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MBB164 > Alvey - Molecular Biology Techniques > Flashcards

Alvey - Molecular Biology Techniques Flashcards

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1
Q

name the reagents required for PCR

A 
template (DNA or cDNA) 
primers (need to know DNA seq for these)
enzyme (Taq polymerase)
dNTPs
Buffer - MgCl2
Appropriate temps - thermocycler
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2
Q

name the stages in a PCR reaction and the temps they occur at

A

starts at 25°C
DNA denaturation - 95°C
Primer annealing - 50°C
Primer extension - 72°C

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3
Q

how is it ensured that DNA doesn’t reanneal in a PCR reaction?

A

primers anneal to DNA as they’re added in excess - DNA doesn’t reanneal

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4
Q

name the 2 DNA polymerases that can be used in PCR

A

Taq polymerase - Thermo aquaticus

PFU - high fidelity and proofreading ability

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5
Q

how can PCR results be analysed (hint: we have done this in labs)

A

Run gels – look at band size (how much PCR product you have)/specificity (how many bands you have)

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6
Q

name 7 applications of PCR

A

Genetic screening – modify primers to pick out certain mutations
Pathogen detection – viral DNA in blood sample
DNA fingerprinting - genetic makeup of a organism
Gene expression analysis
Sequencing
Template generation for cloning
Gibson Assembly

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7
Q

what is RT-PCR? why is it used?

A 
reverse transcription - PCR
used to find out:
how much a certain gene is being expressed
where the gene is being expressed
splice variants in a population
how much mRNA is being produced
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8
Q

what are the steps in RT-PCR?

A

mRNA + reverse transcriptase + primer = cDNA
RNAse removes RNA – (ss)cDNA left
Add primers and polymerase = dsDNA formed
DNA can now be amplified in PCR

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9
Q

what is q-PCR? why is it used?

A

Quantitative real-time PCR

Used to measure amount of template

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10
Q

describe the steps in qPCR

A

1) Denaturation step
2) Annealing step
3) Extension step + SYBR green (fluoresces when attached to dsDNA)

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11
Q

name a specific type of qPCR and explain how it works

A

Allele-specific PCR:
Specificity comes from primers
SYBR green is non-specific, binds to any dsDNA
Amount of fluorescence proportional to amount of dsDNA produced
At end of extension step is when fluorescence is measured
qPCR is measured in cycles using an arbitrary unit called cycle Threshold (cT)
Measure background level of fluorescence by not adding the template immediately (compare fluorescence readings to this value)
After each extension step the PCR machine measures fluorescence

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12
Q

what does a low cT number indicate in qPCR?

A

a low cT number = a higher copy number of template DNA (it takes fewer cycles of qPCR to overcome the threshold value of fluorescence)

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13
Q

what is site-directed mutagenesis?

A

In-vitro mutagenesis technique
Used to introduce specific changes into a DNA sequence (you have to know the original sequence for this) to find out whether an a/a or protein is particularly important in an organism you are studying

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14
Q

which amino acid is used to replace a suspected important a/a in site directed mutagenesis?

A

alanine - v boring a/a

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15
Q

describe the steps of site-directed mutagenesis

A

1) Design primers that work in both directions, with one deliberate mismatch in middle of the primer
2) PCR amplifies all around the plasmid
3) Dpn1 digestions of template (dsPlasmids are mainly made up of two newly synthesised strands, however ones that are made up of a new strand and an old strand are digested by endonucleases to remove the mismatched base pair)
4) Transformation of mutant plasmid (added to E. coli - this repairs the nicks in the DNA where the newly synthesised DNA meets the primer)

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16
Q

what is a methylation-dependent restriction enzyme? give and example of one and state why is it used in site-directed mutagenesis

A

a methylation-dependent restriction enzyme is a restriction enzyme that only digests methylated DNA.
example: DpnI
why is it used?: in a PCR reaction the DNA that is amplified isn’t methylated. Therefore the only methylated DNA present in a sample is the DNA that you started with (which doesn’t contain the mutation you have added with your primers). This means that the methylated DNA doesn’t contain your deliberate mutation, so you want it gone - DpnI degrades this DNA for you

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17
Q

what is sanger sequencing?

