deck_4986910-2 Flashcards
(139 cards)
1
Q
what are 7 advantages to using a mouse as a model organism?
A
- easy to breed and house ( but expensive) - easy to control the environment that the mice are in - physiological similarities with humans - well developed physiological techniques for mousegenetically homogenous (inbred) strains available - deposition of the vaginal plug - strains can be used across labs across the world
2
Q
when do mice reach sexual maturity?
A
around week 4 post birth
3
Q
what is the litter size of mice?
A
up to 10 pups
4
Q
is the physiology of the mouse strain dependent?
A
yes
5
Q
what about the genetics of the mouse makes them suitable for human studies?
A
all human genes have counterparts in the mouse genome. thus cloning of a human gene leads directly to cloning of a mouse homologue which can be used for genetic, molecular and biochemical studies that can then be extrapolated back to an understanding of the function of the human gene. In only a subset of cases are mammalian genes conserved within a genome of the flies of worms
6
Q
what is the most commonly used mouse strain?
A
C57BL/6
7
Q
why is it important to keep buying new mice from your original source when keeping a mouse strain?
A
it is important to keep repopulating your strain with founders because genetic drift can occur within the populations causing th strain to diverge from the original, meaning that there is less similarity between strains used by other labs and less reliable and comparable phenotypes therefore
8
Q
why is it important to always get the same strain from the same supplier even though they are the same strain?
A
because there can always be divergence between two sub strains due to genetic drift
9
Q
why is homogeneity so important in mouse strains?
A
normal development and physiology can vary significantly from one strain of mice to the next and in the analysis of, mutants, it is often not possible to distinguish subtle effects due to the mutation itself, from effects dye to other genes within the background of the mutant strain. to make this distinction, it is essential to be able to compare animals with the same genetic background. Phenotypic differences that persist between these a mutant and a wild type from a strain must be a consequence of the mutant allele.
10
Q
what species of mouse are normally used in mouse experiments?
A
musculus musculus domesticus
11
Q
how many generations of inbreeding do you need to ensure the strain is 98.7% isogenic?
A
20
12
Q
how many generations of inbreeding do you need to do to ensure your strain is 99.9999% isogenic?
A
150
13
Q
what is important to remember about different strains (inbred lines) of mice that are used. give 2 examples of this.
A
they all have different characteristics and are different genetically. For example the C57BL/6 is seizure resistant so epilepsy analysis in this mouse would not be good. The C3H has poor spatial learning so experiments relating to this would not be suitable
14
Q
what is the C3H inbred line of mouse bad at?
A
spatial learning
15
Q
what is the C57BL/6 mouse resistant to?
A
seizures
16
Q
in an ideal world, how would mutations occur in mouse strains and why?
A
they would occur spontaneously in strains, the way you would know that the phenotype observed from the mutation would only be due to it, and not from any other genetic background
17
Q
Due to the fact it is unlikely for interesting mutations to occur spontaneoulsy, how can mutations be induced and what is the protocol for ensuring the mice are coisogenic?
A
You can induce mutations and then either view them in heterozygous animals if they are dominant, or in homozygous animals if they are recessive. They can then be continuously inbreed to create its own strain that is homozygous at all loci. Or if it cannot be maintained as a homozygous then it can be maintained as a heterozygous by continuous backcrossing into the original strain. In both cases they are coisogenic meaning that they are genetically identical to the sister strain expect at the mutant loci
18
Q
what is genetic drift?
A
the constant tendency for genes to evolve. This is the introduction of new alleles into the strain. This can result in one strain being developed into many different strains.
19
Q
after how many generations of a colony being separated from its parent colony should it be considered a sub colony and what is the mechanism behind this?
A
20 generations. This is due to genetic drift
20
Q
how common are spontaneous mutations?
A
5 X 10^-6 per gene per generations
21
Q
how many nucleotides are there in the mouse genome?
A
3 x 10^9 nucleotides
22
Q
after how many generations is there a 90% chance that two substring differ at one or more locus?
A
16
23
Q
after 16 generations, what is the probability that two substring differa t one or more loci?
A
0.9
24
Q
how can substring arise from bad strain maintenance? (2)
A
substring can arise fi there is residual heterzygosity within a strain caused by incomplete inbreeding at the time of separation from progenitor strain. This means that not all loci were homozygous for an allele and so the diffferent allele could replace the normally and cause a sublime. this is why it is important to ensure inbreeding is carried out extensively. 2. substring can also arise from genetic drift if the strain is not constantly replenished by the progenitor strain
25
how do you maintain in bred lines?
you cross brother and sisters
26
how can you get rid of spontaneous mutations which could eventually be incorporated into genetic drift and cause the formation of a substrain?
