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10: Amplifying and Manipulating DNA Fragments - Biology

10: Amplifying and Manipulating DNA Fragments - Biology


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  • 10.1: Prelude to Molecular Genetics
    Today, classical genetics is often complemented by molecular biology, to give molecular genetics, which involves the study of DNA and other macromolecules that have been isolated from an organism. Usually, molecular genetics experiments involve some combination of techniques to isolate and analyze the DNA or RNA transcribed from a particular gene.
  • 10.2: Isolating Genomic DNA
    DNA purification strategies rely on the chemical properties of DNA that distinguish it from other molecules in the cell, namely that it is a very long, negatively charged molecule. To extract purified DNA from a tissue sample, cells are broken open by grinding or lysing in a solution that contains chemicals that protect the DNA while disrupting other components of the cell (Figure 8.2). These chemicals may include detergents, which dissolve lipid membranes and denature proteins.
  • 10.3: Isolating or Detecting a Specific Sequence by PCR
    The Polymerase Chain Reaction (PCR) is a method of DNA replication that is performed in a test tube (i.e. in vitro). Here “polymerase” refers to a DNA polymerase enzyme extracted and purified from bacteria, and “chain reaction” refers to the ability of this technique produce millions of copies of a DNA molecule, by using each newly replicated double helix as a template to synthesize two new DNA double helices. PCR is therefore a very efficient method of amplifying DNA.
  • 10.4: Origins of Molecular Polymorphisms
    Some of mutations occur during DNA replication processes, resulting in an insertion, deletion, or substitution of one or a few nucleotides. Larger mutations can be caused by mobile genetic elements such as transposons, which are inserted more or less randomly into chromosomal DNA, sometimes occurring in clusters.
  • 10.5: Classification and Detection of Molecular Markers
    Mutations that do not affect the function of protein sequences or gene expression are likely to persist in a population as polymorphisms, since there will be no selection either in favor or against them (i.e. they are neutral). Note that the although the rate of spontaneous mutation in natural populations is sufficiently high so as to generate millions of polymorphisms that accumulate over thousands of generations, the rate of mutation is slow.
  • 10.6: Cutting and Pasting DNA- Restriction Digests and DNA Ligation
    Many bacteria have enzymes that recognize specific DNA sequences and then cut the double stranded DNA helix at this sequence. These enzymes are called site-specific restriction endonucleases, or more simply “restriction enzymes”, and they naturally function as part of bacterial defenses against viruses and other sources of foreign DNA. To cut DNA at known locations, researchers use restriction enzymes from various bacterial species, and which can be purchased from various commercial sources.
  • 10.7: Make and Screen a cDNA Library
    The first step in making a cDNA library is to isolate cellular mRNA. This mRNA extract should represent all of the transcripts in the cells at the time of isolation, or the cell’s transcriptome. This term is used by analogy to genome. However, a genome is all of the genetic information of an organism. In contrast, a transcriptome (usually eukaryotic) reflects all of the genes expressed in a given cell type at a moment in time.
  • 10.8: DNA Sequencing
    RNA sequencing came first, when Robert Holley sequenced a tRNA in 1965. The direct sequencing of tRNAs was possible because tRNAs are small, short nucleic acids, and because many of the bases in tRNAs are chemically modified after transcription. An early method for DNA sequencing developed by Walter Gilbert and colleagues involved DNA fragmentation, sequencing of the small fragments of DNA, and then aligning the overlapping sequences of the short fragments to assemble longer sequences.
  • 10.9: Genomic Libraries
    A genomic library might be a tube full of recombinant bacteriophage. Each phage DNA molecule contains a fragmentary insert of cellular DNA from a foreign organism. The library is made to contain a representation of all of possible fragments of that genome. The need for vectors like bacteriophage that can accommodate long inserts becomes obvious from the following bit of math.
  • 10.10: The Polymerase Chain Reaction (PCR)
    The polymerase chain reaction (PCR) can amplify a region of DNA from any source, even from a single cell’s worth of DNA or from fragments of DNA obtained from a fossil. This amplification usually takes just a few hours, generating millions of copies of the desired target DNA sequence. The effect is to purify the DNA from surrounding sequences in a single reaction!

Manipulating the hydrophobicity of DNA as a universal strategy for visual biosensing

Current visual biosensing methods, including colorimetric-based, fluorescence-based and chemiluminescence-based methods, are inappropriate for the hundreds of millions of people affected by color blindness and color weakness. Compared with these available methods, a droplet motion-based strategy might be a promising protocol for extension to a wider user base. Here we report a protocol for manipulating the hydrophobicity of DNA, which offers a droplet motion-based biosensing platform for the visual detection of small molecules (ATP), nucleic acids (microRNA) and proteins (thrombin). The protocol starts with target-triggered rolling-circle amplification that can readily generate short single-stranded DNA (ssDNA) fragments or long ssDNA. By exploiting macroscopic wetting behavior and molecular interaction, one can tailor the conformation of ssDNA on the water-oil interface to control the relevant DNA hydrophobicity. The wettability of DNA can be translated into visual signals via reading the sliding speed or the critical sliding angle. The time range for the entire protocol is ∼1 d, and the detection process takes ∼1 min.


Nucleic acid isothermal amplification technologies: a review

Nucleic acid amplification technologies are used in the field of molecular biology and recombinant DNA technologies. These techniques are used as leading methods in detecting and analyzing a small quantity of nucleic acids. The polymerase chain reaction (PCR) is the most widely used method for DNA amplification for detection and identification of infectious diseases, genetic disorders and other research purposes. However, it requires a thermocycling machine to separate two DNA strands and then amplify the required fragment. Novel developments in molecular biology of DNA synthesis in vivo demonstrate the possibility of amplifying DNA in isothermal conditions without the need of a thermocycling apparatus. DNA polymerase replicates DNA with the aid of various accessory proteins. Recent identification of these proteins has enabled development of new in vitro isothermal DNA amplification methods, mimicking these in vivo mechanisms. There are several types of isothermal nucleic acid amplification methods such as transcription mediated amplification, nucleic acid sequence-based amplification, signal mediated amplification of RNA technology, strand displacement amplification, rolling circle amplification, loop-mediated isothermal amplification of DNA, isothermal multiple displacement amplification, helicase-dependent amplification, single primer isothermal amplification, and circular helicase-dependent amplification. In this article, we review these isothermal nucleic acid amplification technologies and their applications in molecular biological studies.


Results

Strategy

In vitro selection requires iterated rounds of three steps: conversion of genes to gene products, selection of gene products, and gene amplification (Figure 1). The last two steps, selection and amplification, are similar between all forms of in vitro selection. However, conversion of genes to gene products poses a unique problem for the in vitro selection of small molecules. Whereas enzymes convert genetic material into the natural biopolymers, no machinery exists to directly translate genes into small molecules.

Experiments are initiated with a nucleic acid library (colored DNA). The sequence of each gene directs the synthesis of a corresponding gene product (colored ball) that is physically linked to its encoding nucleic acid. The gene products are subjected to selection, for example, through binding to an immobilized macromolecule (cyan widget at bottom). The nucleic acid encoding selected gene products is amplified and used as input for a subsequent cycle.

In general, small-molecule libraries are synthesized by the split-and-pool method which is illustrated in Figure 2 (Furka et al. 1991 Thompson and Ellman 1996). A mixture of supports (the inert material on which small molecules are built, typically polystyrene beads) is randomly split into subpools. A distinct chemical building block is then coupled to the supports in each subpool, after which the supports are pooled together and mixed. Splitting, coupling, and pooling are repeated until the library synthesis is complete. The series of subpools into which a support partitions determines what chemical building blocks are added to the support. Thus, the trajectory that a support takes through a split-and-pool synthesis is essentially a molecular recipe. If a support could predetermine its own trajectory, it would encode the synthesis of the small molecule ultimately attached to it. Predetermining support trajectories can be accomplished by using a DNA library as the support material, and by directing the splits through hybridization. The DNA sequence of each support then governs its subpool path, and acts as a genetic blueprint for a small molecule.

A mixture of solid supports (balls with rotated “L” at top) is randomly split into subpools. A distinct chemical building block (red, green, or blue ball) is coupled to the supports in each subpool. The supports are repooled and mixed. This process of splitting, chemistry, and pooling is iterated until the library synthesis is complete. The small molecules ultimately synthesized are combinations of the different building blocks (colored circles, squares, and diamonds). As highlighted by the black bead, the path taken by a support through the split-and-pool synthesis (right, middle, left) determines the small molecule synthesized on it (blue ball, green square, red diamond). The number of reactions performed is the sum of the number of subpools in each split (3 + 3 + 3 = 9). The number of unique small molecules generated is the product of the number of subpools in each split (3 × 3 × 3 = 27).

The construct we chose for our DNA support library is shown in Figure 3A. The single-stranded DNA (ssDNA) includes a unique reactive site at its 5′ end, upon which a small molecule is synthesized. The DNA sequence contains 20-base “codons” flanked by 20-base noncoding regions. Within the DNA support library, sequence degeneracy exists at the coding positions. The set of codons in each DNA support specifies a small-molecule synthesis by directing the splitting of the ssDNA into appropriate subpools. The noncoding regions enable genetic recombination of support sequences by PCR (Halpin and Harbury 2004).

(A) Schematic showing the structure of the DNA support library. Small molecules are synthesized at the 5′ end of 340-base ssDNA genes. The ssDNA consists of 20-base noncoding regions (black lines labeled Z1–Z7) and 20-base coding positions (colored bars labeled [a–j]1–6). All library members contain the same seven DNA sequences at the seven noncoding regions. At each of the six coding positions, ten mutually exclusive DNA codons, (a–j)n, are present, for a total of 60 different sequences. Each coding region specifies the addition of a single subunit to a growing small molecule. A unique reactive site (in this case a primary amine) for small-molecule synthesis is attached to the 5′ end of the ssDNA through a polyethylene glycol linker (squiggly line). Resin beads coated with an oligonucleotide complementary to one codon (anticodon beads, gray ball at right) capture by hybridization ssDNAs containing the corresponding codon.

