Week 1: Plasmid DNA Isolation and Quantification; Restriction Digestion; Sample Preparation for DNA Sequencing

Introduction

Plasmid DNA isolation from Escherichia coli (E. coli) is a routine procedure in research laboratories. Two standard methods for plasmid DNA isolation are boiling or alkaline lysis.^9,10 The former is quick and recommended for isolating small plasmids (< 10 kb, kilobase) but not for plasmids > 10 kb. Although boiling lysis may yield more plasmids than alkaline lysis, it has an inconsistent yield and impurity, making it less desirable for research-scale preparations.¹¹

This lab exercise employs a commercially available miniprep kit for plasmid DNA isolation based on the alkaline lysis method. An overnight bacterial culture containing the cloned recombinant plasmid is lysed with an alkaline lysis buffer consisting of a detergent sodium dodecyl sulfate (SDS) and NaOH (a strong base).¹⁰ The NaOH ruptures cells and denatures the DNAs and proteins that maintain cell membrane structure. The SDS breaks open the phospholipid bilayer of the membrane. Then, a neutralizing step follows to allow the plasmid and bacterial DNA to reanneal. However, the bacterial chromosomal DNA reannealing is incomplete and tangled with cell debris, which is then removed through precipitation and centrifugation. The plasmid in the supernatant is obtained and purified by mixing and precipitating it with salt and ethanol, followed by centrifugation. The preparation yields relatively clean DNA quickly, ready for sequencing or other downstream manipulations, including restriction digestion.

Restriction enzymes made in bacteria and archaea provide a defense mechanism against bacteriophage, where they selectively cut up invading foreign DNAs. Meanwhile, endogenous methylases methylate the host’s DNAs to protect them from degradation (restriction-modification system). The name of each enzyme contains one or more capital letters (genus initials) and two small letters (species) followed by a Roman numeral (the order of enzymes discovered in the organism). Additional information may be added as a letter. For example, in EcoRI, the RI indicates E. coli strain RY13, the first restriction enzyme isolated from E. coli.

Four major restriction enzyme types (I-IV) are classified based on subunit composition, recognition specificity, cleavage position, and cofactors. Among them, type II restriction enzymes recognize a specific site of 4, 6, or 8 base pair (bp) and cleave within the sequence. We can estimate the cutting frequency of a specific restriction enzyme recognition site in a genome based on the genome’s  guanine-cytosine (GC) content and the number of nucleotides covered by the recognition site. For example, assume a genome has a GC content of 50% (%GC = % adenine-thymine (AT), an equal % of four nucleotides). The estimated occurrence frequency of the restriction enzyme HindIII (A^AGCTT: recognition site, ^: cut site) is (1/4)(1/4)(1/4)(1/4)(1/4)(1/4) = 1/4096. The estimated average length of HindIII-cut DNA fragments is 1/(1/4096) = 4096 bp.

The recognition site is generally palindromic (the same sequence read on the two strands in reverse directions) (Figure 2-2). After cleaving the phosphodiester bonds to produce 5′ phosphate (Pi-) and 3′ hydroxy (-OH) in each strand of double-stranded DNA (dsDNA), the enzyme can produce a “blunt end,” or a 3′- or 5′- overhang of each strand called a “sticky end.” The blunt ends can anneal to digested DNA fragments with blunt ends, whereas sticky ends can only anneal to the other DNA fragments with complementary ends.

Restriction enzymes are essential in constructing recombinant DNA molecules, such as those used in restriction mapping of DNA molecules and gene cloning, as summarized in Figure 2-3, top.¹² Before starting, you need to choose your cloning backbone carefully. It needs to have compatible cut sites for restriction enzymes, allowing your insert to be placed into the backbone in the proper orientation. For example, if you want to clone a gene into an expression vector, you will want the start of the gene to be just downstream of the promoter in the backbone. Ideally, the backbone contains various restriction enzyme cut sites (restriction sites) downstream of the promoter as a so-called multiple cloning site (MCS), which allows you to properly orient your gene with the insert to express it. For example, the plasmid backbone contains a promoter (blue arrow) followed by three restriction sites: EcoRI, XhoI, and HindIII (Figure 2-3, bottom).¹² To place your insert in the proper orientation downstream of the promoter, you can cut the plasmid backbone and insert with EcoRI at the 5′ of the insert and HindIII at the 3′ end. After mixing the cut products from the vector and the DNA to be cloned (insert), the two EcoRI-digested ends and the two HindIII-digested ends anneal, respectively. As a result, the 5′ end of the insert is downstream of the promoter to place the gene in the proper orientation with respect to the promoter. After adding ligase to the mixture to covalently link the backbone and insert, a plasmid is ready to express the insert.