Once you have identified a mouse that has a phenotypic mutation, you then "inheritance test" them. This involves backcrossing the mice and scoring the ratios of the offspring. if the homo mouse is crossed with a het then the ratio will be different than if it was crossed with a homo non mutant (all wild type if recessive). Once identified, you then get rid of the het rogue mice.
27
what is another word for inheritance testing?
test crossing
28
although spontaneous mutations can be damaging, how can they also be useful? give 3 examples
They can be useful if they give rise to interesting phenotypes. reduced insulin secretion, impaired memory and retinal degeneration have manifested themselves in spontaneous mutations
29
how can you reduce genetic drift?
obtain mice from a reputable source, maintain good records to enable mutations to be bred out, cryropreserve a large number of embryos from your colony and return them every 10 generations to replenish, replenish from the original source
30
what is quantitative genetics?
it is the study of phenotypes that are continuous rather than discrete and the genes associated with them
31
what are the 3 approaches to quantitative genetics in rodents (and their subtypes is relevant)?
- artificial selection - inbreeding - crosses (recombinant inbred strains, consomics, outbred (heterogeneous stock).
32
what is the process of artificial selection when carrying out quantitative genetic study?
- you choose a trait of interest and repeatedly mate animals with that trait of interest (for example, long and short tail length). You then end up getting a segregated population. You can then use sequence the animals (how) and identify differences in alleles or point mutations when you compare. This should allow you to identify regions of the genome that contain genes for the trait of interest
33
how can recombinant inbred strains be used to identify genes for traits of interest?
recombinant strains are generated by crossing two or more strains that are very polymorphic. due to recombination events, F2s are inbred repeatedly for about 20 or more generations and produce a strain that is a random mixture of the different strains. different F2 crosses are then used to produced different strains. Traits that are restricted to certain IRS and not others are of interest. The genome of the strains are compared and SNPs that are exclusive to the trait in interesting strains are considered to be within areas of the genome where the gene for the trait is. The sequence of SNPs will belong to one of therapist strains and can be identified via sequencing. The area of interest can then be looked at the candidate genes can be proposed. These can then be subject to further testing
34
what are the downsides of using recombinant inbred strains?
it is very hard to refine the candidate site down to a sufficiently small size that candidate genes can be extrapolated very well- specific genes being derived from these studies are very unlikely
35
what is a recombinant inbred strain?
an organism with chromosomes that incorporate an essentially permanent set of recombination events between chromosomes inherited from two or more inbred strains. F1 and F2 generations are produced by intercrossing the inbred strains; pairs of the F2 progeny are then mated to establish inbred strains through long-term inbreeding.
36
how do you produce inbred recombinant strains
you cross two strains that produce a heterozygous F1 that has no recombination between the two strains. You then cross these F1s together and produce an F2 which is recombinant. You then take two F2s and inbred for generations. eventually, the strain will be homogenous at all loci. after around 20 generations.
37
what is a consomic strain?
chromosome substitution strain- a strain that is homogenous to its sister strain bar one chromosome.
38
what are consomic strains used for?
they are used pinpoint which chromosomes are important for certain phenotype- whether a trait is encoded predominately by a single chromosome.
39
how are consomic strains formed?
you identify many SNPs that are present on a single chromosome. around 233 micro satellite markers have been used before. You can backcross f1 females arising from a interstrain cross, into one of the strains, and continuoulsy check for the SNPs or microsatellites and select this for the backcross, and so on and so forth for 7-16 generations.
40
what is a congenic strain?
a strain that is isogenic to its sister strain bar a region containing the gene which encodes the phenotype of interest
41
how are congenic strains formed and how can it be used to identify genomic regions of interest?
generated in the laboratory by mating two inbred strains (usually rats or mice), and backcrossing the descendants 5-10 generations with one of the original strains, known as the recipient strain. Typically selection for either phenotype or genotype is performed prior to each backcross generation. So you us this to map by backcrossing until you have the entire genome of one strain bar the region which contains the gene of interest- you can then look for SNPs that are not associated with the majority strain and are instead related to the other strain and you will know this is where the gene is interest is. you then input this into a database of the mouse genome and identify which genes lie in this region
42
after how many generations of back crossing is a strain considered congenic?
