Subcloning Notes

Learning Objectives

• Differentiate between:

  • Restriction enzyme cloning: Traditional method using restriction enzymes to cut and ligate DNA fragments into a vector.

  • PCR-mediated restriction enzyme cloning: Involves using PCR to amplify a DNA fragment, then using restriction enzymes for cloning into a vector.

  • TOPO-TA cloning: A rapid cloning technique that utilizes topoisomerase I to directly ligate PCR products into a vector.

• Describe the mechanism of ligation with T4 DNA ligase as it relates to cloning: T4 DNA ligase catalyzes the formation of a phosphodiester bond between the 3'-OH and 5'-phosphate ends of DNA, requiring ATP as a cofactor.

• Describe the mechanism of ligation with topoisomerase as it relates to TOPO-TA cloning: Topoisomerase I catalyzes the transfer of the DNA strand to the enzyme, allowing for the insertion of PCR products directly into the vector, effectively joining them without the need for additional ligation steps.

• Understand the difference between directional cloning and non-directional cloning:

  • Directional cloning: Inserts a DNA fragment into a vector in a specific orientation using two different restriction enzymes.

  • Non-directional cloning: Inserts a DNA fragment into a vector in either orientation, often using a single restriction enzyme or blunt-end ligation.

• Describe how complementation enables blue/white screening of transformants: Blue/white screening uses the lacZ gene to identify recombinant colonies. Disruption of the lacZ gene (by insertion of a DNA fragment) results in white colonies, while functional lacZ leads to blue colonies in the presence of X-gal and IPTG.

Cloning Methods

• Cloning Solely with Restriction Enzymes

• Cloning with PCR and Restriction Enzymes

• Cloning with TOPO-TA

T4 DNA Ligase

• T4 DNA ligase catalyzes a phosphodiester bond between 5’ phosphate and 3’ hydroxyl groups. Requires ATP as a cofactor. This enzyme is commonly used to ligate DNA fragments with compatible ends.

Vector Backbone and Insert

• Draw out the polarity and location of phosphate groups and hydroxyl groups on a vector backbone and insert. Understanding the position of these chemical groups is crucial for successful ligation.

Ligase-Catalyzed Reactions

• How many ligase-catalyzed reactions are required to ligate a single insert into a single vector? Two ligase-catalyzed reactions are required.

• What if the vector was treated with a phosphatase prior to ligation? Treatment with a phosphatase removes the 5’ phosphate groups, preventing self-ligation of the vector.

Topoisomerase

• Topoisomerase creates single-strand breaks and re-ligates the breaks back together. This enzyme is used for TOPO cloning.

PCR4 TOPO Vector

• Is the PCR4 TOPO vector used in lab this week circular or linear? The PCR4 TOPO vector is linear.

TOPO Cloning

• If your forward primer has a 5’ phosphate group and your reverse primer does not have a 5’ phosphate group, would you expect TOPO cloning to work? TOPO cloning does not require phosphate groups on the primers. It relies on the single 3' deoxyadenosine (A) overhangs for ligation.

Digestion and Ligation Products

• Diagram out the products you would get if you digested the two plasmids below with BamHI and mixed the digests together in the presence of T4 DNA ligase. One plasmid contains GFP and Ampicillin resistance. The other plasmid contains Cherry and Kanamycin resistance. Both plasmids have BamHI sites. Products would include:

  • GFP-Ampicillin plasmid recircularized

  • Cherry-Kanamycin plasmid recircularized

  • Linear concatemers of both plasmids

• Diagram out the products you would get if you digested the two plasmids below with BamHI and EcoRI and mixed the digests together in the presence of T4 DNA ligase. One plasmid contains GFP and Ampicillin resistance. The other plasmid contains Cherry and Kanamycin resistance. Both plasmids have BamHI and EcoRI sites. Products would include:

  • GFP-Ampicillin plasmid recircularized

  • Cherry-Kanamycin plasmid recircularized

  • Hybrid plasmids with GFP-Ampicillin fragments ligated to Cherry-Kanamycin fragments

TOPO Vector and Lac Repressor

• A TOPO vector that does not contain a Lac repressor binding site would have the ability to produce blue colonies regardless of whether IPTG was present. This is because the lacZ gene would be constitutively expressed.

