molecular methods sessions 1-3

DNA EXTRACTION

- Isolating DNA by disrupting cell wall/cell membrane and a nuclear membrane is called a DNA extraction - The cell is made up of cytoplasm, and cell membrane/wall. The cytoplasm contains several organelles such as mitochondria, ribosomes, nucleus, endoplasmic reticulum - Animal does not have cell wall but plant cells and most bacterial cells contain the cell wall - So if we want to isolate DNA we have to break cell wall/membrane and nuclear envelope . we also have to remove other cell organelle debris. In the final step we precipitate and purify the DNA

northern blotting

- Detects specific RNA molecules among a mixture of rna or a particular tissue in order to measure rna expression - probes are available, hybridised UTR to mRNA will enable you to look at any splicing events. multiple bands will occur - northern blotting doesn't tell us what part of tissue it's in so in-situ hybridisation is used to see where a specific gene is expressed. it is used for localisation The northern blot, or RNA blot,[1] is a technique used in molecular biologyresearch to study gene expression by detection of RNA (or isolated mRNA) in a sample.[2][3] With northern blotting it is possible to observe cellular control over structure and function by determining the particular gene expression rates during differentiation and morphogenesis, as well as in abnormal or diseased conditions.[4] Northern blotting involves the use of electrophoresis to separate RNA samples by size, and detection with a hybridization probecomplementary to part of or the entire target sequence. The term 'northern blot' actually refers specifically to the capillary transfer of RNA from the electrophoresis gel to the blotting membrane. However, the entire process is commonly referred to as northern blotting.[5] The northern blot technique was developed in 1977 by James Alwine, David Kemp, and George Stark at Stanford University,[6] with contributions from Gerhard Heinrich. Northern blotting takes its name from its similarity to the first blotting technique, the Southern blot, named for biologist Edwin Southern.[2] The major difference is that RNA, rather than DNA, is analyzed in the northern blot A general blotting procedure[5] starts with extraction of total RNA from a homogenized tissue sample or from cells. Eukaryotic mRNA can then be isolated through the use of oligo (dT) cellulose chromatography to isolate only those RNAs with a poly(A) tail.[8][9] RNA samples are then separated by gel electrophoresis. Since the gels are fragile and the probes are unable to enter the matrix, the RNA samples, now separated by size, are transferred to a nylon membrane through a capillary or vacuum blotting system. A nylon membrane with a positive charge is the most effective for use in northern blotting since the negatively charged nucleic acids have a high affinity for them. The transfer buffer used for the blotting usually contains formamide because it lowers the annealing temperature of the probe-RNA interaction, thus eliminating the need for high temperatures, which could cause RNA degradation.[10] Once the RNA has been transferred to the membrane, it is immobilized through covalent linkage to the membrane by UV light or heat. After a probe has been labeled, it is hybridized to the RNA on the membrane. Experimental conditions that can affect the efficiency and specificity of hybridization include ionic strength, viscosity, duplex length, mismatched base pairs, and base composition.[11] The membrane is washed to ensure that the probe has bound specifically and to prevent background signals from arising. The hybrid signals are then detected by X-ray film and can be quantified by densitometry. To create controls for comparison in a northern blot, samples not displaying the gene product of interest can be used after determination by microarrays or RT-PCR. Gels: The RNA samples are most commonly separated on agarose gels containing formaldehyde as a denaturing agent for the RNA to limit secondary structure.[11][12] The gels can be stained with ethidium bromide (EtBr) and viewed under UV light to observe the quality and quantity of RNA before blotting.[11] Polyacrylamide gel electrophoresis with urea can also be used in RNA separation but it is most commonly used for fragmented RNA or microRNAs.[13] An RNA ladder is often run alongside the samples on an electrophoresis gel to observe the size of fragments obtained but in total RNA samples the ribosomal subunits can act as size markers.[11]Since the large ribosomal subunit is 28S (approximately 5kb) and the small ribosomal subunit is 18S (approximately 2kb) two prominent bands appear on the gel, the larger at close to twice the intensity of the smaller probes: Probes for northern blotting are composed of nucleic acids with a complementary sequence to all or part of the RNA of interest, they can be DNA, RNA, or oligonucleotides with a minimum of 25 complementary bases to the target sequence.[5] RNA probes (riboprobes) that are transcribed in vitro are able to withstand more rigorous washing steps preventing some of the background noise.[11] Commonly cDNA is created with labelled primers for the RNA sequence of interest to act as the probe in the northern blot.[15] The probes must be labelled either with radioactive isotopes (32P) or with chemiluminescence in which alkaline phosphatase or horseradish peroxidase (HRP) break down chemiluminescent substrates producing a detectable emission of light.[16] The chemiluminescent labelling can occur in two ways: either the probe is attached to the enzyme, or the probe is labelled with a ligand (e.g. biotin) for which the ligand (e.g., avidinor streptavidin) is attached to the enzyme (e.g. HRP).[11] X-ray film can detect both the radioactive and chemiluminescent signals and many researchers prefer the chemiluminescent signals because they are faster, more sensitive, and reduce the health hazards that go along with radioactive labels.[16] The same membrane can be probed up to five times without a significant loss of the target RNA

sequence comparison

- conservation - determine genomes of different species - evolution

why determine expression of certain genes?

- determine expression levels - biological problems - gene expression levels allow to determine onset age of disease characterization of novel proteins based on sequence search in database determines mrna/protein sequence and generated antibodies against peptide

molar ratio insert- vector

- enough insert should be added so no re-ligation occurs - for ligations of 5' or 3' overhang restriction fragments molar-ratio should be 2:1 - for plasmid ligations a maximum concentration of DNA per ligation reaction should be 500 ng

what uses does sequencing have?

- pathogenic variations - evolutions - exon sequencing makes disease mutations easier to work with than genome sequencing

cDNA Synthesis and Reverse Transcriptase

- synthesizes double stranded DNA from RNA template (mRNA) - used to construct complementary DNA (cDNA) - if the mrna has a 3' poly-a-tail then an oligodt primer can be used to prime all mRNAs - if you want to produce cDNA from a subset of all mRNAs, then a sequence specific primer could be used that'll bind to one mRNA sequence - if you want to produce pieces of cDNA that were scattered all over the mRNA then a random primer cocktail that would produce cDNA from all mRNAS. cDNA would not be in full length cDNA synthesis, also known as reverse transcription, generates complementary DNA (cDNA) from an RNA template. Unsatisfactory cDNA yields can arise from low RNA concentration, target gene complexity, and reagent inconsistencies. Various kits and reagents can help address these issues. cDNA synthesis kits are inclusive of necessary reaction components, including polymerase, primers, buffers, and dNTPs, along with optimized protocols for streamlined sample preparation. Custom primers can be used in place of the kit-supplied random primers or oligo(dT), allowing researchers to manipulate cDNA strand synthesis for maximum yield and accuracy. For RNA starting material, RNA extraction kits are available, designed to produce high-quality RNA yields from a variety of cell and tissue sources. Other reagents such as RNase inhibitors and nuclease-free water can ensure that that the starting samples and the end products are protected from degradation. Browse through our directory of cDNA synthesis products and find solutions for applications ranging from library construction to quantitative reverse transcription PCR (RT-qPCR).

southern blotting

- used to detect specific DNA sequences in DNA samples, it combines the transfer of electrophoresis separated DNA fragments to a filter membrane and fragment detection by probe hybridisation A Southern blot is a method used in molecular biology for detection of a specific DNA sequencein DNA samples. Southern blotting combines transfer of electrophoresis-separated DNA fragments to a filter membrane and subsequent fragment detection by probe hybridization. The method is named after the British biologistEdwin Southern, who first published it in 1975.[1]Other blotting methods (i.e., western blot,[2]northern blot, eastern blot, southwestern blot) that employ similar principles, but using RNA or protein, have later been named in reference to Edwin Southern's name. As the label is eponymous, Southern is capitalised, as is conventional of proper nouns. The names for other blotting methods may follow this convention, by analogy

random prime labelling

1) DS DNA denatured to 2 single strands 2) add hexamers ( 6 nonspecific nucleotide primers), gives an idea where they'll anneal 3) polymerise ( DNA pol 1) with radioactive nucleotides, new radioactive DNA is produced which is then denatured 4) PCR product + radioactive copy of template = labelled probe

labelling of probes

1) random prime labelling 2) radioactive PCR 3) End-labelling of oligo ( PNK)

