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MBB164 > Alvey > Flashcards

Alvey Flashcards

1
Q

What is the process of studying genes?

A
  • isolate single gene
  • amplify it
  • modify it
  • analyse expression
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2
Q

What is the role of DNA ligase

A
  • joins cos sites and allows rejoining of genomic DNA after recombination
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3
Q

What is DNA ligase req for?

A
  • covalent bond formation
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4
Q

Where is DNA ligase most commonly obtained from?

A
  • bacteriophage T4
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5
Q

How does DNA ligase join ends?

A
  • adding adenylate group to Lys
  • transferring adenylate to terminal 5’ phosphate group
  • phosphodiester bond formation
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6
Q

What co-factor does DNA ligase req?

A
  • ATP co-factor
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7
Q

What does restriction mean in terms of phage?

A
  • phages grown in 1 host failed to grow in new host
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8
Q

What does modification mean in terms of phage?

A
  • rare progeny phages able to grow in new host
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9
Q

Why are phage restricted?

A
  • due to nuclease that degrades foreign DNA

- mechanism to protect against viral infection

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10
Q

How are phage modified?

A
  • methylation of host DNA at sites otherwise sensitive to attack by restriction endonuclease
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11
Q

What was the 1st restriction endonuclease discovered, and what type of DNA did it destroy/not destroy?

A
  • K-12 in E. Coli
  • λK DNA not destroyed by K-12 host enzymes
  • λC DNA destroyed by K-12 host enzymes
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12
Q

How do the 3 classes of restriction enzymes differ in role and application?

A
  • 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
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13
Q

How are restriction enzymes named?

A
  • 1st 3 letters abbreviation of bacteria isolated from
  • 4th represents strain of bacteria
  • no.s indicate which of multiple enzymes identified from given strain
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14
Q

What is molecular cloning?

A
  • creation of recombinant DNA molecules and rep in host organism
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15
Q

How is molecular cloning carried out? (overview)

A
  • run agarose gel
  • put PCR product into plasmid vector
  • amplify DNA, eg. transfer into E. Coli, kept separate from genome
  • select transformed bacteria
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16
Q

What can be used as a vector for molecular cloning?

A
  • plasmid
  • phage
  • cosmid
  • BAC
  • YAC
  • MAC
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17
Q

What does a vector req to be used for molecular cloning?

A
  • selectable marker
  • restriction sites for cloning fragment into
  • own origin of rep (need 2 in mammals)
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18
Q

Why are complementary overhangs useful?

A
  • any 2 sites cut w/ same enzyme can be joined
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19
Q

What are problems w/ use of restriction enzymes in molecular cloning?

A
  • self ligation of vector possible

- any complementary overhangs compatible, don’t get restriction site back

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20
Q

How can the problem of self ligation of the vector be solved in molecular cloning?

A
  • 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
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21
Q

What are the problems w/ blunt end cloning?

A
  • inefficient

- lots of false positives

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22
Q

What enzymes can be used for addition/removal of phosphate groups?

A
  • polynucleotide kinase

- phosphatase

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23
Q

How can an overhang be converted to a blunt end?

A
  • 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
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24
Q

What happens during TA cloning?

A
  • 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
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25
Q

What is bacterial transformation?

A
  • inserting recombinant plasmid into cell
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26
Q

How can DNA be forced into cell during bacterial transformation?

A
  • make cells competent w/ CaCl2, creates pores, v effective w/ heat shock
  • electroporation
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27
Q

What is blue-white screening and how does it work? (selection and amplification)

A
  • 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
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28
Q

What are the next steps carried out in molecular cloning after selection and amplification?

A
  • further screening
  • larger scale culture
  • plasmid purification
  • expression analysis
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29
Q

What is PCR?

A
  • method of amplifying specific DNA seq
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30
Q

What is needed for PCR?

A
  • template DNA/cDNA w/ free 3’ OH
  • primers (need knowledge of seq)
  • enzyme (pol)
  • dNTPs
  • buffer (MgCl2)
  • approp temp (thermocycler)
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31
Q

What are the stages of PCR?

A
  • ~92ºC = DNA denaturation
  • ~50ºC = primer annealing
  • ~70ºC = primer extension
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32
Q

What affects the temps PCR is carried out at?

A
  • higher temps if GC rich
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33
Q

Why is PCR only have exponential amplification in theory?

A
  • reagents start to run out near the end
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34
Q

What are the applications of PCR?

