Unit 3

To create gRNAs for AAV Cas9 SmartNuclease Plasmids

**look at slides The workflow at-a-glance 1. Design two DNA oligonucleotides that are sense and antisense sequences of the target DNA and are immediately upstream of a PAM sequence (see above section on creating gRNAs for AAV Cas9 SmartNuclease Plasmids) 2. Anneal the two oligonucleotides to generate a duplex 3. Ligate the duplex into the pre-linearized All-in-one AAV Cas9 SmartNuclease Plasmid 4. Transform into competent cells and grow in LB/Kanamycin plate (50 µg/ml) 5. Confirm positive clones by direct sequencing 6. Transfect sequence-verified All-in-one construct into AAV packaging cells 7. Isolate packaged All-in-one AAV Cas9 SmartNuclease Plasmid using AAVanced AAV Concentration Reagent for easy, high-titer preparations Your AAV Cas9/gRNA virus is ready for your genome editing project.

Figure 9.13 The structure of the siRNA.

- Antisense (guide strand): complementary to RNAi - Sense (passenger strand): direct copy of RNA???? --> double check this The active molecule for RNAi is double-stranded siRNA with a two-base overhang at each end, as shown here. The passenger strand is degraded, and the guide strand, which is complementary to the target mRNA remains to make the dsRNA. It seems likely that which strand in an individual siRNA is degraded is random, but that only the ones capable of forming a double-stranded hybrid are recognized and protected. In a subsequent step in the RISC, one strand is degraded to leave the guide strand, which targets RISC to the mRNA by base pairing.

Figure 9.12 Introducing dsRNA for RNAi experiments: RNAi is performed by feeding.

- Most commonly used in c. elegans because transfecting is easy for bacteria and c. elegans eat bacteria - Eats knockdown pool - Bacteria that express the RNAi It will be recalled from Chapter 3 that the worm C. elegans eats E. coli, and molecules introduced into the gut have a systemic effect throughout the worm. The cDNA for the target gene is first cloned into a plasmid that has T7 promoters on each side. This plasmid is transformed into an E. coli strain that has a T7 polymerase gene under the control of the lacZ promoter integrated into the chromosome. The relevant genes on the plasmid (on the left) and the chromosome (on the right) are shown. Both plasmid and chromosome have other selectable markers, which have been omitted for simplicity. The bacteria with the plasmid are induced by IPTG, which results in the production of T7 polymerase. T7 polymerase works on the plasmid promoters and transcribes both strands of the cDNA for the target gene. These two complementary RNA strands can anneal with each other inside the bacterial cell, to make a dsRNA molecule. The bacteria making the dsRNA are spread onto plates, and worms are placed on the spread plates. As the worms eat the bacteria, they ingest the dsRNA, which then induces RNAi in the target gene. The worms reproduce, and some of their offspring exhibit the RNAi phenotype, here shown as a dumpy worm. A library of E. coli strains with each worm gene cloned into the plasmid vector is commercially available, and genome-wide screens in C. elegans are usually done using this feeding method.

Figure 9.8 One postulated mechanism of antisense experiments.

- RNAi: 20 nucleotides that is complementary to mRNA?? - mRNA level stays the same (sometimes can be knocked down) - *Protein level is repressed* A single-stranded nucleic acid, either RNA or DNA or a synthetic molecule, is made that is complementary in sequence to the target mRNA. The single-stranded molecule is shown in red; the target mRNA is shown in blue. This single-stranded molecule is introduced into the cell, where it is postulated to form a double-stranded hybrid with the mRNA. This double-stranded hybrid is then postulated to be blocked in translation, as shown at the bottom of the figure. Although antisense experiments of this type do block gene expression, the actual mechanism is rarely explored. It seems likely that many of these antisense experiments work by triggering RNA degradation via the RNAi pathway.

figure 7.7 The URA3 gene inserted into the TUB2 gene

- TUB2 is an essential gene that is critical for cell division, rare event as selectable marker - plates without uracil will grow mutations with URA3 mutation - can be done to diploid cells The tubulin protofilament shown in Part A consists of α-tubulin and β-tubulin. In yeast, α-tubulin is encoded by the TUB1 and TUB3 genes, while β-tubulin is encoded by the TUB2 gene. In part B, the wild-type URA3 gene has been cloned into the TUB2 gene with flanking regions of TUB2 on each side. This construct is transformed into a ura3 auxtrophic mutant that cannot grow in the absence of uracil. A double crossover between the chromosomal TUB2 gene and the disrupted TUB2 gene, indicated by the dashed lines, replaces the chromosomal TUB2 gene with the disrupted gene. This disruption can be selected by the ability to grow in the absence of uracil. Since TUB2 is an essential gene, the disruption would involve only one of the two copies in a diploid cell; only one copy is shown in the figure, since the other copy of the gene is not disrupted or replaced. Panel A is reproduced from Meneely et al., Genetics: Genes, Genomes and Evolution (Oxford University Press, 2017), Figure 6.4.

Figure 8.17 The structure of an sgRNA

- crRNA and tracrRNA combined The function of Cas9 from S pyogenes requires an additional RNA molecule known as the tracrRNA, as well as the crRNA that locates the position to be edited. For genome editing, these two RNA molecules are often combined into a single long transcript known as the single guide or sgRNA; the structure of such an sgRNA is shown with the phosphate backbone in purple. The sequences of tracrRNA vary, but the diagram uses a sequence common in S pyogenes. The tracrRNA forms a hairpin by intrastrand base pairing; some mismatches occur in the hairpin.