(B) Chemical translation is a split-and-pool synthesis, with splitting directed by DNA hybridization. A ssDNA library is hybridized to a set of anticodon columns (gray balls) corresponding to the set of codons present at a single coding position. The ssDNA genes partition into subpools based on sequence identity. Distinct chemical subunits (colored balls) are coupled to the DNA in each subpool. Finally, the DNA is repooled, completing the encoded addition of one subunit to the growing small molecule. The process of hybridization splitting, chemistry, and pooling is repeated for all subsequent coding regions.

(C) Schematic product of chemical translation. The sequence of the small-molecule subunits (colored balls) corresponds to the sequence of codons (colored bars) in the ssDNA gene.

Our scheme for DNA-directed split and pool synthesis is shown in Figure 3B. The library is first split by hybridization to a set of anticodon columns complementary to the different 20-base sequences present at the first coding position. Distinct chemical building blocks are coupled to each subpool, and the library is repooled. The process is repeated, but the splitting is directed by a subsequent coding position. Each coding position comprises a set of codons that differ in sequence from the codons at all other coding positions. Consequently, splitting is always directed by hybridization at one intended coding region, and not by codons elsewhere. Small molecules are synthesized directly on their encoding DNAs, maintaining the physical linkage between gene and gene product (Figure 3C). Direct conversion of genes into small-molecule gene products, combined with selection and amplification steps, enables the in vitro selection of small-molecule libraries.

Reduction to Practice

We first developed a Sepharose-based resin derivatized with anticodon oligonucleotides complementary to codon sequences (Halpin and Harbury 2004). We tested the resin by hybridizing a library consisting of seven ssDNA sequences to a corresponding set of seven different anticodon columns (Figure 4). There was little crosshybridization, which ensures that DNA genes will be accurately translated. Analysis of splitting efficiencies by a scintillation counting assay of radiolabeled ssDNA showed that 90% or more of the ssDNA inputs were recovered from the correct hybridization columns for all tested sequences (Halpin and Harbury 2004). The resin is also robust. We have not observed any loss in efficiency with over 30 cycles of hybridization and elution.

Seven serially truncated ssDNAs differing in sequence at one coding position (illustrated at left of gel, number of bases indicated) were hybridized to seven anticodon columns (cylinders at top of gel). The load (lane 1), flow through (lane 2), and column elutes (lanes 3–9) were analyzed by denaturing polyacrylamide gel electrophoresis.

We next addressed chemical synthesis on unprotected DNA. Use of a solid phase in small-molecule synthesis allows for the application of excess reagents, to drive reactions to completion, and simplifies product purification (Merrifield 1963). To realize these advantages, we carried out synthetic steps while DNA was noncovalently bound to diethylaminoethyl (DEAE) Sepharose resin (Halpin et al. 2004). DEAE Sepharose was chosen for solid-phase synthesis because it adsorbs DNA reversibly and in a sequence-independent manner and because it behaves well in organic solvents. Incubation of immobilized DNA with the appropriate reagents results in addition of a building block, completing one step in the synthesis of a small molecule. Following the chemical step, DNA is eluted from the solid phase and manipulated in solution.

As an initial chemistry, we chose 9-Fluorenylmethoxycarbonyl [Fmoc]–based peptide synthesis. Figure 5 shows the results of solid-phase peptide synthesis on DNA using Fmoc-protected succinimidyl esters (Anderson et al. 1963 Carpino and Han 1970 Halpin et al. 2004). Synthesis of the [Leu]enkephalin pentapeptide on an aminated 20-base oligonucleotide (Figure 5B) yielded a highly pure [Leu]enkephalin-DNA conjugate. A nonaminated oligonucleotide internal control was not altered by the chemistry, ruling out nonspecific chemical modification of DNA. Over 90% of the recovered nucleic acid was the intended [Leu]enkephalin-DNA conjugate (the overall recovered yield was 60%). The results correspond to a 98% efficiency for each amino acid coupling step.

(A) Structure of the [Leu]enkephalin–DNA conjugate.

(B) High performance liquid chromatography chromatogram of the [Leu]enkephalin peptide synthesized using succinimidyl ester chemistry on a 20-base oligonucleotide modified with a 5′ primary amine (20mer). A 10-base oligonucleotide without the 5′ primary amine (10mer) was included in the reactions as a control for nonspecific DNA modification. The red and blue traces are the DNA before and after chemistry, respectively. The mass of the major product peak (42-min retention time) matches the expected mass of the [Leu]enkephalin–DNA conjugate.

(C) Electromobility shift assay of peptides synthesized on 340-base ssDNA. Conjugates were eletrophoresed on a native agarose gel in the absence (lanes 1, 3, 5, and 7) or presence (lanes 2, 4, 6, 8, and 9) of the [Leu]enkephalin-binding antibody 3-E7. [Leu]enkephalin (L) or a scrambled sequence (S) was synthesized on a 5′ amino-modified 20-base oligonucleotide, which was subsequently used as a primer for PCR (lanes 1–4), or directly on 5′ amino-modified 340-base ssDNA, which was subsequently converted to dsDNA (lanes 5–9). Addition of free [Leu]enkephalin peptide (lane 9) competes away binding.

Synthesis of [Leu]enkephalin on a 340-base ssDNA support, capable of encoding an eight-step synthesis, was analyzed using an electromobility shift assay and the enkephalin-specific 3-E7 antibody (Hwang et al. 1999). Figure 5C shows that 3-E7 shifts the majority of the [Leu]enkephalin-DNA (approximately 85% when standardized to a positive control), showing that the biological activity of the peptide is maintained while attached to DNA. The 3-E7 antibody does not shift a scrambled-DNA peptide conjugate containing the same amino acids as [Leu]enkephalin but in a different order. Finally, free [Leu]enkephalin peptide eliminates the shifting of [Leu]enkephalin-DNA by 3-E7, demonstrating the specificity of the shift.

Our chemical translation strategy requires repeated hybridization-directed splitting and coupling of chemical building blocks to DNA. Two different solid phases were utilized for these tasks. To efficiently transfer DNA from anticodon columns to DEAE Sepharose columns, we cyclically pumped 50% dimethylformamide (DMF) over the columns connected in series (Figure 6). Conversely, to transfer DNA from DEAE Sepharose columns back to anticodon columns, we used a high salt buffer in a closed system. In both cases, a large effective buffer volume flows over each column, which allows the DNA transfer processes to approach thermodynamic equilibrium. These column-to-column transfers remove intermediate storage tubes and require little solvent, minimizing loss of DNA.

Chemical translation requires iteration of a chemistry step and two column-transfer steps. ssDNA is transferred from anticodon columns to DEAE Sepharose columns by cyclically pumping 50% DMF through a pair of columns (one hybridization, one DEAE) attached in series for 1 h at 45 °C. Chemistry is performed on ssDNA bound to each DEAE column. ssDNA is transferred from DEAE columns to anticodon columns by cyclically pumping a 1.5-M NaCl buffer through all DEAE columns and all anticodon columns associated with the next coding position for 1 h at 70 °C and 1 h at 46 °C. Efficiencies for each step are indicated in red.

In Vitro Selection of a Chemically Synthesized Library

To test and validate our general strategy, we applied in vitro selection to a primarily nonnatural peptide library, with the goal of identifying a high-affinity ligand for the monoclonal antibody 3-E7 (Meo et al. 1983). Isolation of 3-E7 ligands is a well-defined in vitro selection problem characterized previously (Cwirla et al. 1990 Barrett et al. 1992). We designed our library to contain at least one known 3-E7 ligand, [Leu]enkephalin. The [Leu]enkephalin peptide binds to 3-E7 with an affinity of 7.1 nM, and its size (five residues) was well-suited for our experiments.

An initial DNA support library consisting of ten distinct sequences (“all a,” “all b,” etc.) was diversified 10 5 -fold by PCR recombination to generate a support library with a complexity of one million, as verified by DNA sequencing (Halpin and Harbury 2004). This library was chemically translated into acylated pentapeptides using Fmoc-protected succinimidyl esters. The peptide library included ten different monomers at each position (Figure 7A). The first five positions comprised one of ten amino acids (β-alanine, D-alanine, D-leucine, D-tyrosine, 4-nitro-phenylalanine, glycine, leucine, norleucine, phenylalanine, or tyrosine). The N-terminus was left unmodified or was acylated with one of nine acids (acetic, benzoic, butyric, caproic, glutaric, isobutyric, succinic, trimethylacetic, or valeric). After library synthesis and conversion of the ssDNA into duplex form, the library was subjected to selection using the 3-E7 antibody. The selected DNA was PCR amplified and used as input for the subsequent round of synthesis and selection.

(A) Library building blocks. Proteinogenic building blocks are shown in green.

(B) Approximately 70 DNA genes from each round of selection were sequenced, and the results are summarized as a histogram plot. The x-axis indicates the number of amino acid residue matches to [Leu]enkephalin encoded by a library sequence. The y-axis indicates the library generation (0, starting material 1, after round one selection 2, after round two selection). The z-axis indicates the number of sequences encoding a particular number of matches (x-axis) in a particular round (y-axis).

(C) The top row reports the round two library consensus sequence, which matches [Leu]enkephalin. The second row reports the percentage of round two library clones that encode the [Leu]enkephalin amino acid at each residue position. The third row reports the identity and frequency of the most commonly occurring non-[Leu]enkephalin subunit at each position.

