Common tools I & II

Question 1

What is cloning?

Answer 1

Cloning is the creation of identical copies of organisms, cells (or DNA fragments).

Question 2

What is molecular cloning?

Answer 2

Molecular cloning usually means “to ligate a piece of DNA into a plasmid or vector”.

Question 3

Provide two examples of reasons to clone DNA.

Answer 3

1. Produce a lot of one protein: for example insulin which is hard to extract from natural sources.
2. Mutate or tag protein: For example to fuse the coding region to something that can be detected later.
3. Create transgenic organisms: for example inserting genes for antibiotic resistance into a strain to be able to select for it later.

Question 4

In cloning, vectors are often used. What are vectors?

Answer 4

A vector is a carrier of whatever is cloned, usually used to transfer it into a host.

Question 5

Name two common types of vectors and what type of host they’re usually used for.

Answer 5

Common vectors are plasmids or viruses. Plasmids are commonly used to transform bacteria and viruses are typically used to transfect eukaryotic (or only mammalian?) cells.

Question 6

Explain what three characteristics are important for a good vector and why.

Answer 6

The requirements of a good vector is:
1. The ability to replicate: contains an origin of replication/initiation sequence.
2. Selection: for example containing antibiotic resistance or autotrophy genes (a way to know which hosts have taken up the vector as transformation/transduction is far from 100% effective).
3. Small size: less important but the smaller the size the higher efficiency of transformation/transduction.

Question 7

Explain how cloning into a E. Coli host works in short.

Answer 7

You take DNA from any organism, ligate it into a plasmid (containing AB resistance for example), transform it into E. Coli and let the transformed E.coli incubate on medium containing AB to select for successfully transformed bacteria –> many clones containing the DNA of choice!

Question 8

When inserting DNA into a plasmid, the plasmid needs to be cleaved. Name two types of enzymes used for this and how they work.

Answer 8

Nucleases and Phosphatases.

• Nucleases are enzymes that degrade nucleic acids, the opposite function of polymerases. They hydrolyze, or break, an ester bond in a phosphodiester linkage between adjacent nucleotides in a polynucleotide chain. Nucleases can be specific for DNA, as DNases, or RNA, as RNases, or even be specific for a DNA/RNA hybrid as RNaseH (which cleaves the RNA strand of a hybrid duplex).
• Phosphatases are: An enzyme that can break a phosphomonoester bond, cleaving a terminal phosphate. They can be very useful before ligation to remove a phosphate to enable ligation.

Question 9

There are two groups of nucleases, which groups? What is their respective mode of action?

Answer 9

Endo- and Exonucleases.

• Endonucleases cleave inside of the DNA molecule, can be non specific or recognize specific sequences –> restriction endonucleases.
• Exonucleases cleave in free ends of the DNA molecule, hydrolyzing the polynucleotide chain either in a 5’-3’ direction or in a 3’-5’ direction.

Question 10

What is a restriction endonuclease and which type is the most commonly used one?

Answer 10

Restriction endonucleases are endonucleases that recognize a specific sequence and cleave only there (very specific). Usually derived from bacteria or archaea.

The most commonly used ones are type II restriction endonucleases which recognize palindromic sequences (seq. that is the same in the other direction but on opposite strand, such as -GATATC-) of around 4-8 bp that produce blunt ends (or staggered ends overhang or sticky). The type II restriction enzymes have the same recognition site and cleavage site, which the other types don’t, which make them easy to use.

Question 11

In what way does restriction enzyme cleavage leave the sugar-phosphate backbone ends?

Answer 11

Restriction endonucleases produce 5’P and 3’OH ends.

Question 12

What does “sticky” vs “blunt” ends of the cleavage site mean?

Answer 12

• Sticky ends result from a staggered cut, leaving an overhang of a few nucleotides on one of the strands, 3’ end or 5’ end. This is more common and leads to more efficient ligation bc of base pairing, but all ends are not compatible.
• Blunt ends means there’s no overhang. All blunt ends are compatible but the ligation is less efficient.