Reverse Genetic Analysis (figure 7.2)

- easier if you know gene of interest - start with DNA, designing gene to be studied - introduce a mutation: promoter (knockout) or change coding sequence (more direct, disrupts reading frame) - most common strategy: create transgene and add it to the system (dominant allele added in) or knock in (replaced WT with mutated novel) - end result: most likely a null allele Often the DNA sequence corresponding to a gene is isolated before the biological role of the gene is known. In reverse genetic analysis, the DNA sequence is altered or edited in vitro, shown here as an insertion into exon 3 that disrupts the reading frame for the coding region. The edited gene is transformed into one or a few cells, and a transgenic organism is grown from these transformed cells.

figure 7.11 targeting vector used for gene disruption

- homologous region needs to be a lot of DNA (8kb from both ends) to allow homologous recombination to occur to get it into the genome - neomycin or G418: resistance gene, selectable marker - HSV-tk: counter selection, sensitive to gancyclovir, needed to target one copy WT gDNA The mouse gene is shown in blue, with light blue representing introns and the dark blue exons numbered. The neoR gene has been inserted into the third exon. The vector also includes the thymidine kinase gene Herpes simplex virus (HSV-tk). The HSV-tk gene confers sensitivity to gancyclovir, whereas the neoR gene confers resistance to neomycin or to its more stable analogue G418. At least 8 kb of homology are needed for homologous recombination between the targeting vector sequence and the chromosomal gene.

Making mouse mutant: obtaining embryonic stem cells (ES cells)/mouse embryogenesis and the generation of the inner cell mass (figure 7.8)

- in order to do knock in/knock outs in mice, you need create ES cells (harvested from mouse embryo) from super ovulated female --> mate mice to be fertilized, at embryogenesis female is sacrificed. harvest uterine horns and separate out each embryos or harvest blastocyst and separate out inner cell mass and put in culture dish (these are ES cells) Ovulation releases an oocyte that moves down the oviduct where it can be fertilized. After fertilization and the completion of meiosis, the embryo begins to divide mitotically. The dividing embryo reaches the blastocyst stage as it enters the uterus, approximately 3-4 days after fertilization, and implantation in the uterine wall follows. An isolated blastocyst is shown at the bottom as a diagram. A large cavity called the blastocoel forms, with the inner cell mass at one side. The cells of the inner cell mass will give rise to all the cells of the embryo and adult mouse

in this case, cre has what kind of promoter? figure 7.16

- insulin promoter - cells that turn on when insulin is needed will also turn on and make cre - pancreas makes insulin - ex. in pancreas we have pten with exon 5 deleted by the two floxed p sites, every time cre is expressed, econ 5 is removed, knocking out pten in pancreas cells

Gene disruptions/Gene replacement by homologous recombination figure 7.4

- make mutation from WT genomic sequence (in vitro) then introduce into the endogenous genomic sequence and replace WT sequence with new mutation (in vivo) - this is done by homologous recombination/crossing over - happens if there is homology at critical points - reporter gene included (GFP) tagged to show which cells have the mutation - can be knock out or knock in Sequences in the genome can be replaced with introduced exogenous DNA whenever a double crossover occurs between the introduced DNA and the genomic target site. This usually depends on the homology directed recombination and repair system in cells. The introduced DNA sequence can then replace the copy in the genome

universal primer

- no longer have to design multiple primers - helps with amplification/giving sequence of the barcode

Figure 7.17 A *knock-in* of the CFTR deltaF508 allele.

- similar with knock out: still using homologous recombination, target is making piece of DNA in vitro to introduce into ES cells - difference: introducing/introducing mutation into knock in mouse, target depends on what the mutation is, HPRT gene instead of neomycin, HAT media instead of G418 Most cases of cystic fibrosis among people of European descent are due to a specific three-base deletion in exon 10 (point mutation) of the CFTR gene, referred to as delta F508 (delta = delete). This mutation was made in vitro in the mouse CFTR gene, as shown in the dark blue exon with the red star that indicates the mutation. The mutant gene was cloned by conventional molecular methods into a targeting vector, as shown at the top. In this particular case, the selection marker was the HPRT gene (similar to neoR), which can be selected in HAT media. The HPRT gene was cloned into an intron (not disrupting gene) in reverse orientation, so its expression would not interfere with normal CFTR expression in lung cells. Homologous recombination inserted this construct into the chromosomal CFTR gene in ES cells, as shown. These ES cells were used to make chimeric mice that carried the mutant CFTR gene.

slide 11

- spacer gives loop - Target sequence and antisense target sequence will sit on top of each other

figure 7.6: Selecting appropriately transformed cells

- tissue culture cells + DNA molecule with selectable marker - selectable maker transforming into cells through homologous recombination (rare event) - cells with the marker will proliferate once selection is applied A DNA construct with a drug or a nutritional gene that can be used in a selection (shown in purple) is made in vitro and transformed into cells in culture. Even if only a few cells are transformed, those will be the only ones to proliferate once selection is applied. These cells can be used to produce a transgenic organism or tissue.

Figure 9.3 A flow chart of a typical genome-wide mutant screen.

1) Identify Genes: The first step is to identify all genes in the genomes using gene prediction methods such as similarity to other genes, the presence of certain recognizable features, for example start and stop codons, and transcript annotated. - Entire sequence needs to be sequenced and annotated ---> Be able to identify every gene in organism ---> Annotation: knowing the structure of the genes on the genome ======> Knowing where introns, exons, promoters, codons, open reading frame, homology, etc 2) Mutate Genes: Then genes are perturbed or mutated by gene disruptions, RNAi, or CRISPR. - Every gene must be mutated once --->Null gene: CRISPR (double strand break --> indel --> frameshift --> premature stop codon) ======> Some genes are essential, null can only be expressed in certain cells --> Knockdown: RNAi ======> pool CRISPR/RNAi mutations 3) Find Phenotype: Third, phenotypes are found by a range of different assays. - Every mutant from the pool must be added to the organism and see which mutant caused each mutation ======> Morphological: dumpy gene ======> Biochemical: ======> Genetic assays: 4) Refine Phenotype: Finally, the role of single genes involved in the process is confirmed and refined, usually by methods used for other mutants.

Two main parts of CRISPR

1. Cas9: enzyme that does the cut 2. SgRNA: unique targeting RNA designed by scientist a. crRNA: unique to what you're targeting b. tracrRNA: hairpin, loaded up into CRISPR enzyme

How does RNAi work?

