Alvey Flashcards
What is the process of studying genes?
- isolate single gene
- amplify it
- modify it
- analyse expression
What is the role of DNA ligase
- joins cos sites and allows rejoining of genomic DNA after recombination
What is DNA ligase req for?
- covalent bond formation
Where is DNA ligase most commonly obtained from?
- bacteriophage T4
How does DNA ligase join ends?
- adding adenylate group to Lys
- transferring adenylate to terminal 5’ phosphate group
- phosphodiester bond formation
What co-factor does DNA ligase req?
- ATP co-factor
What does restriction mean in terms of phage?
- phages grown in 1 host failed to grow in new host
What does modification mean in terms of phage?
- rare progeny phages able to grow in new host
Why are phage restricted?
- due to nuclease that degrades foreign DNA
- mechanism to protect against viral infection
How are phage modified?
- methylation of host DNA at sites otherwise sensitive to attack by restriction endonuclease
What was the 1st restriction endonuclease discovered, and what type of DNA did it destroy/not destroy?
- K-12 in E. Coli
- λK DNA not destroyed by K-12 host enzymes
- λC DNA destroyed by K-12 host enzymes
How do the 3 classes of restriction enzymes differ in role and application?
- I and III cleave DNA sites away from recognition seq, so no good for most mol bio applications
- II cleave both DNA strands at specific recognition site, most abundant and widely used in mol bio
How are restriction enzymes named?
- 1st 3 letters abbreviation of bacteria isolated from
- 4th represents strain of bacteria
- no.s indicate which of multiple enzymes identified from given strain
What is molecular cloning?
- creation of recombinant DNA molecules and rep in host organism
How is molecular cloning carried out? (overview)
- run agarose gel
- put PCR product into plasmid vector
- amplify DNA, eg. transfer into E. Coli, kept separate from genome
- select transformed bacteria
What can be used as a vector for molecular cloning?
- plasmid
- phage
- cosmid
- BAC
- YAC
- MAC
What does a vector req to be used for molecular cloning?
- selectable marker
- restriction sites for cloning fragment into
- own origin of rep (need 2 in mammals)
Why are complementary overhangs useful?
- any 2 sites cut w/ same enzyme can be joined
What are problems w/ use of restriction enzymes in molecular cloning?
- self ligation of vector possible
- any complementary overhangs compatible, don’t get restriction site back
How can the problem of self ligation of the vector be solved in molecular cloning?
- phosphate treatment –> removes 5’ phosphate, stopping self ligation, insert donates 5’ phosphate
- use of 2 diff enzymes –> vector doesn’t have complementary sticky ends and insert always orientated in same way, this is directional cloning
What are the problems w/ blunt end cloning?
- inefficient
- lots of false positives
What enzymes can be used for addition/removal of phosphate groups?
- polynucleotide kinase
- phosphatase
How can an overhang be converted to a blunt end?
- T4 DNA pol or Klenow fragment of DNA pol I
- fill in 5’ overhang in presence of dNTPs (5’ to 3’ pol activity)
- 3’ to 5’ exonuclease activity will remove 3’ overhang
What happens during TA cloning?
- Taq adds 3’ overhang to DNA products
- can buy vectors linearised w/ T overhang or make it
- T overhang added by terminal transferase
- DNA Taq pol lacks 3’ to 5’ proofreading activity, adds 3’ adenine
- prepare vector by blunt end activity
- ddTTP addition using terminal transferase
What is bacterial transformation?
- inserting recombinant plasmid into cell
How can DNA be forced into cell during bacterial transformation?
- make cells competent w/ CaCl2, creates pores, v effective w/ heat shock
- electroporation
What is blue-white screening and how does it work? (selection and amplification)
- tell difference between recombinant and non-recombinant
- insert interrupts lacZ gene = white colonies
- no insert = blue colonies
- doesn’t show orientation or what insert is
What are the next steps carried out in molecular cloning after selection and amplification?
- further screening
- larger scale culture
- plasmid purification
- expression analysis
What is PCR?
- method of amplifying specific DNA seq
What is needed for PCR?