In order to monitor library convergence, DNA from the starting material (round 0) and from after one (round 1) or two (round 2) selection generations was subcloned, and approximately 70 different isolates from each round were sequenced (Figure 7B). In round 0, none of the sequences encoded more than three residues in common with [Leu]enkephalin. Two sequences from round 1 encoded five [Leu]enkephalin residues, and one sequence encoded four residues. Of the round 2 sequences, twenty encoded full-length [Leu]enkephalin, thirty-four encoded single mutants, and eleven encoded double mutants. Only three round 2 sequences encoded less than four [Leu]enkephalin residues. The round 2 consensus peptide sequence matched [Leu]enkephalin (Figure 7C). Previous work has shown that the N-terminal residues (Tyr-Gly-Gly-Phe) are responsible for most of [Leu]enkephalin's affinity for the 3-E7 antibody (Meo et al. 1983 Cwirla et al. 1990). We observed high sequence conservation at these residues, recapitulating the earlier results.

To assess generality, we carried out a second [Leu]enkephalin in vitro selection experiment using a peptide library of the same size but constructed with a completely different “genetic code.” Every codon in the alternate library coded for an amino acid different from the one it coded for in the first library. The [Leu]enkephalin codon series in the first library was b1-j2-b3-c4-h5-i6, whereas in the alternate library it was d1-b2-g3-g4-i5-f6. Two rounds of selection enriched the alternate [Leu]enkephalin DNA gene 10 5 -fold (data not shown). The data suggest that little, if any, DNA sequence encoding bias exists in our system. Further, they illustrate the reproducibility of the technology. Together, the results demonstrate conclusively that the DNA display strategy can be used for the in vitro selection of synthetic chemical libraries.


Protocol

1. Designing Primers

Designing appropriate primers is essential to the successful outcome of a PCR experiment. When designing a set of primers to a specific region of DNA desired for amplification, one primer should anneal to the plus strand, which by convention is oriented in the 5' → 3' direction (also known as the sense or nontemplate strand) and the other primer should complement the minus strand, which is oriented in the 3' → 5' direction (antisense or template strand). There are a few common problems that arise when designing primers: 1) self-annealing of primers resulting in formation of secondary structures such as hairpin loops (Figure 1a) 2) primer annealing to each other, rather then the DNA template, creating primer dimers (Figure 1b) 3) drastically different melting temperatures (Tm) for each primer, making it difficult to select an annealing temperature that will allow both primers to efficiently bind to their target sequence during themal cycling (Figure 1c) (See the sections CALCULATING MELTING TEMPERATURE (Tm) and MODIFICATIONS TO CYCLING CONDITIONS for more information on Tms).

Below is a list of characteristics that should be considered when designing primers.

Primer length should be 15-30 nucleotide residues (bases).

Optimal G-C content should range between 40-60%.

The 3' end of primers should contain a G or C in order to clamp the primer and prevent "breathing" of ends, increasing priming efficiency. DNA "breathing" occurs when ends do not stay annealed but fray or split apart. The three hydrogen bonds in GC pairs help prevent breathing but also increase the melting temperature of the primers.

The 3' ends of a primer set, which includes a plus strand primer and a minus strand primer, should not be complementary to each other, nor can the 3' end of a single primer be complementary to other sequences in the primer. These two scenarios result in formation of primer dimers and hairpin loop structures, respectively.

Optimal melting temperatures (Tm) for primers range between 52-58 ଌ, although the range can be expanded to 45-65 ଌ. The final Tm for both primers should differ by no more than 5 ଌ.

Di-nucleotide repeats (e.g., GCGCGCGCGC or ATATATATAT) or single base runs (e.g., AAAAA or CCCCC) should be avoided as they can cause slipping along the primed segment of DNA and or hairpin loop structures to form. If unavoidable due to nature of the DNA template, then only include repeats or single base runs with a maximum of 4 bases.

Notes:

There are many computer programs designed to aid in designing primer pairs. NCBI Primer design tool http://www.ncbi.nlm.nih.gov/tools/primer-blast/ and Primer3 http://frodo.wi.mit.edu/primer3/ are recommended websites for this purpose.

In order to avoid amplification of related pseudogenes or homologs it could be useful to run a blast on NCBI to check for the target specificity of the primers.

2. Materials and Reagents

When setting up a PCR experiment, it is important to be prepared. Wear gloves to avoid contaminating the reaction mixture or reagents. Include a negative control, and if possible a positive control.

Arrange all reagents needed for the PCR experiment in a freshly filled ice bucket, and let them thaw completely before setting up a reaction (Figure 2). Keep the reagents on ice throughout the experiment.

Standard PCR reagents include a set of appropriate primers for the desired target gene or DNA segment to be amplified, DNA polymerase, a buffer for the specific DNA polymerase, deoxynucleotides (dNTPs), DNA template, and sterile water.

Additional reagents may include Magnesium salt Mg 2+ (at a final concentration of 0.5 to 5.0 mM), Potassium salt K + (at a final concentration of 35 to 100 mM), dimethylsulfoxide (DMSO at a final concentration of 1-10%), formamide (at a final concentration of 1.25-10%), bovine serum albumin (at a final concentration of 10-100 μg/ml), and Betaine (at a final concentration of 0.5 M to 2.5 M). Additives are discussed further in the trouble shooting section.

Organize laboratory equipment on the workbench.

Materials include PCR tubes and caps, a PCR tube rack, an ethanol-resistant marker, and a set of micropipettors that dispense between 1 - 10 μl (P10), 2 - 20 μl (P20), 20 - 200 μl (P200) and 200 - 1000 μl (P1000), as well as a thermal cycler.

When setting up several PCR experiments that all use the same reagents, they can be scaled appropriately and combined together in a master mixture (Master Mix). This step can be done in a sterile 1.8 ml microcentrifuge tube (see Notes).

To analyze the amplicons resulting from a PCR experiment, reagents and equipment for agarose gel electrophoresis is required. To approximate the size of a PCR product, an appropriate, commercially available molecular weight size standard is needed.

3. Setting up a Reaction Mixture

Start by making a table of reagents that will be added to the reaction mixture (see Table 1).

Next, label PCR tube(s) with the ethanol-resistant marker.

Reaction volumes will vary depending on the concentrations of the stock reagents. The final concentrations (CF) for a typical 50 μl reaction are as follows.

X buffer (usually supplied by the manufacturer of the DNA polymerase may contain 15 mM MgCl2). Add 5 μl of 10X buffer per reaction.

200 μM dNTPs (50 μM of each of the four nucleotides). Add 1 μl of 10 mM dNTPs per reaction (dATP, dCTP, dTTP and dGTP are at 2.5 mM each).

1.5 mM Mg 2+ . Add only if it is not present in the 10X buffer or as needed for PCR optimization. For example, to obtain the 4.0 mM Mg 2+ required for optimal amplicon production of a conserved 566 bp DNA segment found in an uncharacterized Mycobacteriophage add 8 μl of 25 mM MgCl2 to the reaction (Figure 3).

20 to 50 pmol of each primer. Add 1 μl of each 20 μM primer.

Add 10 4 to 10 7 molecules (or about 1 to 1000 ng) DNA template. Add 0.5 μl of 2ng/μl genomic Mycobacteriophage DNA.

Add 0.5 to 2.5 units of DNA polymerase per 50 μl reaction (See manufacturers recommendations) For example, add 0.5 μl of Sigma 0.5 Units/μl Taq DNA polymerase.

Add Q.S. sterile distilled water to obtain a 50 μl final volume per reaction as pre-determined in the table of reagents (Q.S. is a Latin abbreviation for quantum satis meaning the amount that is needed). Thus, 33 μl per reaction is required to bring the volume up to 50 μl. However, it should be noted that water is added first but requires initially making a table of reagents and determining the volumes of all other reagents added to the reaction.

4. Basic PCR Protocol

Place a 96 well plate into the ice bucket as a holder for the 0.2 ml thin walled PCR tubes. Allowing PCR reagents to be added into cold 0.2 ml thin walled PCR tubes will help prevent nuclease activity and nonspecific priming.

Pipette the following PCR reagents in the following order into a 0.2 ml thin walled PCR tube (Figure 4): Sterile Water, 10X PCR buffer, dNTPs, MgCl2, primers, and template DNA (See Table 1). Since experiments should have at least a negative control, and possibly a positive control, it is beneficial to set up a Master Mix in a 1.8 ml microcentrifuge tube (See explanation in Notes).

In a separate 0.2 ml thin walled PCR tubes (Figure 4) add all the reagents with the exception of template DNA for a negative control (increase the water to compensate for the missing volume). In addition, another reaction (if reagents are available) should contain a positive control using template DNA and or primers previously known to amplify under the same conditions as the experimental PCR tubes.

Taq DNA polymerase is typically stored in a 50% glycerol solution and for complete dispersal in the reaction mix requires gentle mixing of the PCR reagents by pipetting up and down at least 20 times. The micropipettor should be set to about half the reaction volume of the master mix when mixing, and care should be taken to avoid introducing bubbles.

Put caps on the 0.2 ml thin walled PCR tubes and place them into the thermal cycler (Figure 5). Once the lid to the thermal cycler is firmly closed start the program (see Table 2).

When the program has finished, the 0.2 ml thin walled PCR tubes may be removed and stored at 4 ଌ. PCR products can be detected by loading aliquots of each reaction into wells of an agarose gel then staining DNA that has migrated into the gel following electrophoresis with ethidium bromide. If a PCR product is present, the ethidium bromide will intercalate between the bases of the DNA strands, allowing bands to be visualized with a UV illuminator.