It’s important to know which type of ends a restriction enzyme produces because of the pros and cons with each type.

Question 13

So called “new generation restriction enzymes” are faster, more efficient and commercially available. One of the pros with these are that there is “no” star activity, what does this mean?

Answer 13

Star activity is when cleavage occurs in other sites than where it “should” cleave based on sequence specificity. If these are left too long some star activity can still happen. To get complete cleavage you might need to incubate longer, but that can risk star activity (balance needed).

Question 14

Subcloning was the first cloning made. Explain subcloning in short.

Answer 14

Subcloning is used to move a small part of donor DNA into a vector. First, the GOI (gene of interest)/small DNA fragment is cleaved with restriction enzymes and separated (usually by gel purification) to only get the GOI. The same restriction enzymes are then used to cleave the recipient plasmid to ensure compatible ends. (this is usually done in the “multiple cloning site” of the vector). Then the two fragments are ligated (with DNA ligase) and the result is a recipient plasmid containing the GOI.

Question 15

A common problem in subcloning is re-circularization of the vector as it has compatible ends, how is this usually solved?

Answer 15

By dephosphorylating the 5’-P ends produced by restriction enzymes, which makes the ends incompatible. Because this is not done to the donor DNA, its phosphorylated ends can attack and ligate properly.

Question 16

When transforming E. Coli cells, they need to be “competent”. What does this mean and how is it done?

Answer 16

Making cells competent means that we make them susceptible to receive/take up the plasmid. E. Coli usually do not take up DNA from environment.

The two most common ways of doing this is by heat shock (easier but less efficient) or electroporation (harder but more efficient), both opening up pores in the membrane to force DNA in to the cell.

Question 17

After plating transformed host on selective media, and re-streaking, describe how you get to a lot of cloned gene of interest.

Answer 17

Since the colonies that grow on selective media can contain the plasmid but without the insert:

1. you can either:
   - digest the hosts with same restriction enzymes and perform gel electrophoresis to see band of expected length.
   - digest with one restriction enzyme and sequence.
2. Once you have identified positive clones, perform PCR ro amplify product.

Question 18

Explain PCR in short.

Answer 18

1. Denaturation: from dsDNA to 2 ssDNA (first cycle longer time in this step)
2. Annealing: Primers binding to either ssDNA
3. Extension: DNA polymerase taking ssDNA to dsDNA –> double the amount of DNA (final extension in the end that everyone do but not sure if necessary)

Then cycle starts over and going for about 30 cycles, resulting in an exponential increase in DNA!

Question 19

What six components do you need in the mix to perform a PCR?

Answer 19

• Template DNA
• Primers (oligonucleotides)
• dNTPs
• Salts (in buffer)
• Mg2+ (usually already in buffer)
• Heat-stable DNA polymerase

Question 20

When designing primers, it’s important to keep track of the melting temperature (Tm) of the primer, why?

Answer 20

The annealing temp of the PCR should be set to be as close to the Tm as possible for an effective reaction.

Question 21

What four factors can influence the Tm of primers and how?

Answer 21

• Mg2+ concentration
• Salt concentration:
• Additives (e.g. DMSO): lower the melting temp
• GC content: more GC –> more hydrogen bonds –> hinger Tm

Question 22

What five factors are important when designing primers?

Answer 22

• Primer length: usually 18-25 nt
• GC content: 30-60% - avoid long stretches (>3 nt) of GCs
• Make sure your primer pair have matching Tm and GC content
• Avoid repeats and hairpin formation (also primer dimers)
• Specificity: BLAST your primers! Not good if the primers can bind to multiple sites leading to the formation of products similar to the one you want.

Question 23

Name three factors that can affect the success of your PCR.