1. Creation of double-stranded RNA either experimentally or naturally within the organism triggers the mechanism 2. Specific endonuclease known as the Dicer complex chops up dsRNA into 20-25mers—called siRNAs for small interfering RNAs 3. Another enzyme complex called the RISC (for RNA Induced Silencing Complex) picks up siRNAs and pulls the two strands apart - leaves guide RNA that will target RNA to only have one strand? 4. This activates the RISC which binds the siRNA (now single-stranded) to it's complementary mRNA 5. mRNA degradation ensues, ensuring that protein is not made

miRNA processing

1. Drosha excises stem-loop from primary miRNA (pri-miRNA) to generate pre-miRNA of ~ 70 nt - in nucleus (where miRNA is 1st transcribed) - crops extra RNA off 2. Dicer processes pre-miRNA to a mature duplex miRNA One strand is incorporated into miRNA-induced silencing complex (RISC) - in cytoplasm - cuts off loop; makes pre-miRNA into miRNA

steps to NHEJ

1. Target location of insertion by DSB 2. Donor DNA invades DSB 3. Holiday junction (3' and 5') will repair DSB using donor DNA template 4. Full intact piece of DNA 5. Opposite strand will make complimentary strand 6. End with chromosome with inserted nonhomologous sequence

knocking out a gene in ES cells

1. harvest blastocyst, pull out inner cell mass, ES cells in culture dish 2. make mutated construct via homologous recombination, introduce marker in knock out gene 3. take manipulated constructed piece of DNA and transfect into culture of cells, some rare cells will pick up and incorporate mutated DNA 4. finding the cell with the knockout allele, cells without selectable marker will die (survivor cells have knockout allele of 1% or less) 5. survivor cells will begin new culture

Steps of reverse genetic analysis

1. identify and isolate gene 2. edit gene 3. transform cells with edited gene 4. grow into transgenic organism - be able to label parts of a sequence (promoter, enhancer, etc)

miRNA regulation/interference

1. mRNA cleavage - perfect complementarily - mRNA is degraded --> no proteins made - miRISC binds to another target mRNA 2. translational repression - incomplete complementarily (enough homology, some indels/mispairing) - mRNA is blocked by ribosomes slide 8

steps to make chimeric/knockout mice using ES cells

1. super ovulated WT female crossed with WT male 2. 3-4 days after, sacrifice female and harvest uterine horns and pull out blastocysts to get Inner Cell Mass cells and put them in dish (ES cells) 3. go through selection/targeted integration, to have knock out gene construct in the DNA 4. transfect DNA into ES cells, then add G418 in dish to select for ES cells that have the DNA 5. Add gancyclovir to dish to select for knockout cells that have both neomycin resistance and HSV end product: knock out cells 6. mate two mice (typically a different color to make identification easier), inject knocked out ES cells 7. mate female with sterile male to become a pseudo pregnant female, inject blastocyst and wait for pups to be born 8. end product: chimeric mice with original coat color from original ICM mouse. ES cells were integrated and hopefully integrated in germ cells (check by mating again)

steps to knock in mouse

1. super ovulated WT female crossed with WT male 2. 3-4 days after, sacrifice female and harvest uterine horns and pull out blastocysts to get Inner Cell Mass cells and put them in dish (ES cells) 3a. Make mutation for target gene to knock in the DNA construct 4. transfect DNA into ES cells, then add HAT media in dish to select for ES cells that have the DNA with HPRT resistance 5. mate two mice (typically a different color to make identification easier), inject knocked in ES cells 6. mate female with sterile male to become a pseudo pregnant female, inject blastocyst and wait for pups to be born 7. end product: chimeric mice with original coat color from original ICM mouse. ES cells were integrated and hopefully integrated in germ cells of mouse **** WT pten is not knocked out & has functional coding sequence, HPRT was integrated in introns 10. mate knock in mouse to check if integrated into germ cells

steps to loxp sites (similar to making knockout mouse) -- difference: introduce cre to take out neomycin & have the functional gene to make conditional or inducable knockout mouse

1. super ovulated WT female crossed with WT male 2. 3-4 days after, sacrifice female and harvest uterine horns and pull out blastocysts to get Inner Cell Mass cells and put them in dish (ES cells) 3a. Make target vector with lox p sites/floxed sites - this step helps get rid of neomyocin in later step (benefit of lox p site) - some cells do not want neomycin resistance to reproduce 3b. Insert targeting vector in between introns with lox p sites - remember floxing occurs in gDNA NOT in RNA end product: selected knock out gene construct in gDNA 4. transfect DNA into ES cells, then add G418 in dish to select for ES cells that have the DNA with neomycin resistance 5. Add gancyclovir to dish to counter select for knock outES cells 6. add CRE recombinase to get rid of neomycin, selecting for cells sensitive to G418 - will still have floxed sites that have important DNA, but removes neomycin gene 7. mate two mice (typically a different color to make identification easier), inject knocked out ES cells 8. mate female with sterile male to become a pseudo pregnant female, inject blastocyst and wait for pups to be born 9. end product: chimeric mice with original coat color from original ICM mouse. ES cells were integrated and hopefully integrated in germ cells of "knockout" mouse **** WT pten is not knocked out & has functional coding sequence, neomycin was integrated in introns 10. mate "knockout" mouse with cre mouse to only knockout the gene of the you care about in the cell type you want, can be inducible (via feeding)

four types of mutating the core region/coding sequence

1. what is the null mutant phenotype?: Mutate starting ATG: change expression via null allele 2. What are the effects of splicing variants? Mutate splice site variants: splice out intron/splice cites. Changes transcripts and the genome sequence that comes out. 3. What is the hypomorphic phenotype?: Leave most of the gene as is: must understand protein folding. Hypermorphic mutation/tempsensitive 4. Is this gene regulated by microRNAs?: Mutate 3' UTR: will not change protein sequence, if 3' UTR is regulated by microRNA

Figure 9.11 Introducing dsRNA for RNAi experiments: RNAi is performed by direct injection of the dsRNA.

A cDNA from the gene of interest is cloned into a vector with different transcriptional promoters on each side of the cloning site. A vector with SP6 and T7 promoters is commercially available, but other promoters can be used. Two reactions are done in vitro in separate vials, one with SP6 polymerase added and the other with T7 polymerase added. Each reaction produces one single stranded RNA molecule, which are annealed in vitro to make dsRNA. As in Figure 9.7, the dsRNA is injected into the animal, which lays eggs with the dsRNA. The eggs hatch, and some of the offspring show the RNAi phenotype, here a dumpy worm. Similar procedures are performed by injecting directly into eggs of Drosophila. - Two different promoters that will show specific sequence for top strand and bottom strand respectively - One tiny cDNA - From knocking down gene with RNAi

Figure 9.4 A schematic diagram of the PCR primers and the insertion cassettes used in genome-wide gene disruption in yeast.

