Week 6 HW: Genetic Circuits Part I: Assembly Technologies
DNA Assembly Part
1. What are some components in the Phusion High-Fidelity PCR Master Mix and what is their purpose?
The Phusion High-Fidelity PCR Master Mix key components include:
Phusion DNA Polymerase: This is a fusion protein comprising a traditional Pfu-like DNA polymerase and a processivity-enhancing domain. Its purpose is to synthesize new DNA strands with exceptionally high fidelity (due to its inherent 3’→5’ proofreading exonuclease activity) and high speed.
Deoxynucleotide Triphosphates (dNTPs): These are the blocks that the polymerase uses as substrates to construct the new complementary DNA strands.
Reaction Buffer: The buffer maintains the optimal pH for enzyme activity. The magnesium ions in it act as an essential cofactor for polymerase function and influence primer annealing by stabilizing the interaction between the primer and the DNA template.
Stabilizers and Enhancers: These proprietary compounds (sugars or detergents) help maintain enzyme stability during thermal cycling and can aid in amplifying difficult templates by reducing secondary structures or preventing non-specific primer binding.
2. What are some factors that determine primer annealing temperature during PCR?
Melting Temperature: It is the temperature at which half of the DNA duplex dissociates. It is calculated based on the primer’s nucleotide composition, length, and concentration.
Primer Length and GC Content: Longer primers and those with a higher proportion of guanine (G) and cytosine (C) bases (which form three hydrogen bonds) have higher Tm values, thus generally requiring higher annealing temperatures.
Salt Concentration: The ionic strength of the reaction buffer significantly impacts Tm. Higher salt concentrations shield the negative charges of the DNA backbone, stabilizing the duplex and raising the Tm.
Primer-Template Complementarity: Perfectly matched primers will anneal at a higher, more predictable temperature. Mismatches, especially at the 3’ end, will destabilize binding and necessitate a lower Ta, though this is often avoided to maintain specificity.
3. There are two methods from this class that create linear fragments of DNA: PCR, and restriction enzyme digests. Compare and contrast these two methods, both in terms of protocol as well as when one may be preferable to use over the other.
Protocol Comparison
Source: PCR amplifies a specific sequence from a small amount of template DNA (e.g., genomic DNA, plasmid, or cDNA), exponentially increasing its copy number. Restriction digestion starts with a large, purified DNA molecule (like a plasmid or a PCR product) and cuts it at specific recognition sites, producing a finite number of fragments without amplification.
Mechanism: PCR uses thermal cycling and a DNA polymerase to synthesize new DNA. Restriction digestion is an incubation at a constant temperature where endonucleases hydrolyze the phosphodiester backbone at specific palindromic sequences.
Product: PCR products are defined by the primers used, resulting in fragments with specific, known ends (which can be blunt or, if designed with overhangs, effectively sticky after processing). Restriction digestion yields fragments whose ends are dictated by the location and type of restriction site (blunt or sticky overhangs).
Preferential Usage
PCR is preferable when: The goal is to amplify a gene from a genome or cDNA for cloning, to introduce specific mutations or sequences (like restriction sites or tags) via the primers, or when the starting template is scarce. It is also the method of choice for diagnostic screening (colony PCR).
Restriction Digest is preferable when: The goal is to prepare a vector (plasmid backbone) for cloning by creating compatible ends, to sub-clone a fragment from an existing plasmid, to analyze the size or orientation of an insert (diagnostic digest), or when the source DNA is already abundant and contains the fragment of interest within convenient restriction sites.
4. How can you ensure that the DNA sequences that you have digested and PCR-ed will be appropriate for Gibson cloning?
Ensure that the ends of your linear fragments share complementary sequences. Typically, overlaps of 20–40 base pairs (bp) are designed. The PCR primers used to amplify an insert must have a 5’ extension that is complementary to the end of the adjacent fragment (e.g., the vector). Therefore, the PCR product itself will have these overhangs built into its ends.
The DNA fragments must have clean, blunt ends or ends compatible with the assembly mechanism. Gibson Assembly relies on the 3’→5’ exonuclease activity of the T5 exonuclease to chew back one strand, creating single-stranded 3’ overhangs that allow complementary overlaps to anneal. Therefore, your fragments must be free of damaged ends or unusual secondary structures at the termini.
Following PCR or restriction digest, the DNA must be purified (via gel extraction or column cleanup) to remove enzymes, primers, dNTPs, and buffer components. Residual polymerase or restriction enzymes could interfere with the Gibson Assembly master mix’s enzymes (exonuclease, polymerase, ligase).
While not part of the physical prep, in silico validation is crucial. Use software to confirm that the designed overlaps are unique and have a suitable $T_m$ (~50°C, the Gibson reaction temperature) to ensure proper annealing during the 50°C incubation step.
5. How does the plasmid DNA enter the E. coli cells during transformation?
Chemical Transformation (Heat Shock): Cells are treated with ice-cold $CaCl_2$ (or other divalent cations), which makes the cell membrane more permeable by neutralizing the repulsion between the negatively charged DNA backbone and the negatively charged lipopolysaccharides on the cell surface. The cations are thought to create patches of positive charge and induce structural changes in the membrane. A brief heat shock (usually 42°C) creates a thermal gradient that is believed to either create pores in the membrane or trigger an influx of buffer into the periplasm, effectively dragging the DNA associated with the membrane into the cell.
Electroporation: This method uses a high-voltage electrical pulse to temporarily destabilize the phospholipid bilayer. The electrical current creates transient pores (electropores) in the cell membrane through which the negatively charged DNA is driven by the electrical potential. Once the pulse subsides, the membrane pores reseal, trapping the plasmid inside.
- Describe another assembly method in detail (such as Golden Gate Assembly)
- Explain the other method in 5 - 7 sentences plus diagrams (either handmade or online).
Golden Gate Assembly is a powerful, seamless DNA cloning method that utilizes Type IIS restriction enzymes to digest and ligate DNA fragments in a one-tube reaction. Unlike traditional enzymes, Type IIS enzymes (such as BsaI or BsmBI) cut outside their recognition sequence, generating short, unique, single-stranded overhangs of 4 base pairs. Because the recognition site is removed from the final assembly junction, the reaction can be programmed to cut, ligate, and re-cut until the correct, seamless product is formed.
In a typical protocol, DNA fragments (PCR products or vectors) are designed with flanking Type IIS sites that generate complementary overhangs. These fragments, along with the destination vector, are mixed in a single tube with the Type IIS enzyme and T4 DNA Ligase. The reaction is incubated in a thermal cycler, alternating between the optimal temperature for the enzyme (e.g., 37°C) and for the ligase (e.g., 16°C). During the digestion phase, fragments are released with specific sticky ends. If they are designed to ligate together correctly, they form a stable assembly that lacks the original enzyme recognition sites. Incorrect assemblies or uncut fragments retain the recognition sites and are subject to re-digestion. This “cut-ligate” cycling drives the reaction towards the desired final construct with very high efficiency, making it ideal for modular cloning (MoClo) and assembling multiple parts simultaneously.