SYBR Green steps

1) reaction setup : the SYBR Green dye fluoresces when bound to dsDNA 2) denaturation: when the dye is denatured, the SYBR Green dye Is released and the fluorescence is decreased 3) polymerisation: during extension, primers anneal and pcr products are generated 4) polymerisation completed: when polymerisation is complete, the SYBR binds to DSDNA resulting in an increase in fluorescent detected by the system ( light excitation)

DNA extraction methods

1. Chemical-based DNA extraction method 2. Solid-phase DNA extraction method.

Sanger sequencing

A procedure in which chemical termination of daughter strands help in determining the DNA sequence. - method of DNA sequencing based on the incorporation of chain terminating di-deoxynucleotides (ddNTP's) by DNA polymerase during in vitro DNA replication. it requires a single strand template, dna primer , DNA polymerase, dntp's, ddntps ( terminate DNA strand elongation ), they lack an 3' OH group required for the formation of a phosphodiester bond between 2 nucleotides. these may be radioactively labelled or fluoroscently for detection in automated sequencing machines - the dna sample is divided into 4 separate sequencing reactions containing all 4 dntps ( dATP/ dGTP/dCTP/dTTP) and dna pol each tube contains 1 of the dNTPs ( 4 in total )

assessing global mRNA expression- Microarrays

A microarray is a laboratory tool used to detect the expression of thousands of genes at the same time. DNA microarrays are microscope slides that are printed with thousands of tiny spots in defined positions, with each spot containing a known DNA sequence or gene. Often, these slides are referred to as gene chips or DNA chips. The DNA molecules attached to each slide act as probes to detect gene expression, which is also known as the transcriptome or the set of messenger RNA (mRNA) transcripts expressed by a group of genes. To perform a microarray analysis, mRNA molecules are typically collected from both an experimental sample and a reference sample. For example, the reference sample could be collected from a healthy individual, and the experimental sample could be collected from an individual with a disease like cancer. The two mRNA samples are then converted into complementary DNA (cDNA), and each sample is labeled with a fluorescent probe of a different color. For instance, the experimental cDNA sample may be labeled with a red fluorescent dye, whereas the reference cDNA may be labeled with a green fluorescent dye. The two samples are then mixed together and allowed to bind to the microarray slide. The process in which the cDNA molecules bind to the DNA probes on the slide is called hybridization. Following hybridization, the microarray is scanned to measure the expression of each gene printed on the slide. If the expression of a particular gene is higher in the experimental sample than in the reference sample, then the corresponding spot on the microarray appears red. In contrast, if the expression in the experimental sample is lower than in the reference sample, then the spot appears green. Finally, if there is equal expression in the two samples, then the spot appears yellow. The data gathered through microarrays can be used to create gene expression profiles, which show simultaneous changes in the expression of many genes in response to a particular condition or treatment. The core principle behind microarrays is hybridization between two DNA strands, the property of complementary nucleic acid sequences to specifically pair with each other by forming hydrogen bonds between complementary nucleotide base pairs. A high number of complementary base pairs in a nucleotide sequence means tighter non-covalent bonding between the two strands. After washing off non-specific bonding sequences, only strongly paired strands will remain hybridized. Fluorescently labeled target sequences that bind to a probe sequence generate a signal that depends on the hybridization conditions (such as temperature), and washing after hybridization. Total strength of the signal, from a spot (feature), depends upon the amount of target sample binding to the probes present on that spot. Microarrays use relative quantitation in which the intensity of a feature is compared to the intensity of the same feature under a different condition, and the identity of the feature is known by its position. mRNA is an intermediary molecule which carries the genetic information from the cell nucleus to the cytoplasm for protein synthesis. Whenever some genes are expressed or are in their active state, many copies of mRNA corresponding to the particular genes are produced by a process called transcription. These mRNAs synthesize the corresponding protein by translation. So, indirectly by assessing the various mRNAs, we can assess the genetic information or the gene expression. This helps in the understanding of various processes behind every altered genetic expression. Thus, mRNA acts as a surrogate marker. Since mRNA is degraded easily, it is necessary to convert it into a more stable cDNA form. Labeling of cDNA is done by fluorochrome dyes Cy3 (green) and Cy5 (red). The principle behind microarrays is that complementary sequences will bind to each other. The unknown DNA molecules are cut into fragments by restriction endonucleases; fluorescent markers are attached to these DNA fragments. These are then allowed to react with probes of the DNA chip. Then the target DNA fragments along with complementary sequences bind to the DNA probes. The remaining DNA fragments are washed away. The target DNA pieces can be identified by their fluorescence emission by passing a laser beam. A computer is used to record the pattern of fluorescence emission and DNA identification. This technique of employing DNA chips is very rapid, besides being sensitive and specific for the identification of several DNA fragments simultaneously. The study of the expression of most, if not all, genes in a specimen is not hypothesis-driven as most of the studies used to be,[4] but is instead referred to as "discovery-type research" or in a less flattering description as "fishing expeditions." Whereas cDNA derived from a tumor is hybridized to a chip to study gene expression levels, alterations in DNA copy number (gene amplification or deletion) can be measured by hybridizing fluorescently labeled DNA from a tumor specimen to these chips.[5,6] TMAs are constructed by transferring cores of paraffin-embedded tissue to pre-cored holes in a recipient paraffin block.[7] Over 500 cores can be placed in a single block by this technique. Sections cut from TMA blocks can then be used for immunohistochemistry (IHC) or in situ hybridization studies. TMAs are similar to gene expression microarrays in having samples arrayed in rows and columns on a glass slide; they differ in that each element on the TMA slide corresponds to a single patient sample, allowing multiple patient samples to be assessed for a single molecular marker in one experiment, while gene expression arrays allow assessment of thousands of molecular markers on a single patient sample per experiment.

components of a plasmid

A plasmid is an accessory chromosomal DNA that is naturally present in bacteria. Some bacteria cells can have no plasmids or several copies of one. They can replicate independently of the host chromosome. Plasmids are circular and double stranded. They carry few genes and their size ranges from 1 to over 200 kilobase pairs. Some functions of their genes include: providing resistance to antibiotics, producing toxins and the breakdown of natural products. However, plasmids are not limited to bacteria; they are also present in some eukaryotes (e.g., circular, nuclear plasmids in Dictyostelium purpureum). A plasmid is a circular, double stranded DNA that is usually found in bacteria (however it does occur in both eukarya and prokarya). It replicates on its own (without the help of chromosomal DNA)and are used frequently in recombinant DNA research in order to replicate genes of interest. Some plasmids can be implanted into a bacterial or animal chromosome in which it becomes a part of the cell's genome and then reveals the gene of interest as a phenotype. This is how much research is done today for gene identification. Plasmids contain three components: an origin of replication, a polylinker to clone the gene of interest (called multiple cloning site where the restriction enzymes cleave), and an antibiotic resistance gene (selectable marker). Plasmids are usually isolated before they are used in recombinant techniques. Alkaline lysis is the method of choice for isolating circular plasmid DNA. This process is quick and reliable. You first obtain the cell that has the plasmid of interest and lyse it with alkali. This step is then followed by extracting the plasmid. The cell fragments are precipitated by using SDS and potassium acetate. This is spun down, and the pellet (cell/cell fragments) is removed. Next, the plasmid DNA is precipitated from the supernatant with the use of isopropanol. The plasmid is then suspended in buffer. Akaline lysis can give you different amounts of plasmid depending if it's a mini-, midi-, or maxi- prep. Plasmids can be related to viruses because they can be independent life-forms due to their ability to self-replicate inside their host. Though they may be viewed as independent life-forms, they have a sense of dependency on their host. A plasmid and its host tend to have a symbiotic relationship. Plasmids can give their hosts needed packages of DNA carrying genes that can lead to mutual survival during tough times. Providing its host with such genetic information, plasmid allows the host to survive and at the same time allows itself to continue living in the host for generations. Plasmids are used as vectors to clone DNA in bacteria. One example of a plasmid used for DNA cloning is called pBR322 Plasmid. The pBR322 plasmid contains a gene that allow the bacteria to be resistant to the antibiotics tetracycline and amipicillin. To use pBR322 plasmid to clone a gene, a restriction endonuclease first cleaves the plasmid at a restriction site. pBR322 plasmid contains three restriction sites: PstI, SalI and ecoRI. The first two restriction sites are located within the gene that codes for ampicillin and tetracycline resistance, respectively. Cleaving at either restriction site will inactivate their respective genes and antibiotic resistance. The target DNA is cleaved with a restriction endonuclease at the same restriction site. The target DNA is then annealed to the plasmid using DNA ligase. After the target DNA is incorporated into the plasmid, the host cell is grown in a environment containing ampicillin or tetracycline, depending on which gene was left active. Many copies of the target DNA is created once the host is able to replicate. Another plasmid used as a vector to clone DNA is called pUC18 plasmid. This plasmid contains a gene that makes the host cell ampicillin resistant. It also contains a gene that allows it produce beta-galactosidase, which is an enzyme degrades certain sugars. The enzyme produces a blue pigment when exposed to a specific substrate analog. This allows the host to be readily identified. The gene for beta-galactosidase contains a polylinker region that contains several restriction sites. The pUC18 plasmid can be cleaved by several different restriction endonucleases which provide more versatility. When the polylinker sequence is cleaved and the target DNA is introduced and ligased, this inactivates the gene that codes for beta-galactosidase and the enzyme will not be produced. The host cell will not produce a blue pigment when exposed to the substrate analog. This allows the recombinant cells to be readily identified and isolated. - Antibiotic resistance gene - origin of replication - inserted gene - restriction enzymes - 5' site , 3' site - promoter