A collection of PCR primer pairs is made, one upstream and downstream primer pair for each gene in the genome. Five pairs of primers are shown at the top. Each primer is seventy bases long and has four sequence components, two of which are specific to that gene and two of which are common to all primers. The common components are shown in the same color in each primer, whereas the gene-specific components are shown as different shades of the same color. The color key for the primer components is shown at the top right. Each primer pair has sequences corresponding to the target gene in the genome, here shown in shades of blue. The gene-specific sequence is indexed with a specifically defined sequence known as molecular bar code, shown in shades of red. Note that each gene-specific sequence in blue is associated with one specific bar code in red. By knowing the bar code sequence, the investigator could determine which gene has been disrupted. The components that are common to all primer pairs are the sequences to amplify the kanomycin-resistance gene (in purple) and the sequence used to amplify the molecular bar code (in green). By using an adjacent universal primer, all bar codes could be amplified without regard to their own sequence. Each primer pair is used to amplify the kanomycin-resistance gene, which results in a collection of insertion cassettes specific to each gene in the genome. Although five cassettes are shown here, the actual genome-wide screen had nearly 6,000 different insertion cassettes, one for each gene in the yeast genome - site directed incorporation

Figure 9.9 Post-transcriptional gene silencing in petunias. ***Dr. Ostler said this was a harder figure to understand

A petunia with light purple flowers is shown at the left. The flower is lightly pigmented because it has a hypomorphic mutation in the blue pigmentation gene shown beneath the flower; the gene is transcribed and protein is produced, but in reduced amounts by comparison to darkly pigmented petunias. Additional copies of the pigmentation gene were introduced into the plant. The top half of the figure summarizes the expected result of introducing additional copies of the gene, while the bottom half of the figure shows the actual result. Increasing the copy number of a hypomorphic mutation is expected to result in more pigmentation, a more nearly wild-type phenotype, as described for an allelic series in Chapter 4. Contrary to expectations, gene expression was decreased rather than increased and a white flower resulted. Further investigation showed that the effect occurred post-transcriptionally, through RNA degradation. Post-transcriptional gene silencing in petunias was one of the first demonstrations of the RNAi pathway - Organism is protecting itself from RNA viruses - Silenced by dsRNA being produced - More of pink gene--> hypermorph --> white flower

Figure 7.16 Cre-lox deletion of PTEN in the pancreas

A targeting vector is inserted in the PTEN gene by homologous recombination. In the targeting vector both the neo gene in the intron and exon 5 of the PTEN gene are flanked by loxP sites. Although recombination can occur at many locations, the recombinants with crossovers at the locations of the dashed lines are screened for; these have replaced the chromosomal PTEN gene with the edited gene. The actual PTEN gene has nine exons, of which seven are shown here. With transient Cre expression in the ES cells, the neo gene is deleted. These events can be detected in ES cells by appropriate molecular probes. The ES cells are used to make a transgenic mouse strain that also has the Cre gene expressed under the regulatory region from an insulin gene. Under the control of this regulatory region, Cre recombinase is expressed only in the pancreas, and exon 5 is deleted in those cells of the pancreas to produce a tissue-specific gene deletion that is to be analysed for the role of PTEN in pancreatic cells. In other cells Cre is not expressed, so the mouse can survive

CRISPR

Clustered Regular Interspaced Short Palindromic Repeats a collection of sequences in a bacterial genome separated by highly similar palindromic repeats. "CRISPR" is an acronym for "clustered interspersed short palindromic repeat." While the term specifically refers to the array of repeats, it is often used more generally to refer to the process of genome editing using this array of repeats and the associated enzyme Cas9.

Figure 7.14 The Cre-lox system for site-specific recombination

Cre recombinase recognizes the specific sequence of the loxP site shown and cleaves it at the sites indicated by the arrow heads. Note that the lox site has palindromic arms, shown by the green boxes, flanking a unique sequence core. The center core sequence is asymmetric and confers directionality to a loxP site. In subsequent figures, this directionality is represented by a red arrow head. - allow for downstream recombination - using homologous recombination, insert pieces of DNA into genome of ES cells ---> what is being inserted is different compared to previous methods - *targets introns* - simple cre-lox system: adding two loxp sites

Figure 7.10 ES cells and chimeric mice

ES cells are used for targeted insertions with a selection for particular genetic events. The selected ES cells can be assayed molecularly, to ensure that the recombination event of interest has occurred. Those cells with the proper insert are injected into the blastocoel of an isolated blastocyst, and the injected blastocyst is implanted into a pregnant female host. In this example, the ES cells were derived from a white mouse and the recipient blastocyst was derived from a brown mouse, but other coat color combinations are often used. The pregnant female gives birth to a litter with some mice from her own embryos and others from the chimeric embryos. Mice derived from the chimeric embryos are recognized by the variegated coat color and are presumed to be chimeric in all cell types. The chimeric mice are mated to produce a mouse that derives entirely from the ES cells with the targeted insertion. Here those mice will be white

Figure 9.7 An overview of RNAi.

Effect of RNAi is temporary - dsRNA does not last very long - Some cases RNAi is inheritable The example here uses a muscle myosin gene from C. elegans. dsRNA is made in vitro, corresponding to a portion of the coding region of the myosin gene. This dsRNA is injected directly into the worm gonad. The injected worm lays eggs, some of which have gotten the injected dsRNA. The affected egg hatches and the resulting worm is paralysed, as if it had a mutation in the myosin gene. The gene itself is not structurally altered in the paralysed worm, so in most cells and organisms its offspring will not have the dsRNA and will not show an RNAi phenotype. C. elegans is unusual among animals, since the RNAi phenotype can be heritable and some offspring will continue to be affected.

Figure 9.2 Saturation in traditional genetic screens and genome-wide mutant screens.