Notes:

When setting up multiple PCR experiments, it is advantageous to assemble a mixture of reagents common to all reactions (i.e., Master Mix). Usually the cocktail contains a solution of DNA polymerase, dNTPs, reaction buffer, and water assembled into a 1.8 ml microcentrifuge tube. The amount of each reagent added to the Master Mix is equivalent to the total number of reactions plus 10% rounded up to the nearest whole reaction. For instance, if there are 10 x 0.1 = 1 reaction, then (10 + 1) x 5 μl 10X buffer equals 55 μl of 10X buffer for the Master Mix. The reagents in the Master Mix are mixed thoroughly by gently pumping the plunger of a micropipettor up and down about 20 times as described above. Each PCR tube receives an aliquot of the Master Mix to which the DNA template, any required primers, and experiment-specific reagents are then added (see Tables 1 and 7).

The following website offers a calculator for determining the number of copies of a template DNA (http://www.uri.edu/research/gsc/resources/cndna.html). The total number of copies of double stranded DNA may be calculated using the following equation: Number of copies of DNA = (DNA amount (ng) x 6.022x10 23 ) / (length of DNA x 1x10 9 ng/ml x 650 Daltons) Calculating the number of copies of DNA is used to determine how much template is needed per reaction.

False positives may occur as a consequence of carry-over from another PCR reaction which would be visualized as multiple undesired products on an agarose gel after electrophoresis. Therefore, it is prudent to use proper technique, include a negative control (and positive control when possible).

While ethidium bromide is the most common stain for nucleic acids there are several safer and less toxic alternatives. The following website describes several of the alternatives including Methylene Blue, Crystal Violet, SYBR Safe, and Gel Red along with descriptions of how to use and detect the final product (http://bitesizebio.com/articles/ethidium-bromide-the-alternatives/).

While most modern PCR machines use 0.2 ml tubes, some models may require reactions in 0.5 ml tubes. See your thermal cyclers manual to determine the appropriate size tube.

5. Calculating Melting Temperature (Tm)

Knowing the melting temperature (Tm) of the primers is imperative for a successful PCR experiment. Although there are several Tm calculators available, it is important to note that these calculations are an estimate of the actual Tm due to lack of specific information about a particular reaction and assumptions made in the algorithms for the Tm calculators themselves. However, nearest-neighbor thermodynamic models are preferred over the more conventional calculation: Tm ≈ 4(G-C) + 2(A-T). The former will give more accurate Tm estimation because it takes into account the stacking energy of neighboring base pairs. The latter is used more frequently because the calculations are simple and can be done quickly by hand. See Troubleshooting section for information about how various PCR conditions and additives affect melting temperature. For calculating the Tm values by nearest-neighbor thermodynamic models, one of the following calculators is recommended: http://www6.appliedbiosystems.com/support/techtools/calc/ http://www.cnr.berkeley.edu/

6. Setting Up Thermal Cycling Conditions

PCR thermal cyclers rapidly heat and cool the reaction mixture, allowing for heat-induced denaturation of duplex DNA (strand separation), annealing of primers to the plus and minus strands of the DNA template, and elongation of the PCR product. Cycling times are calculated based on the size of the template and the GC content of the DNA. The general formula starts with an initial denaturation step at 94 ଌ to 98 ଌ depending on the optimal temperature for DNA polymerase activity and G-C content of the template DNA. A typical reaction will start with a one minute denaturation at 94 ଌ. Any longer than 3 minutes may inactivate the DNA polymerase, destroying its enzymatic activity. One method, known as hot-start PCR, drastically extends the initial denaturation time from 3 minutes up to 9 minutes. With hot-start PCR, the DNA polymerase is added after the initial exaggerated denaturation step is finished. This protocol modification avoids likely inactivation of the DNA polymerase enzyme. Refer to the Troubleshooting section of this protocol for more information about hot start PCR and other alternative methods.

The next step is to set the thermal cycler to initiate the first of 25 to 35 rounds of a three-step temperature cycle (Table 2). While increasing the number of cycles above 35 will result in a greater quantity of PCR products, too many rounds often results in the enrichment of undesirable secondary products. The three temperature steps in a single cycle accomplish three tasks: the first step denatures the template (and in later cycles, the amplicons as well), the second step allows optimal annealing of primers, and the third step permits the DNA polymerase to bind to the DNA template and synthesize the PCR product. The duration and temperature of each step within a cycle may be altered to optimize production of the desired amplicon. The time for the denaturation step is kept as short as possible. Usually 10 to 60 seconds is sufficient for most DNA templates. The denaturation time and temperature may vary depending on the G-C content of the template DNA, as well as the ramp rate, which is the time it takes the thermal cycler to change from one temperature to the next. The temperature for this step is usually the same as that used for the initial denaturation phase (step #1 above e.g., 94 ଌ). A 30 second annealing step follows within the cycle at a temperature set about 5 ଌ below the apparent Tm of the primers (ideally between 52 ଌ to 58 ଌ). The cycle concludes with an elongation step. The temperature depends on the DNA polymerase selected for the experiment. For example, Taq DNA polymerase has an optimal elongation temperature of 70 ଌ to 80 ଌ and requires 1 minute to elongate the first 2 kb, then requires an extra minute for each additional 1 kb amplified. Pfu DNA Polymerase is another thermostable enzyme that has an optimal elongation temperature of 75 ଌ. Pfu DNA Polymerase is recommended for use in PCR and primer extension reactions that require high fidelity and requires 2 minutes for every 1 kb to be amplified. See manufacturer recommendations for exact elongation temperatures and elongation time indicated for each specific DNA polymerase.

The final phase of thermal cycling incorporates an extended elongation period of 5 minutes or longer. This last step allows synthesis of many uncompleted amplicons to finish and, in the case of Taq DNA polymerase, permits the addition of an adenine residue to the 3' ends of all PCR products. This modification is mediated by the terminal transferase activity of Taq DNA polymerase and is useful for subsequent molecular cloning procedures that require a 3'-overhang.

Termination of the reaction is achieved by chilling the mixture to 4 ଌ and/or by the addition of EDTA to a final concentration of 10 mM.

7. Important Considerations When Troubleshooting PCR

If standard PCR conditions do not yield the desired amplicon, PCR optimization is necessary to attain better results. The stringency of a reaction may be modulated such that the specificity is adjusted by altering variables (e.g., reagent concentrations, cycling conditions) that affect the outcome of the amplicon profile. For example, if the reaction is not stringent enough, many spurious amplicons will be generated with variable lengths. If the reaction is too stringent, no product will be produced. Troubleshooting PCR reactions may be a frustrating endeavor at times. However, careful analysis and a good understanding of the reagents used in a PCR experiment can reduce the amount of time and trials needed to obtain the desired results. Of all the considerations that impact PCR stringency, titration of Mg 2+ and/or manipulating annealing temperatures likely will solve most problems. However, before changing anything, be sure that an erroneous result was not due to human error. Start by confirming all reagents were added to a given reaction and that the reagents were not contaminated. Also take note of the erroneous result, and ask the following questions: Are primer dimers visible on the gel after electrophoresis (these run as small bands 𼄀 b near the bottom of the lane)? Are there non-specific products (bands that migrate at a different size than the desired product)? Was there a lack of any product? Is the target DNA on a plasmid or in a genomic DNA extract? Also, it is wise to analyze the G-C content of the desired amplicon.

First determine if any of the PCR reagents are catastrophic to your reaction. This can be achieved by preparing new reagents (e.g., fresh working stocks, new dilutions), and then systematically adding one new reagent at a time to reaction mixtures. This process will determine which reagent was the culprit for the failed PCR experiment. In the case of very old DNA, which often accumulates inhibitors, it has been demonstrated that addition of bovine serum albumin may help alleviate the problem.

Primer dimers can form when primers preferentially self anneal or anneal to the other primer in the reaction. If this occurs, a small product of less than 100 bp will appear on the agarose gel. Start by altering the ratio of template to primer if the primer concentration is in extreme excess over the template concentration, then the primers will be more likely to anneal to themselves or each other over the DNA template. Adding DMSO and or using a hot start thermal cycling method may resolve the problem. In the end it may be necessary to design new primers.

Non-specific products are produced when PCR stringency is excessively low resulting in non-specific PCR bands with variable lengths. This produces a ladder effect on an agarose gel. It then is advisable to choose PCR conditions that increase stringency. A smear of various sizes may also result from primers designed to highly repetitive sequences when amplifying genomic DNA. However, the same primers may amplify a target sequence on a plasmid without encountering the same problem.

Lack of PCR products is likely due to reaction conditions that are too stringent. Primer dimers and hairpin loop structures that form with the primers or in the denatured template DNA may also prevent amplification of PCR products because these molecules may no longer base pair with the desired DNA counterpart.

If the G-C content has not been analyzed, it is time to do so. PCR of G-C rich regions (GC content 㹠%) pose some of the greatest challenges to PCR. However, there are many additives that have been used to help alleviate the challenges.

8. Manipulating PCR Reagents

Understanding the function of reagents used on conventional PCR is critical when first deciding how best to alter reaction conditions to obtain the desired product. Success simply may rely on changing the concentration of MgCl2, KCl, dNTPs, primers, template DNA, or DNA polymerase. However, the wrong concentration of such reagents may lead to spurious results, decreasing the stringency of the reaction. When troubleshooting PCR, only one reagent should be manipulated at a time. However, it may be prudent to titrate the manipulated reagent.

