Week 2: Agarose Gel Electrophoresis; Purification and Quantification of PCR Products; Degenerate PCR Primer Design

Introduction

The most common method of visualizing DNAs of different sizes is to separate them by agarose gel electrophoresis. Agarose is a neutral linear polysaccharide purified from agar-agar of red algae. It is composed of repeating sugar units, linked by alternating D-galactose and 3,6-anhydro-L-galactopyranose with α-(1→3) and β-(1→4) glycosidic bonds.¹⁰ Agarose is melted in hot water. When the melted agarose solution cools, a matrix forms through cross-linking of hydrogen bonds, creating agarose polymer chains that have a double-helical structure and form a web of pores (a gel). The higher the agarose concentration in the gel, the more fibers per cm³, resulting in smaller pore sizes.

The two most common running buffers for agarose gel electrophoresis are tris-acetate-EDTA (TAE) and tris-borate-EDTA (TBE). Both buffers contain the corresponding acetate or borate ions that move to the anode to donate electrons, which go through the gel’s wire to provide electrons to the cathode. Thus, H⁺ moves to the cathode to receive electrons from the anode (via wire) to produce H₂ gas. EDTA is a chelator. It forms strong complexes with divalent (e.g., magnesium) and trivalent (e.g., aluminum, iron) metal ions. Mg²⁺ or Ca²⁺ are required by DNases for optimal enzymatic activity. Without them, the DNA degrading enzymes are inactive. Tris maintains the proper pH of the buffer. Meanwhile, DNA in water ionizes at a neutral pH, releasing H⁺ from the phosphate (acidic nature), which leaves the DNA with a negative charge on each nucleotide. As a result, the highly negatively charged DNA molecule moves toward the anode in an electrical field. If the DNA is separated in water or a buffer, all the DNAs move together since they all have the same charge-to-mass ratio. However, if the DNAs must also move through a molecular sieve, e.g., an agarose gel, the smaller DNAs move more quickly than the larger ones. As a result, differently sized DNA fragments separate, with the smallest migrating the furthest distance from the starting position and the largest being the closest to the starting point.

For each agarose gel, a calibration curve can be constructed by measuring the distances of known DNA fragments, such as DNA molecular weight markers which have moved through the gel, and then plotting log₁₀ of DNA sizes in base pairs (bp) (Y-axis) versus their migration distances in cm (X-axis) (Figure 1-4). Each gel has an effective separation range for DNA fragments based on its agarose concentration. For example, the effective separation range for a 1.5% agarose gel is 200–7,000 bp. The sizes of unknown linear DNAs in the same gel can be extrapolated using the equation derived from the standard curve within the effective separation range or estimated by directly comparing them with the DNA markers of known sizes.

Agarose gels with different pore sizes corresponding to their agarose concentrations are used to separate DNA of different sizes. As agarose concentration increases, the effective separation ranges and sizes of DNA fragments to be separated decrease (Table 1-3).¹⁰ Since DNA is only visible after intercalating with a fluorescence dye, e.g., ethidium bromide under UV light, tracking dyes, e.g., bromophenol blue and xylene cyanole that do not interact with the DNA are used to monitor electrophoresis progress and ensure DNA samples to be retained in the gel while maintaining good separation. Buffer type, TAE or TBE, and agarose concentration determine the migration of either tracking dye (Table 1-3).

In addition to the size of linear DNA and agarose concentration, the mobility of DNA is affected by applied current/voltage, temperature, and salt. Increasing voltage and temperature increases mobility. We cannot separate DNA in water because ions are the driving force of the electric current. Although salt increases gel pore size, significantly increasing the ionic strength of the running buffer decreases mobility (extra ions shield DNA from the electric field). DNA conformation also affects the mobility of DNA. For example, the mobility of intact super-coiled (or covalently closed-circular) plasmid DNA, which has a compact form due to a built-in twist, is higher than that of the same-size linear DNA.

Since DNA is invisible to the naked eye, the common fluorescent dye, ethidium bromide, has been used to visualize DNA in the gel (Figure 1-5 left). However, the dye is highly toxic. In contrast, GelRed™, made of two ethidium subunits bridged by a linear oxygenated spacer, is a sensitive, stable, and relatively safe fluorescent nucleic acid dye (Figure 1-5 right).¹¹ It is designed to replace the highly toxic ethidium bromide for staining dsDNA, ssDNA, or RNA in agarose gels or polyacrylamide gels. Like EtBr, GelRed binds DNA as it passes through the gel, staining the DNA and making it bulkier and thus move slower, but it does not require a destaining step.