An enzymatic method of DNA extraction:

Actually, this method is a combination of a salt method as well as enzymatic method. Here the extraction buffer is used before going further on enzymatic digestion. The extraction buffer composition may vary from lab to lab, however, the major components are Tris, EDTA, NaCl, sodium lauryl and SDS. Here phenol, chloroform or isoamyl alcohol is not used. Instead, the enzyme proteinase K is utilized for digesting the sample. The sample is incubated with proteinase K for 2 hours this will digest all the protein present inside the sample. Immediately after the proteinase K digestion, the sample is precipitated by chilled alcohol. By centrifuging sample, all other cell debris are removed. Finally, the DNA pellet is dissolved in TE buffer. This method of DNA extraction is rapid and easy. We can use ready to used DNA extraction buffer. Even the yield is very high. However, the quality of DNA is a major concern for this method.

Steps of DNA cloning

DNA cloning is used for many purposes. As an example, let's see how DNA cloning can be used to synthesize a protein (such as human insulin) in bacteria. The basic steps are: Cut open the plasmid and "paste" in the gene. This process relies on restriction enzymes (which cut DNA) and DNA ligase (which joins DNA). Insert the plasmid into bacteria. Use antibiotic selection to identify the bacteria that took up the plasmid. Grow up lots of plasmid-carrying bacteria and use them as "factories" to make the protein. Harvest the protein from the bacteria and purify it.

nick translation

DNA to be processed is treated with DNAase to produce ss nicks is a tagging technique in molecular biology in which DNA Polymerase I is used to replace some of the nucleotides of a DNA sequence with their labeled analogues, creating a tagged DNA sequence which can be used as a probe in fluorescent in situ hybridization (FISH) or blotting techniques. It can also be used for radiolabeling This process is called nick translation because the DNA to be processed is treated with DNAase to produce single-stranded "nicks". This is followed by replacement in nicked sites by DNA polymerase I, which elongates the 3' hydroxyl terminus, removing nucleotides by 5'-3' exonuclease activity, replacing them with dNTPs. To radioactively label a DNA fragment for use as a probe in blotting procedures, one of the incorporated nucleotides provided in the reaction is radiolabeled in the alpha phosphate position. Similarly, a fluorophore can be attached instead for fluorescent labelling, or an antigen for immunodetection. When DNA polymerase I eventually detaches from the DNA, it leaves another nick in the phosphate backbone. The nick has "translated" some distance depending on the processivity of the polymerase. This nick could be sealed by DNA ligase, or its 3' hydroxyl group could serve as the template for further DNA polymerase I activity. Proprietary enzyme mixes are available commercially to perform all steps in the procedure in a single incubation. Nick translation could cause double-stranded DNA breaks, if DNA polymerase I encounters another nick on the opposite strand, resulting in two shorter fragments. This does not influence the performance of the labelled probe in in-situ hybridization.

Transfection and Selection

Following ligation, the recombinant DNA is placed into a host cell, usually bacterial, in a process called transfection or transformation. Finally, the transfected cells are cultured. Many of these cultures may not contain a plasmid with the target DNA as the transfection process is not usually 100% successful, so the appropriate cultures with the DNA of interest must be selected. Many plasmids/vectors include selectable markers - usually some sort of antibiotic resistance (Figureabove). When cultures are grown in the presence of an antibiotic, only bacteria transfected with the vector containing resistance to that antibiotic should grow. However, these selection procedures do not guarantee that the DNA of insert is present in the cells. Further analysis of the resulting colonies is required to confirm that cloning was successful. This may be accomplished by means of a process PCR or restriction fragment analysis, both of which need to be followed by gel electrophoresis and/or DNA sequencing (DNA sequence analysis). DNA sequence analysis, PCR, or restriction fragment analysis will all determine if the plasmid/vector contains the insert. Restriction fragment analysis is digestion of isolated plasmid/vector DNA with restriction enzymes. If the isolated DNA contains the target DNA, that fragment will be excised by the restriction enzyme digestion. Gel electrophoresis will separate DNA molecules based on size and charge.

Gel Electrophoresis

Gel electrophoresis is an analytical technique used to separate DNA fragments by size and charge. Notice in Figure below that the "gels" are rectangular in shape. The gels are made of a gelatin-like material of either agarose or polyacrylamide. An electric field, with a positive charge applied at one end of the gel, and a negative charge at the other end, forces the fragments to migrate through the gel. DNA molecules migrate from negative to positive charges due to the net negative charge of the phosphate groups in the DNA backbone. Longer molecules migrate more slowly through the gel matrix. After the separation is completed, DNA fragments of different lengths can be visualized using a fluorescent dye specific for DNA, such as ethidium bromide. The resulting stained gel shows bands correspond to DNA molecules of different lengths, which also correspond to different molecular weights. Band size is usually determined by comparison to DNA ladders containing DNA fragments of known length. Gel electrophoresis can also be used to separate RNA molecules and proteins.

summary

Gene cloning, also known as molecular cloning, refers to the process of isolating a DNA sequence of interest for the purpose of making multiple copies of it. Classic gene cloning involves the following steps: - Restriction enzyme digestion and ligation Isolation of DNA - Ligation - Transfection and Selection - Gel electrophoresis

Cutting and pasting DNA

How can pieces of DNA from different sources be joined together? A common method uses two types of enzymes: restriction enzymes and DNA ligase. A restriction enzyme is a DNA-cutting enzyme that recognizes a specific target sequence and cuts DNA into two pieces at or near that site. Many restriction enzymes produce cut ends with short, single-stranded overhangs. If two molecules have matching overhangs, they can base-pair and stick together. However, they won't combine to form an unbroken DNA molecule until they are joined by DNA ligase, which seals gaps in the DNA backbone. Our goal in cloning is to insert a target gene (e.g., for human insulin) into a plasmid. Using a carefully chosen restriction enzyme, we digest: The plasmid, which has a single cut site The target gene fragment, which has a cut site near each end Then, we combine the fragments with DNA ligase, which links them to make a recombinant plasmid containing the gene.

protein interactions- western blotting SDS page

In brief, the sample undergoes protein denaturation, followed by gel electrophoresis. A synthetic or animal-derived antibody (known as the primary antibody) is created that recognises and binds to a specific target protein. The electrophoresis membrane is washed in a solution containing the primary antibody, before excess antibody is washed off. A secondary antibody is added which recognises and binds to the primary antibody. The secondary antibody is visualised through various methods such as staining, immunofluorescence, and radioactivity, allowing indirect detection of the specific target protein.

isolation of DNA

Initially, the DNA fragment to be cloned must be isolated. This DNA of interest may be a gene, part of a gene, a promoter, or another segment of DNA, and is frequently isolated by the polymerase chain reaction (PCR) or restriction enzyme digestion. As discussed above, a restriction enzyme is an enzyme that cuts double-stranded DNA at a specific sequence. The enzyme makes two incisions, one through each strand of the double helix, without damaging the nitrogenous bases. This produces either overlapping ends (also known as sticky ends) or blunt ends. The 1978 Nobel Prize in Medicine was awarded to Daniel Nathans and Hamilton Smith for the discovery of restriction endonucleases. The first practical use of their work was the manipulation of E. coli bacteria to produce human insulin for diabetics.

