purify the gel band using an QIAquick gel extraction kit (Qiagen).

Image described by caption.

Figure 4.3 SnapGene simulated agarose gel of pSpliceExpress plasmid following digestion with restriction enzymes NheI and XbaI. The band visible at approximately 4.2 kb (shown in red) relates to the portion of plasmid needed for subsequent Gibson assembly.

Carry out the Gibson assembly by incubating digested pSpliceExpress plasmid with both Fragment‐1 and Fragment‐2, or with WT‐Fragment alone. A total of 0.02–0.5 pmol of DNA fragments should be used with a 2–3‐fold excess of fragments to vector.

Mix DNA components with 10 µl of 2× Gibson assembly Master Mix and adjust the reaction volume to 20 µl with deionised water. Incubate samples in a thermocycler at 50 °C for 60 minutes and store on ice or at −20 °C for subsequent transformation.

4.9.1.6 Bacterial Transformation

Transform Gibson assemblies into recommended competent cells provided with Gibson assembly kit (New England Biolabs) using the manufacturer's recommended protocol. Briefly, thaw an aliquot(s) of NEB 5‐alpha competent E. coli (NEB) on ice. Add 2 µl of chilled Gibson assembly to the competent cells and mix gently (do not vortex). Incubate the mixture on ice for 30 minutes. Heat shock at 42 °C for 30 seconds and place back on ice for 2 minutes. Add 950 µl of room‐temperature SOC (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4 and 20 mM glucose) media and incubate at 37 °C for at least one hour. Shake vigorously or rotate. Warm selection plates (e.g. Carbenicillin containing Luria‐Bertani [LB] agar) to 37 °C. Spread 100 µl of cells on to the plates. Briefly centrifuge the remaining 900 µl of cell culture, aspirate the media and re‐suspend the pellet in 100 µl of fresh media. Spread the remaining 100 µl of cell culture on to an additional plate. This plate serves as a concentrated replicate in case transformation efficiency is low. Incubate the plates overnight at 37 °C.

4.9.1.7 Plasmid Propagation/Purification

Inoculate 5 ml of LB media (10 g Bacto‐tryptone, 5 g yeast extract, 10 g NaCl per litre) containing an appropriate antibiotic (e.g. Carbenicillin) with a single colony for each sample. Incubate cultures for 12–18 hours at 37 °C shaking vigorously. Carry out a mini‐prep on overnight cultures using GenElute Plasmid Miniprep Kit (Sigma) using the manufacturer's recommended protocol. Briefly, pellet 2 × 2 ml of overnight culture in a 2 ml Eppendorf tube, aspirate media following each spin and finally re‐suspend in 200 µl of re‐suspension solution. Lyse cells by adding 200 µl of Lysis solution. Terminate the lysis step after five minutes by adding 350 µl of neutralisation solution. Invert to mix. Centrifuge the lysate for 10 minutes at 12 000 g to pellet debris. Add 500 µl of column preparation mix to the binding column and briefly centrifuge, discard the flow‐through. Transfer the cleared lysate to the binding column and centrifuge for one minute at 12 000 g, discard the flow‐through. Add 750 µl of wash solution to the column, centrifuge for one minute at 12 000 g and discard the flow‐through. Spin the column for an additional one minute to remove excess ethanol from the column. Transfer the column to a fresh collection tube and add 50–100 µl of elution solution to the centre of the column in a dropwise fashion. Allow to stand at room temperature for one minute. Spin the column for one minute. Measure the concentration of purified plasmid with a spectrophotometer measuring absorbance at 260 nm, using the kit elution solution as a blank. Estimate the plasmid concentration given that 1 A260 unit = approximately 50 µg of dsDNA. Plasmid concentrations should be 50–100 ng/µl. Confirm the sequence of cloned constructs by Sanger sequencing using the sequence 5′‐CTACTCAGGAGAGCGTTCAC as a bespoke sequencing primer.