Magnesium salt Mg 2+ (final reaction concentration of 0.5 to 5.0 mM) Thermostable DNA polymerases require the presence of magnesium to act as a cofactor during the reaction process. Changing the magnesium concentration is one of the easiest reagents to manipulate with perhaps the greatest impact on the stringency of PCR. In general, the PCR product yield will increase with the addition of greater concentrations of Mg 2+ . However, increased concentrations of Mg 2+ will also decrease the specificity and fidelity of the DNA polymerase. Most manufacturers include a solution of Magnesium chloride (MgCl2) along with the DNA polymerase and a 10X PCR buffer solution. The 10 X PCR buffer solutions may contain 15 mM MgCl2, which is enough for a typical PCR reaction, or it may be added separately at a concentration optimized for a particular reaction. Mg 2+ is not actually consumed in the reaction, but the reaction cannot proceed without it being present. When there is too much Mg 2+ , it may prevent complete denaturation of the DNA template by stabilizing the duplex strand. Too much Mg 2+ also can stabilize spurious annealing of primers to incorrect template sites and decrease specificity resulting in undesired PCR products. When there is not enough Mg 2+ , the reaction will not proceed, resulting in no PCR product.

Potassium salt K + (final reaction concentration of 35 to 100 mM) Longer PCR products (10 to 40 kb) benefit from reducing potassium salt (KCl) from its normal 50 mM reaction concentration, often in conjunction with the addition of DMSO and/or glycerol. If the desired amplicon is below 1000 bp and long non-specific products are forming, specificity may be improved by titrating KCl, increasing the concentration in 10 mM increments up to 100 mM. Increasing the salt concentration permits shorter DNA molecules to denature preferentially to longer DNA molecules.

Deoxynucleotide 5'-triphosphates (final reaction concentration of 20 and 200 μM each) Deoxynucleotide 5'-triphosphates (dNTPs) can cause problems for PCR if they are not at the appropriate equivalent concentrations (i.e., [A] = [T] = [C] = [G]) and/ or due to their instability from repeated freezing and thawing. The usual dNTP concentration is 50 μM of EACH of the four dNTPs. However, PCR can tolerate concentrations between 20 and 200 μM each. Lower concentrations of dNTPs may increase both the specificity and fidelity of the reaction while excessive dNTP concentrations can actually inhibit PCR. However, for longer PCR-fragments, a higher dNTP concentration may be required. A large change in the dNTP concentration may necessitate a corresponding change in the concentration of Mg 2+ .

Thermal stable DNA polymerases PCR enzymes and buffers associated with those enzymes have come a long way since the initial Taq DNA polymerase was first employed. Thus, choosing an appropriate enzyme can be helpful for obtaining desired amplicon products. For example the use of Taq DNA polymerase may be preferred over Pfu DNA polymerase if processivity and/or the addition of an adenine residue to the 3' ends of the PCR product is desired. The addition of a 3' adenine has become a useful strategy for cloning PCR products into TA vectors whit 3' thymine overhangs. However, if fidelity is more important an enzyme such as Pfu may be a better choice. Several manufactures have an array of specific DNA polymerases designed for specialized needs. Take a look at the reaction conditions and characteristics of the desired amplicon, and then match the PCR experiment with the appropriate DNA polymerase. Most manufactures have tables that aid DNA polymerase selection by listing characteristics such as fidelity, yield, speed, optimal target lengths, and whether it is useful for G-C rich amplification or hot start PCR.

Template DNA DNA quality and purity will have a substantial effect on the likelihood of a successful PCR experiment. DNA and RNA concentrations can be determined using their optical density measurements at 260 nm (OD260). The mass of purified nucleic acids in solution is calculated at 50 μg/ml of double stranded DNA or 40 μg/ml for either RNA or single stranded DNA at an OD260 =1.0. DNA extraction contaminants are common inhibitors in PCR and should be carefully avoided. Common DNA extraction inhibitors of PCR include protein, RNA, organic solvents, and detergents. Using the maximum absorption of nucleic acids OD260 compared to that of proteins OD280 (OD260/280), it is possible to determine an estimate of the purity of extracted DNA. Ideally, the ratio of OD260/280 is between 1.8 and 2.0. Lower OD260/280 is indicative of protein and/ or solvent contamination which, in all probability, will be problematic for PCR. In addition to the quality of template DNA, optimization of the quantity of DNA may greatly benefit the outcome of a PCR experiment. Although it is convenient to determine the quantity in ng/μl, which is often the output for modern nanospectrophotometers, the relevant unit for a successful PCR experiment is the number of molecules. That is, how many copies of DNA template contain a sequence complementary to the PCR primers? Optimal target molecules are between 10 4 to 10 7 molecules and may be calculated as was described in the notes above.

9. Additive Reagents

Additive reagents may yield results when all else fails. Understanding the reagents and what they are used for is critical in determining which reagents may be most effective in the acquisition of the desired PCR product. Adding reagents to the reaction is complicated by the fact that manipulation of one reagent may impact the usable concentration of another reagent. In addition to the reagents listed below, proprietary commercially available additives are available from many biotechnology companies.

10. Additives That Benefit G-C Rich Templates

Dimethylsulfoxide (final reaction concentration of 1-10% DMSO) In PCR experiments in which the template DNA is particularly G-C rich (GC content 㹠%), adding DMSO may enhance the reaction by disrupting base pairing and effectively lowering the Tm. Some Tm calculators include a variable entry for adding the concentration of DMSO desired in the PCR experiment. However, adding more than 2% DMSO may require adding more DNA polymerase as it has been demonstrated to inhibit Taq DNA polymerase.

Formamide (final reaction concentration of 1.25-10%) Like DMSO, formamide also disrupts base pairing while increasing the stringency of primer annealing, which results in less non-specific priming and increased amplification efficiency. Formamide also has been shown to be an enhancer for G-C rich templates.

7-deaza-2'-deoxyguanosine 5'-triphosphate (final reaction concentration of dc 7 GTP 3 dc 7 GTP:1 dGTP 50 μM) Using 3 parts, or 37.5 μM, of the guanosine base analog dc 7 GTP in conjunction with 1 part, or 12.5 μM, dGTP will destabilize formation of secondary structures in the product. As the amplicon or template DNA is denatured, it will often form secondary structures such as hairpin loops. Incorporation of dc 7 GTP into the DNA amplicon will prohibit formation of these aberrant structures.

dc 7 GTP attenuates the signal of ethidium bromide staining which is why it is used in a 3:1 ratio with dGTP.

Betaine (final reaction concentration of 0.5M to 2.5M) Betaine (N,N,N-trimethylglycine) is a zwitterionic amino acid analog that reduces and may even eliminate the DNA melting temperature dependence on nucleotide composition. It is used as an additive to aid PCR amplification of G-C rich targets. Betaine is often employed in combination with DMSO and can greatly enhance the chances of amplifying target DNA with high G-C content.

11. Additives That Help PCR in the Presence of Inhibitors

Non ionic detergents function to suppress secondary structure formation and help stabilize the DNA polymerase. Non ionic detergents such as Triton X-100, Tween 20, or NP-40 may be used at reaction concentrations of 0.1 to 1% in order to increase amplicon production. However, concentrations above 1% may be inhibitory to PCR. The presence of non ionic detergents decreases PCR stringency, potentially leading to spurious product formation. However, their use will also neutralize the inhibitory affects of SDS, an occasional contaminant of DNA extraction protocols.

Addition of specific proteins such as Bovine serum albumin (BSA) used at 400 ng/μl and/ or T4 gene 32 protein at 150 ng/μl aid PCR in the presence of inhibitors such as FeCl3, hemin, fulvic acid, humic acid, tannic acids, or extracts from feces, fresh water, and marine water. However, some PCR inhibitors, including bile salts, bilirubin, EDTA, NaCl, SDS, or Triton X-100, are not alleviated by addition of either BSA or T4 gene 32 protein.

12. Modifications to Cycling Conditions

Optimizing the annealing temperature will enhance any PCR reaction and should be considered in combination with other additives and/ or along with other modifications to cycling conditions. Thus, in order to calculate the optimal annealing temperature the following equation is employed: Ta OPT = 0.3 Tm Primer + 0.7 Tm Product -14.9 Tm Primer is calculated as the Tm of the less stable pair using the equation: Tm Primer = ((ΔH/(ΔS+R x ln(c/4)))-273.15 + 16.6 log[K + ] Where ΔH is the sum of the nearest neighbor enthalpy changes for hybrids ΔS is the sum of the nearest neighbor entropy changes R is the Gas Constant (1.99 cal K-1 mol-1) C is the primer concentration and [K + ] is the potassium concentration. The latter equation can be computed using one of the Tm calculators listed at the following website: http://protein.bio.puc.cl/cardex/servers/melting/sup_mat/servers_list.html Tm Product is calculated as follows: Tm Product = 0.41(%G-C) + 16.6 log [K + ] - 675/product length For most PCR reactions the concentration of potassium ([K + ]) is going to be 50 mM.

Hot start PCR is a versatile modification in which the initial denaturation time is increased dramatically (Table 4). This modification can be incorporated with or without other modifications to cycling conditions. Moreover, it is often used in conjunction with additives for temperamental amplicon formation. In fact, hot start PCR is increasingly included as a regular aspect of general cycling conditions. Hot start has been demonstrated to increase amplicon yield, while increasing the specificity and fidelity of the reaction. The rationale behind hot start PCR is to eliminate primer-dimer and non-specific priming that may result as a consequence of setting up the reaction below the Tm. Thus, a typical hot start reaction heats the sample to a temperature above the optimal Tm, at least to 60 ଌ before any amplification is able to occur. In general, the DNA polymerase is withheld from the reaction during the initial, elongated, denaturing time. Although other components of the reaction are sometimes omitted instead of the DNA polymerase, here we will focus on the DNA polymerase. There are several methods which allow the DNA polymerase to remain inactive or physically separated until the initial denaturation period has completed, including the use of a solid wax barrier, anti-DNA polymerase antibodies, and accessory proteins. Alternatively, the DNA polymerase may simply be added to the reaction after the initial denaturation cycle is complete.