Answer 23

• Primers: everything with primer design matters.
• Impurities in template (e.g. colony PCR, dirty genomic preps) –> often less template is more!
• EDTA in your template buffer (TE buffer) – chelates Mg2+
• Too much dNTPs (or DNA) can soak up Mg2+
• Optimization usually includes varying Ta,
  sometimes Mg2+ concentration, or simply ordering new primers
• Positive and negative controls crucial!

Question 24

What is the most important quality of the DNA polymerase used in a PCR?

Answer 24

That it’s thermostable! If not, it will degrade during the cycle and not able to perform multiple cycles of the PCR. The most commonly used one is Taq polymerase but there are other ones used to, like proof reading DNA polymerases with lower error rate but more sensitive than Taq.

Question 25

Name three applications for using PCR.

Answer 25

• PCR based cloning
• sequence analysis
• forensics
• disease diagnostics
• RT-PCR
• qPCR

And much more!

Question 26

What is reverse transcription (RT)-PCR and what is it used for?

Answer 26

RT-PCR is using PCR amplification using RNA (less stable) as template get amplified cDNA (more stable). It’s used to investigate the functions of RNA. Often, the RNA in question is in a limited amount or expressed in low abundance, and RT-PCR is solving that issue.

Common applications of RT-PCR include detection of expressed genes, examination of transcript variants (diversity), and generation of cDNA templates for cloning and sequencing - which is using DNA and not RNA usually.

Question 27

Explain how RT-PCR works in short.

Answer 27

In RT-PCR you use DNA primers that anneal to RNA and regular deoxynucleotides, and let reverse transcriptase (RTase) synthesize a complementary DNA (cDNA) which creates a RNA:cDNA hybrid, which is then denatured and the cDNA can be used as template in a regular PCR to amplify the number of cDNA copies.

Question 28

The primers used in RT-PCR are DNA oligonucleotides, what four types of primers can be used?

Answer 28

• Gene specific primers: primers complementary to gene. Useful when you already know gene sequence.
• Standard: a 3’-TTTTTTT-5’seq. binding to the poly(A) tail. Good when you want all to amplify all mRNA in a cell.

When you have no or too little info about mRNA:

• Anchored: a 3’-VTTTTTTT-5’ seq. with an addition of one nucleotide V= a mix of G, A or C.
• Random primers: hexamers (6 dTs) in random order. Shouldn’t work but does!

Question 29

What is qPCR and what is it used for?

Answer 29

Quantitative real-time PCR is a method for measuring relative DNA abundance during the exponential phase of PCR using florescence based detection.

Question 30

What is RT-qPCR and what is it used for?

Answer 30

RT-qPCR is the same as qPCR but measures relative RNA abundance instead of DNA.

Question 31

Explain how qPCR with the most common detection system, CYBR Green, works in short.

Answer 31

CYBR green binds to dsDNA, its florescent. During denaturation –> low/no florescence, during extension and after amplification more florescence. So by measuring the florescence, you can quantify the amount of DNA!

Question 32

When performing qPCR, the output is a cycle threshold (Ct) value, what does this value mean and what information does it give you?

Answer 32

The Ct value represents the number of cycles needed before the sample’s reaction crosses a florescence threshold, indicating the detection of the target amount of sample DNA (or RNA). A lower Ct value means that you have a higher amount of starting template (higher expression) and higher Ct means you have lower amount of starting template (less expressed) or that you have issues if it’s too high. Needs to be checked against reference values though.

Question 33

RT-PCR and RT-qPCR can be one-step or two-step reactions, what is the differences and applications?

Answer 33

one-step = one tube for both RT and PCR: faster, less pipetting needed, minimizes contamination risk (and improves reproducibility).

Two-step = separate tubes for RT and PCR reaction: good when you want to save the cDNA for future use or when you want to analyze multiple targets from sample RNA in separate reactions.

Question 34

Before processing DNA and RNA further, you often need to purify the sample. Explain the three major steps in DNA/RNA purification from intact cells.

