Week 1: Genomic DNA Isolation and Quantification; Degenerate PCR

Introduction

The core of genomic DNA isolation is to purify DNA from cellular material and prevent its degradation. Procedures for DNA extraction from plant tissue vary depending on the source materials. Nevertheless, all involve the mechanical breakdown of membranes and cell walls to allow access to nuclear material without degrading DNA. Similarly, in plants, an initial grinding with liquid N₂ breaks down cell walls and membranes to allow the release of DNA. Meanwhile, harmful cellular enzymes and chemicals remain inactivated by the cold. The ground tissue is then resuspended in a proper buffer containing detergent, i.e., CTAB/SDS, to destroy plasma, nuclear, and organelle membranes to release DNAs. To obtain pure DNA, unwanted insoluble materials are removed through centrifugation, while soluble proteins and other materials are eliminated from DNA through further purification steps. DNA is then precipitated from the aqueous phase using ethanol and salt or by binding to a silica-based membrane. The DNA pellet and those attached to the membrane are washed to remove the excess salt. Finally, the purified DNA is resuspended or eluted from the membrane and stored in tris-EDTA (TE) buffer or sterile distilled water. During isolation, RNases can be added to the lysis buffer to remove RNAs.

After obtaining genomic DNA in the solution, we cannot tell if DNA is present by visualizing it. However, DNA and RNA bases absorb UV light at 260 nm, measured as an optical density (OD). Their presence and concentration in a solution can be indicated by the absorbance at 260 nm (A260). NanoDrop One UV-Vis spectrophotometer is used to analyze DNA quantity and purity.⁵ The DNA solution is placed on the stage connected to the end of the unit’s fiber optic cable, and a 2^nd cable makes contact with the solution to make a “bridge” between the two cables. UV light (260 nm) is passed through the sample, and the absorbance is recorded.

According to the Beer Lambert’s law, when light passes through a solution, it is absorbed by particles or molecules in it. The amount of light absorbed is directly proportional to the concentration (c) and path length (l).

We can express this relationship mathematically as: A = εcl, where

A: absorbance (dimensionless quantity)
ε: molar absorptivity (L mol^-1 cm^-1). (A higher ε value means stronger absorption at that wavelength.)
c: concentration (molar mass mol L^-1)
l: path length (cm)

• • It is the physical distance that the light beam travels through the absorbing medium. The longer the path length, the more opportunities there are for the light to interact with the sample molecules, leading to higher absorbance.

An A260 of 1 equals 50 µg/ml (50 ng/µl) double-stranded DNA (dsDNA). A typical UV spectrophotometer can also determine the concentration of the nucleic acid sample based on OD readings obtained and the equation:

DNA concentration (µg/ml) = (A260 – A320) x 50 µg/ml per A260 unit (dsDNA) x dilution.

(1 A260 = 50 µg/ml dsDNA; 40 µg/ml RNA; 33 µg/ml single-stranded DNA (ssDNA))

In addition to quantifying nucleic acids, ODs at various wavelengths indicate impurities in the samples (Table 1-1). An OD ratio of A260/A280 for pure RNA and DNA should be ~2.0 and 1.8, respectively.⁵ A lower ratio indicates protein contamination, which can affect downstream manipulations. Small changes in the pH of the solution can also cause the A260/A280 ratio to vary. Acidic solutions decrease the A260/A280 ratio by 0.2–0.3, while a basic solution increases by the same magnitude. The OD ratio of A260/A230 should be 1.8–2.2 in a pure nucleic acid sample.⁵ A ratio < 1.8 indicates significant organic compound contamination, which can affect downstream manipulations such as in vitro transcription. Also, possible contamination from isolation reagents, i.e., phenol, TRIzol, and chaotropic salt, can result in a high shoulder at 220 nm and a bulge on the peak shoulder at 270 nm, thus affecting the OD ratios of A260/A280 and A260/A230, as well as DNA concentration based on OD. In plants, carbohydrate contamination is often problematic, resulting in a lower A260/A230 ratio.

Since its discovery in 1985, the polymerase chain reaction (PCR) has been used in a wide variety of studies in molecular biology. In comparison with screening cDNA or genomic library, PCR cloning is a simple alternative with its straightforward procedure and diverse applications. Each DNA to be cloned requires a pair of primers flanking the sequence to be amplified, which goes through 30–40 cycles of amplification.⁶ Each repeating cycle has three stages: 1) Denaturation, the temperature increases to ~95–98°C for 15–30 sec, and the high temperature causes the double-stranded DNA template to denature, forming two single-stranded DNA templates; 2) Annealing, the temperature is lowered to ~50–56°C for specific amplification, and the primers bind to the single-stranded DNA templates to provide a starting point for the polymerase enzyme; 3) Extension, the temperature is raised to ~72–75°C for 15–30 seconds, and the Taq polymerase extends the primers by adding nucleotides to the 3′ end of the DNA template, creating a new dsDNA molecule.⁵ Hot-start Taq requires an extra denaturation step at ~95–98°C for 1–5 minutes for Taq to activate before the cycle starts. Degenerate PCR is a modified PCR technique, where a mixture of primers with all possible nucleotide sequences at positions of corresponding amino acid sequences are used due to the redundancy of codons. It is used to clone or analyze families of related genes. Like regular PCR, the primer design of degenerate PCR is crucial to the success of cloning and will be discussed further in the next lab exercise.