A
  • genetic screening (eg. specific mutation)
  • pathogen detection
  • DNA fingerprinting
  • gene expression analysis
  • sequencing
  • template gen for cloning
  • Gibson Assembly
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35
Q

How are PCR results analysed?

A
  • stained w/ ethidium bromide
  • single band = pure product
  • brightness = efficiency
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36
Q

How can PCR results be used for cloning?

A
  • cut out of gel and extract DNA, so not cloning any impurities from other products on gel
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37
Q

Why is PFU better than Taq pol for PCR?

A
  • any early errors are amplified and present in more DNA

- PFU has much lower error rate

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38
Q

How is direct cloning of PCR products carried out?

A
  • 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
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39
Q

What is the alternative to direct cloning of PCR products?

A
  • TA cloning of Taq modified products
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40
Q

What is RT-PCR used for?

A
  • find out if certain gene being expressed
  • find where gene is expressed
  • investigate splice variants of pop
  • find how much RNA prod
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41
Q

How does RT-PCR work?

A
  • 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
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42
Q

What does qPCR do?

A
  • monitors amplification in real time by monitoring light emitted
  • estimates amount of specific template DNA present in sample
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43
Q

What are the 2 variations of qPCR?

A
  • 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
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44
Q

What do the results of qPCR show?

A
  • amount of DNA quantified as cycle threshold

- the higher the Ct value, the less DNA present and lower the expression level of that gene

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45
Q

What is the disadvantage of qPCR?

A
  • 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
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46
Q

What is site directed mutagenesis used for?

A
  • for intro of specific mutations into DNA (see if protein changes w/ AA)
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47
Q

What are the 3 steps of site directed mutagenesis?

A
  • mutant strand synthesis
  • DpnI digestion of template
  • transformation
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48
Q

What primers are designed for site directed mutagenesis?

A
  • primers designed in both directions w/ 1 deliberate mismatch
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49
Q

What is the cycle threshold in qPCR?

A
  • point fluorescence level enters exponential phase (and exceeds background level)
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50
Q

What is Sanger sequencing?

A
  • variation of PCR technique
  • DNA synthesis reaction
  • allows seq of 1000-1500 bp of good quality seq
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51
Q

What do you need to know for Sanger sequencing, and why?

A
  • length of fragment and base at last position, can deduce seq by adding 1 letter at a time
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52
Q

How does chain termination occur?

A
  • 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
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53
Q

What are the components of sequencing reaction? (Sanger sequencing)

A
  • 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)
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54
Q

How can fluorescence detection used in Sanger sequencing?

A
  • 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
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55
Q

What are the limitations of Sanger sequencing?

A
  • can only seq 1 template at a time

- need some knowledge of seq to design primer

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56
Q

How is Sanger sequencing diff now and what is it used for?

A
  • now automated

- used to seq plasmids, PCR products etc.

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57
Q

What is next gen seq (NGS) used for?

A
  • genomes

- expression profiles

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58
Q

What are some of the common features of NGS technologies?

A
  • millions of reactions run in parallel
  • reactions spatially separated
  • no seq knowledge of template req, can seq from adaptor into unknown
59
Q

What is GIbson Assembly?

A
  • 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
60
Q

What are the limitations of traditional cloning?

A
  • restricted to single insertion per cloning cycle
  • inefficient for large inserts
  • plasmid design can be restricted by restriction site availability
61
Q

What are the benefits of Gibson Assembly?

A
  • 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
62
Q

What are the 5 principles of Gibson Assembly

A
  • imagine plasmid design
  • design primers
  • PCR
  • assembly
  • transformation
63
Q

What occurs during imagining plasmid design? (Gibson Assembly)

A
  • place insert(s) into vector back bone
  • draw out seq of vector and insert
  • draw out plasmid seq to prod
64
Q

What occurs during design of primers? (Gibson Assembly)

A
  • 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’
65
Q

What occurs during PCR? (Gibson Assembly)

A
  • cut bands from gel and extract linear DNA fragment

- now have overlapping linear DNA for whole plasmid inc vector backbone

66
Q

What occurs during assembly? (Gibson Assembly)

A
  • 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
67
Q

What occurs during transformation? (Gibson Assembly)

A
  • 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)
68
Q

When is Gibson Assembly most useful?

A
  • when used to gen batches of related constructs, eg. adding fluorescent tag to all genes of interest
69
Q

What is Golden Gate Assembly (Type IIS)?