A few buffers are used for most restriction enzymes, but no single buffer provides optimal activity for all enzymes. Buffers are usually provided at 10X concentration. Since restriction enzymes require different buffer conditions, various strategies are used to set up a “double digest.” It is best to simultaneously digest both enzymes in a compatible “universal” buffer if available despite one enzyme not being fully active, e.g., < 100% active, because more units of one enzyme (1 U of enzyme A plus 1.33 U of enzyme B) can be added for equal cutting efficiency. Nevertheless, there are limits to adding excess enzymes due to increased glycerol, a cryoprotectant of proteins, reducing the specificity of some enzymes. Alternatively, it can be set up to digest with the “low salt” enzyme and then add more buffer or salt for the “high salt” enzyme to complete the digestion. In extreme cases, the DNA is precipitated with ethanol and salt after one digestion and then dissolved in the second buffer for the subsequent enzyme digestion. Most digestions are carried out at 37°C unless specified otherwise.

This lab exercise isolates plasmid DNA from an overnight bacterial culture, which is then prepared for outsourcing DNA sequencing after quantification. Meanwhile, the isolated plasmid and a known grape NBS-containing plasmid, pSCA7 (T1-T3-W6),⁸ are digested with EcoRI and PstI. In the next lab (Week 2), agarose gel electrophoresis will separate the restriction-digested DNA samples to verify the size of the DNA insert. The band corresponding to the NBS sequence from pSCA7(T1-T3-W6) will be excised from the gel, and the rest will be used for Southern transfer (Figure 2-1).

Procedure

A. Growing Overnight Bacterial Culture the night before the lab

1. Using one sterile micropipette tip, touch one white colony from your Amp/X-Gal plate, twirl it in one 50 ml conical tube containing 2 ml LB/Amp+ medium, and label the tube with your group number.
2. Repeat step 1 using a new sterile pipette tip with a known NBS-containing bacterial colony containing the pSCA7 (T1-T3-W6) provided by the instructor.
3. Label the tube “NBS and pSCA7 (T1-T3-W6)” with your group number.
4. Next morning, add 1 ml of new culture medium into each culture tube and put it back on the shaker.

B. Plasmid DNA Isolation (NEB Monarch Plasmid Miniprep Kit)

For this part of the experiment, each group will isolate a duplicate of their cloned plasmid DNA to ensure sufficient purified plasmid for the next experiment.

1. For the bacterial culture containing your cloned plasmid, pipette 1.2 ml (600 µl twice)  into each of two 1.5 ml centrifuge tubes and centrifuge both at the top speed for 30 seconds. Remove the supernatant, which will be collected and autoclaved before being discarded. (The plasmid isolation kit is designed to isolate plasmid from 1−5 ml, not to exceed 1.5 optical density (OD) units; 12−16 hours of growth is ideal.)
2. Repeat step 1 so that each 1.5 ml microcentrifuge tube contains a bacterial pellet from a total of 2.4 ml liquid culture.
3. Each pair of groups repeats steps 1 and 2 to obtain two 1.5 ml microcentrifuge tubes, each containing a bacterial pellet harboring the NBS-containing plasmid, pSCA7 (T1-T3-W6). After this step, each group should have three 1.5 microcentrifuge tubes, two containing bacterial pellets for isolating the group’s cloned plasmid DNA and one for isolating pSCA7 (T1-T3-W6) DNA.

The steps below are carried out for the pellet in each 1.5 ml microcentrifuge tube. 

4. Suspend each pellet in 200 µl of Plasmid Resuspension Buffer (B1). Vortex or pipet to ensure cells are completely resuspended without visible clumps. Change tip for resuspending each pellet.
5. Add 200 µl of Plasmid Lysis Buffer (B2), gently invert the tube 5-6 times, and incubate at room temperature for 1 minute. Do not vortex, which may nick strands of plasmid DNA due to the high pH of the alkaline lysis solution. The high pH solution lyses the bacteria and denatures both chromosomal and plasmid DNAs. The solution should become transparent, viscous, and dark pink.
6. Add 400 µl of Plasmid Neutralization Buffer (B3), gently invert the tube until neutralized, and incubate at room temperature for 1 minute. Do not vortex. The sample is neutralized when the color is uniformly yellow and precipitate forms.
7. Centrifuge lysate for 5 minutes. The pellet should be compact; spin longer if needed.
8. Carefully transfer the supernatant to the spin column and centrifuge for 1 minute. Discard the flow-through. This step binds the nucleic acid to the column and is critical in determining the purity of the plasmid obtained.
9. Re-insert the column in the collection tube and add 200 µl of Plasmid Wash Buffer 1.
10. Centrifuge for 1 minute and discard the flow-through.
11. Add 400 µl of Plasmid Wash Buffer 2 and centrifuge for 1 minute.
12. Transfer the column to a clean 1.5 ml microfuge tube. Use care to ensure the column tip does not come into contact with the flow-through. If in doubt, re-centrifuge the column for 1 minute after removing the filtrate.
13. Add 40 µl of DNA Elution Buffer to the center of the matrix. Wait for 1 minute, then centrifuge for 1 minute to elute DNA.

    Note to the Instructor: Nuclease-free water (pH 7–8.5) can also be used to elute the DNA. Although the yield may slightly increase if a larger volume of DNA Elution Buffer is used, the DNA is less concentrated. For larger DNA (≥ 10 kb), heating the elution buffer to 50°C before use can improve yield.