Four different genes are involved in this hypothetical process, as indicated by the four different colors in the gene structures diagrammed on the left. In the traditional mutant screen, the first 1,000 samples yield a mutant in the blue gene, the second 1,000 samples yield one in the brown gene, and so on. After five 5,000 samples, the blue gene has been mutated three times, the brown gene twice, the green gene once, and the purple gene has not been mutated at all. Thus the role of the purple gene in the process has not been recognized and the gene remains undetected. In a genome-wide screen, all genes are mutated by directed methods first, without regard to what process they may affect. Each gene is tested as part of a pool with other genes in the genome. Because all genes are mutated, the role of the purple gene in this process would be detected. Furthermore, no gene needs to be mutated more than once.

Forward Genetics

Function to Gene -What genes are required/involved in my biological question of interest -Mutants

Figure 8.3 Some questions addressed by gene and genome editing.

Gene and genome editing allows for very specific questions to be addressed. This schematic of a eukaryotic gene with four exons shows some of its functional components and the locations of edits that could be made. Null and hypomorphic alleles can be made for any part of the protein-coding region of the gene, so these locations are representative. In animal genes the target sequence is typically found in the 3´UTR, as suggested in this diagram.

Reverse Genetics

Gene to Function -What does my gene of interest do? -Knock-out (or knock-down) the gene. •Many different approaches. -RNA interference (RNAi), knockdown -Site-specific homologous recombination, knockout, Yeast -Mutate a domain or specific site (knock-in) •Site-directed mutagenesis. •CRISPR -Over-express the gene via transgene or by changing transcription factor •Risk problems with novel or ectopic expression.

Figure 8.1 Strategies for finding a mutation in a gene of interest

Genetic analysis relies on having a mutant for the gene to be studied, so a mutant and its corresponding DNA sequence can be found by different methods. In this hypothetical example, a gene responsible for a blue pigment in purple flowers is under study, so the mutant has only the red pigment. A. In traditional mutagenesis screens, a strain that is of the wild type for the gene is treated with a mutagen in order to induce mutations randomly throughout the genome. Other than the mutations that are induced, the offspring are genetically uniform, which is shown here by the flowers with uniform purple colors. After several generations, a mutant with a phenotype of interest is found and isolated, as described in Chapter 4. From this, the gene is then cloned as described in Chapter 5. B. With naturally occurring variants, much more diversity is present in the original population, which is shown here in different shades of purple. The individual specimen with the phenotype of interest is found and isolated so as to identify the corresponding DNA sequence, similarly to what is done in traditional mutagenesis screens. - If same gene: phenotype same as parents - If different gene: will have one functional copy to show WT phenotype C. In genome-wide mutant screens, to be described in Chapter 9, every gene in the genome is mutated first, and the mutants are stored as a library, regardless of their phenotypes. The library of gene disruptions is screened for those with the phenotype of interest. As seen here, many different phenotypes will be found and disruptions for some genes will result in no mutant phenotype. The clone encoding the gene of interest is identified by the phenotype. D. With genome editing, a mutation is created by a directed change in the DNA sequence of the gene of interest. In this example, the insertion of two bases changes the reading frame and disrupts the gene, so that no functional protein is likely to be made. The phenotype arising from the edited gene is then analysed. *most edits are either null or hypomorph --> this might not be what we want in a mutation Ex. If we want a temperature gradient allele we would have to do a missense mutation instead of inserting a reading frameshift to slightly change protein folding

Figure 8.5 Homology-directed repair

Homology-directed repair of a DSB requires a template DNA molecule with a highly similar sequence to the chromosome; the donor template is shown here in red, with the regions of homology between the donor template and the chromosome flanking the DSB on each side. Because it relies on sequence homology between the two DNA molecules, homology-directed repair is usually considered to be precise. Donor template DNA first "invades" between the strands of the chromosomal DNA at the region of the DSB, where it provides the template sequence for the repair of the DSB; the repair DNA synthesis is illustrated by a dashed red line. Once this repair from the donor template is completed, the chromosomal DNA will have replaced its own sequence or inserted the sequence from the template. ==> Invaded strand will be copied - Two pieces of DNA and the other strand will be copied as well - Similar to knock in/knock out from chapter 7

forward genetics cons

If multiple gene mutated, must find which gene is what you want --> long process - needle in haystack

Figure 9.1 A comparison of forward and reverse genetic analysis with genome-wide mutant screens.

In forward genetics, as described in Chapter 4, a wild-type organism is treated with a mutagen, mutant organisms are found, and the gene is cloned. The mutant phenotype is found before the gene is identified. In reverse genetics, as described in Chapter 7 for yeast and mice, the gene is identified first and is mutated in vitro. The mutant gene is introduced back into the organism and the phenotype is scored. In genome-wide screens, all genes are altered in vitro and reintroduced individually into the organism or cell without regard to their possible phenotypes or functions. The phenotypes are then scored

Figure 8.7 Non-homologous end joining

In most eukaryotes, the more common type of DSB repair is NHEJ. With no homologous sequence to use as a template, the DSB is repaired by joining the ends of two different DNA molecules. While the joining may be precise such that no changes occur, it is more often the case that the process is imprecise and one or a few bases are either inserted or deleted as a result. If the DSB is made within the protein-coding region of gene, the small indels arising from the imprecision of NHEJ produce a frameshift mutation and disrupt the function of the gene good to knows: - DSB must be present - Bases will have over hang that can be repaired (if they fit together) if they do not fit correctly, it can add or cleave off the overhang - 25% of mutations are usually fixed with no proof reading/homology - Imprecise insertions and deletions - This is how we get chromosome translocations - sloppy

Primary transcripts containing miRNA

Most miRNAs are transcribed by RNA polymerase II from noncoding DNA regions that generate short dsRNA hairpins - some in coding sequence - some not in coding sequence (introns)

mutating the promoter/enhancer/cis-regulatory module

Mutating promoter: null allele Mutating enhancer: TF binding sites, leaves gene alone--> modifies expression Consensus binding sequences, binding motifs Different TF will bind at different binding sequences, can vary

is RNA made double stranded?

No ma'am

Figure 9.6 The use of the molecular bar code.