A

essentially a DNA synthesis reaction by the addition of ddNTPs to create fragments increasing in length
Allows DNA sequencing of 1000-1500 base pairs of good quality sequence
Still most accurate sequencing method

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18
Q

what 2 pieces of information do you have to know in order to carry out sanger sequencing?

A

the length of the fragment and the base at the last position

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19
Q

list the reagents required for sanger sequencing

A 
Template DNA
Oligonucleotide primer
Buffer (including Mg)
dNTP
ddNTP (small amount)
DNA polymerase (Taq)
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20
Q

how do you analyse the results of sanger sequencing using a polyacrylamide gel?

A

you put each of your 4 tubes of sanger sequences (ddATP/ddDTP/ddCTP/ddGTP) into 4 separate wells at the top of the gel. at the 5’ end (furthest away from the wells) is your smallest fragment, and at the 3’ end (closest to the wells) is your largest fragment. as you have made a complementary strand to your template strand you have to use complementary-base pairing rules to work out your original sequence

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21
Q

describe an easier way of analysing sanger sequences than using polyacrylamide gels

A

Fluorescent detection
Each ddNTP is tagged with a fluorescent marker (4 different colours) which a machine picks up and analyses. The last base is determined by the colour of the marker.
Can run the reaction at the same time, instead of four separate reactions.

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22
Q

if in your fluorescent analysis of you sanger sequenced DNA there is a double peak in
a single base position what does this indicate about your DNA sequence?

A

If in a chromatogram there is a double peak at one base position, then:

  • The organism the template is from is a heterozygote
  • This means there were 2 templates present in the reaction mix (one from each chromosome)
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23
Q

name 2 limitations of sanger sequencing

A

Only sequence one template at a time

Need to have some knowledge of the sequence to design the primer

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24
Q

genomes and expression profiles are sequenced using NGS. what is NGS? what are the common features of it?

A

next generation sequencing
common features:
- Millions of reacts occur in parallel
- These reactions are spatially separated
- No sequence knowledge of the template is necessary (this is achieved by adding adapters to the template in which DNA is synthesised from)