Answer 34

1. Break open cells by lysis: can be mechanical or chemical (chaotropic salts, lysozyme, proteinase K or with detergents for genomic RNA/DNA, and alkaline for plasmid DNA). How you lyse depends on organism, plant and yeast cells have cell walls, so more “rough” methods are needed, like mortar and pestle.
2. Enrich for desired nucleic acid (from proteins, other nucleic acids, carbohydrates, lipids…): Chemical: precipitation, physical: centrifugation or enzymatical: Proteinase K, RNase A or DNase I. Method depending on what you want to enrich for, DNA and RNA are more homogenous than proteins, so proteins is easier to separate from nucleic acids than it is to separate RNA and DNA.
3. Remove contaminants (Silica column purification or Phenol/chloroform extraction + precipitation) and and increase concentration of nucleic acid by centrifuging and switching to less buffer.

Methods of choice depends on source and form of DNA, and desired purity. Commercial kits are available for easier purification, but it’s important to know what happens when using those too.

Question 35

How does Phenol(/chloroform) extraction work?

Answer 35

Phenol extraction is used to separate DNA and proteins. When having a mix of proteins and DNA in aqueous mix, the protein is folded in a way that puts the polar amino acids outwards.

Then you add phenol (non-polar) and mix thoroughly to force phenol droplets into the water. The protein will then fold in a way that puts the non-polar residues outwards to handle being in non-polar environment.

After that you centrifuge the sample which will result in the denser phenol in the bottom, with the proteins in it, and aqueous solution with DNA in the top –> separated! (this works with chloroform to separate RNA and proteins too.

Question 36

One of the most common quantification methods for nucleotides are using a spectrophotometer. How does quantification of DNA/RNA with a spectrophotometer work?

Answer 36

Nucleic acids absorb UV light (260 nm), and dsDNA, ssDNA and ssRNA have slightly different absorbance at 260nm so by illuminating the sample and measuring absorbance, we get a concentration.
10 mm cuvette: A260 = 1 means
dsDNA: 50 μg/ml
ssRNA: 40 μg/ml
ssDNA: 33 μg/ml

Proteins absorb UV light maximally at 280 nm, so looking at the A260/A280 ratio, we can see purity. If ratio is low –> less pure (more protein). High ratio–> high purity. A260/A230 ratios also estimate purity of DNA/RNA solution, mainly for detecting presence of phenols/carbohydrates that absorb maximally at 230nm. Basically, a peak at anything else than 260 indicates that you have something else in your sample, and not pure nucleic acid.

Remember: absorbance says nothing about quality of DNA/RNA!

Question 37

If you’ve done a PCR and want to determine the concentration, can you do it directly?

Answer 37

No! You need to purify the sample first, to get contaminants and PCR reagents out. To find out quality and or separate just the fragment of choice, you can run a gel electrophoresis to see if degraded or intact and cut out the gel piece to get only the separated fragment.

Question 38

Gel electrophoresis and blotting is commonly used for separation of DNA/RNA/protein, what are the differences in these processes for detecting DNA/RNA vs protein?

Answer 38

Separation by size:
- horizontal agarose gel electrophoresis is commonly used for large molecules like nucleic acids. The concentration of the gel determines how fine the pores are, the higher the conc, the smaller the pores. usually in the range of 0.8%–3% agarose)
- vertical polyacrylamide gel (PAGE) is commonly used for smaller molecules like proteins, different sorts for proteins with different properties.

Native/denaturing conditions: Native=nothing added, denaturing=adding something to change the sample eg.
- Denaturing nucleic acids with denaturants such as urea to make them linear and from there being able to separate by only size (even small differences).
- Proteins usually have varying charge based on side chains, denaturing conditions can help separation.

Question 39

How does gel electrophoresis work?

Answer 39

Alkaline buffer ensures that the nucleic acids have a uniform negative charge –> migrating towards the positive pole. Usually the samples are stained to be seen with the naked eye or by UV-illumination and used together with a size ladder to compare to.

Same principle for both PAGE and regular horizontal gel electrophoresis.