10 generations
43
what are the requirements in terms of the types of strains that can be used when forming a recombinant inbred strain for quantitative genetic analysis?
the strains used must be polymorphic and significantly different to each other
44
describe how you would make a congenic strain for a mutation that was recessive or one that was dominant.
following the crossing of two strains, the differing phenotype may be able to be viewed in the F2. If this is the case, then you can carry out SNP PCR analysis and find out which regions of chromosomes is linked to the phenotype. You then carry out progressive back crosses but select for the genotype of phenotype of interest each time. It may be easier to select for the genotype as then you dont have to expose the mice to whatever it is that you are looking for. you then repeatedly backcross the heterzygotes. Once this has occurred you can eventually keep the allele as a homozygous by sister brother mating after 10 back crosses. 2. if the trait is homozygous then you can also hopefully view in the F2 in 25% of offspring. You can then use SNP PCR to identify the region and then continue to backcross as heterozygous if you can detect the QTL every time.
45
Is it preferable to maintain congenic strains in het or homo and why?
in het because then ever time you backcross the heterozygosity decreases by 50%. also this may not be a choice if the allele can't be maintained as a homozygous if it is lethal or causes sterility etc.
46
what is the mendelian principle behind congenic backcrosses?
for every backcross, the heterozygosity should decrease by 50% at all loci that aren't linked. This means that after 10 generations, the strain will be 99.8% isogenic for the recipient strain. But, due to linkage, after 10 generations of back crossing there will be 20cM of linked donor strain.
47
in terms of making a congenic strain, after 10 generations of back crossing, what percentage will be isogenic to the recipient strain and what will the length of linked donor strain DNA?
99.8% and 20 cM of linked
48
what 2 things should always be considered when comparing congenic to coisogenic strains?
First, congenic strains, especially those that have undergone only a minimum number of backcrosses, will have small random remnants of the donor strain — so-called passenger loci — scattered throughout the genome. In congenic strains maintained by inbreeding, the same passenger genes will be present in all members of the strain. In rare instances, traits attributed to the selected donor allele may actually result from one of these cryptic passenger genes. Such effects can be sorted out by breeding the congenic strain back to its original inbred partner. If a trait is due to a passenger gene, it will assort independently of the donor locus in subsequent backcrosses.The second difference between a congenic strain and a coisogenic strain is in the chromosomal vicinity of the differential locus. Congenic strains will always differ from their inbred partner along a significant length of chromosome flanking the differential locus; coisogenic strains will only differ at the differential locus itself and nowhere else. Thus, there is always the possibility that phenotypic differences between the two members of a congenic pair are actually caused by a closely linked gene rather than the selected differential locus. This potential problem is much more difficult to resolve by simple breeding protocols.
49
what is the process of genotyping SNPs using PCR?
the process relies on allele specific primers. You use for primers, two outer primers (one 5' up from the SNP site on the 5' to 3' strand and one 5' up from the SNP site in the 3' to 5' strand) You then have two inner primers, one that matches one SNP allele on the 5' to 3' strand and another which matches the other SNP allele on the 3' to 5' strand. When you PCR with all of these primers and with the two alleles, you will get a non-speific allele product which is very long and connects the two outer primers. You will also get two products, one for each allele, they are designed in such a way that the length is easily distinguishable
50
draw a picture of how SNP genotyping works when using PCR primers
http://bmcbioinformatics.biomedcentral.com/articles/10.1186/1471-2105-9-253
51
how do you genotype SNPs?
using PCR primer technique
52
what are heterogenous stocks?
you can create recombinant inbred strains from 8 different strains
53
what is the advantage of using more strains when creating recombinant inbred strains?
in enables high resolution link mapping. 2cM between each recombinant
54
when you use 8 strains for RIS, what is the distance between the recombinants?
2cM
55
If you you want the highest resolution linkage mapping using inbred strains, what strains can you use?
outbred stock. These are colonies descended from a small number of inbreds. This means that 9%% sequence variants.
56
what percentage sequence variance do you have when using outbred stock?