Degenerate PCR

Unlike standard PCR, which has a specific primer pair to amplify target DNA, degenerate PCR uses a mixture of primers with similar sequences that cover all possible nucleotide combinations for a given protein sequence to amplify unknown DNA sequences in a PCR.^12,13 The primer degeneracy is the number of potential unique primer sequences corresponding to a protein sequence/stretch of amino acid sequence used in a PCR mix. Like standard PCR, primer design is a critical step in degenerate PCR. Since the exact target sequence is unknown, the two primer sequences for degenerate PCR cloning are deduced from the corresponding conserved amino acid sequences at the target sites. The corresponding nucleotide sequences may vary at the > 1 position because of genetic codon degeneracy.¹²

Steps for designing a degenerate primer are to i)  align multiple known amino acid sequences of the target site using free online software such as Clustal Omega, ii) target an area ~200–500 bp for optimal PCR amplification, iii)  design forward and reverse primers, each covering 6–7 amino acids (~20 bases) in conserved regions—the less degeneracy, the better.^12,13 For example, a degenerate forward primer encoding a peptide composed of the seven amino acids Thr-Tyr-Pro-His-Glu-(Asp/Glu)-Met is “5′- ACI TA(T/C) CCI CA(C/T) GA (A/G) GAI ATG-3′.” I is inosine, representing all four possible nucleotides (G, A, T, C) at the position. The primer contains a mixture of 8 (2 x 2 x 2) primers (degeneracy = 8) with 21 nucleotides. The degeneracy in the three nucleotides at the 3′ end should be avoided for primer design. Met- or Trp-encoding triplets at the 3′ end should be used if possible.^12,13 It may be necessary to reduce degeneracy by allowing some mismatches between the primer and template to increase the primer-template binding efficiency, especially toward the 5′ end rather than the 3′ end.

For starting, it is recommended to start PCR with 0.2 µM of primer concentration and then increase primer concentrations in increments of 0.25 µM until satisfactory results are achieved.¹¹ Try to design primers with < 4-fold degeneracy with any nucleotide. If possible, include the amino acids methionine and tryptophan, both of which are encoded by a single codon. Exclude leucine, serine, and arginine since each is encoded by six possible codon combinations and avoid degeneracy in the three nucleotides at the 3′ end. For subsequent cloning and increasing primer length (thus annealing temperature), tails (6–9 bases) containing a restriction enzyme site can be added. If no bases match, consider using “inosine,” which is structurally similar to guanine and can pair with all four bases, cytosine preferentially. Alternatively, a nitrogenous (Ny) base can be added to ensure equimolar concentrations of each base at that position in your primer mix.

It can be challenging to optimize degenerate PCR because within a degenerate primer mixture, i) not all primers correspond to the template, ii) the melting temperature (Tm) of different primers may differ significantly, and iii) there is a higher probability of primers complementing/annealing to each other or non-target templates than the regular PCR with a single pair of primers, and thus requires effort to minimize non-specific primer-template and primer-primer interactions during amplification.¹² Nevertheless, degenerate PCR cloning is extremely sensitive and effective in isolating conserved sequences. It is also valuable for identifying new members of a gene family or orthologous genes (genes of shared ancestry between species) from different organisms.

Most cloning reactions perform better with purified DNA fragments. PCR often requires a high salt condition that may interfere with the cloning reaction. The fastest and most common way to clean up PCR products is the use of PCR clean-up kits/columns. They will remove most, if not all, enzymes, salts, and remaining dNTPs in the PCR product. Nevertheless, it is not crucial to purify the PCR product in this experiment as long as only < 2 µl of the PCR reaction without further dilution is used. In addition to validating the size of the expected PCR product by agarose gel electrophoresis, this lab exercise designs degenerate primers from known amino acid sequences.