SUBCLONING (TAKING A DNA FRAGMENT FROM ONE VECTOR AND INSERTING IT INTO ANOTHER)

• A typical procedure in a molecular biology lab involves moving an insert, or a defined portion of an insert, from one vector to another. There are lots of reasons for doing this. If the insert is a cDNA, it might have arrived in your lab cloned into a very minimal vector, designed specifically to maintain it in a bacterial stock. But perhaps you want to generate RNA or protein from this insert - so you need to move the cDNA into a different vector that possesses suitable promoter sequences for expression. Or perhaps, and this is very common, you want to cut out a selected piece of the cDNA, a coding sequence for a DNA binding domain, say, and fuse it to another coding sequence like that of the green fluorescent protein (GFP) to enable in situ localization experiments. Very typically, you need to put the cDNA into a specialized vector that allows you to make a transgenic animal or plant - these vectors are pretty unique to each area of research, and so subcloning is almost inevitable in any area of molecular biology.

• There are a great many protocols and kits that are used in subcloning, but the first step is to purify the insert away from the vector it came in. The insert can be separated from the vector by restriction digestion, or it could be amplified out of the insert by PCR. If necessary, you can purify the insert DNA using a kit.

• The next stage in subcloning is essentially the same as cloning you need to place your insert into the multiple cloning site of the new vector using a ligase enzyme (usually either T4 ligase or topoisomerase), and transform the new construct into bacteria for propagation ("cloning"). Again, there has been much diversification of protocol, and we are going to employ a method that will introduce you to the important parameters that are shared by many cloning methodologies. We will be describing these extensively in lecture, but to summarize, there are two main cloning techniques that are in wide use today.

1. Traditional, or classical cloning. Don't be fooled by the old fashioned name! Molecular biology labs often perform this kind of cloning, in which the same restriction enzymes are used to cut both the insert and the vector. Then, in a subsequent reaction, we use ligase to join them together. The simplest form of traditional cloning involves cutting the insert and the vector with the same restriction enzyme and mixing them in defined proportions to maximize the likelihood that they will come together in a ligation reaction. There are several problems with this method, however. First, you have to find a way to prevent linearized vector cut with a single enzyme from recircularizing. Because the recircularized vector is smaller than the insert plus vector would be, it will transform into bacteria more efficiently, reducing your odds of obtaining a successful ligation. You can solve this problem by treating the linearized vector with a phosphatase (typically calf intestinal phosphatase (CIP) or shrimp alkaline phosphatase (SAP) are employed). A phosphatase removes the 5' phosphates necessary for ligation. The resulting ligated product thus contains single stranded "nicks" on the 3' ends of each strand of the insert. These nicks are repaired by enzymes endogenous to E. coli, after transformation.

   • What would happen if you dephosphorylated both your insert and your vector prior to ligation and transformation?

   • Another problem with using a single enzyme in traditional cloning is orientation - if the insert and vector are cut with the same enzyme, there is no way to control which way the insert goes into the vector both orientations are typically equally possible. (It should be noted that sometimes one orientation is favored over another, for reasons which may be due to inadvertent expression of a toxic product or the appearance of a cryptic promoter sequence that causes the bacteria containing the construct with a specific orientation to die.) You can solve this problem by employing a version of traditional cloning called "directional cloning," in which the insert and vector each are cut with two different restriction enzymes that leave cohesive ("sticky") ends. The incompatibility of the sticky ends prevents the vector from recircularizing and forces a particular orientation in the ligation reaction. The difficulties associated with this methodology are largely strategic: do you happen to have the right combination in the right 5' to 3' order - of sticky-end restriction sites in both your insert and vector? Again, a work-around exists - it is always possible to add restriction sites to your inserts by using PCR, or by ligating linkers that contain restriction sites to your insert.

   • Finally, it is possible to use enzymes that do not leave complementary overhangs to clone. This methodology is called blunt cloning, and is much less efficient, and will be discussed in class as an option.

2. PCR-based cloning. Taq polymerase has revolutionized molecular biology as a robust way of amplifying DNA. But it has another trick that turns out to be highly useful. Taq possesses a terminal transferase activity that adds single adenines to the 3' ends of a substantial proportion of the amplified product. This single 3' A overhang is sufficient to generate a sticky-end strategy for cloning into vectors that possess a single 3' overhanging thymine. The method is often called TA cloning for this reason. Because the overhanging T's are incompatible, there is no danger of the vector recircularizing, BUT do you think we can solve the insert orientation problem we described earlier?