- template DNA/cDNA w/ free 3’ OH
- primers (need knowledge of seq)
- enzyme (pol)
- dNTPs
- buffer (MgCl2)
- approp temp (thermocycler)
What are the stages of PCR?
- ~92ºC = DNA denaturation
- ~50ºC = primer annealing
- ~70ºC = primer extension
What affects the temps PCR is carried out at?
- higher temps if GC rich
Why is PCR only have exponential amplification in theory?
- reagents start to run out near the end
What are the applications of PCR?
- genetic screening (eg. specific mutation)
- pathogen detection
- DNA fingerprinting
- gene expression analysis
- sequencing
- template gen for cloning
- Gibson Assembly
How are PCR results analysed?
- stained w/ ethidium bromide
- single band = pure product
- brightness = efficiency
How can PCR results be used for cloning?
- cut out of gel and extract DNA, so not cloning any impurities from other products on gel
Why is PFU better than Taq pol for PCR?
- any early errors are amplified and present in more DNA
- PFU has much lower error rate
How is direct cloning of PCR products carried out?
- design primer complementary to 3’ end of template strand
- restriction enzymes find hard to cut ends and add nucleotides to end of seq
- DNA amplified using primers inc suitable restriction sites
- PCR product cloning after restriction digestion
What is the alternative to direct cloning of PCR products?
- TA cloning of Taq modified products
What is RT-PCR used for?
- find out if certain gene being expressed
- find where gene is expressed
- investigate splice variants of pop
- find how much RNA prod
How does RT-PCR work?
- RNA ss, so don’t need 1st denaturation step
- use polyA tail to design primer w/ run of Ts
- add RT (from virus), zips along gene to end
- results in ds RNA/DNA hybrid
- use RNAse to remove RNA, leaving ss cDNA strand
- then normal PCR
What does qPCR do?
- monitors amplification in real time by monitoring light emitted
- estimates amount of specific template DNA present in sample
What are the 2 variations of qPCR?
- multiple probes –> each have own fluorescent tag, so can measure many genes at once
- allele specific –> uses SYBR green which -fluoresces only when bound to ds DNA, so measured at end of extension step
What do the results of qPCR show?
- amount of DNA quantified as cycle threshold
- the higher the Ct value, the less DNA present and lower the expression level of that gene
What is the disadvantage of qPCR?
- only estimated relative amounts so need control (‘housekeeping’ gene) with constant expression level
- if investigating gene expression level, 1st need to convert mRNA molecule into cDNA molecule before performing qPCR, using RT
What is site directed mutagenesis used for?
- for intro of specific mutations into DNA (see if protein changes w/ AA)
What are the 3 steps of site directed mutagenesis?
- mutant strand synthesis
- DpnI digestion of template
- transformation
What primers are designed for site directed mutagenesis?
- primers designed in both directions w/ 1 deliberate mismatch
What is the cycle threshold in qPCR?
- point fluorescence level enters exponential phase (and exceeds background level)
What is Sanger sequencing?
- variation of PCR technique
- DNA synthesis reaction
- allows seq of 1000-1500 bp of good quality seq
What do you need to know for Sanger sequencing, and why?
- length of fragment and base at last position, can deduce seq by adding 1 letter at a time
How does chain termination occur?
- fragments added by 2’3’ dideoxy analogue of dNTP
- dideoxy analogue can’t be extended (ddNTPs)
- used to radiolabel ddNTPs, everytime got a fragment, knew it was that letter
- new fragments separated by gel electrophoresis
What are the components of sequencing reaction? (Sanger sequencing)
- template DNA
- oligonucleotide primer
- buffer for pol, inc Mg
- all dNTPs
- small amount ddNTPs
- DNA pol (Taq) –> not proofreading enzyme as would correct last base (which is incorrect)
How can fluorescence detection used in Sanger sequencing?
- would need 4 separate reactions w/ ddATP, ddCTP, ddGTP, ddTTP
- but labelled w/ diff colours so can be carried out together
- know by colour which is last base
What are the limitations of Sanger sequencing?