4.9.1.8 HEK Cell Transfection

Start with a 10 cm cell culture dish of HEK 293 cells in an appropriate culture media (DMEM +10% Fetal calf serum) near full confluency. Wash twice with 10 ml of phosphate buffered saline and detach cells with a trypsin/EDTA solution. Count cells using a haemocytometer or coulter counter and seed a six‐well culture dish with 2.5 × 105 cells in a total volume of 1.5 ml. Incubate cells overnight at 37 °C + 5% CO2 to reach 40–60% confluency. Per sample, combine 2.5 µl of Lipofectamine (Thermo Fisher) with 250 µl of Opti‐MEM media (Thermo Fisher). Combine 200 ng of plasmid construct with 250 µl Opti‐MEM media. Combine both mixes together and allow to stand for 25 minutes at room temperature. Gently add to the cells and incubate the cells for 48 hours at 37 °C + 5% CO2.

4.9.1.9 RNA Extraction

Extract RNA from cells using an appropriate protocol such as Trizol/chloroform extraction. Briefly, aspirate the media and add 1 ml of Trizol reagent (Sigma) per well of a six‐well plate. Homogenise the solution by pipetting up and down or by passing the solution several times through a 20‐gauge needle. Transfer to an Eppendorf tube. Add 200 µl of chloroform and shake vigorously for 15 seconds. Allow to stand at room temperature for five minutes. Centrifuge at 4 °C for 15 minutes at 12 000 g. Carefully remove the aqueous phase into a fresh tube containing 500 µl of isopropanol. Allow to stand at room temperature for five minutes. Centrifuge at 4 °C for 10 minutes at 12 000 g. Aspirate the supernatant taking care not to disturb the pellet. Wash the pellet with 70–75% ethanol. Centrifuge at 4 °C for five minutes at 12 000 g. Remove the ethanol and allow the pellet to briefly air dry. Resuspend the pellet in 100 µl of nuclease water. Clean up the RNA using an RNeasy column kit and include the DNase digestion step. Measure the concentration of RNA using a spectrophotometer measuring absorbance at 260 nm.

4.9.1.10 cDNA Conversion

Convert all RNA samples to cDNA using the Superscript IV kit (Invitrogen). Use this step to normalise all sample concentrations by adding an equal amount of RNA into each sample reaction. Briefly, add 1 µg of RNA (in a volume not exceeding 11 µl) to 1 µl of dNTP mix (10 mM) and 1 µl of Oligo d(T)20 (50 µM). Adjust the volume to 13 µl with nuclease‐free water. Incubate the samples at 65 °C for five minutes followed by one minute on ice. Add to the samples, 4 µl of 5× SSIV buffer, 1 µl DTT (100 mM), 1 µl of RNaseOUT RNase inhibitor and 1 µl of Superscript IV. Incubate the samples in a thermal cycler for 10 minutes at 55 °C followed by 10 minutes at 80 °C to inactivate enzyme activity. Use cDNA immediately for PCR amplification or store at −20 °C long‐term.

4.9.1.11 PCR Amplification of Spliced Products

Amplify the spliced mini‐gene products using primers designed to target the first and last exon of the pSpliceExpress mini‐gene construct. Forward primer (5′‐TGCTGGCCCTGCTCATCCTCTG) and reverse primer (5′‐TGGACAGGGTAGTGGTGGGCCT). Use 2 µl of cDNA sample as a template in a 25 µl reaction containing 0.5 µl of dNTPs (10 µM), 0.5 µl of forward primer (10 µM), 0.5 µl of reverse primer (10 µM), 5 µl of 5× HF buffer and 0.25 µl of Phusion polymerase, made up to 25 µl with nuclease free water. Use the following thermal cycler settings. Initial denaturation at 98 °C for 1 minute, followed by 35 cycles of 98 °C for 30 seconds, annealing/extension at 72 °C for 1 minute, followed finally by a further extension at 72 °C for 10 minutes. If the fragment is >1 kb in length then the annealing/extension step may need to be increased. Use SnapGene software to carry out a silico prediction of product sizes of the most likely splice products, e.g. endogenous vector exons only (mini‐gene spliced out) or a fully spliced product (mini‐gene spliced in). Run PCR products on a suitable percentage agarose gel to visualise the splicing outcome from a mini‐gene assay. Possible outcomes could be exon inclusion, intron inclusion, exon skipping or activation of cryptic splice sites.