95%
57
what is the mendelian principles behind the resulting homozygous it that results from sister brother mating of F2s during the production of recombinant inbred strains?
there is a 12.5% chance that both F2 progenitors are identically homozygous at any one locus, then approximately 12.5% of all loci in the genome will fall into this state at random. The consequence for these loci is dramatic: all offspring in the following F3 generation, and all offspring in all subsequent filial generations will also be homozygous for the same alleles at these particular loci. Another way of looking at this process is to consider the fact that once a starting allele at any locus has been lost from a strain of mice, it can never come back, so long as only brother-sister matings are performed to maintain the strain.At each filial generation subsequent to F3, the class of loci fixed for one parental allele will continue to expand beyond 12.5%. This is because all fixed loci will remain unchanged through the process of incrossing, while all unfixed loci will have a certain chance of reaching fixation at each generation. At each locus which has not been fixed, matings can be viewed as backcrosses, outcrosses, or intercrosses, which are all inherently unstable since they can all yield offspring with heterozygous genotypes- after 20 generations, 98.7% of the loci in the genome of each animl should be homozygous, at 40 generations, 99.98% will be homozygous. However, chromosomes do not assort randomly and some are linked- inherited in junks. e, there is an ever-increasing chance of complete homozygosity as mice pass from the 30th to 60th generation of inbreeding (Bailey, 1978). In fact, by 60 generations, one would be virtually assured of a homogeneous homozygous genome if it were not for the continual appearance of new spontaneous mutations (most of which will have no visible effect on phenotype). However, every new mutation that occurs will soon be fixed or eliminated from the strain through further rounds of inbreeding. Thus, for all practical purposes, mice at the F60 generation or higher can be considered 100% homozygous and genetically indistinguishable from all siblings and close relatives
58
what are the downsides of congenic strains?
infertility and bad health can occur
59
why do some people use outbred stocks for quantitative trait mapping
Outbred mice are used for the same reasons as F1 hybrids — they exhibit hybrid vigor with long life spans, high disease resistance, early fertility, large and frequent litters, low neonatal mortality, rapid growth, and large size. However, unlike F1 hybrids, outbred mice are genetically undefined. Nevertheless, outbred mice are bought and used in large numbers simply because they are less expensive than any of the genetically-defined strains.Outbred mice are useful in experiments where the precise genotype of animals is not important and when they will not contribute their genome toward the establishment of new strains. They are often ideal as a source of material for biochemical purification and as stud males for the stimulation of pseudo-pregnancy in females to be used as foster mothers for transgenic or chimeric embryos. It is unwise to use outbred males as progenitors for any strain of mice that will be maintained and studied over multiple generations; the random-bred parent will contribute genetic uncertainty which could result in unexpected results down-the-road.
60
when creating transgenic mice, why is it important to do the same experiment in 3 or more founder lines?
potential problem can result from the insertion of the transgene into a normally-functioning endogenous locus with unanticipated consequences. In approximately 5-10% of all cases studied to date, homozygosity for a particular transgene locus has been found to cause lethality or some other phenotypic anomaly. (Palmiter and Brinster, 1986). These recessive phenotypes are most likely due to the disruption of some normal vital gene. In less frequent cases, a transgene may land at a site that is flanked by an endogenous enhancer which can stimulate gene activity at inappropriate stages or tissues. This can lead to the expression of dominant phenotypes that are not strictly a result of the transgene itself. 39 For all of these reasons, it is critical to analyze data from three or more founder lines with the same transgene construct before reaching conclusions concerning the effect, or lack thereof, on the mouse phenotype.
61
what is the process of calculating linkage using markers?
...
62
what is a quantitative trait locus?
an area of the genome that's been implicated in containing a gene that of interest for a particular trait
63
what are the most commonly used two strains for recombinant inbred mice strains?
C57BL/6J and DBA/2J
64
describe the drug and alcohol sensitivity experiment using recombinant mice
..
65
how can reduce the number of backcrossing generations needed for congenic production from 10 to 5?
by not only mapping for the SNP of interest but also mapping for the greatest loss of the donor strain
66
what does ENU do?
The chemical is an alkylating agent, and acts by transferring the ethyl group of ENU to nucleobases (usually thymine) in nucleic acids. It changed GC pairings to AT pairings and AT to GC pairings
67
who determined the mutation rate of ENU?
Bill russel
68
how did bill Brussel work out the mutation rate of different mutagens?
he used a mouse from the T-stock which have 7 mutations across different chromosomes. They are all recessive mutations. He then applied the mutagen to wild type mice and then crossed with these T -stock. If the mutagen mutagenised one oft the loci then then the phenotype would be exposed in the cross. He could then work out the rate of mutation rate per locus.