- can only seq 1 template at a time
- need some knowledge of seq to design primer
How is Sanger sequencing diff now and what is it used for?
- now automated
- used to seq plasmids, PCR products etc.
What is next gen seq (NGS) used for?
- genomes
- expression profiles
What are some of the common features of NGS technologies?
- millions of reactions run in parallel
- reactions spatially separated
- no seq knowledge of template req, can seq from adaptor into unknown
What is GIbson Assembly?
- alt cloning technique
- relies on recombination
- efficient for gen large multi-part constructs
- assembly occurs in single reaction
- collection of promoters, terminators, selectable markers or tags generated w/ compatible overlaps
- allows rapid gen of series of constructs w/ diff properties
What are the limitations of traditional cloning?
- restricted to single insertion per cloning cycle
- inefficient for large inserts
- plasmid design can be restricted by restriction site availability
What are the benefits of Gibson Assembly?
- scarless cloning –> can directly add tag so no restriction site left over after cloning
- can assemble multiple components in single reaction, eg. ORF , purification tag, promoter and vector
- can accom v long inserts and 4-6 in single step
- restriction digest (and therefore restriction sites) not necessary
What are the 5 principles of Gibson Assembly
- imagine plasmid design
- design primers
- PCR
- assembly
- transformation
What occurs during imagining plasmid design? (Gibson Assembly)
- place insert(s) into vector back bone
- draw out seq of vector and insert
- draw out plasmid seq to prod
What occurs during design of primers? (Gibson Assembly)
- must overlap junctions in both directions
- half homologous to insert and half complementary to vector seq (so half amplify vector and half amplify insert)
- always written 5’ to 3’
What occurs during PCR? (Gibson Assembly)
- cut bands from gel and extract linear DNA fragment
- now have overlapping linear DNA for whole plasmid inc vector backbone
What occurs during assembly? (Gibson Assembly)
- 5’ to 3’ exonuclease activity creates 3’ overhangs
- these complementary seq anneal, forming ds DNA
- DNA pol extends 3’ end and DNA ligase seals remaining nicks
- resulting in fully sealed ds DNA
What occurs during transformation? (Gibson Assembly)
- can be directly transformed into E. Coli for amplification and storage
- transformed colonies screened for correct plasmid
- plasmids routinely seq after PCR to check for errors (uncommon)
When is Gibson Assembly most useful?
- when used to gen batches of related constructs, eg. adding fluorescent tag to all genes of interest
What is Golden Gate Assembly (Type IIS)?
- restriction enzyme and ligase cloning
What are the advantages of Golden Gate Assembly?
- don’t need repeat elements, as doesn’t use homology like Gibson
- seamless (restriction sites lost)
- reaction can occur in single step
- can assemble multiple fragments at once
What are the disadvantages of Golden Gate Assembly?
- have to avoid recognition seq w/in insert DNA
- although theoretically 256 (NNNN) distinct flanking seqs (sticky ends), seqs differing by 1 base may result in unintended ligation products
What are principles of Golden Gate Assembly? eg. BsaI
- type IIS restriction enzymes cleave away from non-palindromic recognition sites
- can choose sticky ends (at least 2 diff)
- incorp into insert in correct orientation, to ensure lost during cloning
- PCR fragments and vector assembled in single tube reaction containing BsaI and T4 DNA ligase
- cleavage w/ BsaI exposes complementary seq for fragment assembly
- nicks sealed w/ DNA ligase
- correctly assembled products lack BsaI sites
- add 55ºC incubation step, BsaI digests incorrect fragments
- resulting in correctly assembled products, which can be directly cloned into E. Coli
What is gene expression?
- prod of functional RNA or protein from genetic info encoded by genes
How can gene expression be analysed at the level of transcrip?
- look at level of transcrip by analysing mRNA levels
What does differential expression of genes result in?
- diff cells, tissues and organs
What is hybridisation?
- 2 single NA strand coming together and binding
What do hybridisation techniques involve?
- once seq of gene known, can be used to make probe
- bind in seq specific manner using bping rules
- eg. northern blots, in-situ hybridisation, microarrays
How is northern blotting carried out?