4.10 Case Study: Epitope Tagging of a Yeast Gene

When investigating the function of a novel protein, or a protein you do not have an antibody to, epitope tagging can be a quick and convenient method of providing a tool for a range of experimental techniques. Epitope tagging allows the potential for the researcher to carry out western blotting, immunolocalisation, immunoprecipitation and affinity purification with the tagged protein. Epitope tagging involves cloning of the DNA sequence containing the open reading frame of the target protein in frame with the epitope tag into a plasmid vector appropriate for the host cell type. This plasmid could then be introduced into cultured higher eukaryotic cells by transfection, into yeast cell by transformation or into bacteria cells by transformation. The following case study will describe in full the methods that may be used to clone a yeast gene into a plasmid vector containing an epitope tag that can express the tagged protein in yeast cells.

4.10.1 Methods

4.10.1.1 Yeast Genomic DNA Preparation

Genomic DNA can either be purchased or can be easily prepared from most yeast strains using the following protocol. Grow a 2 ml yeast culture in YPD (10% BactoYeast extract, 20% BactoPeptone, 20% Dextrose) broth from a single colony. Spin down 1.5 ml of cells at 2000 rpm for five minutes in an Eppendorf tube. Discard the supernatant and resuspend cells in 500 µl of nuclease free water. Spin down cells at 2000 rpm for five minutes. Add 200 µl of lysis buffer (10 mM Tris‐Cl pH 8, 1 mM EDTA, 100 mM NaCl, 1% sodium dodecyl sulfate [SDS], 2% Triton‐X 100), 200–300 µl of acid washed glass beads (Sigma G8772), and 200 µl of phenol:chloroform:isoamyl alcohol (25 : 24 : 1 saturated with 10 mM Tris, pH 8 and 1 mM EDTA). Vortex vigorously for three minutes. Spin for five minutes at top speed in a microfuge. Carefully remove 150 µl of the top aqueous phase to a new tube and add 300 µl of 100% ethanol and mix well. Spin for five minutes at top speed in a microfuge. Remove the supernatant and then wash the DNA pellet with 95% ethanol. Remove all the ethanol and let the DNA pellet air dry for 30 minutes. Resuspend the DNA pellet in 100 µl of Tris‐Cl pH 8. Measure the concentration of the genomic DNA using a spectrophotometer measuring absorbance at 260 nm.

4.10.1.2 Amplification of Target Yeast Gene

Use the Saccharomyces Genome Database (SGD – https://www.yeastgenome.org) to obtain the DNA sequence of the open reading frame for your GoI. In this case study we will be using the small yeast gene DIB1/YPR082C as an example for epitope tagging. Search for DIB1 on the SGD webpage; then from the DIB1 ‘Summary’ page select the ‘Sequence’ tab where, if you scroll down the page, you will find a pull‐down menu in the ‘Sequence’ section where you can select the DNA for the ‘Coding DNA’. It is important to only download/copy the coding DNA as selecting genomic DNA, for some genes in yeast, will include intron sequences that are not needed. In the case of genes with introns, cDNA would need to be used as a template for PCR amplification instead of genomic DNA.

Load the DIB1 coding DNA sequence into SnapGene (or an equivalent sequence viewer). Remove the stop codon (TAA) at the 3′ end of the DIB1 coding sequence to allow in frame fusion of the DIB1 coding sequence with the tag (Figure 4.4).

Image described by caption.

Figure 4.4 Sequence of the yeast DIB1 gene with added EcoRI and KpnI restriction enzyme sites and the TAA stop codon removed. The Dib1 start codon is indicated with a red box and the position where the stop codon of Dib1 was removed is indicated with a red line.