69
what strain did bill russel use to estimate the mutation rate per nucleus for different mutagens?
t-stock- 7 recessive mutant alleles loci with phenotypes
70
what is the most powerful mutagen? (rate)
ENU fractionated dose (1/750)
71
list the success of mutagens in order
ENU fractionated dose, ENU, procarbazine, x-irradiation, sponatenous
72
does the mutation rate vary from locus to locus?
yes
73
how did bill Russel use his t-stock experiment to work out the types of mutation that ENU and x-irradaition induced?
Two of the 7 loci in the strain were present on the same chromosome (9), he noticed that when irradiaition they were always both present and they were separate when used ENU. This was because irradiation caused deletions but ENU cause base pair mutations
74
what are the 5 classes of genetic mutation?
amorph, hypomorph, hypermorph, antimorph, neomorph
75
what is an amoprh?
complete loss of function (null), generally recessive unless haploinsufficient
76
what is a hypomorph?
partial loss of function, usually recessive,
77
what is a hypermorph?
increase a normal gene function, normally dominant or semi dominant,
78
what is an antimorph?
dominant negatie opposes normal function
79
what is a neo morph?
dominant and gives a completely new function
80
what is an allelic series?
Allelic series describe different mutant alleles of a gene that cause a range of phenotypes, whereby each one carries a single point mutation within different regions of the same gene. They uncover new functional domains
81
give an example of ENU uncovering an allelic series
the trembler (PMP22) locus,a point mutation in the transmembrane section caused a very severe loss of myelin phenotype whereas the intracellular membrane mutation cause a less severe phenotype
82
what is a nonsense mutation?
a mutation which results in a the insertion of a premature stop codon
83
what is a missense mutation?
a mutation which results in a different amino acid
84
how can ENS mutations induce the same phenotypes as knockouts?
- missense- nonsense - splice site mutatnt - mutations in promoters and other control elements - dominant negative mutations
85
when can ENU phenotypes be preferential to knock out phenotypes and why?
when the knockout produces embryonic lethality but the ENU doesn't because it only targets a specific functional domain rather than the entire gene.
86
what is the process of ENU mutagenesis?
• Inject male mice with ENU• Fractionated dose more efficient (3 x 100 mg/kg)• ENUinducestemporaryperiodofsterility• New germ cells re-populate testis• Sperm carry independent mutations• Generation of approximately 50 progeny per injected male
87
what is the process of a dominant screen strategy?
there is a single founder individual (male of female) that is mutagenised and then crossed with a wild type, you then screen the F1. Conduct whole-genome scan of affected individuals, Successive out crossing of affected individuals for fine mapping (more recombination events), Candidate gene(s)/ sequencing
88
what are the two options for carrying out a screen for recessive mutant alleles?
cross a male mouse that has been mutagenised with a female normal mouse. This will result in each F1 mouse being heterozygous for a different mutation. So each F1 male mouse is then crossed with a wild type and produces a variety of offspring, some will be heterozygous. The F2s can then either be crossed back with the het F1 or they can be intercrossed (inbred or backcrossed) to produce F3s, some of which will be homozygous mutant. If you have identifies G3 mutants, you can then cross the G2 or G2/G1s again and get more mutants
89
how have dominant screen strategies been revolutionised?
Once you have performed a few backcrossed of your identified mutant, you dont have to do lots of linkage mapping, you can simply send it off for next generation sequencing and they will identify potential missions mutations.
90
what are the differences between the procedure for a dominant screen and for a recessive screen?
- you have to screen more animals with recessive screens- recessive screens require an F3- more expensive and time consuming
91
what is a modifier screen and give an example
when you carry out a screen on a certain genetic background. for example you could have a mouse that is a carrier for a huntington disease and then you would cross with a mutagenise mouse and then look for F1s that have a phenotype that modifies the carrier phenotype- so for example fives a later or earlier insert than normal
92
are modifier screens difficult to carry out in mice?
yes
93
what are challenger screens?
this involves challenging the mutant with environmental stresses, drugs, disease or ageing etc and then carrying out a screen to see if there is a progression of phenotypes with age or under the other stresses- then do congenics and mapping etc
94
what is a sensitised screen?
where you carry out a screen with animals that have a predisposition for a certain disease- you would do this to see what other genes may be acting to produce diseases
95
what are region specific saturation screens?
where you look for all potential mutations within a small region of genome. This is done using various mutations that have
96
how can you carry out a saturation screen for a certain loci?