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25
describe the steps in traditional cloning
- Chose restriction enzyme that cuts vector once - Digest insert and vector with same enzyme (or enzyme that creates complementary sticky ends) - Ligate insert into vector - Transform into E. coli to amplify and maintain the plasmid
26
name 3 limitations of traditional cloning
Restricted to single insertion per cloning cycle Inefficient for large inserts Plasmid design can be restricted by restriction site availability (ie enzyme may cut gene in middle)
27
what is gibson assembly?
a scar-less cloning method which relies upon recombination rather than restriction to join fragments
28
give 4 benefits of using gibson assembly
Can assemble multiple components in a single reaction (eg ORF + purification tag + promoter + vector) Can accommodate v long inserts Can accommodate up to 4-6 fragments in a single step Restriction digest (therefore restriction sites) aren’t necessary -results in seamless cloning
29
describe the steps in gibson assembly
Imagine and design plasmid Design primers – the primer has to amplify the inserts and be complementary to the next bit of insert/plasmid PCR – amplifies the fragments. Cut the bands from the gel (post-PCR) and extract linear DNA fragments Assemble – add Gibson enzyme mix to create plasmid Transform into E. coli/PCR – stores or amplifies plasmid (plasmids are routinely sequenced after PCR to check for errors
30
describe how the primers create scar-less cloning
overlapping DNA fragments are created with primers that are complementary to both the insert and the vector. 5' --> 3' exonuclease activity then creates single stranded 3' overhangs allowing the fragments (with the complementary primers) to anneal DNA pol extends the 3' ends from after the primer and DNA ligase anneals them.
31
give 4 advantages of golden gate assembly
No need to have repeat elements (doesn’t use homology unlike Gibson) Seamless (restriction sites are lost during cloning) Reaction can occur in a single step Can assemble multiple fragments at once
32
give 2 disadvantages of golden gate assembly
Have to avoid the recognition sequence within your insert DNA Although there are theoretically 256 (NNNN) distinct sticky ends, sequences that differ by only one base may result in unintended ligation products
33
explain the steps in goldengate assembly
GG assembly uses type IIS restriction enzymes which cleave away from their recognition site. BsaI is a type of type IIS restriction enzyme - BsaI sites are introduced into the vector (2 in the vector) and insert DNA by primer design (flank the insert) and create 4 NTP overhangs (the sticky ends can be designed into any seq of 4NTPs therefore they can be a continuation of the insert fragment (leaves no scar) as the vector contains 2 recognition sites the recognition sequence can be within the part of the vector that the inserted fragments replace (and are therefore lost = scarless)
34
which analytical methods are used in analysing RNA expression and localisation
``` used for analysing gene expression at the level of transcription RT-PCR Northern blots in situ hybridisation microarrays ```
35
which analytical methods are used in analysing protein expression and translation
western blots | reporter genes
36
name 4 reporter genes
green fluorescent protein luciferase beta-glucoronidase beta-galactosidase
37
name 3 methods for analysing interactions between proteins and other molecules
``` Chromatin immunoprecipitation (DNA:Protein) Pull down assay (protein:protein in vitro) Yeast two-hybrid (protein:protein in vivo) ```
38
describe northern blotting
RNA extracted and run on gel then transferred to membrane and hybridised with a labelled DNA gene-specific probe N blots reveal size and abundance of your specific gene can reveal tissue-specific expressions if you extract the total RNA from different tissues/developmental stages can reveal splice variants
39
describe in situ hybridisation
binding labelled probe to thin slice fixed tissue | FISH can show specific mRNA accumulation patterns in a tissue/organ/organism
40
describe microarray transcriptional profiling
2 different mRNAs mRNA ---> cDNA cDNA attached to label mix 2 samples together and then hybridise to surface DNA microarrays monitor expression 1000s genes simultaneously describe expression profile of particular tissue/organ
41
describe western blotting
proteins extracted and resolved by SDS PAGE (separates protein by size) proteins transferred to mem incubate with specific antibody (against target protein) incubated with secondary antibody to visualise 1st antibody
42
what are reporter genes
Reporter genes are used to analyse gene expression. They are known genes whose RNA or protein levels can be measured easily
43
what is co-localisation?
the overlap of 2 images with different fluorescent markers - allows comparison between where genes are (both) expressed
44
describe chromatin immunoprecipitation
study of DNA-protein interactions in vivo (within context living cell) DNA-protein incubated with primary and secondary antibodies the secondary antibody is prebound to a resin (which enters the pellet) DNA-protein complexes are immunoprecipitated out of solution unbound DNA remains in supernatant Bound and unbound DNA are analysed and compared
45
describe a pull-down assay
protein-protein interactions in vitro uses GST-tagged protein 'bait' to identify new protein partners the 'prey' protein can be from a cell lysate proteins are eluted and visualised by SDS-PAGE (see which 'prey' proteins have bound the 'bait' and which have been eluted new partners identified by w. blotting or mass spec
46
describe yeast two-hybrid analysis
protein-protein interactions in vivo 'bait' fused to DNA binding domain 'prey' fused to an activating region if the 'bait' and 'prey' bind together (and with their respective domains), a reporter gene is switched on
47
what are DNA libraries?