14. Measure OD ratios and concentrations of plasmid DNA samples, following the steps described in Procedure B, Analysis of Concentration and Purity of DNA using NanoDrop One UV-Vis Spec in Week 1 of Module 1, and fill in the results in the table below.

C. Prepare Plasmid Sample for DNA sequencing

1. Label a 0.5 ml PCR tube with your group number, T1, T2, T3, etc.
2. Dilute your DNA sample to ~150 ng/µl. A concentration of less than 150 ng/µl is rare. Still, if it happens, the DNA sample is concentrated by placing it in Savant™ SpeedVac™ DNA130 Integrated Vacuum Concentrator Systems to dry or by ethanol precipitation (see below) and then bringing it to 150 ng/µl. However, this is generally not an issue because at least one of the two plasmid DNA samples contains DNA concentration ≥ 150 ng/µl.
3. Add 20 µl of your diluted DNA sample to the 0.5 ml PCR tube according to the instructions provided by the commercial DNA sequencing lab.

The instructor collects all the prepared DNA samples and mails them out for sequencing.

Ethanol Precipitation

1. Measure the volume of the DNA sample.
2. Add 1/10 volume of 3M sodium acetate, pH 5.2 (final concentration of 0.3M).
3. Mix well.
4. Add 2 to 2.5 volumes of cold 100% ethanol (calculated after salt addition).
5. Mix well.
6. Place on ice or at -20°C for > 20 minutes.
7. Spin at maximum speed in a microcentrifuge for 15 min at 4°C.
8. Carefully remove supernatant.
9. Add 1 ml 70% ethanol. Mix. Spin at maximum speed in a microcentrifuge for 5 min at 4°C. Carefully remove supernatant.
10. Air dry, briefly vacuum dry, or spin briefly in a room-temperature centrifuge with the cap open to dry the pellet.
11. Resuspend pellet in less volume of water or elution buffer.

D. Restriction Endonuclease Digestion of DNA

Change the pipette tip for each solution/enzyme used.

1. Calculate the volume required to make up 0.5 µg of your isolated plasmid DNA samples and then calculate the volume of sterile water needed to add in each tube shown in Table 2-2.
2. Label four 1.5 ml microcentrifuge tubes on ice (make sure no ice gets into your tube). You are going to set up four 20 µl restriction digestion reactions (Tubes 1−4 in Table 2-2).
   1. Add 2 µl of the corresponding 10X reaction buffer into each tube.
   2. Add the 0.5 µg of DNA into each of Tubes 1−4 based on the calculation from step 1 and with additional H₂O to make the total volume of 14−18 µl indicated in Table 2-2.
   3. Add the restriction enzyme last.
3. Half of the groups will also prepare an additional digest in Tube 5 with 2 µg of pSCA7 (T1-T3-W6) to be used for probe labeling for the entire class.
4. Mix by pipetting up and down several times without making bubbles.
5. Incubate the tubes at 37°C for 1−2 hr.
6. Heat at 65°C for 20 minutes to inactivate the enzymes.
7. Add 5X loading buffer and store at -20°C for agarose gel electrophoresis to isolate DNA insert for labeling (Tube 5) and to set up Southern blot (Tubes 1−4) for the next lab exercise.

Data Analysis and Discussion

1. You make a genomic library to clone a 3 kb gene. The organism’s genome size is 1 x 10¹² base pair, and its GC content is 50%.
   1. To ensure successful cloning of the gene (keeping the coding sequence intact), it is necessary to use a proper restriction enzyme to cut the genomic DNA before making the library. Will you use a restriction enzyme with 3, 4, 5, or 6 nucleotides as its recognition site? Explain.
   2. How many colonies do you need to screen to ensure you cover the organism’s entire genome to get the clone? Show your calculation.
2. Draw a restriction map of linear DNA using the following data. Indicate the number of base pairs between every two restriction sites on the linear DNA.
   

   DNA	Sizes of Fragments (bp)
   uncut DNA	600
   DNA cut with EcoRI	400, 200
   DNA cut with HindIII	450, 150
   DNA cut with BamHI	300, 250, 50
   DNA cut with EcoRI + HindIII	400, 150, 50
   DNA cut with EcoRI + BamHI	300, 150, 100, 50
   DNA cut with HindIII + BamHI	300, 150, 100, 50

3. Recognition sites for two restriction enzymes are XbaI: 5′-T^CTAGA-3′ and NheI: 5′-G^CTAGC-3′. You digest a DNA sample with these two enzymes separately and then mix the digested products with DNA ligase and ATP.
   1. Like those shown in Figure 2-2, show the two cut products made by each enzyme (also indicate the Pi- and -OH groups on each), using different colors for the four cut products.
   2. Similarly, show the possible ligated products, including at the ligation site (Give four but exclude the ligate back original DNAs). Use the same colors for the ligated products as in answer (a), which should be dsDNAs with 5′ and 3′ ends indicated on both strands.