Procedure

A. Agarose Gel Electrophoresis

1. Seal a gel tray with masking tape and place a comb on it.
2. Heat 0.72 g of agarose plus 45 ml of 1X TAE (0.9 ml 50X TAE, brought up to 45 ml with RO water) in a 125 ml flask until the agarose completely melts. What is the percentage of agarose in the gel?
3. After cooling to ~65°C, add 4.5 µl of GelRed and mix well (avoid making bubbles in the flask).
4. Pour the agarose solution into a gel tray and allow it to solidify.
5. Place the gel tray in the electrophoresis box, add 300 ml of 1X TAE buffer, and remove the comb.
6. Add 2 µl of 6X DNA loading buffer and 10 µl of PCR product on a parafilm strip, and mix the aliquot by pipetting it up and down on the parafilm. This step can be skipped if you use the RED JumpStart Taq (5 U/µl).
7. Load the entire aliquot into the well.
8. Load 5 µl of DNA molecular weight size marker.
9. Electrophorese samples at a constant voltage of 8V/cm in 1X TAE until the bromophenol blue is approximately 3/4 of the distance through the gel (~1 1/2 hrs at 100V).
10. While waiting for the gel electrophoresis, quantify your PCR sample. Important: You must first measure each absorbance, and then calculate OD ratios and record them in Table 1-4 below. These results will be compared to readings taken after further purification of the PCR products in section B.
11. After completing gel electrophoresis, visualize DNA fragments in the gel under a Biospectrum UVP GelDoc-It^®2 Imager and photograph.

B. Purification of PCR Product

To assess whether the purification step improves cloning efficiency, half of the groups should further purify their PCR product using a PCR cleanup kit. The remaining groups may either help with the purification step or take a break to observe the process, as the procedure takes only 5–10 minutes.

The protocols are adapted from QIAquick PCR Purification Kit.¹⁴

1. Add 5 volumes of Buffer PB to 1 volume of the PCR sample and mix. If the mixture is orange or violet, add 10 µl of 2M sodium acetate (pH 5.0) and mix. The mixture will turn yellow.
2. Obtain a QIAquick spin column in a provided 2 ml collection tube.
3. To bind DNA, apply the sample to the QIAquick column and centrifuge for 1 min. Discard the flow-through. Place the QIAquick column back into the same tube.
4. Add 750 µl of Buffer PE to the QIAquick column and centrifuge for 1 min to wash.
5. Discard the flow-through and place the QIAquick column back in the same tube.
6. Centrifuge the column again in the same collection tube for 1 min to remove the residual wash buffer. Note: Residual ethanol from Buffer PE is not entirely removed unless the flow-through is discarded before this additional centrifugation.
7. Place the QIAquick column in a clean 1.5 ml microcentrifuge tube.
8. To elute DNA, add 25 µl of Buffer EB (10 mM Tris·HCl, pH 8.5) or water (pH 7.0–8.5) to the center of the QIAquick membrane, then centrifuge the column for 1 min. Note: Ensure that the elution buffer is dispensed directly onto the QIAquick membrane for complete elution of bound DNA.
9. Transfer the sample into a 0.5 ml PCR tube and measure the OD and record the results in Table 1-4 above.

Data Analysis and Discussion

1. a.   Based on what you learned from this lab, how do you prepare 100 ml of 1.5% TAE agarose gel with GelRed nucleic acid binding stain using 50X TAE buffer and 10,000X GelRed? How many different primers (degeneracy) are in the LM637 primer pool that we used?
   2. How many possible “TARGETS” exist for the LM637? Show your calculation. (Hint: “I” can form H-bonds with A, T, C, and G equally.)
2. a.   What is the amino acid sequence corresponding to the reverse primer sequence LM 637: 5′-A(A/G)IGCTA(A/G)IGG IA(A/G)ICC-3′? (You need to consider how the reverse primer is designed and the relationship of the amino acid sequence to the corresponding DNA coding sequence.)
   2. Design degenerate PCR forward primer(s) for the amino acid sequence “Trp-(ser/thr)-ala-gly-his-gly-thr.”
   3. How many different primers are in the primer mix for (b)? (Remember that “I” is expensive, and you should try to get the shortest primer possible.)
3. a.   Make a calibration curve based on the log molecular weight of the DNA marker log DNA in bp (Y-axis) versus migration distance in cm (X-axis). Make sure axes are properly labeled.
   2. Use the calibration curve to estimate the size of your PCR product(s).
4. After genomic DNA isolation, the quality of DNA can be checked by agarose gel electrophoresis to see if the DNA is intact. Suppose we had examined the isolated genomic DNA prior to PCR by agarose gel electrophoresis last week. What would you expect to see in the gel, assuming there is no DNase activity in the sample? (Hint: Consider the size of grape genomic DNA, % of agarose concentration, and the isolation procedure.)