Touchdown PCR (TD-PCR) is intended to take some of the guess work out of the Tm calculation limitations by bracketing the calculated annealing temperatures. The concept is to design two phases of cycling conditions (Table 5). The first phase employs successively lower annealing temperatures every second cycle (traditionally 1.0 ଌ), starting at 10 ଌ above and finishing at the calculated Tm or slightly below. Phase two utilizes the standard 3-step conditions with the annealing temperature set at 5 ଌ below the calculated Tm for another 20 to 25 cycles. The function of the first phase should alleviate mispriming, conferring a 4-fold advantage to the correct product. Thus, after 10 cycles, a 410-fold advantage would yield 4096 copies of the correct product over any spurious priming.

Stepdown PCR is similar to TD-PCR with fewer increments in the first phase of priming. As an example, the first phase lowers annealing temperatures every second cycle by 3 ଌ, starting at 10 ଌ above and finishing at 2 ଌ below the calculated Tm. Like TD-PCR, phase two utilizes the standard 3-step conditions with the annealing temperature set at 5 ଌ below the calculated Tm for another 20 to 25 cycles. This would allow the correct product a 256-fold advantage over false priming products.

Slowdown PCR is simply a modification of TD-PCR and has been successful for amplifying extremely G-C rich (above 83%) sequences (Table 6). The concept takes into account a relatively new feature associated with modern thermal cyclers, which allows adjustment of the ramp speed as well as the cooling rate. The protocol also utilizes dc 7 GTP to reduce 2 °structure formation that could inhibit the reaction. The ramp speed is lowered to 2.5 ଌ s -1 with a cooling rate of 1.5 ଌ s -1 for the annealing cycles. The first phase starts with an annealing temperature of 70 ଌ and reduces the annealing temperature by 1 ଌ every 3 rounds until it reaches 58 ଌ. The second phase then continues with an annealing temperature of 58 ଌ for an additional 15 cycles.

Nested PCR is a powerful tool used to eliminate spurious products. The use of nested primers is particularly helpful when there are several paralogous genes in a single genome or when there is low copy number of a target sequence within a heterogeneous population of orthologous sequences. The basic procedure involves two sets of primers that amplify a single region of DNA. The outer primers straddle the segment of interest and are used to generate PCR products that are often non-specific in 20 to 30 cycles. A small aliquot, usually about 5 μl from the first 50 μl reaction, is then used as the template DNA for another 20 to 30 rounds of amplification using the second set of primers that anneal to an internal location relative to the first set.

Other PCR protocols are more specialized and go beyond the scope of this paper. Examples include RACE-PCR, Multiplex-PCR, Vectorette-PCR, Quantitative-PCR, and RT-PCR.

13. Representative Results

Representative PCR results were generated by following the basic PCR protocols described above. The results incorporate several troubleshooting strategies to demonstrate the effect of various reagents and conditions on the reaction. Genes from the budding yeast Saccharomyces cerevisiae and from an uncharacterized Mycobacteriophage were amplified in these experiments. The standard 3-step PCR protocol outlined in Table 2 was employed for all three experiments described below.

Before setting up the PCR experiment, the genomic DNA from both S. cerevisiae and the Mycobacteriophage were quantified and diluted to a concentration that would allow between 10 4 and 10 7 molecules of DNA per reaction. The working stocks were prepared as follows. A genomic yeast DNA preparation yielded 10 4 ng/μl. A dilution to 10 ng/μl was generated by adding 48 μl into 452 μl of TE pH 8.0 buffer. Since the S. cerevisiae genome is about 12.5 Mb, 10 ng contain 7.41 X 10 5 molecules. The genomic Mycobacteriophage DNA preparation yielded 313 ng/μl. A dilution to 2 ng/μl was generated by adding 6.4 μl into 993.6 μl of TE pH 8.0 buffer. This phage DNA is about 67 Kb. Thus, 1 ng contains 2.73 X 10 7 molecules, which is at the upper limit of DNA generally used for a PCR. The working stocks were then used to generate the Master Mix solutions outlined in Table 7. Experiments varied cycling conditions as described below.

In Figure 3a, genomic DNA from S. cerevisiae was used as a template to amplify the GAL3 gene, which encodes a protein involved in galactose metabolism. The goal for this experiment was to determine the optimal Mg 2+ concentration for this set of reagents. No MgCl2 was present in the original PCR buffer and had to be supplemented at the concentrations indicated with a range tested from 0.0 mM to 5.0 mM. As shown in the figure, a PCR product of the expected size (2098 bp) appears starting at a Mg 2+ concentration of 2.5 mM (lane 6) with an optimal concentration at 4.0 mM (lane 9). The recommended concentration provided by the manufacturer was 1.5 mM, which is the amount provided in typical PCR buffers. Perhaps surprisingly, the necessary concentration needed for product formation in this experiment exceeded this amount.

A different DNA template was used for the experiment presented in Figure 3b. Genomic DNA from a Mycobacteriophage was used to amplify a conserved 566 bp DNA segment. Like the previous experiment, the optimal Mg 2+ concentration had to be determined. As shown in Figure 3b, amplification of the desired PCR product requires at least 2.0 mM Mg 2+ (lane 5). While there was more variability in the amount of product formed at increasing concentrations of MgCl2, the most PCR product was observed at 4 mM Mg 2+ (lane 9), the same concentration observed for the yeast GAL3 gene.

Notice that in the experiments presented in Figures 3A and 3B, a discrete band was obtained using the cycling conditions thought to be optimal based on primer annealing temperatures. Specifically, the denaturation temperature was 95 ଌ with an annealing temperature of 61 ଌ, and the extension was carried out for 1 minute at 72 ଌ for 30 cycles. The final 5 minute extension was then done at 72 ଌ. For the third experiment presented in Figure 3c, three changes were made to the cycling conditions used to amplify the yeast GAL3 gene. First, the annealing temperature was reduced to a sub-optimal temperature of 58 ଌ. Second, the extension time was extended to 1 minute and 30 seconds. Third, the number of cycles was increased from 30 to 35 times. The purpose was to demonstrate the effects of sub-optimal amplification conditions (i.e., reducing the stringency of the reaction) on a PCR experiment. As shown in Figure 3c, what was a discrete band in Figure 3a, becomes a smear of non-specific products under these sub-optimal cycling conditions. Furthermore, with the overall stringency of the reaction reduced, a lower amount of Mg 2+ is required to form an amplicon.

All three experiments illustrate that when Mg 2+ concentrations are too low, there is no amplicon production. These results also demonstrate that when both the cycling conditions are correctly designed and the reagents are at optimal concentrations, the PCR experiment produces a discreet amplicon corresponding to the expected size. The results show the importance of performing PCR experiments at a sufficiently high stringency (e.g., discreet bands versus a smear). Moreover, the experiments indicate that changing one parameter can influence another parameter, thus affecting the reaction outcome.

Table 1. PCR reagents in the order they should be added.

*Units may vary between manufacturers

** Add all reagents to the Master Mix excluding any in need of titration or that may be variable to the reaction. The Master Mix depicted in the above table is calculated for 11 reactions plus 2 extra reactions to accommodate pipette transfer loss ensuring there is enough to aliquot to each reaction tube.

 Standard 3-step PCR Cycling 
Cycle stepTemperatureTimeNumber of Cycles
Initial Denaturation94 ଌ to 98 ଌ1 minute1
Denaturation Annealing Extension94 ଌ 5 ଌ below Tm 70 ଌ to 80 ଌ 10 to 60 seconds 30 seconds Amplicon and DNA polymerase dependent 25-35
Final Extension70 ଌ to 80 ଌ5 minutes1
Hold*4 ଌ1

Table 2. Standard 3-step PCR Cycling.

* Most thermal cyclers have the ability to pause at 4ଌ indefinitely at the end of the cycles.

 2-step PCR Cycling 
Cycle stepTemperatureTimeNumber of Cycles
Initial Denaturation94 ଌ to 98 ଌ1 minute1
Denaturation Annealing/Extension94 ଌ 70 ଌ to 80 ଌ10 to 60 seconds Amplicon and DNA polymerase dependent 25-35
Final Extension70 ଌ to 80 ଌ5 minutes1

Table 3. 2-step PCR Cycling.

 Hot Start PCR Cycling 
Cycle stepTemperatureTimeCycles
Initial Denaturation60 ଌ to 95 ଌ5 minute then add DNA polymerase1
Denaturation Annealing Extension94 ଌ 5 ଌ below Tm 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and DNA polymerase dependent 25-35
Final Extension70 ଌ to 80 ଌ5 minutes1

Table 4. Hot Start PCR Cycling.

 Touchdown PCR Cycling 
Cycle stepTemperatureTimeCycles
Initial Denaturation94 ଌ to 98 ଌ1 minute1
Denaturation Annealing Extension94 ଌ X =10 ଌ above Tm 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and DNA polymerase dependent 2
Denaturation Annealing Extension94 ଌ X-1 ଌ reduce 1 ଌ every other cycle 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and polymerase dependent 28
Denaturation Annealing Extension94 ଌ 5 ଌ below Tm 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and DNA polymerase dependent 20-25
Final Extension70 ଌ to 80 ଌ5 minutes1

Table 5. Touchdown PCR Cycling.

 Slowdown PCR Cycling 
Cycle stepTemperatureTimeCycles
Initial Denaturation94 ଌ to 98 ଌ1 minute1
Denaturation Annealing Extension94 ଌ X ଌ =10 ଌ above Tm 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and polymerase dependent 2
Denaturation Annealing Extension94 ଌ X-1 ଌ reduce 1 ଌ every other cycle 70 ଌ to 80 ଌ*10 to 60 seconds 30 seconds Amplicon and polymerase dependent 28
Denaturation Annealing Extension94 ଌ 5 ଌ below Tm 70 ଌ to 80 ଌ10 to 60 seconds 30 seconds Amplicon and polymerase dependent 20-25
Final Extension70 ଌ to 80 ଌ5 minutes1

Table 6. Slowdown PCR Cycling.