LYSIS OF CELL WALL/CELL MEMBRANE

Lysis of cell wall/ cell membrane: Chemical disruption enzymatic disruption Mechanical disruption Lysis of nuclear membrane: Chemical lysis Enzymatic lysis Removing cell debris • Centrifugation

Next Generation sequencing

Multiple technologies available (454, Illumina, Iontorrent etc) Shorter reads (up to 100bp) but up tens of millions reads per experiment - PacBio sequenecing now allows for long reads Much cheaper :$ 48,000 genome vs $ 300 million Human Genome Project (2004) or $10 million in 2008.

TaqMan

Most popular method of quantitative PCR; two primers and one probe; probe has a fluorescent (reporter) dye on 5' end and a quencher molecule on 3' end; as long as probe is intact, fluorescence is quenched; during DNA synthesis, Taq polymerase cleaves the probe, releasing the dye TaqMan probes are hydrolysis probes that are designed to increase the specificity of quantitative PCR The TaqMan probe principle relies on the 5´-3´ exonuclease activity of Taqpolymerase to cleave a dual-labeled probe during hybridization to the complementary target sequence and fluorophore-based detection As in other quantitative PCRmethods, the resulting fluorescence signal permits quantitative measurements of the accumulation of the product during the exponential stages of the PCR; however, the TaqMan probe significantly increases the specificity of the detection TaqMan probes consist of a fluorophore covalently attached to the 5'-end of the oligonucleotide probe and a quencher at the 3'-end. Several different fluorophores (e.g. 6-carboxyfluorescein, acronym: FAM, or tetrachlorofluorescein, acronym: TET) and quenchers (e.g. tetramethylrhodamine, acronym: TAMRA) are available. The quencher molecule quenches the fluorescence emitted by the fluorophore when excited by the cycler's light source via Förster resonance energy transfer(FRET).As long as the fluorophore and the quencher are in proximity, quenching inhibits any fluorescence signals. TaqMan probes are designed such that they anneal within a DNA region amplified by a specific set of primers. (Unlike the diagram, the probe binds to single stranded DNA.) TaqMan probes can be conjugated to a minor groove binder (MGB) moiety, dihydrocyclopyrroloindole tripeptide (DPI3), in order to increase its binding affinity to the target sequence; MGB-conjugated probes have a higher melting temperature (Tm) due to increased stabilisation of van der Waals forces. As the Taq polymerase extends the primer and synthesizes the nascent strand (again, on a single-strand template, but in the direction opposite to that shown in the diagram, i.e. from 3' to 5' of the complementary strand), the 5' to 3' exonucleaseactivity of the Taq polymerase degrades the probe that has annealed to the template. Degradation of the probe releases the fluorophore from it and breaks the proximity to the quencher, thus relieving the quenching effect and allowing fluorescence of the fluorophore. Hence, fluorescence detected in the quantitative PCR thermal cycler is directly proportional to the fluorophore released and the amount of DNA template present in the PCR. 1) polymerisation: a fluorescent reporter dye and a quencher are attached to the 5' and 3' ends of a taqman probe 2) strand displacement: when the probe is intact, the reported dye emission is quenched 3) cleavage: During each extension cycle, the DNA polymerase cleaves the reporter from the probe 4) polymerisation completed: once separated from the quencher, the reporter dye emits its characteristic fluorescence. an oligonucleotide probe is constructed containing a fluorescent reporter dye on the 5' end and a quencher dye on the 3' end. while the probe is intact, the proximity of the quencher dye greatly reduces the fluorescence resonance energy transfer ( FRET) if the target sequence is present, the probe anneals downstream from one of the primer sites and is cleaved by the 5' nuclease activity of Taq DNA polymerase as this primer is extended cleavage of the probe separates the reporter dye from the quencher dye increasing the reporter dye signal

quantitative gene expression analysis - SYBR Green

Non-specific intercalation into the minor groove of dsDNA, can be used in qPCR Sso7d is a double-stranded (ds) DNA-binding protein that increases speed and processivity and increases tolerance to PCR inhibitors. SYBR® Green is a dsDNA-binding dye that intercalates nonspecifically into dsDNA, allowing measurement of the amount of PCR product It is used in quantitative PCR because the fluorescence can be measured at the end of each amplification cycle to determine, relatively or absolutely, how much DNA has been amplified.

Solid-phase DNA extraction method:

Nowadays all the DNA extraction kits available are based on the unique chemistry of the solid/ liquid phase DNA extraction.The silica is the solid substance which binds with DNA during purification along with it, different solutions are used to purify the DNA. The solid phase silica is one of them.The main advantage of silica gel- based DNA extraction is that it is rapid and gives "PCR ready DNA" for the downstream applications. No extraction or precipitation steps are required therefore this method is superior among all.

Ligation

Once the DNA of interest is isolated, a ligation procedure is necessary to insert the amplified fragment into a vector to produce the recombinant DNA molecule. Restriction fragments (or a fragment and a plasmid/vector) can be spliced together, provided their sticky ends are complementary. Blunt end ligation is also possible. The plasmid or vector (which is usually circular) is digested with restriction enzymes, opening up the vector to allow insertion of the target DNA. If the isolated DNA of interest and the plasmid or vector are digested with the same restriction enzyme, their sticky ends will be complementary. The two DNAs are then incubated with DNA ligase, an enzyme that can attach together strands of DNA with double strand breaks. This produces a recombinant DNA molecule. below depicts a plasmid with two additional segments of DNA ligated into the plasmid, producing the recombinant DNA molecule.

Protein production

Once we have found a bacterial colony with the right plasmid, we can grow a large culture of plasmid-bearing bacteria. Then, we give the bacteria a chemical signal that instructs them to make the target protein. The bacteria serve as miniature "factories," churning out large amounts of protein. For instance, if our plasmid contained the human insulin gene, the bacteria would start transcribing the gene and translating the mRNA to produce many molecules of human insulin protein. Once the protein has been produced, the bacterial cells can be split open to release it. There are many other proteins and macromolecules floating around in bacteria besides the target protein (e.g., insulin). Because of this, the target protein must be purified, or separated from the other contents of the cells by biochemical techniques. The purified protein can be used for experiments or, in the case of insulin, administered to patients.

DNA extraction by magnetic beads:

Positively charged magnetic beads attract the negatively charged DNA. The DNA is separated under the magnetic field. DNA extraction buffer is needed in this technique also