Save the sequence of a yeast epitope tagging vector into a separate file accessible by SnapGene. For example, Oxford Genetics (https://www.oxfordgenetics.com) sell a wide range of plasmid vectors for use in yeast, including plasmid vectors for epitope tagging. In this case study we will be using the pSF‐TEF1‐COOH‐3C‐FLAG vector, which has the strong yeast TEF1 promoter sequence, a multiple cloning site with a range of unique restriction enzyme recognition sites, a sequence for the 3C Protease/PreScission protease to allow the removal of the tag following protein purification or isolation, if required, and a FLAG tag sequence to C‐terminally tag the inserted open reading frame. The plasmid also contains a URA3 marker and a 2 µm origin of replication for selection and maintenance in yeast as well as a kanamycin resistance gene and pMB1 bacteria replicon for selection and maintenance in bacteria.

Identify restriction enzyme recognition sites in the pSF‐TEF1‐COOH‐3C‐FLAG vector that are not found in the DIB1 open reading frame sequence. In this case EcoRI and KpnI are two restriction enzyme recognition sites that are not within the DIB1 coding sequence and are unique restriction enzyme recognition sites within the multicloning site upstream of the FLAG tag in the pSF‐TEF1‐COOH‐3C‐FLAG vector (Figure 4.5).

Image described by caption and surrounding text.

Figure 4.5 Multicloning site region of the pSF‐TEF1‐COOH‐3C‐FLAG vector.

Design PCR primers to amplify the DIB1 coding sequence from genomic DNA. The primers should include the EcoR1 and KpnI restriction enzyme recognition sites and allow in frame fusion with the FLAG tag. Example primers for this case would be a forward primer (5′–GGGAATTCATGGCTAGTGTTTTGTTGCC with the EcoRI site underlined) and a reverse primer (5′‐GGGGTACCTGAAACACGCTTATGATTATAATCG with the KpnI site underlined). Extra sequences have been added to the 5′ ends of the primers to allow a more efficient restriction digest.

Amplify the DIB1 coding sequence with the designed primers using a high fidelity polymerase such as Phusion. Use 50–250 ng of yeast genomic DNA as a template in a 50 µl reaction containing 1 µl dNTPs (10 µM), 1 µl forward primer (1 µM), 1 µl reverse primer (10 µM), 10 µl 5× HF buffer and 0.5 µl Phusion polymerase, made up to 50 µl with nuclease free water. Use the following thermocycler settings. Initial denaturation at 98 °C for 1 minute, followed by 35 cycles of 98 °C for 30 seconds, calculated annealing temperature (55–72 °C) for 20 seconds and extension at 72 °C for 30 s/kb of amplicon, followed finally by a further extension at 72 °C for 5 minutes.

Run a small volume (5–10 µl) of the PCR on an agarose gel (at an appropriate percentage to check that only a single product has been amplified in each reaction). Use PrimerBlast or SnapGene to calculate the product size and check band heights against expected product size; in this case it should be 445 bp. Clean up the remaining PCR reaction using a column cleanup kit (i.e. Qiagen PCR purification kit) and measure the concentration of the PCR product using a spectrophotometer.

4.10.1.3 Restriction Digestion of PCR Product

Usually up to 1 µg of the cleaned‐up PCR product is then digested with both EcoRI and KpnI. A typical restriction enzyme digestion reaction in this case contains up to 1 µg of PCR product, 5 µl of 10× CutSmart buffer (New England Biolabs), 10 units of EcoRI‐HF (New England Biolabs) and 10 units of KpnI‐HF (New England Biolabs) made up to 50 µl with nuclease free water. The reaction is then incubated at 37 °C for one hour. The restriction digest reaction is then cleaned up using a column cleanup kit (i.e. Qiagen PCR purification kit) and the concentration of the digested PCR product is measured using a spectrophotometer.