Once disruptions are made for each gene the mutants are pooled and tested for a mutant phenotype. In our drawing, each yeast cell at the top left has a single gene disruption. - Every mutation in the pool is represented (Barcode helps identify) The pool of mutants is grown in media that lack uracil. Once the culture has grown, the cells are removed and assayed by PCR. The universal primer will amplify each bar code in the culture, as shown. - Whittled down to four mutants (Have the ability to make uracil----> grows) The collection of bar codes is hybridized to a microarray that has all of the input bar codes in a defined pattern, here the same one as shown in the pool of mutants. The bar code for any gene that is needed for growth in the absence of uracil will not hybridize in the output array. This is often referred to as a dropout screen, since the desired mutants are the ones that failed to grow and are not present in the final population - Instead of sequencing they use microarray (hybridizing) - The five genes are represented as 5 samples on the microarray - Shows if gene is needed to make uracil (missing samples are uracil auxotrophs) - BARCODE is linked back to the gene you designed it too

figure 7.12 selection and counter-selection for targeted integrations

Once the targeting vector has been introduced into ES cells, one of three events can occur. 1. No integration: no recombining, If the vector fails to integrate into the chromosome in the ES cells, the cells will not grow in the presence of the neomycin analogue G418. G418-resistant cells can arise either from integration of the targeting vector at random sites (suggested by the light purple) or from targeted integration via homology-directed repair with the chromosomal copy of the gene of interest. These ES cells can be distinguished by a selection that is based on gancyclovir. - cells are sensitive to neomycin, cassette integrated, they will die if added in G418 2. Random integration: If the vector inserts at random locations without relying on homologous recombination, the HSV-tk gene, which confers sensitivity to gancyclovir, will also be integrated. - integrates randomly from NHEJ, contains HSV gene, in random location, resistant to G418 but sensitive to gancyclovir 3. Targeted integration: If the vector inserts by homologous recombination with the target gene, the HSV-tk gene is excluded (since it lies outside the region of homology) and the cells are resistant to ganciclovir. - resistant to both, will survive

RNAi

RNA interference - in vitro in a lab - cytoplasm - injecting double stranded RNA into a cell turns off expression of a gene with the same sequence as the RNA - dsRNA is synthesized that is complementary to the mRNA sequence of interest. When transfected into human cells, dsRNA separates and promotes degradation of target mRNA, knocking down gene expression.

Figure 8.6 Inserting a non-homologous sequence using homology-directed repair.

Since the regions of homology (upstream and downstream) where the crossover occurs flank the DSB, sequences from the donor template that lie between the homology regions but are not similar to the chromosome can also be inserted; the non-homologous region, such as a reporter gene or some other construct, is shown in blue. When the donor DNA molecule is used as a template, the non-homologous region is also copied and is inserted into the chromosomal DNA, along with flanking sequences from the donor.

Genome wide mutant screens

Studying the function of many genes by RNAi

Figure 8.13 The domains of Cas9 nuclease.

The Cas9 nuclease from S. pyogenes is the most commonly used enzyme to make the double-stranded break in the target DNA sequence. Cas9 has two separate domains that target the individual DNA strands and make the single-stranded cuts at approximately the locations indicated by the red arrow heads. Note the CGG sequence on the target DNA just downstream of the cut site by the RuvC domain; Cas9 from S. pyogenes depends on the presence of NGG as its PAM sequence to distinguish foreign invading DNA from sequences in the bacterial genome. - RuvC & HNH domain

figure 7.9 Inner cell mass and ES cells

The cells in the inner cell mass of the blastocyst will give rise to all cells and tissues in the adult. Cells of the inner cell mass can be isolated from blastocysts and grown indefinitely in culture, where they are referred to as embryonic stem or ES cells - cells from the inner cell mass are isolated and grown in culture as ES cells - used for knock ins and know outs - ES cells can arise and create every cell in adult mice (including germ cells!)

Figure 9.14 One plasmid used for shRNA libraries in mammalian cells.

The sequence corresponding to the short interfering RNA (siRNA) is shown at the top; the siRNA sequence corresponds to the two arms flanking the loop. The loop sequence will be cleaved and is not important. The sequence corresponding to the short hairpin RNA (shRNA) is cloned into the expression vector plasmid in the middle of the drawing. In this particular library, the shRNA sequence is until the control of the promoter from the U6 gene, which encodes an RNA involved in splicing, so the shRNA will be constitutively transcribed. Other promoters can also be used. Drug markers are included on the plasmid for growth in bacteria (ampicillin) and in mammalian cells (puromycin). The dashed arrow indicates the direction of transcription, and the viral LTRs are involved in transcriptional regulation and termination. When the sequence corresponding to the shRNA is transcribed, the transcript forms the short hairpin shown at the bottom of the figure by intrastrand base pairing. Cleavage of the loop produces the siRNA involved in RNA interference.

Figure 8.15 The origins of CRISPR spacers.

The spacers in the CRISPR array are sequences from foreign genomes that have recently invaded the bacteria. In this example, the foreign genome comes from bacteriophage (DNA or RNA), shown infecting the bacterium at the top. The phage genome is light brown. Most of the bacteria infected by the phage are lysed but a few cells survive the infection. These bacteria acquire sequences from the phage genome immediately upstream of the PAM sequence (in red) characteristic for those bacteria; for S. pyogenes, the PAM sequence is NGG. The fragment of the phage genome with its PAM sequence is referred to as the protospacer. When these protospacers are incorporated into the CRISPR array as spacers, the PAM sequence is not included. Sequences from the most recent infections are incorporated near the beginning of the CRISPR arrays, as shown here. - Repeats do not do anything, but they were identified first - Spacer used as guides - PAM sequence: unique to each cas9 enzyme

Figure 9.10 Introducing dsRNA for RNAi experiments: dsRNA is transfected into mammalian cells as a hairpin.