southern blotting method

Restriction endonucleases are used to cut high-molecular-weight DNA strands into smaller fragments. The DNA fragments are then electrophoresed on an agarose gel to separate them by size. If some of the DNA fragments are larger than 15 kb, then prior to blotting, the gel may be treated with an acid, such as dilute HCl. This depurinates the DNA fragments, breaking the DNA into smaller pieces, thereby allowing more efficient transfer from the gel to membrane. If alkaline transfer methods are used, the DNA gel is placed into an alkaline solution (typically containing sodium hydroxide) to denature the double-stranded DNA. The denaturation in an alkaline environment may improve binding of the negatively charged thymine residues of DNA to a positively charged amino groups of membrane, separating it into single DNA strands for later hybridization to the probe (see below), and destroys any residual RNA that may still be present in the DNA. The choice of alkaline over neutral transfer methods, however, is often empirical and may result in equivalent results.[citation needed] A sheet of nitrocellulose (or, alternatively, nylon) membrane is placed on top of (or below, depending on the direction of the transfer) the gel. Pressure is applied evenly to the gel (either using suction, or by placing a stack of paper towels and a weight on top of the membrane and gel), to ensure good and even contact between gel and membrane. If transferring by suction, 20X SSC buffer is used to ensure a seal and prevent drying of the gel. Buffer transfer by capillary action from a region of high water potential to a region of low water potential (usually filter paper and paper tissues) is then used to move the DNA from the gel onto the membrane; ion exchange interactions bind the DNA to the membrane due to the negative charge of the DNA and positive charge of the membrane. The membrane is then baked in a vacuum or regular oven at 80 °C for 2 hours (standard conditions; nitrocellulose or nylon membrane) or exposed to ultraviolet radiation (nylon membrane) to permanently attach the transferred DNA to the membrane. The membrane is then exposed to a hybridization probe—a single DNA fragment with a specific sequence whose presence in the target DNA is to be determined. The probe DNA is labelled so that it can be detected, usually by incorporating radioactivity or tagging the molecule with a fluorescent or chromogenic dye. In some cases, the hybridization probe may be made from RNA, rather than DNA. To ensure the specificity of the binding of the probe to the sample DNA, most common hybridization methods use salmon or herring sperm DNA for blocking of the membrane surface and target DNA, deionized formamide, and detergents such as SDS to reduce non-specific binding of the probe. After hybridization, excess probe is washed from the membrane (typically using SSC buffer), and the pattern of hybridization is visualized on X-ray film by autoradiography in the case of a radioactive or fluorescent probe, or by development of colour on the membrane if a chromogenic detection method is used. Result: Hybridization of the probe to a specific DNA fragment on the filter membrane indicates that this fragment contains DNA sequence that is complementary to the probe. The transfer step of the DNA from the electrophoresis gel to a membrane permits easy binding of the labeled hybridization probe to the size-fractionated DNA. It also allows for the fixation of the target-probe hybrids, required for analysis by autoradiography or other detection methods. Southern blots performed with restriction enzyme-digested genomic DNA may be used to determine the number of sequences (e.g., gene copies) in a genome. A probe that hybridizes only to a single DNA segment that has not been cut by the restriction enzyme will produce a single band on a Southern blot, whereas multiple bands will likely be observed when the probe hybridizes to several highly similar sequences (e.g., those that may be the result of sequence duplication). Modification of the hybridization conditions (for example, increasing the hybridization temperature or decreasing salt concentration) may be used to increase specificity and decrease hybridization of the probe to sequences that are less than 100% similar. - detection genomic re-arrangment - checking recombination in generating KO mouse

DNA extraction using the anionic resins:

The Chelex the product of Bio-Rad laboratories is the best example of the anionic resin used in the DNA extraction. The positively charged chelex binds to the negatively charged phosphate of DNA and helps in the extraction of DNA. However, sometimes several negatively charges contaminants also interfere with DNA extraction by binding with the anion. The chelex is made up of the styrene-divinylbenzene copolymers. The method for using chelex as an anionic resin for DNA extraction was first described by Walsh et al., in the year 1991, the sample they had used was the blood. Another resin used by Seligson et al., was the diethylaminoethyl. In this method, the column of the tube is filled with positive resins. The cell lysate passes through the matrix and DNA binds to the positively charged resins. By applying the low concentration salt buffer, proteins and other impurities or debris are washed off and only DNA remains into the matrix. In the final step filling the matrix with the high concentration salt buffer the DNA is eluted from the resins and precipitated using the alcohol.The method is also called as DNA extraction through anion exchange chromatography.

PNK

The DNA 5′ End-Labeling System is a complete system for phosphorylating both oligonucleotides and DNA fragments using T4 polynucleotide kinase (PNK), which catalyzes the transfer of the terminal [γ-32P]phosphate of ATP to the 5′-hydroxyl terminus of a DNA molecule

classic Sanger sequencing

The classical chain-termination method requires a single-stranded DNA template, a DNA primer, a DNA polymerase, normal deoxynucleotidetriphosphates (dNTPs), and modified di-deoxynucleotidetriphosphates (ddNTPs), the latter of which terminate DNA strand elongation. These chain-terminating nucleotides lack a 3'-OH group required for the formation of a phosphodiester bond between two nucleotides, causing DNA polymerase to cease extension of DNA when a modified ddNTP is incorporated. The ddNTPs may be radioactively or fluorescently labelled for detection in automated sequencing machines. The DNA sample is divided into four separate sequencing reactions, containing all four of the standard deoxynucleotides (dATP, dGTP, dCTP and dTTP) and the DNA polymerase. To each reaction is added only one of the four dideoxynucleotides (ddATP, ddGTP, ddCTP, or ddTTP), while the other added nucleotides are ordinary ones. The dideoxynucleotide concentration should be approximately 100-fold higher than that of the corresponding deoxynucleotide (e.g. 0.5mM ddTTP : 0.005mM dTTP) to allow enough fragments to be produced while still transcribing the complete sequence(but the concentration of ddNTP also depends on the desired length of sequence).[2] Putting it in a more sensible order, four separate reactions are needed in this process to test all four ddNTPs. Following rounds of template DNA extension from the bound primer, the resulting DNA fragments are heat denatured and separated by size using gel electrophoresis. In the original publication of 1977,[2] the formation of base-paired loops of ssDNA was a cause of serious difficulty in resolving bands at some locations. This is frequently performed using a denaturing polyacrylamide-urea gel with each of the four reactions run in one of four individual lanes (lanes A, T, G, C). The DNA bands may then be visualized by autoradiography or UV light and the DNA sequence can be directly read off the X-ray film or gel image. Part of a radioactively labelled sequencing gel In the image on the right, X-ray film was exposed to the gel, and the dark bands correspond to DNA fragments of different lengths. A dark band in a lane indicates a DNA fragment that is the result of chain termination after incorporation of a dideoxynucleotide (ddATP, ddGTP, ddCTP, or ddTTP). The relative positions of the different bands among the four lanes, from bottom to top, are then used to read the DNA sequence.

Silica column-based DNA extraction method

The silica column-based DNA extraction method is very unique and different from other DNA extraction methods. In the PCI or proteinase K method we have to centrifuge sample many times and have to collect aqueous phase or pellets depending upon the step of extraction. Sometimes we have to collect aqueous phase or sometimes we have to collect pellets. The silica-based DNA extraction method works on the unique chemistry of interaction between silica and DNA. A positively charged silica particles bind with the negatively charged DNA and hold it during centrifugation. The silica-based solid-phase DNA extraction method is now commercially available and it is most routinely used in diagnostic laboratories. Because of its good quality DNA yield and minimal simple operating system, it is widely accepted. The lysis buffer breaks the cell membrane and the nuclear envelope. The proteinase K digests all the protein. In the very first step, the sample is incubated with a cell lysis buffer or called a DNA extraction buffer.Along with it, a small amount of proteinase k is added to the sample. All the other impurities are removed by centrifugation. Here the DNA remains bounded with silica and other impurities passes through the silica column. Now the DNA can be washed twice for improving the purity in it. The aqueous phase contains the impurities are discarded by discarding the collection tube. Finally, the DNA is dissolved into the TE buffer. The method is fast reliable, accurate and consumes less time as compared to other methods. Other DNA extraction methods are DNA extraction using the anionic resins, magnetic bead DNA extraction method and CsCl density gradient DNA extraction method.

Phenol-chloroform method of DNA extraction:

This method is one of the best methods of DNA extraction. The yield and quality of DNA obtained by the PCI method is very good if we perform it well. The method is also called as a phenol-chloroform and isoamyl alcohol, PCI method of DNA extraction. The major chemicals of PCI DNA extraction methods are lysis buffer, Phenol and chloroform. The lysis buffer contains Tris, EDTA, MgCl2, NaCl, SDS, and other salts. Here the components of lysis buffer help in lysis of cell membrane as well as the nuclear envelope. The protein portion of the cell denatured with the help of chloroform and phenol which are organic in nature.