A
  • restriction enzyme and ligase cloning
70
Q

What are the advantages of Golden Gate Assembly?

A
  • 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
71
Q

What are the disadvantages of Golden Gate Assembly?

A
  • 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
72
Q

What are principles of Golden Gate Assembly? eg. BsaI

A
  • 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
73
Q

What is gene expression?

A
  • prod of functional RNA or protein from genetic info encoded by genes
74
Q

How can gene expression be analysed at the level of transcrip?

A
  • look at level of transcrip by analysing mRNA levels
75
Q

What does differential expression of genes result in?

A
  • diff cells, tissues and organs
76
Q

What is hybridisation?

A
  • 2 single NA strand coming together and binding
77
Q

What do hybridisation techniques involve?

A
  • 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
78
Q

How is northern blotting carried out?

A
  • extract RNA and resolve through gel

- transferred to membrane, then hybridised w/ gene specific probe

79
Q

What do results of northern blotting show?

A
  • 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
80
Q

How is in situ hybridisation carried out?

A
  • involves binding of labelled probe to thin slice of fixed tissue
  • FISH can show specific mRNA accum patterns of tissue, organ or whole organism
81
Q

What are microarrays?

A
  • DNA microarrays monitor expression of 1000s of genes simultaneously
  • describes expression of particular organ/tissue
  • use hybridisation on large scale
82
Q

How are microarrays carried out?

A
  • 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)
83
Q

What is a southern blot?

A

-probe DNA blot w/ labelled DNA probe

84
Q

What is the difference between the 3 blotting techniques?

A
  • northern = probe RNA blot w/ labelled DNA probe
  • southern = probe DNA blot w/ labelled DNA probe
  • western = use protein-specific antibodies to detect proteins
85
Q

How can gene expression be analysed at level of transcrip?

A
  • 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
86
Q

How is western blotting carried out?

A
  • 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
87
Q

Why does SDS PAGE have smaller holes than agarose gel?

A
  • proteins smaller
88
Q

Why were engineered fluorescence proteins needed?

A
  • needed more colours to look at more than 1 protein at once
89
Q

What are reporter genes and what are they used for?

A
  • to analyses gene expression
  • known genes whose RNA or protein levels can be measured easily
  • commonly used allow protein visualisation in vivo
90
Q

What is co-localisation and why is it needed?

A
  • 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
91
Q

Why are tool for analysing protein interactions needed?

A
  • 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
92
Q

What is chromatin immunoprecipitation (ChIP) and how is it carried out?

A
  • 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)
93
Q

What is a pull-down assay and how is it carried out?

A
  • 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)
94
Q

What is yeast 2-hybrid and how is it carried out?

A
  • ‘bait’ fused to DNA binding domain
  • prey fused to activating regions
  • if bait and prey interact, reporter gene switched on
95
Q

What are DNA libraries a resource for?

A
  • gene cloning
96
Q

What are DNA libraries?

A
  • collection of diff genomic DNA fragments into same vector type
97
Q

What is needed for a library to have good coverage?

A
  • contain several genome equivalents (GEs)
98
Q

How are DNA libraries constructed?

A
  • 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
99
Q

What are the types of libraries?

A
  • genomic
  • cDNA (easier for large genomes)
  • seq (M13 phage as vector
100
Q

How are genomic libraries made?

A
  • 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
101
Q

How is the correct enzyme chosen to use to digest genome?

A
  • must give approp size fragments and gen sticky ends complementary to vector
102
Q

For humans the no. fragments is too large for a genomic library, so what is used instead?

A
  • diff vector

- cDNA library

103
Q

What are YACs and why are the used?

A
  • 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
104
Q

When are cDNA libraries used?

A
  • for looking at expressed genes
105
Q

What needs to be done in cDNA libraries before vector can be cloned?

A
  • need to copy ss RNA into ds DNA
106
Q

What are the main steps of generating a cDNA library?

A
  • extract RNA from cells of interest
  • 1st strand synthesis
  • 2nd strand synthesis
  • cloning PCR product into vector
107
Q

What happens during extraction of RNA from cells of interest? (generating a cDNA library)

A
  • 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
108
Q

What type of RNA is most abundant?