The target mRNA is shown in blue at the top of the figure; the dsRNA to create RNAi is shown with the complementary strands in red and blue, based off sequence in exon. This is cloned, as DNA, into a vector whose backbone structure is based on a microRNA gene. A RNA hairpin is synthesized in vitro, with a stem of twenty-two nucleotides, including a two base overhang at the 3´ end. The loop is usually about ten nucleotides long. The dashed line indicates the site of future cleavage by Dicer. The hairpin RNA is transfected into mammalian cells in culture, where it is cleaved by Dicer to make an siRNA, which triggers RNAi. The RNAi effect in the cultured cells is transient. - Uses one strand of RNA to target mRNA What is injected/transfected into the cells? - dsRNA as a hairpin (shRNA) - 10- 20 nc Gives enough specificity

Traditional Mutant Screen vs Genome-Wide Screen

Traditional: - No control with what will be mutated --> EMS mutation hits randomly Genome-Wide: - Every single gene has one mutation, no genes are missed - Finds importance of the gene - Not looking at just a subset - Looking globally - Pool together: Sifting through different phenotypes

Figure 7.15 Floxed sites and deletions

Two loxP sites (orange arrow heads) in the same orientation are in introns and flank exons 2, 3, and 4; the sequence between the loxP sites is said to be floxed. The DSBs made by Cre at these lox sites allow recombination to occur, which deletes the sequence between them. Since both molecules retain a loxP site, Cre-mediated insertion can also occur in the reverse reaction. Flanking lox sites in other orientations can be used to produce other types of rearrangement of floxed regions.

Figure 9.5 The insertion cassettes are used to disrupt the genes.

Two of the insertion cassettes from Figure 9.5 are shown here; they correspond to different genes in different yeast strains. The insertion cassettes are transformed at random into yeast cells. Homologous recombination between the ends of the insertion cassette and the target gene inserts the kanomycin-resistance gene and the specific molecular bar code into each gene. Kanomycin is used to select for yeast cells that have an insertion, and the bar code will be used in Figure 9.6 to determine which gene has been disrupted. Transfect into normal yeast cells - Homology in blue region (18bp), endogenous to the yeast cells - Forces them to take up kan MX gene in order to survive ===> In doing so it disrupts function of the gene ===>If phenotype observed --> then sequence and find barcode (using universal primers when sequencing)

Figure 8.4 Pathways to repair double-stranded breaks

When a double-stranded break (DSB) in a DNA sequence occurs, two different pathways are used to repair it. NHEJ: In the absence of a homologous sequence to serve as a donor template. NHEJ often results in small insertions or deletions Homology directed: When a sequence with sufficient homology is present to serve as the donor, homology-directed repair can occur, as shown in the two examples: A. The donor template (represented by the gray line in the backbone) need not match the original sequence precisely. The locations of possible crossover are indicated by the dashed lines. In one example, the donor has the sequence CCC, while the original sequence is TCA; the sequence from the donor can replace the sequence in the original. B. The donor has a novel donor sequence (such as a reporter gene) flanked by sequences that match the original genome sequences. By homologous recombination, this donor sequence can be inserted into the original. The altered sequence is shown with the red backbone and shaded boxes. --> CRISPR cuts double strand breaks then donor DNA can come in and force in DNA

why use a virus?

Why use virus? ---> depends on your cell!! Transfection: allows plasmid to go into complicated cell that have 293T, will go into cells (has phospholipid bilayer) using AAV packaging cells Most benign Virus: lentivirus Very strong Infection: can easily infect normal cells

Figure 8.12 Using CRISPR to target a sequence for editing.

With CRISPR, the location to be edited is targeted by an crRNA sequence ( shown with the purple phosphate backbone) that can base-pair with one strand of the DNA sequence, as shown here. The crRNA itself is 20 bases or more. The diagram shows that each base is paired between the crRNA and the DNA target, but the precise rules for targeting are not known and some mismatches do occur.

homology-directed repair

a DNA repair process within a cell in which a sequence that is highly similar to the target site is used as a template to make the repair.

single guide RNA (sgRNA)

a RNA molecule that includes both the crRNA (20 nuc complimentary to target RNA) and the tracrRNA needed by Cas9 for genome editing. - specifically targets RNA that you would like to mutant (40-50 nucleotides)

knockout mutation

a mutant allele in which the wild-type allele has been severely disrupted or deleted. A knockout mutation is a *null allele*

knock-in mutation

a procedure in which the wild-type gene on the chromosome is replaced by a version that has been engineered in vitro with a specific mutation. The procedure is widely used in mice

Cre

a recombination enzyme from the bacteriophage P1 that catalyzes site-specific crossovers at the nucleotide sequences known as loxP sites. The Cre-lox recombination system is widely used to insert sequence or produce recombination at specific sites.

nonhomologous end joining (NHEJ)

a way of repairing a DNA double-strand break and is often error-prone - happens when there are free ends of DNA - happens when there is no counter selection gene (no specific target)

How to edit a genome instead of indels (no frameshift/mutation)

add DNA donor to initiate homologous recombination: - Inject CAS9 mRNA, sgRNA to your target, or DNA as donor - Can also use single stranded oligo as DNA donor

primer for KanR gene

antibiotic resistance, disrupts target gene, selects for insertion

double-stranded breaks and their repair

cells correctly repair double strand breaks 99.9% of the time, but when not repaired there will be small insertions/deletions

Need to deliver CAS9 protein and your sgRNA into your cells?

clone a short guide sequence into a Cas9 All in one vector ex. Zebrafish: Inject sgRNA and mRNA of CAS9 into 1 celled embryo Two different promoters on opposite strands: One for sgRNA One for Cas9 - Kanamycin resistance - select against - SV 40 ori - allows DNA to replicate on itself - Antibody to myc - shows if you're expressing cas9 correctly - WPRE - regulatory element that helps increase expression of sgRNA - polyA tail - makes it a protein

genomic wide mutant screens

every single gene/mutant identified More directed way - Every gene mutated once - Introduce mutation into WT organism and find mutant phenotype - Be able to identify the genes responsible for mutant phenotype - End up identifying unsuspecting phenotypes for multiple genes because of pleiotropy Method used: 1. Knockdown via RNAi 2. Null mutation via CRISPR

molecular barcode

gene specific sequence, tells what gene is being sequenced

CRISPR-associated (Cas) genes

genes located adjacent to the CRISPR array, and involved with its function to degrade foreign DNA sequences, such as from viruses. Cas9 is CRISPR associated protein 9 in S. pyogenes. - causes double strand breaks

most common method for yeast to repair DSB?