Illumina method

This sequencing method is based on reversible dye-terminators that enable the identification of single nucleotides as they are washed over DNA strands. It can also be used for whole-genome and region sequencing, transcriptome analysis, metagenomics, small RNA discovery, methylation profiling, and genome-wide protein-nucleic acid interaction analysis Illumina sequencing technology works in three basic steps: amplify, sequence, and analyze. The process begins with purified DNA. The DNA is fragmented and adapters are added that contain segments that act as reference points during amplification, sequencing, and analysis. The modified DNA is loaded onto a flow cell where amplification and sequencing will take place. The flow cell contains nanowells that space out fragments and help with overcrowding.[6]Each nanowell contains oligonucleotides that provide an anchoring point for the adaptors to attach. Once the fragments have attached, a phase called cluster generation begins. This step makes about a thousand copies of each fragment of DNA and is done by bridge amplification PCR. Next, primers and modified nucleotides are washed onto the chip. These nucleotides have a reversible 3' fluorescent blocker so the DNA polymerase can only add one nucleotide at a time onto the DNA fragment.[6] After each round of synthesis, a camera takes a picture of the chip. A computer determines what base was added by the wavelength of the fluorescent tag and records it for every spot on the chip. After each round, non-incorporated molecules are washed away. A chemical deblocking step is then used to remove the 3' fluorescent terminal blocking group.. The process continues until the full DNA molecule is sequenced.[5] With this technology, thousands of places throughout the genome are sequenced at once via massive parallel sequencing. - Genomic Library : After the DNA is purified a DNA library, genomic library, needs to be generated. There are two ways a genomic library can be created, sonification and tagmentation. With tagmentation, transposases randomly cuts the DNA into 500 bp fragments and adds adaptors simultaneously.[6] A genetic library can also be generated by using sonification to fragment genomic DNA. Sonification fragments DNA into similar sizes using ultrasonic sound waves. Right and left adapters will need to be attached by T7 DNA Polymerase and T4 DNA ligase after sonification. Strands that fail to have adapters ligated are washed away Adapters: Adapters contain three different segments: the sequence complementary to solid support (oligonucleotides on flow cell), the barcode sequence (indices), and the binding site for the sequencing primer.[6] Indices are usually six base pairs long and are used during DNA sequence analysis to identify samples. Indices allow for up to 96 different samples to be run together, this is also known as multiplexing. During analysis, the computer will group all reads with the same index together. Illumina uses a "sequence by synthesis" approach.This process takes place inside of an acrylamide-coated glass flow cell.The flow cell has oligonucleotides (short nucleotide sequences) coating the bottom of the cell, and they serve as the solid support to hold the DNA strands in place during sequencing. As the fragmented DNA is washed over the flow cell, the appropriate adapter attached to the complementary solid support. Bridge amplification: Once attached, cluster generation can begin. The goal is to create hundreds of identical strands of DNA. Some will be the forward strand; the rest, the reverse. This is why right and left adapters are used. Clusters are generated through bridge amplification. DNA polymerase moves along a strand of DNA, creating its complementary strand. The original strand is washed away, leaving only the reverse strand. At the top of the reverse strand there is an adapter sequence. The DNA strand bends and attaches to the oligo that is complementary to the top adapter sequence. Polymerases attach to the reverse strand, and its complementary strand (which is identical to the original) is made. The now double stranded DNA is denatured so that each strand can separately attach to an oligonucleotide sequence anchored to the flow cell. One will be the reverse strand; the other, the forward. This process is called bridge amplification, and it happens for thousands of clusters all over the flow cell at once clonal amplification: Over and over again, DNA strands will bend and attach to the solid support. DNA polymerase will synthesize a new strand to create a double stranded segment, and that will be denatured so that all of the DNA strands in one area are from a single source (clonal amplification). Clonal amplification is important for quality control purposes. If a strand is found to have an odd sequence, then scientists can check the reverse strand to make sure that it has the complement of the same oddity. The forward and reverse strands act as checks to guard against artefacts. Because Illumina sequencing uses DNA polymerase, base substitution errors have been observed,[12] especially at the 3' end.[13] Paired end reads combined with cluster generation can confirm an error took place. The reverse and forward strands should be complementary to each other, all reverse reads should match each other, and all forward reads should match each other. If a read is not similar enough to its counterparts (with which it should be a clone), an error may have occurred. A minimum threshold of 97% similarity has been used in some labs' analyses Sequence by synthesis: At the end of clonal amplification, all of the reverse strands are washed off the flow cell, leaving only forward strands. A primer attaches to the forward strands adapter primer binding site, and a polymerase adds a fluorescently tagged dNTP to the DNA strand. Only one base is able to be added per round due to the fluorophore acting as a blocking group; however, the blocking group is reversible.[6] Using the four-color chemistry, each of the four bases has a unique emission, and after each round, the machine records which base was added. Once the color is recorded the fluorophore is washed away and another dNTP is washed over the flow cell and the process is repeated. dATPs, dTTPs, dGTPs, and dCTPs are washed over the cell separately so each nucleotide is able to be identified. Starting with the launch of the NextSeq and later the MiniSeq, Illumina introduced a new two-color sequencing chemistry. Nucleotides are distinguished by either one of two colors (red or green), no color ("black") or combining both colors (appearing orange as a mixture between red and green).Once the DNA strand has been read, the strand that was just added is washed away. Then, the index 1 primer attaches, polymerizes the index 1 sequence, and is washed away. The strand forms a bridge again, and the 3' end of the DNA strand attaches to an oligo on the flow cell. The index 2 primer attaches, polymerizes the sequence, and is washed away. A polymerase sequences the complementary strand on top of the arched strand. They separate, and the 3' end of each strand is blocked. The forward strand is washed away, and the process of sequence by synthesis repeats for the reverse strand. Data analysis: The sequencing occurs for millions of clusters at once, and each cluster has ~1,000 identical copies of a DNA insert. The sequence data is analyzed by finding fragments with overlapping areas, called contigs, and lining them up. If a reference sequence is known, the contigs are then compared to it for variant identification. This piecemeal process allows scientists to see the complete sequence even though an unfragmented sequence was never run; however, because Illumina read lengths are not very long (HiSeq sequencing can produce read lengths around 90 bp long), it can be a struggle to resolve short tandem repeat areas.Also, if the sequence is de novo and a reference doesn't exist, repeated areas can cause a lot of difficulty in sequence assembly.Additional difficulties include base substitutions (especially at the 3' end of reads) by inaccurate polymerases, chimeric sequences, and PCR-bias, all of which can contribute to generating an incorrect sequence

bacterial transformation and screening

after ligating the DNA into the plasmid, you'll have to transform by either : 1) electroporation ( heat-shock) 2) chemically competent cells ( CACL2) to facilitate the attachment of plasmid dna to competent cell membranes, alternatively heated in a water bath opens pores of cell membranes allowing entry of the plasmid

check the notebook for other types of interactions

check the notebook for other types of interactions

how to analyse gene expression

proteins: - western blotting - immunohistochemistry - insitu hybridisation - northern blotting -immunofluroscence DNA/RNA: PCR/ q-PCR/ RT-PCR/microarray

Silica basedDNA/RNA Purification: Kits ( QiaAmp spin)

spin columns contain silica resin that selectively binds DNA/RNA depending on salt conditions influenced by extraction methods which results in high quality material for cloning sample - lyse - bind - wash buffer- elute - DNA

Molecular mechanism

β-galactosidase is a protein encoded by the lacZ gene of the lac operon, and it exists as a homotetramer in its active state. However, a mutant β-galactosidase derived from the M15 strain of E. coli has its N-terminal residues 11—41 deleted and this mutant, the ω-peptide, is unable to form a tetramer and is inactive. This mutant form of protein however may return fully to its active tetrameric state in the presence of an N-terminal fragment of the protein, the α-peptide. The rescue of function of the mutant β-galactosidase by the α-peptide is called α-complementation. In this method of screening, the host E. coli strain carries the lacZ deletion mutant (lacZΔM15) which contains the ω-peptide, while the plasmids used carry the lacZαsequence which encodes the first 59 residues of β-galactosidase, the α-peptide. Neither is functional by itself. However, when the two peptides are expressed together, as when a plasmid containing the lacZα sequence is transformed into a lacZΔM15 cells, they form a functional β-galactosidase enzyme. The blue/white screening method works by disrupting this α-complementation process. The plasmid carries within the lacZα sequence an internal multiple cloning site (MCS). This MCS within the lacZα sequence can be cut by restriction enzymes so that the foreign DNA may be inserted within the lacZα gene, thereby disrupting the gene that produces α-peptide. Consequently, in cells containing the plasmid with an insert, no functional β-galactosidase may be formed. The presence of an active β-galactosidase can be detected by X-gal, a colourless analog of lactose that may be cleaved by β-galactosidase to form 5-bromo-4-chloro-indoxyl, which then spontaneously dimerizes and oxidizes to form a bright blue insoluble pigment 5,5'-dibromo-4,4'-dichloro-indigo. This results in a characteristic blue colour in cells containing a functional β-galactosidase. Blue colonies therefore show that they may contain a vector with an uninterrupted lacZα (therefore no insert), while white colonies, where X-gal is not hydrolyzed, indicate the presence of an insert in lacZα which disrupts the formation of an active β-galactosidase. The recombinant clones can be further analyzed by isolating and purifying small amounts of plasmid DNA from the transformed colonies and restriction enzymes can be used to cut the clone and determine if it has the fragment of interest. If the DNA is necessary to be sequenced, the plasmids from the colonies will need to be isolated at a point, whether to cut using restriction enzymes or performing other assays.