You use a mouse that you already have a strain containing a deletion for that gene such as the Del36H deleted region. you expose a mouse to a mutagen and then cross it with another inbred strain. This gives you a het for the mutation and one normal section from the other strain. You then cross this het with a mouse that is het for a deletion at this section. This will give rise to some of the animals in the G2 expressing a recessive phenotype for mutant section as there is a deletion in the other chromosome. This is easier than producing G3 homozygotes and guessing that it is at the same loci because of a similar phenotype- this way you know it is at the deleted loci and it is in the G2.
97
what is the aim of a saturation screen in mice?
Saturation screens aim to identify every possible mutant phenotype within a particular deletion region
98
what are gene driven screen ?
identifying mutations in a specific gene
99
how would you carry out a gene driven screen?
There are sperm banks containing sperm of 10,000 mutagenise mice. if you were interested in a paritcular gene then you design primers for the gene of interest and then amplify the DNA from this particular gene for 10,0000 mutagenised mice and then look for mutations. The number of DNAs screen (mutagenised animals) the more likely you are to find a mutation of the allele- provided that the mutation rate is 1/1000 per locus and the mutation detection rate is 90% then you will find 4 or more alleles with a porbability of 0.9% if you screen 5000 animals:
100
Why would you want to do a gene driven screen? give an example?
If you know that the knock out is lethal you may want to screen for alleles that have mutations in certain domains of the gene. This was used to look at the alleles for the L-type voltage-dependent calcium channel which normally produced cardiac deficits. They wanted to look for hypomoprhic mutations. The Ko was embryonic lethal- so wanted to look for hypomorphic mutations- screened for mutations in 2 regions of the gene- it is an intracellular protein- so they screened for certain exons which they thought would give hypomorphic phneotype- screened for exon 6 which is an rea important for the interaction with the beta subunit of the complex and another region which is inolved in the activation of the channel- using this ENU gene driven screen they found 1 missense and one nonsense (stop) mutation they then resurected the two mouse lines which contained these mutations (using the sperm) and then screened them for phenotypes
101
what is an additive transgenesis?
increase gene dosage or adding a reporter gene
102
how can you produce a loss of function mutation using transgenics? (4)
use homologus recombination to replace DNA with a selection marker or a mutated form of the gene.- gene trap - conditional KO
103
how do you produce transgenic mice?
via pronuclear injection or by homologues recombination in an ES cell
104
how do you carry out transgenics using pronuclear injection?
you inject into a single fertilised cell into the male pronucleus. You can construct any piece of DNA and put it into a vector, then linearise your vector and then inject this into the nucleus and it will randomly insert into the genome. You then implant this egg into a surrogate mouse and one of the offspring will be het for the transgene. You can do DNA analysis to identify which of the offspring contains the construct by using primers within the transgene which will produce a known length of DNA (should normally span the promoter gene junction)
105
what are the downsides of random insertion of the pronuclear injection approach?
- if there is a promoter in the construct then it can affect other gene expression which could cause a phenotype that isn't ue t the expression of your transgene - it could be inserted into a low expression region - it could be inserted into a gene and disrupt the expression of that gene and cause a phenotype unrelated to the injection of your construct- for small DNA constructs you can get tandem insertions so you need to analyse how many genes have inserted and where
106
what is a BAC?
bacterial artificial chromosome- - large pieces of DNA. BAC libraries exist for the human, mice and other species.
107
what are BACs used for?
- they are used to create large constructs inside e.coli. This involves recombining a construct such as one being used for homologues recombination inside an ES cell . This is done inside an e.coli and then the BAC can be removed, the construct removed from the BAC and then injected into the ES cell. - BACS can be used to inject into pronucleus of mice embryos. There are BAC libraries available for the mouse, they can be engineered to put reporters in to report gene expression well. They are linearised and then injected
108
what is the good thing about using BACs?
large and faithful expression of gene or reporter
109
what are the downsides of BACs?
very large and may contain other genes so you also get expression of this gene too which can interfere
110
how can you produce knock out mice?
you can produce a construct with homology arms to the area around the gene you want to kick out, then you replace the gene of interest with a resistance gene that allows the successfully transfected ES cells to be identified. Then you inject these ES cells into a blatsocyst.
111
how do you create a knock in mouse?
same as with the knock out mice but you put an altered allele into the gene area instead but you have also have a resistance gene with its own promoter too. you could then potential surround this resistance gene with a loop site so that it can be flexed out via gene trapping mechanism.