Question 40

How do you see on the gel (after gel electrophoresis) if the sample is intact or degraded?

Answer 40

If clear bands are showing, the sample is intact. If there is no clear bands, but instead a blurry gradient it’s degraded.

Question 41

When would you need to do a blot transfer and how does it work in broad principle?

Answer 41

Since the contents of a gel will diffuse over time and get blurry, you won’t be able to keep it forever. Blotting is done to get a snapshot of a gel, for example when you need to detect what’s in different bands of a gel.

Blotting works by transferring the contents of a gel to a membrane, that’s impenetrable but can attract compounds to stick to it. Either by having a charge on the other side (electroblotting), by capillary transfer, which “sucks up” the contents of the gel and make it stick to the membrane or by vacuum pressure, where it stays in place and can be used for detection or to determine properties of compounds stuck to the membrane.

Question 42

What is a southern and northern blot used for and how do they work?

Answer 42

Southern blot is used to detect DNA, Northern blot is a similar technique but used to detect RNA.

These blotting methods work by transferring the samples in the gel to a membrane (either by capillary transfer or electroblotting) and then adding probes (marked nucleic acids) to the membrane, to see where the probes hybridize (base pair). The membrane is dried/treated with UV to produce cross links between the membrane and the sample, and then the marked probes bound to the target can be detected using for example autoradiography (X-ray) or phosphorimaging.

Question 43

Give two examples of probe labels that can be used in southern/northern blotting.

Answer 43

Internal labeling
- radioactive label
- biotin label
- fluorophore (a fluorescent compound)
- Colorimetric (produce a colored precipitate)
End-labeling also possible.

Question 44

The probes for nucleic acids are made up of nucleic acids that can base pair to the target sequences, but the same thing doesn’t apply to proteins. What is used for probes in specific protein detection?

Answer 44

Antibodies! Antibodies have antigen binding sites that bind to specific domains of proteins also called antigens. Antigens can be used to recognize endogenous proteins or epitope tags
(s a short peptide sequence that is recognized by a commercially available antibody, eg. Myc, FLAG, HA).

Question 45

Antibodies can be mono- or polyclonal, what do these terms mean? How are they produced

Answer 45

Polyclonal antibodies: Produced in animals and bind to several different epitopes of the same protein. When animal dies, you need to start from scratch to produce new ones.

Monoclonal antibodies: produced in vitro after getting base from animal, all antibodies clones of each other that bind the same epitope. Can be produced forever!

Question 46

What is a western blot used for and how des it work?

Answer 46

A western plot is used to detect proteins. The blotting itself is done in the same was as in southern and northern blot but primary antibodies are used as probes instead and membrane is made of other material. Detection usually via colored labels, reagent labeling, or labeled secondary antibodies, eg labeled with enzyme and add substrate to form a compound that can be detected.

Question 47

What are the biggest differences between southern, northern and western blot?

Answer 47

southern (S) - northern (N) - western (W)

• Different targets: DNA - RNA - Protein
• Separation: electrophoresis for all
• Membrane material: Nylon - Nylon - Nitrocellulose/PVDF
• Probe: Nucleic acid with complementary sequence (S) - RNA, DNA or oligodeoxynucleotide (N) - Primary antibody (W)
• Probe label: Radiolabel/enzyme for S and N, enzyme (W)
• Detection method: X-ray film/chemiluminescense for S and N, Film(camera/infrared etc.

Question 48

In proteins, charge is not proportional to the length of the protein as many side chains are uncharged. How is separation by size accomplished for proteins?

Answer 48

By adding the denaturing agent SDS that both denature any secondary structures but the peptide chain itself, which makes it possible to separate by peptide length instead of shape and add a uniform negative charge to all proteins so that they will migrate toward the positive pole of the gel.

(In the absence of SDS, each protein has a specific individual charge at a given pH; it is possible to separate proteins based on these charges, rather
than size, in a technique called isoelectric focusing.)