*For slowdown PCR, the ramp speed is lowered to 2.5 ଌ s -1 with a cooling rate of 1.5 ଌ s -1 for the annealing cycles.

Table 7. Titration of Mg 2+ used in Figure 3.

Figure 1. Common problems that arise with primers and 3-step PCR amplification of target DNA. (a) Self-annealing of primers resulting in formation of secondary hairpin loop structure. Note that primers do not always anneal at the extreme ends and may form smaller loop structures. (b) Primer annealing to each other, rather than the DNA template, creating primer dimers. Once the primers anneal to each other they will elongate to the primer ends. (c) PCR cycles generating a specific amplicon. Standard 3-step PCR cycling include denaturation of the template DNA, annealing of primers, and extension of the target DNA (amplicon) by DNA polymerase.

Figure 2. Ice bucket with reagents, pipettes, and racks required for a PCR. (1.) P-200 pipette, (2.) P-1000 pipette, (3.) P-20 pipette, (4.) P-10 pipette, (5.) 96 well plate and 0.2 ml thin walled PCR tubes, (6.) Reagents including Taq polymerase, 10X PCR buffer, MgCl2, sterile water, dNTPs, primers, and template DNA, (7.) 1.8 ml tubes and rack.

Figure 3. Example of a Mg 2+ titrations used to optimize a PCR experiment using a standard 3-step PCR protocol. (a) S. cerevisiae Yeast genomic DNA was used as a template to amplify a 2098 bp GAL3 gene. In lanes 1 - 6, where the Mg 2+ concentration is too low, there either is no product formed (lanes 1-5) or very little product formed (lane 6). Lanes 7 - 11 represent optimal concentrations of Mg 2+ for this PCR experiment as indicated by the presence of the 2098 bp amplicon product. (b) An uncharacterized mycobacteriophage genomic DNA template was used to amplify a 566 bp amplicon. Lanes 1 - 4, the Mg 2+ concentration is too low, as indicated by the absence of product. Lanes 5 - 11 represent optimal concentrations of Mg 2+ for this PCR as indicated by the presence of the 566 kb amplicon product. (c) . S. cerevisiae Yeast genomic DNA was used as a template to amplify a 2098 bp GAL3 gene as indicated in panel a. However, the annealing temperature was reduced from 61 ଌ to 58 ଌ, resulting in a non-specific PCR bands with variable lengths producing a smearing effect on the agarose gel. Lanes 1 - 4, where the Mg 2+ concentration is too low, there is no product formed. Lanes 5 - 8 represent optimal concentrations of Mg 2+ for this PCR as seen by the presence of a smear and band around the 2098 kb amplicon product size. Lanes 9 - 11 are indicative of excessively stringent conditions with no product formed. (a-c) Lanes 12 is a negative control that did not contain any template DNA. Lane M (marker) was loaded with NEB 1kb Ladder.

Figure 4. Sterile tubes used for PCR. (1.) 1.8 ml tube (2.) 0.2 ml individual thin walled PCR tube, (3.) 0.2 ml strip thin walled PCR tubes and caps.

Figure 5. Thermal cycler. Closed thermal cycler left image. Right image contains 0.2 ml thin walled PCR tubes placed in the heating block of an open thermal cycler.


T5-LIKE PHAGES (SIPHOVIRIDAE)

Cloning Genes from T5 or BF23

Many restriction fragments from T5 or BF23 DNA are not directly clonable because they either code for lethal products or contain such strong promoters that the cells harboring them cannot survive. The strength of some T5 promoters has prompted their use in expression vectors, some of which are commercially available. Fragments that have been cloned are largely from the region from 58 to 92%, which includes mostly late structural genes with some early genes, and from 21 to 36%, which includes all the tRNA genes that are expressed during the early period. A small fragment from 2.1 to 3.4% in the pre-early region has also been cloned.

T5 genes that have been overproduced from an expression vector include gene D7-8-9 (coding for T5 DNA polymerase), gene D15 (coding for T5 5′-exonuclease), and 11p (coding for a lipoprotein that inactivates host cell receptors). BF23 genes 24 and 25 (coding for a minor and major tail protein, respectively) have also been cloned, sequenced, and expressed.


Basic Techniques to Manipulate Genetic Material (DNA and RNA)

To understand the basic techniques used to work with nucleic acids, remember that nucleic acids are macromolecules made of nucleotides (a sugar, a phosphate, and a nitrogenous base) linked by phosphodiester bonds. The phosphate groups on these molecules each have a net negative charge. An entire set of DNA molecules in the nucleus is called the genome. DNA has two complementary strands linked by hydrogen bonds between the paired bases. Exposure to high temperatures (DNA denaturation) can separate the two strands and cooling can reanneal them. The DNA polymerase enzyme can replicate the DNA. Unlike DNA, which is located in the eukaryotic cells’ nucleus, RNA molecules leave the nucleus. The most common type of RNA that researchers analyze is the messenger RNA (mRNA) because it represents the protein-coding genes that are actively expressed. However, RNA molecules present some other challenges to analysis, as they are often less stable than DNA.

DNA and RNA Extraction

To study or manipulate nucleic acids, one must first isolate or extract the DNA or RNA from the cells. Researchers use various techniques to extract different types of DNA (Figure 2). Most nucleic acid extraction techniques involve steps to break open the cell and use enzymatic reactions to destroy all macromolecules that are not desired (such as unwanted molecule degradation and separation from the DNA sample). A lysis buffer (a solution which is mostly a detergent) breaks cells. Note that lysis means “to split”. These enzymes break apart lipid molecules in the cell membranes and nuclear membranes. Enzymes such as proteases that break down proteins inactivate macromolecules, and ribonucleases (RNAses) that break down RNA. Using alcohol precipitates the DNA. Human genomic DNA is usually visible as a gelatinous, white mass. One can store the DNA samples frozen at –80°C for several years.

Figure 2: This diagram shows the basic method of DNA extraction.

Scientists perform RNA analysis to study gene expression patterns in cells. RNA is naturally very unstable because RNAses are commonly present in nature and very difficult to inactivate. Similar to DNA, RNA extraction involves using various buffers and enzymes to inactivate macromolecules and preserve the RNA.

Gel Electrophoresis

Because nucleic acids are negatively charged ions at neutral or basic pH in an aqueous environment, an electric field can mobilize them. Gel electrophoresis is a technique that scientists use to separate molecules on the basis of size, using this charge. One can separate the nucleic acids as whole chromosomes or fragments. The nucleic acids load into a slot near the semisolid, porous gel matrix’s negative electrode, and pulled toward the positive electrode at the gel’s opposite end. Smaller molecules move through the gel’s pores faster than larger molecules. This difference in the migration rate separates the fragments on the basis of size. There are molecular weight standard samples that researchers can run alongside the molecules to provide a size comparison. We can observe nucleic acids in a gel matrix using various fluorescent or colored dyes. Distinct nucleic acid fragments appear as bands at specific distances from the gel’s top (the negative electrode end) on the basis of their size (Figure 3). A mixture of genomic DNA fragments of varying sizes appear as a long smear whereas, uncut genomic DNA is usually too large to run through the gel and forms a single large band at the gel’s top.

Figure 3: a) Shown are DNA fragments from seven samples run on a gel, stained with a fluorescent dye, and viewed under UV light and b) a researcher from International Rice Research Institute, reviewing DNA profiles using UV light. (credit: a: James Jacob, Tompkins Cortland Community College b: International Rice Research Institute)

Nucleic Acid Fragment Amplification by Polymerase Chain Reaction

Although genomic DNA is visible to the naked eye when it is extracted in bulk, DNA analysis often requires focusing on one or more specific genome regions. Polymerase chain reaction (PCR) is a technique that scientists use to amplify specific DNA regions for further analysis (Figure 4). Researchers use PCR for many purposes in laboratories, such as cloning gene fragments to analyze genetic diseases, identifying contaminant foreign DNA in a sample, and amplifying DNA for sequencing. More practical applications include determining paternity and detecting genetic diseases.

Figure 4: Scientists use polymerase chain reaction, or PCR, to amplify a specific DNA sequence. Primers—short pieces of DNA complementary to each end of the target sequence combine with genomic DNA, Taq polymerase, and deoxynucleotides. Taq polymerase is a DNA polymerase isolated from the thermostable bacterium Thermus aquaticus that is able to withstand the high temperatures that scientists use in PCR. Thermus aquaticus grows in the Lower Geyser Basin of Yellowstone National Park. Reverse transcriptase PCR (RT-PCR) is similar to PCR, but cDNA is made from an RNA template before PCR begins.

DNA fragments can also be amplified from an RNA template in a process called reverse transcriptase PCR (RT-PCR) . The first step is to recreate the original DNA template strand (called cDNA) by applying DNA nucleotides to the mRNA. This process is called reverse transcription. This requires the presence of an enzyme called reverse transcriptase. After the cDNA is made, regular PCR can be used to amplify it.


DNA Manipulation

Bsu DNA Polymerase I, Large Fragment retains the 5´→ 3´ polymerase activity of the Bacillus subtilis DNA polymerase I (1), but lacks the 5´→ 3´ exonuclease domain. This large fragment naturally lacks 3´→ 5´ exonuclease activity.

DNA Polymerase I (E. coli) (NEB #M0209)

DNA Polymerase I (E. coli) is a DNA-dependent DNA polymerase with inherent 3´→ 5´ and 5´→ 3´ exonuclease activities (2). The 5´→ 3´ exonuclease activity removes nucleotides ahead of the growing DNA chain, allowing nick-translation.