modification of sites/ blunting sticky ends

5 ́overhangs are filled in by the polymerase activity of Klenow fragment whereas 3 ́overhangs are removed by the 3 ́to 5 ́exonuclease activity of the enzyme. Due to the relatively weak exonuclease activity of Klenow fragment 3 ́overhang repair is much less efficient than 5 ́overhang repair. its the process where the single strand overhand created by restriction digest is either:- 1) 5' overhang = filled in by the addition of nucleortides on the complimentary strand using the overhang as a template for polymerisation ( 5' Klenow, 5' to 3' fill) 2) 3' overhang= chewing back the overhang using an exonuclease activity ( 3' Klenow fragment, dna pol 1 , T4 dna pol/ mung bean nuclease ) , removal of the 5' overhang which is much slower vectors and inserts are blunted to allow non-compatible ends to join. sequence information is lost. blunting a region of translated coding sequence creates a shift in the reading frame DNA polymerase activities: - 5' to 3' polymerase activity - 5' to 3' exonuclease - 3' to 5' exonuclease activity removal of overhang: 3' Klenow fragment/ dna pol 1/ t4 dna pol/ 5' mung bean nuclease fill in an overhang: -3' not possible - 5' Klenow, produced from truncated ecoli pol a gene where 5'-3' exonuclease domain is removed

bacterial transformation screening

Because transformation usually produces a mixture of relatively few transformed cells and an abundance of non-transformed cells, a method is necessary to select for the cells that have acquired the plasmid Antibiotic resistance is the most commonly used marker for prokaryotes. The transforming plasmid contains a gene that confers resistance to an antibiotic that the bacteria are otherwise sensitive to. The mixture of treated cells is cultured on media that contain the antibiotic so that only transformed cells are able to grow - plasmids that contain the ABR gene will be able to survive and therefore grow colonies on plates as antibiotics usually kill bacteria 2) Blue- white screening: Reporter genes can be used as markers, such as the lacZ gene which codes for β-galactosidase used in blue-white screening. This method of screening relies on the principle of α-complementation, where a fragment of the lacZ gene (lacZα) in the plasmid can complement another mutant lacZ gene (lacZΔM15) in the cell. Both genes by themselves produce non-functional peptides, however, when expressed together, as when a plasmid containing lacZ-α is transformed into a lacZΔM15 cells, they form a functional β-galactosidase. The presence of an active β-galactosidase may be detected when cells are grown in plates containing X-gal, forming characteristic blue colonies. However, the multiple cloning site, where a gene of interest may be ligated into the plasmid vector, is located within the lacZα gene. Successful ligation therefore disrupts the lacZα gene, and no functional β-galactosidase can form, resulting in white colonies. Cells containing successfully ligated insert can then be easily identified by its white coloration from the unsuccessful blue ones. The blue-white screen is a screening technique that allows for the rapid and convenient detection of recombinant bacteria in vector-based molecular cloning experiments. DNA of interest is ligated into a vector. The vector is then inserted into a competent host cellviable for transformation, which are then grown in the presence of X-gal. Cells transformed with vectors containing recombinant DNA will produce white colonies; cells transformed with non-recombinant plasmids (i.e. only the vector) grow into blue colonies. This method of screening is usually performed using a suitable bacterial strain, but other organisms such as yeast may also be used. The method is based on the principle of α-complementation of the β-galactosidasegene. This phenomenon of α-complementation was first demonstrated in work done by Agnes Ullmann in the laboratory of François Jacob and Jacques Monod, where the function of an inactive mutant β-galactosidase with deleted sequence was shown to be rescued by a fragment of β-galactosidase in which that same sequence, the α-donor peptide, is still intact.[1] Langley et al. showed that the mutant non-functional β-galactosidase was lacking in part of its N-terminus with its residues 11—41 deleted, but it may be complemented by a peptide formed of residues 3—90 of β-galactosidase.[2] M13 filamentous phage containing sequence coding for the first 145 amino acid was later constructed by Messing et al., and α-complementation via the use of a vector was demonstrated by the formation of blue plaques when cells containing the inactive protein were infected by the phage and then grown in plates containing X-gal

Chemical or solution-based DNA extraction method:

Different types of organic and inorganic solutions are used in the chemical or solution-based DNA extraction method.The steps of the DNA extraction remain the same in all the types of DNA extraction methods. SDS, CTAB, phenol, chloroform, isoamyl alcohol, Triton X100, guanidium thiocyanate, Tris and EDTA are several common chemicals used in the solution based DNA extraction method. The solution-based ( also called chemical) DNA extraction method is subdivided into organic solvent-based DNA extraction and inorganic solvent-based DNA extraction. The organic solvent-based DNA extraction method is based on the use of organic substances such as phenol and chloroform. Due to the harmful nature of the phenol and chloroform, the method is restricted. Nonetheless, phenol-chloroform DNA extraction method is one of the best methods among all.Among the inorganic DNA extraction, two are most popular: use of proteinase K and use of salt. The proteinase K DNA extraction method facilitates high DNA yield but the method is time-consuming. Also, if not maintained well in a cold chain, the proteinase K cannot be utilized for a longer period of time.The lower stability of the enzyme is another major issue in this method. Salting out DNA extraction method is safer than the PCI method. The use of salts such as sodium chloride, potassium acetate and ammonium acetate help in the DNA extraction.However, the method is more aggressive in combination with proteinase K. Use of the different salt in DNA extraction can increase the yield but the purity is not good enough.

Uses of DNA cloning

Gene therapy. In some genetic disorders, patients lack the functional form of a particular gene. Gene therapy attempts to provide a normal copy of the gene to the cells of a patient's body. For example, DNA cloning was used to build plasmids containing a normal version of the gene that's nonfunctional in cystic fibrosis. When the plasmids were delivered to the lungs of cystic fibrosis patients, lung function deteriorated less quickly2 Gene analysis. In basic research labs, biologists often use DNA cloning to build artificial, recombinant versions of genes that help them understand how normal genes in an organism function

Bacterial transformation and selection

Plasmids and other DNA can be introduced into bacteria, such as the harmless E. coli used in labs, in a process called transformation. During transformation, specially prepared bacterial cells are given a shock (such as high temperature) that encourages them to take up foreign DNA. A plasmid typically contains an antibiotic resistance gene, which allows bacteria to survive in the presence of a specific antibiotic. Thus, bacteria that took up the plasmid can be selected on nutrient plates containing the antibiotic. Bacteria without a plasmid will die, while bacteria carrying a plasmid can live and reproduce. Each surviving bacterium will give rise to a small, dot-like group, or colony, of identical bacteria that all carry the same plasmid. Not all colonies will necessarily contain the right plasmid. That's because, during a ligation, DNA fragments don't always get "pasted" in exactly the way we intend. Instead, we must collect DNA from several colonies and see whether each one contain the right plasmid. Methods like restriction enzyme digestion and PCR are commonly used to check the plasmids.

Purpose of DNA extraction or DNA isolation:

The main purpose of DNA extraction or DNA isolation is the same, to provide a pure DNA. Direct bodily cells or tissue can not be used in DNA testing. For doing a DNA test for DNA fingerprinting, DNA heritage testing or DNA diagnosis testing we should require a pure DNA. Using different chemicals and solution, all the other organelles and debris of cells are isolated from the DNA and pure DNA is extracted. The pure DNA must have ~1.80 260/280 ratio. Impurities in DNA may result in PCR inhibition. Thus, the purpose of DNA extraction is to provide a pure, unfragmented and highly concentrated DNA for doing a DNA test or PCR.

..