4.10.1.4 Restriction Digestion of Plasmid Vector

Usually up to 1 µg of the plasmid vector is digested with both EcoRI and KpnI. A typical restriction enzyme digestion reaction in this case contains 1 µg of pSF‐TEF1‐COOH‐3C‐FLAG, 5 µl of 10× CutSmart buffer (New England Biolabs), 10 units of EcoRI‐HF (New England Biolabs) and 10 units of KpnI‐HF (New England Biolabs) made up to 50 µl with nuclease free water. Add 10 units of calf intestinal phosphatase (CIP) to remove phosphates from the ends of the cut plasmid to prevent any re‐ligation of singly cut plasmid. The reaction is then incubated at 37 °C for one hour. The restriction digest reaction is then cleaned up using a column cleanup kit (i.e. Qiagen PCR purification kit) and the concentration of the digested plasmid is measured using a spectrophotometer. The column purification removes the small piece of DNA liberated by the double digestion of the plasmid vector with the two enzymes.

4.10.1.5 Ligation of DIB1 PCR Product into the pSF‐TEF1‐COOH‐3C‐FLAG Plasmid

A typical ligation reaction is carried out in a 20 µl volume consisting of 2 µl of 10× T4 DNA ligase buffer, 50 ng (∼12 fmol of ∼6765 bp) of restriction enzyme digested and purified plasmid vector, 10 ng (∼36 fmol of ∼445 bp) of restriction enzyme digested and purified DIB1 PCR product, made up to 19 µl with nuclease free water and 1 µl (400 units) T4 DNA ligase. The molar ratio given here is 1 : 3 for the amount of vector to insert. For making a vector to insert ratio calculations the NEBioCalculator (http://nebiocalculator.neb.com) is a useful tool. A control reaction is also set up with the vector alone to determine the background of uncut/re‐ligated plasmid. For ligations with compatible overhanging ends the ligation reaction is incubated at room temperature for at least 10 minutes. The resulting ligation of DIB1 into the pSF‐TEF1‐COOH‐3C‐FLAG plasmid to give a fusion of the DIB1 open reading frame with the FLAG tag is shown in Figure 4.6.

Image described by caption.

Figure 4.6 DIB1 gene cloned into the pSF‐TEF1‐COOH‐3C‐FLAG vector in frame with the FLAG tag. Binding sites for the sequencing primers OGP‐F2 and OGP‐R3 are shown.

4.10.1.6 Transformation of Ligation Reaction

A volume of 1–10 µl from the ligation reactions is transformed into an appropriate commercially available competent bacteria strain according to the manufacturer's protocol. An example of the bacteria transformation procedure can be found in Case Study 1 (Section 4.9.1.6). Following transformation the control plate should have no or very few bacteria colonies whereas the ligation reaction with the insert should have significantly more colonies.

4.10.1.7 Plasmid Propagation/Purification

Bacteria colonies from the ligation plate should be cultured and plasmid DNA isolated to determine by DNA sequencing whether the DNA sequence has been inserted correctly and is oriented in frame with the FLAG tag. An example of a typical protocol for the plasmid propagation/purification procedure can be found in Case Study 1 (Section 4.9.1.7).

4.10.1.8 Test Restriction Digest and Sequencing

It is useful to carry out a diagnostic restriction digest of the potential plasmid constructs before sending them off for DNA sequencing. In this case a restriction digestion with the enzyme AccI will produce three bands if the DIB1 sequence has been inserted and only two bands if it has not been inserted (Figure 4.7). Plasmids found by this test restriction digest to have the DIB1 inserted should then be sent for DNA sequencing using the sequencing primers OGP‐F2 (5′‐ TGTCGATCCTACCATCCA) and OGP‐R3 (5′‐ AGCTGAAGGTACGCTGTATC) (Figure 4.6).

Image described by caption.

Figure 4.7 SnapGene simulated agarose gel of test restriction enzyme digests with the enzyme AccI of the pSF‐TEF1‐COOH‐3C‐FLAG vector (Lane 1) or the DIB1 gene cloned into the pSF‐TEF1‐COOH‐3C‐FLAG vector (Lane 2). The successful cloning of DIB1 into the pSF‐TEF1‐COOH‐3C‐FLAG produces an additional band at 1358 bp and the change in mobility of the 6260 bp band to 5316 bp.