DNA Polymerase I, Large (Klenow) Fragment (NEB #M0210)

DNA Polymerase I, Large (Klenow) Fragment is a proteolytic product of E. coli DNA Polymerase I, which retains polymerization and 3'→ 5' exonuclease activity, but has lost 5'→ 3' exonuclease activity (3). Klenow retains the polymerization fidelity of the holoenzyme without degrading 5' termini.

Klenow Fragment (3´→ 5´ exo- ) (NEB #M0212)

Klenow Fragment (3´→ 5´ exo- ) is an N-terminal truncation of DNA Polymerase I, which retains polymerase activity, but has lost the 5´→ 3´ exonuclease activity and has mutations (D355A, E357A), which abolish the 3´→ 5´ exonuclease activity (4).

Phi29 DNA Polymerase (NEB #M0269)

phi29 DNA Polymerase is the replicative polymerase from the Bacillus subtilis phage, phi29 (Φ29) (5). This polymerase has exceptional strand displacement and processive synthesis properties (6). The polymerase has an inherent 3´→5´ proofreading exonuclease activity (7).

T4 DNA Polymerase (NEB #M0203)

T4 DNA Polymerase catalyzes the synthesis of DNA in the 5´→ 3´ direction and requires the presence of template and primer. This enzyme has a 3´→ 5´ exonuclease activity, which is much more active than that found in DNA Polymerase I. Unlike E. coli DNA Polymerase I, T4 DNA Polymerase does not have a 5´→ 3´ exonuclease function.

T7 DNA Polymerase (unmodified) (NEB #M0274)

T7 DNA Polymerase (unmodified) catalyzes the replication of T7 phage DNA during infection. The protein dimer has two catalytic activities: DNA polymerase activity and strong 3´→ 5´ exonuclease activity (8,9,10). The high fidelity and rapid extension rate of the enzyme make it particularly useful in copying long stretches of DNA template.

Terminal Transferase (NEB #M0315)

Terminal Transferase (TdT) is a template-independent polymerase that catalyzes the addition of deoxynucleotides to the 3' hydroxyl terminus of DNA molecules. Protruding, recessed or blunt-ended double or single-stranded DNA molecules serve as a substrate for TdT. The 58.3 kDa enzyme does not have 5' or 3' exonuclease activity. The addition of Co 2+ in the reaction makes tailing more efficient.

References

  1. Okazaki, T. et al. (1964) J. Biol. Chem., 239, 259-268. PMID: 14114852
  2. Lehman, I.R.. (1981) P.D. Boyer (Eds.), The Enzymes, 14A, pp.16-38. San Diego: Academic Press.
  3. Jacobsen, H., Klenow, H. and Overgaard-Hansen, K. (1974) Eur. J. Biochem., 45, 623-7. PMID: 4605373
  4. Derbyshire, V. et al. (1988) Science, 240, 199-201. PMID: 1309614
  5. Blanco, L. and Salas, M. (1984) Proc. Natl. Acad. Sci. USA, 81, 5325-9. PMID: 6433348
  6. Blanco, L. et al. (1989) J. Biol. Chem., 264, 8935-40. PMID: 2498321
  7. Garmendia, C., et al. (1992) J. Biol. Chem., 267, 2594-9. PMID: 1733957
  8. Hori, K. et al. (1979) J. Biol. Chem., 258, 11598-11604. PMID: 227873
  9. Engler, M.J. et al. (1983) J. Biol. Chem., 258, 11165-73. PMID: 6411726
  10. Nordstrom, B. et al. (1981) J. Biol. Chem., 256, 3112-7. PMID: 7009606

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Feature Articles

Read about the relationship between Polymerase structure and function when copying DNA.

  • Random primer labeling
  • Second strand cDNA synthesis
  • Strand displacement DNA synthesis (1)
  • Nick translation of DNA to obtain probes with a high specific activity (2)
  • DNA sequencing by the Sanger dideoxy method (3) • Fill-in of 5´ overhangs to form blunt ends (4) • Second strand synthesis in mutagenesis protocols (5). • Removal of 3´ overhangs to form blunt ends (4)
  • Single strand deletion subcloning (6).
  • Addition of homopolymer tails to the 3' ends of DNA
  • Labeling the 3' ends of DNA with modified nucleotides (e.g., ddNTP, DIG-dUTP)
  • TUNEL assay (in situ localization of apoptosis)
  • TdT dependent PCR

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Polymerase Chain Reaction

Polymerase chain reaction is a technique used to produce multiple copies of DNA fragments. The process of DNA amplification is divided into three stages: the denaturing stage, annealing stage, and extending stage. In the denaturing stage, the DNA molecule is heated up to 95ºC to separate the two strands. Primers then attach to each of the strands in the annealing stage. The extending stage then proceeds to add more nucleotides into the growing chain. The steps are repeated over and over to produce more copies of the DNA.


Supporting information

S1 Fig. BOMB magnetic beads.

(A) Magnetic core particles (see S1 Appendix, BOMB protocol 1.1) in TEM. (B) Silica-coated magnetic beads (see S1 Appendix, BOMB protocol 2.1) in SEM and TEM. (C) Carboxyl-coated magnetic beads (see S1 Appendix, BOMB protocol 3.1) in LM, with and without an applied magnet. BOMB, Bio-On-Magnetic-Beads LM, light microscopy SEM, scanning electron microscopy TEM, transmission electron microscopy.

S2 Fig. Clean-up and size exclusion of DNA.

(A) Size exclusion of GeneRuler DNA Ladder Mix (Thermo) using silica-coated magnetic beads (see S1 Appendix, BOMB protocol 4.1). Different volumes of BB compared to sample volume were used to achieve size exclusion as a comparison, 2 volumes of cBB were used or no binding buffer was included (w/o BB) (B) Size exclusion of GeneRuler DNA Ladder Mix using carboxyl-coated magnetic beads (see S1 Appendix, BOMB protocol 4.2). Different volumes of BB compared to sample volume were used to achieve size exclusion as a comparison, 3 volumes of cBB were used or no beads were included (w/o beads). (C) Total recovery of approximately 6 μg plasmid DNA (input) using either a commercial kit that includes silica-packed columns (kit) or the S1 Appendix, BOMB protocol 4.1 for clean-up with silica-coated beads (BOMB). For binding, either cBB or the BB described in the BOMB protocol above was used. Error bars represent standard deviation, n = 3. Underlying data for S2 Fig can be found in S2 Data. BB, binding buffer BOMB, Bio-On-Magnetic-Beads cBB, commercial binding buffer.

S3 Fig. Optimisation and quality control of BOMB protocol 5.1—BOMB plasmid extraction from E. coli.

(A) Optimisation of reaction volume and media. One colony was picked and used to inoculate a 5 ml preculture containing LB media and ampicillin. The culture was incubated at 37 °C and 250 rpm until an OD of 0.6 was reached 5 μl of the preculture was used to inoculate different volumes (0.5 ml, 1.0 ml, 1.5 ml, and 2.0 ml) of a variety of growth media (LB, TB, SB, SOC, and ZYM505, including the respective antibiotic) in a 2.2 ml 96-well culture plate (Sarstedt). The plate was sealed with the lid of a 6-well cell culture plate, so a decent exchange of air was possible, and incubated at 37 °C and 250 rpm for 22 hours. Plasmid DNA was then isolated with the S1 Appendix, BOMB protocol 5.1, and the resulting concentration (c [ng/μl]) was determined. Error bars represent the standard deviation, n = 3. (B) Comparison of commercially purified plasmid DNA (kit) and BOMB extracted DNA (BOMB) with (+) and without (−) restriction enzyme digestion. MW: GeneRuler DNA Ladder Mix (Thermo Scientific). (C) Exemplary sequencing trace of a BOMB extracted plasmid via Sanger sequencing. A sequencing read length of at least 800–1,000 bp is typically observed. Underlying data for S3 Fig can be found in S3 Data. BOMB, Bio-On-Magnetic-Beads LB, Luria-Bertani Broth TB, Terrific BrothSB, Super Broth SOC, Super Optimal Broth.

S4 Fig. Genomic DNA isolation from various rabbit tissues.

Genomic DNA was isolated from the indicated tissues of a 12-hour deceased rabbit, using S1 Appendix, BOMB protocol 6.3 and (A) Speed Beads or (B) BOMB silica beads. A comparison to (C) phenol-chloroform–based extraction is also shown. MW in all panels represents Hyperladder I (Bioline). Inevitably, some tissues (like bone marrow) produce far greater (D) yields per mg of input material, compared to other tissues. However, the bead-based methods generally outperform phenol-chloroform extractions in our hands. Note: rabbit tissues were not preserved immediately after animal death, hence why tissues like spleen have experienced some DNA degradation. Underlying data for S4 Fig can be found in S4 Data. BOMB, Bio-On-Magnetic-Beads.

S5 Fig. Example genomic DNA isolation from clover leaves (Trifolium repens).

Genomic DNA was isolated from individual leaves (approximately 5 mg of tissue) using S1 Appendix, BOMB protocol 6.4 (high-throughput). A subset of DNA samples (18) from 96 extractions, represented in Fig 2O, were run on an agarose gel. MW: Hyperladder I (Bioline). BOMB, Bio-On-Magnetic-Beads.

S6 Fig. Bisulfite conversion.

(A) Scheme of DNA methylation analysis using bisulfite conversion. (B) Agarose gel after PCR amplification of bisulfite-converted human DNA. Multiple primer pairs with expected product sizes between 221 and 435 bp were tested successfully. MW: GeneRuler DNA Ladder Mix (Thermo Scientific). (C) Sequencing trace of a PCR-amplified bisulfite converted sample, aligned with the original, unconverted sequence. All non-CG cytosines were successfully converted. Conversion rate is approximately 99% as measured by Sanger sequencing after PCR amplification. BOMB, Bio-On-Magnetic-Beads CG, cytosine-guanine dinucleotide.


Watch the video: Gel Electrophoresis (February 2023).