collections of different overlapping fragments within the same vector libraries are made from a specific source (eg yeast genomic or human hepatocyte cell expression library)
48
describe in brief how DNA libraries are constructed
DNA fragments generated by restriction endonuclease activity Fragments ligated into vector Collection recombinant molecules transferred to host cells (eg E. coli) (one molecule per cell) Library screened with molecular probe to identify the clone that contains the gene of interest (GoI)
49
name 3 types of DNA library and explain them
Genomic libraries: whole genome (non-coding incl.) eg yeast cDNA libraries: DNA copy of RNA; only the expressed genome (no introns) eg human Sequencing libraries: libraries used to sequence genomes, used in HGP
50
describe how genomic DNA is extracted when constructing a genomic library
Extract gDNA (genomic DNA) using restriction enzymes Fragments must be appropriate size Generation of compatible sticky ends using BamHI and Sau3A
51
state the size of the recognition site in BamHI and Sau3A and explain why one will be favoured in cutting the plasmid
BamH1 (6 base recognition site) Sau3A (4 base recognition site) BamH1 used in plasmid vector as it has a 6 base recognition site therefore there is less of a chance of it being cut in multiple places
52
describe how and why gDNA is partially digested when constructing a genomic library
Partially digest the gDNA: Complete digestion: the restriction enzyme has cut every restriction site Incomplete digestion: the restriction enzyme has cut some recognition sites - This makes overlapping fragments
53
what is loaded into a gel when constructing a genomic library? which fragment sizes are chosen?
the overlapping fragments produced by the partial digestion of gDNA by Sau3A are loaded into the gel 2-8kb fragments are chosen
54
why are Sau3A and BamHI used in constructing a genomic library?
they generate complementary sticky ends
55
why and when is a plasmid used as a vector? whats its max insert size?
uses multicopy plasmids cDNA cloning and expression libraries max insert size 10kb
56
why and when is a M13 phage used as a vector? whats its max insert size?
double stranded replicative form of M13 genomic libraries and sequencing projects max insert size: 20kb
57
why and when is a yeast artificial chromosome used as a vector? whats its max insert size?
why use it: contain telomeres, centromeres (to ensure segregation into daughter cells) and replicating sequence (ability to replicate) max insert: 1000kb used for: analysis of large genomes - makes library more manageable
58
what is the difference in the M13 genome and replicative form? how can inserts be sequenced in parallel?
M13 has a (+)ssDNA genome but is replicated as a dsDNA replicative form (RF) in E. coli DNA can be cloned in the RF form and isolated as ssDNA from phage particles M13 used in HGP Inserts can be sequenced in parallel using a common universal primer Primer is complementary to vector adjacent to insert
59
describe how RNA is extracted when a cDNA library is being constructed (ie step 1)
rRNA is most common form of RNA, need to collect only mRNA mRNA has 5’ m7G cap and polyA tail: use oligo (dT) affinity chromatography Oligo dT is immobilised and recognises the polyA tail
60
describe how mRNA | is collected from the RNA when a cDNA library is being constructed (ie step 2)
All RNA population from cell loaded in column containing oligo dT mRNA with polyA tail bind to column. Other RNA washed off with NaCl The mRNA can be eluted from the column by using a low [NaCl] buffer
61
describe how cDNA is synthesized from mRNA when a cDNA library is being constructed (ie step 3)
Oligo dT acts as a primer (TTTTTT…) Reverse transcriptase and dNTP's added to extend the TTTT… tail Cap trapping means the full length cDNA is copied (only synthesises things with m7G cap and tail) RNAse I/heat or alkaline denaturation removes the ssRNA
62
describe how the (ss)cDNA's complementary strand is synthesised when a cDNA library is being constructed (ie step 4)
Single stranded cDNA has its 3’ end extended using a terminal deoxynucleotidyl transferase (TdT) This adds a series of G’s to the 3’ end (essentially acting as a known primer) PCR amplifies from this primer as the tails are a series of G’s and T’s (ie there are primers to work from)
63
describe how the dsDNA is transformed into its vector when a cDNA library is being constructed (ie step 5)
cDNA treated with methylase to protect it from the exonuclease activity EcoR1 linker (restriction) sites ligated to the cDNA ends Linkers are digested by EcoR1, cDNA protected by methyl groups Product inserted into EcoR1-ligated vector
64
name 3 methods of finding a clone with your gene of interest in it
synthesised oligonucleotides random priming screening a library by hybridisation
65
describe how synthesised oligonucleotides are used to identify clones containing your gene of interest
MUST know peptide sequence for this Use peptide sequence and codon tables to work out all the possible codon sequences These added to mix where one will anneal
66
describe how random printing is used to identify clones containing your gene of interest
Have sample of DNA but don’t know its sequence Use a mixture of 4096 sequences (4(diff NTPs)^6) Some will anneal Add DNAP which has been labelled with 32P. Denature, hybridise, use labelled probe as primer
67
describe how screening a library by hybridisation is used to identify clones containing your gene of interest
For hybridisation assays DNA must be bound to a membrane surface Colonies transferred from agar plate to membrane and are then lysed After NaOH wash and baking the DNA is now fixed on a membrane Radioactive probe is added and binds to complementary DNA Autoradiography (x-ray) is used to highlight the clones with the correct sequence - This is positive screening

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