   • TA cloning can be carried out using T4 ligase the same ligase used in traditional cloning. There are also kits available that use topoisomerase instead of T4 ligase. Recall that topoisomerase can cut and re-ligate DNA molecules in order to relieve torsional stress in DNA molecules that are undergoing replication. In kits that employ topoisomerase, the enzyme is covalently linked to the open vector, and ligation is achieved by releasing the enzyme. The reaction is very fast and efficient. We will in fact be using topoisomerase to subclone.

LIGATION THEORY

• We will be using a TA system to subclone our gene fragments. You will need to understand how the ligation works and calculate what ingredients you will need based on your yield of PCR-derived insert.

• Like everything in molecular biology, your actual protocol will arise from much troubleshooting and tweaking. In order to optimize ligations, you need to know the length and concentration of the insert and vector, whether the ends are cohesive or blunt, and the flexibility of the molecules (to some extent sequence dependent) among other things. But here are some basic guidelines (we will cover their derivation in more detail in lecture).

1. Determine the concentration of the vector and the insert. TA vector concentrations are known, because it comes as part of a kit. Quantifying the insert can be carried out by using a spectrophotometer, or image analysis relative to a commercial molecular weight ladder with known fragment concentrations for a given volume.

2. Calculate the amount of insert and vector required for the most efficient ligation. Ligation volumes are usually quite small, to keep the molecules involved fairly concentrated.

3. The actual calculation for an optimal amount of insert relative to vector requires that you consider molar ratios rather than absolute concentrations. An insert of 1 kb mixed with a vector of 3 kb would exist in a 3:1 molar ratio of vector to insert if the concentrations of DNA were identical. Generally you want more moles of insert than moles of vector because usually the insert is smaller than the vector (though not always) and ideally you would like to favor intermolecular ligation (insert + vector) over no ligation (ratio of insert to vector is too low) or over concatemerization of the insert (ratio of insert to vector is too high). Typically, for inserts that are 1-3 kb cloned into vectors that are 3-5 kb, a 3:1 insert to vector ratio is a good place to start. But you may have to go up or down depending on your experience with the particular sequences involved. Also, inherently inefficient ligations like blunt ligations require a much higher insert to vector ratio (10:1 is not unusual), to maximize the likelihood of a linkage holding long enough for the ligation to proceed.

4. Determine the temperature and duration for the ligation. Annealing of cohesive ends is most efficient at lower temperatures, and so sticky-end ligations using T4 ligase are typically performed anywhere from 4 °C to 16°C. This is not the best temperature for the T4 ligase, however, and if the ligation is inefficient, like a blunt ligation, the temperature may be raised, usually to no higher than room temperature (about 25°C).

   • Like almost everything nowadays, it is possible to find "ligation calculators" on the web, where you simply plug in your concentrations, guess at the ratio, and an algorithm provides you with a complete protocol. But it is critical you understand how this algorithm works, and its derivation. We will discuss this further in class, but here is the formula that is most widely used to determine an optimal ligation. Since you usually know the sizes and concentrations of your insert and vector, all you need to work out is how much of your insert to use in the reaction. This is the most commonly used formula:

(ng \text{ of vector } \times \text{ kb size of insert) /kb size of vector} \newline \times \text{ insert:vector molar ratio ng of insert }

EXAMPLE

• Suppose I wanted to set up a ligation where I had 140 ng of vector (10 kb) but a 1:3 molar ratio of vector:insert. My insert is 2.5 kb in length. How much (ng) insert DNA would I use in my ligation reaction?

(\frac{140 \text{ ng } \times 2.5 \text{ kb}}{10 \text{ kb}}) \times 3 = 105 \text{ ng of insert}

Sometimes, you don't know the amount of vector that you should add and have to figure it out yourself. In that case, here are some steps to help you.

1. Figure out the concentration of your vector and insert DNA, in ng/µL. That may require you to perform spectrophotometry on your DNA. Remember that an absorbance at 260 nm of 1 is equal to a DNA concentration of 50 ng/µL. Or, you may run your DNA on a gel and compare the brightness of your band to the molecular weight standards to estimate its mass and concentration. If you have made any dilution of your DNA prior to measuring its concentration, make sure to take into account the dilution factor.