homologous recombination

Yeast- homologous recombination

introduce a novel mutation into a system via homologous recombination, have restriction enzymes that will be at the ends of gene of interest 1. clone yeast selectable marker into middle of your favorite gene 2. transform URA3- yeast with construct 3. Select transformants on growth media without uracil. Only URA3+ (uracil auxotroph, selectable marker) cells will grow. These should be your knockouts. 4. Confirm by PCR to ensure no revertants

coat colors...

mark the cells that have been manipulated

RISC complex

miRNA * miRNA duplex - one miRNA will be degraded - one miRNA will be kept -----> single stranded guide miRNA

what is the selectable marker for mice?

neo-mycin - used to *knock out* via homologous recombination - inserted in an exon in the same reading frame - neoR: resistance gene against the neomycin (drug) added to petri dish of cells to select those who have the resistance --> selectable marker: those without this gene will die

question: do knock in mouse have functional gene?

no? bc it is a mutation?

floxed region

region between loxp sites - portion of gene that will be fluxed out and deleted

shRNA plasmid: sense vs antisense

sense: G's and C's antisense: T's and A's shows directionality

Endogenous CRISPR pathway in Bacteria and Archaea - acquired prokaryotic immune system

short 'spacers' are made from invasive DNA - inserted into the CRISPR array. Serves as the 'memory' of infection. Spacers are processed into crRNAs and an RNAi-like process inactivates viral DNA - has a set of palindrome (24-48 bp) on each side of exogenous (30 bp) ** look at slides 10 & 11

shRNA

short hairpin RNA - cytoplasm

siRNA

short interfering RNA - blocks protein synthesis - interferes message - cytoplasm - class of double-stranded RNAs about 23 nucleotides in length that silence gene expression; act by either promoting the degradation of mRNAs with precisely complementary sequences or by inhibiting the transcription of genes containing precisely complementary sequences

making dsRNA

since RNA is not double stranded we need to induce complementary RNA to anneal - RNAi vectors with different promoters will allow pieces of RNA to anneal 1.Use opposing promoters on either side of the sequence to generate both strands at the same time - each strand gets used, no loop/dicer 2.Synthesize ssRNA that can anneal to itself with a few bases in the middle—shRNA (small hairpin RNA) - Easy because One promoter, one direction - Can be more complicated because when it loops it must compliment slide 10 and 11

sgRNA

single guide RNA - CRISPR complex - cuts genomic sequence - *nucleus*

Reverse Genetics: editing genes in yeast and mice

starting with sequence

loxP

the 34 base-pair sequence that is the target for recombination by the Cre recombinase. The Cre-lox recombination system is widely used to insert sequence or produce recombination at specific sites.

homologous recombination

the ability to swap your sequence with another sequence. Create any mutation or modification you want in the endogenous gene - gene knockout via premature stop codon - gene knock ins via introducing novel mutation into WT sequence - tag to your protein via GFP --> problem is too big, can use BFP, iRFP) - site-directed mutagenesis via point mutations ------> this successfully done in bacteria, yeast, and mice

non-homologous end joining

the process by doublestranded break is repaired by attached two DNA sequences not related in sequence.

trans-activating CRISRR RNA (tracrRNA)

the specific RNA molecule required for the activity of Cas9 from S. pyogenes. - hairpin, complimentary sequence of bp from crRNA/target DNA ----> this will get loaded into cas9 that will actually do the editing

crRNA (CRISPR RNA)

the transcript made from the CRISPR array that, in S. pyogenes, directs the Cas9 protein to its target sequence. - 20 nucleotides complimentary to target RNA - gives sequence specificity - like PCR primer - has a set of palindrome (24-48 bp) on each side of exogenous (30 bp)

cre recombinase

triggers recombination between loxp sites - loops DNA on its self to line up two arrows and let crossing over happen between the two - flox out DNA in between the loxp sites

CAS9 is can also be guided by a single guide RNA (sgRNA) to cleave both strands of DNA

• sgRNA is designed to target desired mutation location in genome • CAS9 has two nuclease domains - one for each backbone of dsDNA • Creates double stranded break • Repair via nonhomologous end joining (NHEJ) will sometimes result in insertion or deletion. This is targeting a gene and creating a Knock out: Keeps cutting until indels are present in the sequence - Frameshift = premature stop codon - Result: null allele Relating to bacteria: Instead of cutting gDNA of the cell, it uses spacers and cuts where the bacteriophage came in and infected

RNA interference

•Called RNAi—method for "knocking down" a gene by selectively destroying the mRNA that would translate into that gene product, blocks RNA from translating, stops gene expression -No mRNA = no protein -That's the theory; hard to create a complete "knock-out" with RNAi—some leakage - •Also known as RNA silencing •First studied in C. elegans—now in other organisms -Andrew Fire and Craig Mellow - Nobel Prize - RNAi in C. elegans -Not all organisms can do RNAi—necessary machinery not present -Mammals, Drosophila, yeast also have the machinery •Thought to protect organisms from viral infection and also from endogenous transposable element interference

Applications of RNAi

•Naturally occurring RNAi defends the cell against itself and others -Regulates transposable elements -Can destroy viral DNA •Artificial RNAi can be used to determine the function of a particular sequence—what does "Gene X" do? -Don't need the entire sequence of the gene to do this - only need 20 nucleotides that are specific to gene of interest •Application in disease therapy -Destroy harmful gene products of a mutation - WT gene can be untouched when destroying harmful gene •Kits produced by Invitrogen, Ambion etc. to facilitate quick and easy RNAi - Complication of RNAi but not with CRISPR: off target spots, if the sequence is not 100% the same, it must be homologous enough to

amplification of RNAi

•RNAi seems like it would only go until the original dsRNA was "used up" by binding to mRNA •In fact, RNAi once started can *self-perpetuate* -RISC chops up mRNA into small bits—can be used as siRNAs for other RISC activation ----> degraded mRNA (from knocking down) can be amplified by targeting mRNA system, degrading process helps shut down •Evidence also for the existence of RdRp—RNA dependent RNA polymerase - ** Polymerase that can only use RNA as a template, not DNA -Uses antisense strand of a denatured siRNA and mRNA of the cell to form dsRNA...which continues the cycle of RNAi ----> This is what was not used - makes more double stranded RNA that will repeat the siRNA production once copied by RNA polymerase, RdRp - reamplifies using the 3' end - The other strand was used to target the mRNA