As indicated earlier, partial nbs-lrr genes were cloned from different plant species by PCR using degenerate primers corresponding to the conserved motifs within the NBS domain (Figure 1-1). In this lab exercise, genomic DNA is isolated from grape leaves and PCR-amplified using the degenerate PCR primers LM638 and LM637, which correspond to P-loop and GLPL within the NBS domain. The size(s) of PCR product(s) is then verified by agarose gel electrophoresis (week 2). If more than one PCR product (band) is present, the one similar to the NBS sequence’s known size is extracted, purified from the gel, and then cloned into a plasmid vector to be transformed into E. coli (week 3).

Procedure

A. Plant genomic DNA isolation

Procedure follows the protocols described in DNeasy Plant Handbook⁷ with minor modifications.

1. Retrieve the 1.5 ml microcentrifuge tubes containing the leaf samples from the -70°C freezer and place them in a container filled with liquid N₂.
2. Grind ~80 mg of frozen leaf in each tube to a fine powder with a plastic pestle. Once the sample starts to thaw, continue to step 3 immediately.
3. Add 400 µl of AP1 and then 4 µl of RNase A or RNase mix solution (100 mg/ml) to the powder (do not mix AP1 Buffer and RNase A before use).

   Alternative for steps 1–3 to be performed by the instructor

   For a lab of six groups, 0.5 g of leaf sample is ground into powder in a mortar containing liquid N₂, followed by additions of 2.5 ml of AP 1 and 25 µl of RNase mix solution (100 mg/ ml) to the sample mixture and grounded for another 10 seconds. Each group is provided with 400 µl of the ground mixture in a 1.5 ml microcentrifuge tube before continuing to step 4.

4. Quickly vortex vigorously for 5 seconds until no tissue clumps are visible. Vortex or pipet further to remove any lumps. Clumps of tissue will not lyse properly, resulting in a lower yield of DNA.
5. Incubate the mixture for 10 mins at 65°C. Mix 2–3 times during incubation by inverting the tube. This step lyses the cells.
6. Add 130 µl of P3 Buffer to the lysate, vortex the lysate, and incubate on ice for 5 mins. This step precipitates detergent, proteins, and polysaccharides.
7. Centrifuge the lysate for 5 mins at 20,000 g (14,000 rpm) or the top speed. (Some plant materials can generate very viscous lysates and large quantities of precipitants, resulting in the shearing of DNA in the next step. Optimal results can be obtained if most of these precipitants are removed by centrifugation for 5 mins at 20,000 g.)
8. Pipet the supernatant into the QIAshredder Mini spin column (lilac), place it in a 2 ml collection tube, and centrifuge for 2 mins at 20,000 g (14,000 rpm). This step reduces the viscosity of the homogenate. It may be necessary to cut off the end of the pipet tip to apply the lysate to the QIAshredder Mini spin column. The column removes most precipitates and cell debris, but a small amount passes through and forms a pellet in the collection tube. Do not disturb this pellet in the next step.
9. Transfer the flow-through fraction into a new 1.5 ml microcentrifuge tube without disturbing the cell-debris pellet. Approximately 450 µl of filtrate from the lysate is recovered. For some plant species, less is recovered. In this case, determine the volume of the filtrate for the next step.
10. Add 1.5 volumes of Buffer AW1 to the cleared lysate and mix by pipetting up and down. For example, for 450 µl lysate, add 675 µl Buffer AW1. If the lysate volume is small, reduce the amount of Buffer AW1 accordingly. A precipitate may form after adding Buffer AW1, but this does not affect the procedure.
11. Pipet 650 µl of the mixture, including any precipitant that may have formed, into the DNeasy Mini spin column in a 2 ml collection tube (supplied). Centrifuge for 1 min at ≥ 6000 g (≥ 8000 rpm for most microcentrifuges). We centrifuge at 10,000 rpm unless specified otherwise and then discard the flow-through. Reuse the collection tube in the next step.
12. Repeat the last step by loading the remaining sample and centrifuge. Steps 10–11 precipitate and bind the DNA into the column.
13. Place the DNeasy Mini spin column in a new 2 ml collection tube (provided by the instructor), add 500 µl of Buffer AW2, and centrifuge for 1 min at 10,000 rpm. Discard the flow-through and reuse the collection tube. This step removes excess salt.
14. Add 500 µl of Buffer AW2 to the DNeasy Mini spin column and centrifuge for 2 mins at 20,000 g (14,000 rpm) to dry the membrane. The next step is critical, so please pay attention!
15. After centrifuging, transfer the DNeasy Mini spin column to a 1.5 ml microcentrifuge tube without a cap and prevent the column from contacting the flow-through. If contact is made, it will result in the carryover of ethanol, which will significantly reduce yield. Also, the residual ethanol may obscure quantification and interfere with subsequent reactions. This centrifugation step ensures that no residual ethanol is carried over during elution. Discard the flow-through and collection tube.
16. Pipet 40 µl of Buffer AE onto the center of the Mini spin column without touching the membrane inside. Incubate the Mini spin column for 5 min at room temperature (15–25°C) and centrifuge for 1 min at 10,000 rpm to elute DNA. Transfer the DNA to a 1.5 ml microcentrifuge tube on ice.