2. Divide the size of the insert (in Kb) / the size of the vector (in Kb). Multiply that result by the desired moles of insert/ desired moles of vector. (For a sticky-ended ligation, that's 3 moles insert / 1 mole vector; for a blunt-ended ligation, that/s 10 moles insert/1 mole vector.) That will give you a factor to use below. Let's call this factor Y. Y tells you how many times (more/less) ng of insert to use than ng of vector. At this point, we are working in mass, not in concentration.

3. Usually, we use a total of 250 ng in a ligation. Y(x) + (1x) = 250 ng, so (Y+1)x = 250 ng. Solve for x. The mass (in ng) of vector you wish to use is x. The mass (in ng) of insert that you wish to use is Y(x).

   • Go back to your DNA concentrations, and divide the mass of insert or vector by the concentration in ng/ µL. That should tell you how many µL of your DNA solutions to add to your ligation reaction.

   • While vector:insert ratios should be optimized for an efficient cloning using the enzyme DNA ligase, the biotechnology industry has devised kits that make for quick, easy screening of the recombinant plasmids. PCR4-TOPO is such a plasmid and relies on:

     • a) 3' terminal thymidines on both strands of the vector to prevent self-ligation,

     • b) a vast excess of insert (PCR amplicon).

   • Any self-ligation is detected by blue-white screening.

BLUE-WHITE SCREENING

• Blue-white screening is a common feature of cloning vectors that allows for the rapid (and convenient) detection of recombinant bacteria after a molecular cloning experiment. There are two important components that need to be considered when using blue-white screening: a) the genotype of your E. coli strain and b) the MCS of your cloning vector.

• Bacterial Genotype: ẞ-galactosidase is an enzyme encoded by the lacz gene in the lac operon of many enteric bacteria (like E. coli). Normally, the lac operon is tightly regulated by the Lacl repressor, which binds to the operator sequence within the lac operon promoter sequence. This ensures that the metabolic enzymes (including ẞ-galactosidase) contained within the lac operon are not expressed unless glucose levels are low and lactose levels are high. In the lab, you can induce expression of the lac operon in E. coli by adding Isopropyl ẞ-D-1-thiogalactopyranoside (IPTG) to the culture media. ẞ-galactosidase is a homotetramer in its active state, however, many strains of E. coli have a gene mutation that results in an N-terminal deletion at residues 11-41 of the B- galactosidase protein sequence. This deletion results in an inactive ẞ-galactosidase (the Ω fragment), which is only compensated when the N-terminal fragment of the enzyme (the a fragment) is provided in trans.

• Cloning Vector: Many cloning vectors (including pCR4-TOPO) posses the lacza gene sequence, which overlaps the MCS. Assuming that there are no interruptions to the MCS, any bacteria that are transformed with these types of cloning vectors will express the a fragment of ẞ- galactosidase. As with the & fragment of ẞ-galactosidase, the a fragment of ẞ-galactosidase is inactive on its own.

• Blue-White Screening: In order to utilize blue-white screening, you will need to ensure that your E. coli strain contains the modified lacz gene (i.e. only expresses the Ω fragment of ẞ- galactosidase) and that your cloning vector contains a modified lacZ gene (i.e. only expresses the a fragment of ẞ-galactosidase). If you were to transform your E. coli with an empty cloning vector (like pCR4-TOPO) then both the a and 2 fragments of β-galactosidase will be expressed, forming a functional ẞ-galactosidase enzyme. However, if you clone an insert (generated after PCR and/or enzyme digestion) into the MCS of a cloning vector, you will disrupt the LacZa gene in that cloning vector. This means that your transformed bacteria will only express the fragment of ẞ- galactosidase (which is inactive by itself). You can screen for the subsequent ẞ-galactosidase activity of your transformants by using a lactose analog called, X-gal. When X-gal is present in the culture media, it will be cleaved by active ẞ-galactosidase enzymes to form 5-bromo-4-chloro- indoxyl. This molecule then spontaneously dimerizes and oxidizes to form a bright blue insoluble pigment (5,5'-dibromo-4,4'-dichloro-indigo). This insoluble blue pigment turns the E. coli blue and indicates that these cells were transformed with a cloning vector that had an intact MCS, which resulted in the formation of a fully functional ẞ-galactosidase enzyme. White bacterial colonies result when the MCS in your cloning vector has been disrupted by your insert during your cloning experiment.

• When using blue-white screening, why do you want to see white colonies after ligation/transformation?