- extract RNA and resolve through gel
- transferred to membrane, then hybridised w/ gene specific probe
What do results of northern blotting show?
- shows size (small = fast) and abundance
- can reveal tissue-specific expressions if extract total RNA from diff tissues of diff dev stages
- can reveal splice variants of gene
How is in situ hybridisation carried out?
- involves binding of labelled probe to thin slice of fixed tissue
- FISH can show specific mRNA accum patterns of tissue, organ or whole organism
What are microarrays?
- DNA microarrays monitor expression of 1000s of genes simultaneously
- describes expression of particular organ/tissue
- use hybridisation on large scale
How are microarrays carried out?
- extract mRNA from 2 samples to compare expression profiles of, eg. healthy and cancer cells
- convert mRNA to DNA
- add fluorophore to cDNAs
- mix 2 samples and apply to microarray chip
- wash
- read fluorescence using laser scan (presence of colour indicates dominance)
What is a southern blot?
-probe DNA blot w/ labelled DNA probe
What is the difference between the 3 blotting techniques?
- northern = probe RNA blot w/ labelled DNA probe
- southern = probe DNA blot w/ labelled DNA probe
- western = use protein-specific antibodies to detect proteins
How can gene expression be analysed at level of transcrip?
- req detecting levels of specific proteins
- often uses antibodies to specifically bind to protein of interest
- eg. Western blotting
- use reporter genes, easily measured and analysed
How is western blotting carried out?
- proteins extracted and resolved by SDS PAGE (separates by size, small = bottom/fast)
- proteins transferred to membrane
- incubated w/ labelled 2º antibody to visualise 1st antibody
- vertical gel
Why does SDS PAGE have smaller holes than agarose gel?
- proteins smaller
Why were engineered fluorescence proteins needed?
- needed more colours to look at more than 1 protein at once
What are reporter genes and what are they used for?
- to analyses gene expression
- known genes whose RNA or protein levels can be measured easily
- commonly used allow protein visualisation in vivo
What is co-localisation and why is it needed?
- often wish to know where protein resides in cell w/ respect to something else
- researchers dev strains and cell lines w/ specific cellular compartments labelled w/ certain colour
- confocal microscope software can ‘overlay’ 2 images to highlight where both genes expressed
Why are tool for analysing protein interactions needed?
- proteins rarely function alone, usually part of large complexes
- to study function, need to know what it binds w/
- eg. chromatin immunoprecipitation, pull-down assay, yeast 2-hybrid
What is chromatin immunoprecipitation (ChIP) and how is it carried out?
- DNA-protein in vivo
- DNA-protein complexes immunoprecipitated out solution
- unbound DNA remains in supernatant
- bound and unbound DNA analysed by variety of methods (eg. microarray)
What is a pull-down assay and how is it carried out?
- protein-protein in vitro
- uses GST tagged ‘bait’ to identify new protein partners
- ‘prey’ protein can be from lysate
- proteins eluted and visualised by SDS PAGE
- new partners identified by mass spec (no idea) or western blotting (to confirm idea, need to know antibodies)
What is yeast 2-hybrid and how is it carried out?
- ‘bait’ fused to DNA binding domain
- prey fused to activating regions
- if bait and prey interact, reporter gene switched on
What are DNA libraries a resource for?
- gene cloning
What are DNA libraries?
- collection of diff genomic DNA fragments into same vector type
What is needed for a library to have good coverage?
- contain several genome equivalents (GEs)
How are DNA libraries constructed?
- vectors used to compile library of DNA fragments of genomes
- DNA fragments gen by cutting DNA w/ restriction endonuclease
- fragments ligated into approp vector
- collection of recombinant molecules transferred into host cells, 1 unique molecule in each cell
- library screened w/ mol probe to identify clone containing target DNA of interest
What are the types of libraries?
- genomic
- cDNA (easier for large genomes)
- seq (M13 phage as vector
How are genomic libraries made?
- extract gDNA from organism/tissue/cell line of interest
- choose approp restriction enzyme to cut vector and gDNA
- partial digest
- gel electrophoresis to extract approp size fragments (2-8kb)
- mix w/ vector and ligate w/ DNA ligase
How is the correct enzyme chosen to use to digest genome?