A
  • rRNA

- mRNA only ~5%

109
Q

What happens during 1st strand synthesis? (generating a cDNA library)

A
  • 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)
110
Q

What happens during 2nd strand synthesis? (generating a cDNA library)

A
  • 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
111
Q

What happens during cloning of PCR product into vector? (generating a cDNA library)

A
  • cDNA treated w/ methylase to protect endogenous EcoRI site from digestion
  • linkers digested w/ EcoRI, cDNA protected by methylation
112
Q

How is method to find clone containing gene of interest decided?

A
  • each colony contains many copies of plasmid containing unique DNA insert
  • screening approach chosen based on info available about gene
113
Q

What are the methods for finding clone containing gene of interest?

A
  • 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
114
Q

How is screening by hybridisation carried out?

A
  • 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
115
Q

How is the probe for hybridisation chosen?

A
  • 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
116
Q

How is a synthesised oligonucleotide probe made? (hybridisation)

A
  • 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
117
Q

How does random priming work? (hybridisation)

A
  • 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
118
Q

How do different temps and salt concs affect colony hybridisation?

A
  • for perfect match salt low and temp high

- for some mismatches salt high and temp low

119
Q

How are expression libraries screened and when are they used?

A
  • 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
120
Q

How is immunological screening of cDNA libraries?

A
  • 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
121
Q

How is gene expression and function studied in mice?

A
  • transgenic mice
  • inducible systems
  • gene targeting
122
Q

Why do mice make good models?

A
  • mouse and human genomes show ~90% synteny
  • directly homologous mouse genes for 99% + human genes
  • most protein seqs highly conserved
123
Q

What does transgenic mean?

A
  • organism that carries transferred genetic material (transgene), inserted at random site in genome
124
Q

What is gene targeting?

A
  • disruption or mutation of particular gene (eg. knockout)
125
Q

How can transgenics be generated via micro-injection into fertilised eggs?

A
  • transgenic mice successfully integrated and expressed rat growth hormone
  • gain of function model
126
Q

How can transgenics be generated via pronuclear micro-injection into fertilised eggs?

A
  • mice will be hemizygous and may be mosaics

- prod of stable transgenic lineage req 2nd inbred gen

127
Q

How is transgene expression analysed?

A
  • 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
128
Q

What are the limits of early transgenic tech?

A
  • limited to gain of function studies
  • random insertion
  • no control over expression
  • tissue specific –> spatial control of transgene
  • inducible promoter –> temporal control of transgene
129
Q

How can expression be analysed at level of transcrip?

A
  • northern blots
  • RT-PCR
  • in-situ hybridisation
130
Q

How can expression be analysed at level of translation?

A
  • western blots
  • immunohistochemistry
  • GFP expression
131
Q

Why do inducible systems exist?

A
  • if intro of transgene prevents or limits survival in utero (embryonic lethal)
  • can delay expression of transgene
132
Q

How do inducible systems work?

A
  • use of inducible transcriptional activator

- expression reg in reversible and quantifiable manner

133
Q

What is gene targeting?

A
  • 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
134
Q

What are the types of gene targeting? (knocking)

A
  • knock out = inactivate gene
  • knock in = exogenous gene introd to disrupt targeted endogenous gene
  • knock down = gene expression decreased
135
Q

What does the targeting vector contain? (gene targeting)

A
  • selectable marker
  • regions of homology w/ mouse chromosome
  • planned mods that alter expression of targeted gene
136
Q

What is a commonly used selection marker? (gene targeting)

A
  • neomycin resistance gene

- allows use of neomycin to kill other cells

137
Q

What knock out mice model resulted in adult-onset obesity in mice?

A
  • targeted deletion of neuronal basic helix-loop-helix transcrip factor 2 (NhIh2)
138
Q

What are mice libraries?

A
  • 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
139
Q

What are “floxed” mice?

A
  • site specific recombination to remove targeted gene
140
Q

Why were “floxed” mice created?

A
  • to examine gene only in context of particular organ system or cell type
  • to avoid embryonic lethal mutants
141
Q

How are “floxed” mice created?

A
  • “cre-lox” system

- cre recombinase combines w/ recombinase recognition sites (LoxP sites)

142
Q

How is cre-lox strain made?

A
  • 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
143
Q

How is M13 phage used as a library vector?

A
  • 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
144
Q

What are the steps in cleavage of DNA by restriction enzyme?

A
  • nonspecific binding
  • sliding/hopping/jumping
  • specific binding
  • coupling
  • catalysis
  • product release

MBB164 (7 decks)

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