112
how can you test that your gene has inserted? and where it is inserted?
checking for for resistance given by the resistance gene. You can check it is inserted by using primer for the SNP within the allele and a primer just outside within the homology arms?
113
if a knock out is embryonic lethal, what can you do?
produce a conditional knock out
114
how do you produce a conditional knock out?
using the cre inducible system and inserting a gene via homologues recombination that replaces the gene of interest with one that has loop sites around it so that it can be knocked out when there is Cre expression which causes it to recombine out.
115
when do knock outs normally cause lethality?
when the gene is pleiotropic and has an essential function during development but others later
116
What can cre be used for in terms of gene reporting?
You can have a reporter downstream of a flowed stop codon insetting into the ROSA26 region. Then Cre can be driven by a gene of interest and this will result in the reporter being expressed whether this gene is expressed or has been expressed- can be used for fate mapping
117
what is the tamoxifen method of cre induction?
rather than having genes stimulating Cre expression, you can express cre everywhere but have it as a cre- oestrogen receptor fusion protein, that when activated ny tamoxifen, will enter the nucleus of the cell and drive recombination
118
what is Cre more commonly used that flp/FRT?
Cre historically was much more efficient than FLP recombinase (FLP thermo-unstable).• Consequently, many deletor strains generated in past have been based on cre.• Cre more adaptable due to mutant lox sites.• However, Flp now improved my Flpe and Flpo versions
119
what is the cre multiplex system? and what is it used for?
It is a modified Cre/lox system that relies upon transgenes that contain three pairs of self-compatible lox sites (loxN, lox2272 and loxP). Recombination has three different possible outcomes, each with the same probability. The use of tamoxifen inducible Cre recombinase allows for recombination to be regulated temporally and enables one to limit recombination to few cells. this can be used in the brainbow for example to generate different coloured cells. You can use this in lineage tracing to map the fate of individual cells over time??
120
what should you always do when you are wanting to use a CRe drive? give an example of when this is needed to be done.
check the pattern of expression by using it to drive a reporter. it could look like it was only expressed in a precise area in adulthood when in reality it was expressed everywhere in development.
121
what are the three things you should consider before going ahead with your Cre experiment?
- measure cre activity - understand quirks of cre driver ?? (checking that the BAC you are using doesn't contain other genes that would complicate results) - inducer delivery and side effects
122
what controls should you do before using a Cre experiment?
Check for phenotype in driver lineCheck pattern of cre activity (may be substrate dependent)Check for variability of activity (leakiness may be in the germ line!)
123
how many matings must you do when you are performing a cre loop experiment and why?
you can't just cross a Cre with a homo loxp because you will only get a het loxp chromosome. You must cross and then cross this one with a homo loxp. This will give you a homo loxp cre
124
is it important that both the Cre and floxed mice are from the same line?
yes
125
what 3 controls should you produce when carrying out a cre/loxp experiment? And how do you make these?
homo loxp, cre and het loxp, just cre and no loxp. first you cross a cre with no loxp with a homo loxp. This gives you a het loxp and cre. Then cross this with a homo loxp and you get your desired mouse and your three controls.
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how do you orchestrate directed gene trapping?
by producing your cassette with homologues arms for the exon either side
127
why would you want to carry out directed gene trapping into certain genes.
To cause a truncated protein and to determine where it is that the gene is being perturbed via the reporter. The reporter shows that the gene trap has worked. Depending on what cassette you use you can induce a mutation via the insertion and then flp is back out to look at the control to show that the cassette is causing the mutation and then you can flp it back into again for another control. you want to report where the mutation is occurring because CRe is probably tissue specifically driven
128
what is an example of a gene trap construct that could be inserted?
FRT site- loxp- SA- B-geo-pA-loxp- FRT
129
what does the inserted poly adenylate site of a cassette do?
prematurely stops transcriptions
130
how can you use a gene trap in conditional knock outs?
you can have a FRT'd resistance gene (neo) and a floxed essential exon. This way you can remove the resistance gene as a control to show there is no artefact. Then you can remove the exon for the knock out
131
Describe the knock out conditional first cassette and the 4 genotypes available from using this, including why you would want each of the genotypes.
the cassette in inserted around an exon that is required for the functioning of a gene. it would be as follows: exon1- FRT-lacZ-loxp-promoter driven Neo-FRT-loxP-exon 2-loxp-exon 4. This would be tm1a. tm1b would occur when crossed with Cre and the neo and exon could be removed (this would result in a labeled truncated proteinto show successful cassette and where the gene is being mutated (Cre will be tissue specific). tm1c would be when crossed with Flp. this would remove marker and neo and leave normal gene expression. (this could be done after you know cassette is successfuly inserted as a control). Then tm1d would occur when crossed again with Cre to cause a mutant without the exon and because you have done the controls you know it can only be down to the removal of the exon and nothing else.