- must give approp size fragments and gen sticky ends complementary to vector
For humans the no. fragments is too large for a genomic library, so what is used instead?
- diff vector
- cDNA library
What are YACs and why are the used?
- yeast artificial chromosomes
- can carry most DNA and accom v large inserts (1Mb)
- can be grown in yeast (relatively easy)
- shuttle vectors = can live in more than 1 host (also E. Coli)
- can be digested to prod 2 telomeric seqs of artificial chromosome to be propagated in yeast
- DNA insertion makes artificial chromosome
When are cDNA libraries used?
- for looking at expressed genes
What needs to be done in cDNA libraries before vector can be cloned?
- need to copy ss RNA into ds DNA
What are the main steps of generating a cDNA library?
- extract RNA from cells of interest
- 1st strand synthesis
- 2nd strand synthesis
- cloning PCR product into vector
What happens during extraction of RNA from cells of interest? (generating a cDNA library)
- all RNA species extracted
- use oligo (dT) affinity chromatography to enrich for mRNA from protein-coding genes
- all RNA pop loaded onto column w/ oligo dT immobilised on beads
- mRNA (w/ polyA tail) will bind to column, others washed off w/ NaCl
- mRNA can be eluted and collected from column using low salt buffer
What type of RNA is most abundant?
- rRNA
- mRNA only ~5%
What happens during 1st strand synthesis? (generating a cDNA library)
- prod of ss DNA complementary to ss RNA
- performed by RT
- oligo dT (11Ts) common primer
- cap trapping ensures only full length cDNAs gen (RNAse I treatment)
What happens during 2nd strand synthesis? (generating a cDNA library)
- ss cDNA needs to be converted to ds DNA
- RNA strand denatured during heating
- 3’ cDNA extended using homopolymer tailing, using terminal deoxynucleotidyl transferase
- product amplified by PCR
What happens during cloning of PCR product into vector? (generating a cDNA library)
- cDNA treated w/ methylase to protect endogenous EcoRI site from digestion
- linkers digested w/ EcoRI, cDNA protected by methylation
How is method to find clone containing gene of interest decided?
- each colony contains many copies of plasmid containing unique DNA insert
- screening approach chosen based on info available about gene
What are the methods for finding clone containing gene of interest?
- hybridisation –> exploit that DNA and RNA can bind to specific seq
- immunological techniques –> need antibody for gene product
- complementation –> screen library by protein function, spot cell reverting to WT
How is screening by hybridisation carried out?
- synthesise hybridisation probe
- for hybridisation assays, DNA must be bound to membrane surface
- transfer of colonies –> library transformants replicated from agar plates onto membranes and cells lysed
- DNA fixed onto membrane (NaOH, wash, bake)
- colony hybridisation
- detection of +ve clones –> position probe detected on membrane used to identify colony containing correct seq
How is the probe for hybridisation chosen?
- if know peptide or DNA seq (or part seq) for gene then use synthesised oligonucleotide probe
- if have some DNA, but don’t know seq, then random priming used
How is a synthesised oligonucleotide probe made? (hybridisation)
- peptide seq converted to DNA seq
- ordered as pools of degenerate seq (as don’t always now 3rd base)
- can use coding or template DNA, but always 5’ to 3’ direction
How does random priming work? (hybridisation)
- ds DNA denatured and anneal N6 random primers
- hybridise randomly w/ DNA pol and dNTPs
- denature, hybridise and use as labelled probe
- use radioactive copies to create library
How do different temps and salt concs affect colony hybridisation?
- for perfect match salt low and temp high
- for some mismatches salt high and temp low
How are expression libraries screened and when are they used?
- used when antibody against encoded cDNA or gene available
- antibodies can be used to screen expression libraries
- expression libraries use vectors that permit expression of inserted cDNA seq
- MCS in vector situated downstream of promoter, ribosome binding and operator seqs
How is immunological screening of cDNA libraries?