B. Analysis of Concentration and Purity of DNA using NanoDrop One UV-Vis Spectrophotometer⁵

Before taking measurements, lift the instrument arm and clean the upper and lower pedestals with a new laboratory wipe.

To measure nucleic acid

1. From the Home screen, select the Nucleic Acid and choose dsDNA from the menu of dsDNA, ssDNA, or RNA.
2. Pipette 2 µl of blanking solution onto the lower pedestal and lower the arm, or insert the blanking cuvette into the cuvette holder. Tip: If using a cuvette, make sure to align the light path with the instrument light path.
3. Tap “Blank” and wait for the measurement to complete. Tip: If Auto-Blank is On, the blank measurement starts automatically after you lower the arm (this option is not available for cuvette measurements.)
4. Lift the arm and clean both pedestals with a new laboratory wipe, or remove the blanking cuvette.
5. Pipette 2 µl of sample solution onto the pedestal and lower the arm, or insert the sample cuvette into the cuvette holder.
6. Start the sample measurement. Pedestal: If “Auto-Measure” is off, lower the arm and tap “Measure” or Cuvette: Tap “Measure.”
7. When you finish measuring samples, record the results in Table 1-2 and then tap “End Experiment.” Alternatively, you can export the data into a USB port.
8. Lift the arm and clean both pedestals with a new wipe, or remove the sample cuvette.
9. Label tubes with your group number and store them at -20°C.

C. Set Up PCR (Use filter tips!)

Each group will obtain one 0.2 ml PCR tube from the instructor.

1. Add 10 µl sterile water, 25 µl of HotMaster 2X PCR reaction mix, 2.5 µl of each primer, and 10 µl of your genomic DNA.

*Due to the low Taq polymerase activity of Promega 2X PCR Master Mix, we add 1 µl of JumpStart Taq (5U/µl) per 50 µl PCR reaction to obtain good amplification.

2. Two groups also set up two 25 µl of PCR reactions, each containing either of the two primers.

*Add 0.5µl of JumpStart Taq (5U/µl) per 25µl PCR reaction.

3. Close the caps of the PCR tube tightly and amplify the samples using a thermal cycler.

PCR:

• • 1 cycle of 94°C for 2 mins (with HotStart Taq only)
  • 35 cycles of 94°C for 30 sec, 45°C for 30 sec, 72°C for 1 min

4. The instructor will collect leftover genomic DNA samples and store them in a freezer.

Data Analysis and Discussion

1. a.   Based on the measurement obtained from NanoDrop, does the isolated genomic DNA appear to be pure? Explain.
   2. Can you differentiate successful genomic DNA isolation (pure DNA sample) from one contaminated with RNA by spectrometry? Why or why not? How do you know if the isolated DNA sample is contaminated with RNA? Explain.
2. a.   If today’s lab is a success, based on what you learned about degenerate PCR, do you expect to see single or multiple bands when you separate your PCR product in an agarose gel? Explain.
   2. If you only see one band after agarose gel electrophoresis, does it mean you obtained a single identical PCR product? Explain.
3. Instead of using NanoDrop, you pipette 20 µl of your DNA samples into a Quartz cuvette and bring up the volume to 1 ml before measuring the ODs with a UV spectrophotometer; you obtained A260 = 0.057, A280 = 0.030, A230 = 0.028, and A320 = 0.
   1. What is the concentration (ng/µl) of the DNA sample?
   2. Is your DNA sample pure? Explain and show your calculation.
4. You measure the PCR product and obtain OD readings of A260/A280 = 1.56 and A260/A230 = 1.69, as well as a concentration = 474.3 ng/µl. After further purification, you measure OD readings of A260/A280 = 1.85 and A260/A230 = 2.15, with concentration = 30.3 ng/µl. Explain what causes the change in (a) DNA concentration, (b) A260/A280, and (c) A260/A230. Indicate at least one possible cause for each change if the volume of the DNA sample remains the same before and after purification.

5. To prepare for next week’s degenerate PCR primer design, review the relationships among the DNA coding strand, template strand, mRNA, anticodon on the tRNA, and amino acid codon. What is the sequence relationship of the coding strand to that of its (a) mRNA, (b) anticodon on tRNA, and (c) amino acid codon?

6. As part of the homework, please watch the following videos before next week’s lab: “Making an Agarose Gel”⁸ and “Running an Agarose Gel” from University of Leicester.⁹