4.10.1.9 Yeast Transformation

Once the plasmid sequence has been confirmed to be correct by DNA sequencing it can then be transformed into a yeast strain that contains a deletion/inactivation of the URA3 gene to allow selection of the DIB1‐containing pSF‐TEF1‐COOH‐3C‐FLAG plasmid, which contains the URA3 gene. A general protocol for yeast transformation based on the Geitz method for yeast transformation is described in [15].

Inoculate the single colony into 5 ml YPD (10% BactoYeast extract, 20% BactoPeptone, 20% Dextrose) broth in a 15 ml sterile tube and incubate for 16–20 hours with shaking at 30 °C. Dilute the cells in 25 ml of YPD broth in a 50 ml sterile tube to A600 of 0.5 units (this amount of cells will be enough to carry out four transformation reactions). Incubate with shaking at 30 °C until A600 = 2 (approximately four hours). Pellet the cells at 2000 g for five minutes and then decant the supernatant. Re‐suspend the cell pellet in 25 ml of sterile water, then pellet the cells at 2000 g for five minutes and decant the supernatant. Re‐suspend the cells in 700 µl of sterile 0.1 M lithium acetate and transfer to a sterile 1.5 ml microcentrifuge tube. Pellet the cells at 10 000 g for 10 seconds and then aspirate the supernatant. Re‐suspend the cells in 200 µl of 0.1 M lithium acetate. Make up a transformation mix for each transformation consisting of 36 µl of 1 M lithium acetate, 240 µl of 50% polyethylene glycol 3500, 50 µl of 2 mg/ml of salmon sperm DNA (the salmon sperm DNA is boiled for five minutes and then cooled on ice before use) and 0.2–1.0 µg of plasmid DNA in a volume of 34 µl. Be sure to also set up a negative control reaction with no plasmid DNA. Add 50 µl of cells to each transformation mix and vortex vigorously to mix. Incubate at 30 °C for 30 minutes and then incubate at 42 °C for 30 minutes. Pellet the cells at 2000 g for 15 seconds. Gently re‐suspend the cells in 1 ml of sterile water and gently spread 50–200 µl of aliquots on the appropriate selective plate. In this case the appropriate selective plate would be a synthetic defined plate without URA (SD‐URA). MP Biomedicals (https://www.mpbio.com) distributes a wide variety of media for yeast growth. Grow cells at 30 °C for three days to allow individual yeast colonies to form.

Transformed cells can then be used for western blotting, immunolocalisation, immunoprecipitation or affinity purification of the tagged protein.

References

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  2. 2 Kosuri, S. and Church, G.M. (2014). Large‐scale de novo DNA synthesis: technologies and applications. Nat. Methods 11 (5): 499–507.
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  7. 7 Clark, J.M. (1988). Novel non‐templated nucleotide addition‐reactions catalyzed by procaryotic and eukaryotic DNA‐polymerases. Nucleic Acids Res. 16 (20): 9677–9686.
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Further Reading

  1. Green, M.R. and Sambrook, J. (2012). Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press. ISBN: 978‐1‐936113‐42‐2.

Websites

  1. NCBI (https://www.ncbi.nlm.nih.gov)
  2. UCSC (https://genome.ucsc.edu)
  3. Ensembl (https://www.ensembl.org/index.html)
  4. SGD (https://www.yeastgenome.org)
  5. PomBase (https://www.pombase.org)
  6. ZFIN (https://zfin.org)
  7. Flybase (http://flybase.org)
  8. Benchling (https://benchling.com)
  9. SnapGene (http://www.snapgene.com)
  10. pDraw (http://www.acaclone.com)
  11. UGENE (http://ugene.net)
  12. NEB Tm calculator (https://tmcalculator.neb.com/#!/main)
  13. GatewayTM cloning (https://www.thermofisher.com/uk/en/home/life‐science/cloning/gateway‐cloning.html)
  14. Gibson Assembly® (https://www.neb.com/products/e2611‐gibson‐assembly‐master‐mix#Product%20Information)
  15. Oxford Genetics (https://www.oxfordgenetics.com)
  16. NEBioCalculator (http://nebiocalculator.neb.com)
  17. MP Biomedicals (https://www.mpbio.com)
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