- expression libraries can be screened using protein-specific antibodies
- colony transfer –> replica plated onto filters, cells lysed and protein fixed to membrane
- colony incubation
- membranes incubated w/ protein-specific antibody (1º) then labelled 2º antibody
- detection of +ve clones –> presence of 2º antibodies detected
- more efficient as screening prone to false +ves
How is gene expression and function studied in mice?
- transgenic mice
- inducible systems
- gene targeting
Why do mice make good models?
- mouse and human genomes show ~90% synteny
- directly homologous mouse genes for 99% + human genes
- most protein seqs highly conserved
What does transgenic mean?
- organism that carries transferred genetic material (transgene), inserted at random site in genome
What is gene targeting?
- disruption or mutation of particular gene (eg. knockout)
How can transgenics be generated via micro-injection into fertilised eggs?
- transgenic mice successfully integrated and expressed rat growth hormone
- gain of function model
How can transgenics be generated via pronuclear micro-injection into fertilised eggs?
- mice will be hemizygous and may be mosaics
- prod of stable transgenic lineage req 2nd inbred gen
How is transgene expression analysed?
- is there stable integration of transgene into mouse chromosome?
- if transgene present, is it expressed approp?
- tail biopsies for DNA analysis by southern blot or PCR, integration random and occurs by nonhomologous recombination, more than 1 copy can be integrated
What are the limits of early transgenic tech?
- limited to gain of function studies
- random insertion
- no control over expression
- tissue specific –> spatial control of transgene
- inducible promoter –> temporal control of transgene
How can expression be analysed at level of transcrip?
- northern blots
- RT-PCR
- in-situ hybridisation
How can expression be analysed at level of translation?
- western blots
- immunohistochemistry
- GFP expression
Why do inducible systems exist?
- if intro of transgene prevents or limits survival in utero (embryonic lethal)
- can delay expression of transgene
How do inducible systems work?
- use of inducible transcriptional activator
- expression reg in reversible and quantifiable manner
What is gene targeting?
- alters endogenous genes in mouse ES cells
- homologous recombination w/ foreign DNA seq
- +vely selected cells injected into mouse blastocyst
- mice will be chimeric, further breeding and selection steps req
- ends of vector must have seq homologous to mouse chromosome
- disrupts targeted gene (knockout)
- inserts selectable marker to enable identification
What are the types of gene targeting? (knocking)
- knock out = inactivate gene
- knock in = exogenous gene introd to disrupt targeted endogenous gene
- knock down = gene expression decreased
What does the targeting vector contain? (gene targeting)
- selectable marker
- regions of homology w/ mouse chromosome
- planned mods that alter expression of targeted gene
What is a commonly used selection marker? (gene targeting)
- neomycin resistance gene
- allows use of neomycin to kill other cells
What knock out mice model resulted in adult-onset obesity in mice?
- targeted deletion of neuronal basic helix-loop-helix transcrip factor 2 (NhIh2)
What are mice libraries?
- US based consortium systematically knocking out mouse genes 1 by 1 in ES cells
- European based consortium, engineering knock out containing genes that can be switched on or off at any dev stage in mutant mouse
What are “floxed” mice?
- site specific recombination to remove targeted gene
Why were “floxed” mice created?
- to examine gene only in context of particular organ system or cell type
- to avoid embryonic lethal mutants
How are “floxed” mice created?
- “cre-lox” system
- cre recombinase combines w/ recombinase recognition sites (LoxP sites)
How is cre-lox strain made?
- cre recombinase recognises 34bp site on bacteriophage P1 genome
- catalyses reciprocal recombination between pair of lox sites
- cre and loxP strains dev separately and crossed together to prod cre-lox strain
How is M13 phage used as a library vector?
- ss(+) circular DNA genome
- replicated as ds replicative form in E. Coli
- DNA can be cloned in replicative form and isolated as ss DNA from phage particles
- used extensively in genome seq projects
- inserts can be seq in parallel using common universal primer, complementary to vector adj to insert
What are the steps in cleavage of DNA by restriction enzyme?
- nonspecific binding
- sliding/hopping/jumping
- specific binding
- coupling
- catalysis
- product release