132
what are the three types of gene traps? what is the general thought behind these traps?
enhancer traps, promoter traps and gene traps. The cassettes contain a reporter which will report expression when inserted into the genome.
133
describe the cassette used for an enhancer trap and how it would work.
hsppromoter-lacz-pA-HSV promoter, Neo, pA. When inserted into near a regulatory element, the promoter will be activated and the reporter gene transcribed. and the neo will be transcribed as a separate protein (this is needed for the insertion into the ES cell). This will therefore report an enhancer which can then be identified using 5' RACE
134
describe the cassette used ofr a gene trap and how it works.
SA (splice acceptor)-lacZ-pA-actin promotoer-neo-pA. This will insert into an intron and will induce a truncated protein which will be labelled by the b-gal. The splice acceptor ensures it is attached to the truncated protein. Then the gene that has been perturbed can be identified by 5' RACE
135
describe the cassette used for a promoter trap and how it works.
no-promoter- lacZ pA promoter neo pA. This will insert in frame with a protein and has no promoter so needs to land very close to a promoter. The lacZ can be used to demonstrate protein expression levels. 5' RACE
136
once your gene trap has randomly inserted into a site of interest, how can you then determine where it has inserted?
first you extract the RNA from your organism. Then you put it through electrophoresis, then you apply a probe for lacZ via northern blot. Then you cut out the RNA and perform RNA race : you have the RNA containing the galactosidase gene you use reverse transcriptase to create one strand of cDNA from the RNA and the transcriptse adds 3-5 C’s on the 3’ end of the cDNA> then you add an oligonucleotide which has 3-5 G’s on the 3’ end which will anneal to the Cs of the cDNA with an added known artificial primer site you then use reverse transcriptase to extrend from the C’s and on the primer site you then use primers for the known primer site and for a known sequence within the cDNA (your reporter gene) to amplify the cDNA via PCRthe PCR product can then be sequences and the 5” region of the gene that has been trapped can be identified and compared to the sequenced genome in order to find which gene has been trapped.
137
draw out the process of 5' RACE.
..
138
how can you carry out a knock in using Cre loxp recombination?
A gene of interest (GoI) can be inserted into the ATG position of a targeted gene (Fig. 4), and the GoI is then driven by the endoge- nous promoter of the targeted gene. Design of such a replacement vector is shown in Fig. 4a. Importantly, the selection marker in this vector should not contain a pA signal to provide further selection of the recombination event, as described below. The replacement and Cre expression vectors are coelectroporated into the targeted ES clone in their circular forms. The cre gene is transiently expressed and mediates recombination. Since the replacement plasmid and the targeting vector in the ES genome both carry two lox sites with the same spacer region (RE or LE mutant lox and loxP), it is expected that intramolecular recombination between the two lox sites should occur after coelectroporation, resulting in the production of two intermediate molecules, as shown in Fig. 4b. Integrative recombi- nation then occurs between LE mutant lox sites in the genome and RE mutant lox sites in the intermediate molecule. ES cells in which (RE is the white arrow head, black arrow bod) (Le is the opposite)- SO first the RE or LE site recombine with normal loxP site. then the LE and RE recombine with each other.
139
describe the process of 5' Race
You extract tissue that is expressing the gene trap- you then extract the RNAelectrophores the RNA to seperate by size then transfer RNA to membrane for northern blotting, fix RNA to the membrane by using UV or heat hybiridze membrane with labeled probes for the galactosidase gene which you know is in the gene trap cut out the RNAyou have the RNA containing the galactosidase gene you use reverse transcriptase to create one strand of cDNA from the RNA and the transcriptse adds 3-5 C’s on the 3’ end of the cDNA> then you add an oligonucleotide which has 3-5 G’s on the 3’ end which will anneal to the Cs of the cDNA with an added known artificial primer site you then use reverse transcriptase to extrend from the C’s and on the primer site you then use primers for the known primer site and for a known sequence within the cDNA (your reporter gene) to amplify the cDNA via PCRthe PCR product can then be sequences and the 5” region of the gene that has been trapped can be identified and compared to the sequenced genome in order to find which gene has been trapped.