The nuclear membrane and cell membrane is made up of protein and lipids almost. Hence the same types of chemicals can work for both.Chemicals such as SDS, CTAB, Tris and other detergents can lyse the cell wall/ cell membrane by solubilizing it. Each chemical has a different function to play which we will discuss separately in each method Enzymes such as proteinase K, peptidase, protease disrupt proteins by digesting it. The enzyme works better than any other chemicals because it directly targets bonds between the amino acids and digests the protein. Once the cell wall or cell membrane lysed, there are no compartments inside the cell hence all the cell organelles are mixed into the solution. By doing high-speed centrifugation DNA remains in the solution and the other cell debris settled into the bottom of the tube. Moreover, the DNA extraction method varies depending upon the type of cells. Take look at some example here,The cell having a soft cell wall: some of the bacteria have a very smooth and soft cell wall. For example, M.tuberculosis has a smooth cell wall. By only heating the bacterial solution we can lyse cell wall. The supernatant can directly be used for the PCR. Even, by putting the bacterial culture directly into the PCR tube for 15 minutes at initial denaturation we can directly get the good quality of result in PCR. However, the addition of a simple lysis buffer during heating will increase the yield and quality of DNA. The cell having a harder cell wall: Plant cells have pectin and other polysaccharides present in their cell wall. This pectin protects the cell from mechanical damage. Therefore pectin provides additional strength to the cell wall of the plant. The cell having a cell membrane: The cell wall is not present in animal cells. A combination of the enzymatic and chemical method is most suitable for the DNA extraction from animal cells, however, each method individually performs the best.

role of chemicals

Tris: DNA is pH sensitive, Tris buffer maintains the pH of the solution. Also, it interacts with the lipopolysaccharides of the cell membrane and makes them permeable, this will help in lysis of the cell membrane. EDTA: EDTA is a chelating agent and can be used to block DNase activity. DNase is an enzyme which lyses the DNA. However, every enzyme required cofactor to work properly. The chelator EDTA blocks the activity of DNase by blocking the cofactor binding site. It will work best in combination with Tris. SDS: Sodium dodecyl sulphate is an anionic detergent which helps cell membrane and nuclear envelope to break open. NaCl: the Na+ ion of NaCl creates the ionic bond with the negative charge of DNA and neutralize it. It will help DNA comes together and protect from denaturation. MgCl2: overall, it protects the DNA. MgCl2 block the negative charge of the lipoproteins of the cell membrane. After the lysis of cell, there is no compartment in the cell hence it protects DNA by mixing with other cell organelles. Phenol: it precipitates the protein impurities. The SDS removes the negative charges from the amino acid and disrupts the confirmation of a protein. Therefore, the protein loses its structure and stabilized by using the SDS. The combination of phenol, chloroform and isoamyl alcohol helps in the removal of protein. After centrifugation, the phenol settles in the bottom of the tube and DNA in the aqueous phase while the denatured protein remains between both layers as a whitish cloud. So care must be taken to collect nucleic acid from this method. The collected nucleic acid is precipitated with the help ofchilled alcohol (isoamyl alcohol). We can add salt as well to increase the yield of the DNA. Finally, the DNA is dissolved in TE buffer. Because of the use of the organic solvents, this method is also named as an organic solvent-based DNA extraction method. The PCI method of DNA extraction is widely accepted. Even the forensic departments trust PCI method rather than a Kit method. The quantity ofDNA obtains by PCI method is very high. We can obtain 800 to 900 ng of DNA with great quality. If we prepare all chemicals very well and perform it sincerely we can always get a good result by this method. However, the amount of sample required for PCI DNA extraction is high. It is difficult to isolate DNA from the samples such as hair and nail. Also, the purity of DNA becomes a major issue if not performed well.

how to prevent self- ligation ?

When we try to insert a gene of interest in vector, the most common problem faced is of self-ligation.Hence prevention of self-ligation is the most important parameter in obtaining high transformation efficiency. - Double digestion of DNA vector and insert - Dephosphorylation of 5' ends of vectors - Replacing 2'-deoxyribose with 2',3'-dideoxyribose sugar -dephosphorhylation of the 5' end which stops DNA ligase from catalysing the formation of phosphodiester bonds between 3' end hydroxyl and 5' phosphate residue at the DNA ends - however it cannot be applied to both DNA species to be ligated thus the untreated DNA species remains capable of self-ligation hence, modification by removing phosphate groups required ( alkaline phosphatase) 1) DIGESTION WITH 2 DIFFERENT RE = THE MOST BASIC STEP FOR PREVENTION OF SELF LIGATION IS CUTTING THE INSERT AND VECTOR WITH 2 DIFFERENT RESTRICTION ENZYMES, GENERATING FRAGMENTS WITH 2 DIFFERENT RESTRICTION SITES 2)DEPHOSPHORYLATION OF 5'- ENDS OF VECTORS = Removing 5'-phosphate groups from the vectors using phosphatases (e.g. alkaline phosphatase), prevents self- ligation. 3) REPLACING 2'-DEOXYRIBOSE WITH 2',3'-DIDEOXYRIBOSE SUGAR: We know phosphodiester bond forms between 5'P and 3'OH.In this technique, the 3'OH of the vector, which takes part in phosphodiester bond formation, is altered. 2'deoxyribose at the 3'end of the vector is replaced by 2',3'- dideoxyribose. This prevents self ligation of vector. The insert is dephosphorylated.

Gibson cloning

cloning method which allows multiple dna fragments to be joint in a single reaction ( isothermal reaction ) - robust exonuclease based method to assemble dna in the correct order advantages: 1) no restriction digest of DNA fragments after PCR is necessary 2) cheaper and faster ( fewer steps and reagents) 3) up to 5 DNA fragments can be can be combined simultaneously in a single tube reaction 4) Gibson Assembly method can be used for site-directed mutagenesis to incorporate site-specific mutations ( INDELS) it has 3 enzymatic activities= 1) 5' exonuclease generates long overhangs ( T5 exonuclease ). this chews back DNA from the 5' end , not inhibiting polymerase activity and allowing the reaction to occur in one single process. resulting s-s regions on adjacent DNA fragments can anneal 2) 3' extension activity of DNA pol ( phusion polymerase ). this incorporates nucleotides to fill in any gaps 3) DNA ligase = covalently joins the DNA of adjacent segments thereby removing any nicks in the DNA the result is different DNA fragments joint into one

orientation of ligation

proper orientation requires 2 restriction enzymes

analysis of gene expression RT-PCR

technique combining reverse transcription of RNA into DNA (in this context called complementary DNA or cDNA) and amplification of specific DNA targets using polymerase chain reaction (PCR).[1] It is primarily used to measure the amount of a specific RNA. This is achieved by monitoring the amplification reaction using fluorescence, a technique called real-time PCR or quantitative PCR (qPCR). Combined RT-PCR and qPCR are routinely used for analysis of gene expression and quantification of viral RNA in research and clinical settings. Currently, there are four different fluorescent DNA probes available for the real-time RT-PCR detection of PCR products: SYBR Green, TaqMan, molecular beacons, and scorpion probes. All of these probes allow the detection of PCR products by generating a fluorescent signal. While the SYBR Green dye emits its fluorescent signal simply by binding to the double-stranded DNA in solution, the TaqMan probes', molecular beacons' and scorpions' generation of fluorescence depend on Förster Resonance Energy Transfer (FRET) coupling of the dye molecule and a quencher moiety to the oligonucleotide substrates. - allows detection of low abundance RNAs in a sample and production of cDNA

DNA extraction

the process of separating DNA from the rest of the cell 1- extract the DNA from tissue cells and purify them 2- homogenisation blender ( SDS/Prot K), lysis of cell wall and membrane as well as nuclear membrane 3- add phenol and chloroform to purify samples of nucleic acids taken from cells, it's a non-polar compound so nucleic acids which are highly polar won't dissolve in the presence of phenol which has a higher density than water so when phenolic added to a cell sample, the water and phenol will separate as they are centrifuged - the aqueous phase ( polar ) is the top bit which contains the dna/rna - the organic phase ( non polar) is at the bottom which contains denatured proteins phenol is combined with chloroform to ensure clear separation occurs between aqueous and organic phases. the density of chloroform is higher than phenol so both will create a denser solution than with water benefits : Able to extract lots of DNA at once however this is time consuming and requires multiple centrifugation steps last step is to precipitate the ethanol

DNA cloning

the production of multiple copies of a specific DNA segment 1) target gene is inserted into the plasmid 2) plasmid is introduced into bacteria via transformation and the bacteria carrying the plasmid is selected using antibiotics 3) bacteria with correct plasmid is used to make plasmid DNA or induced to express genes and make proteins DNA cloning is the process of making multiple, identical copies of a particular piece of DNA. In a typical DNA cloning procedure, the gene or other DNA fragment of interest (perhaps a gene for a medically important human protein) is first inserted into a circular piece of DNA called a plasmid. The insertion is done using enzymes that "cut and paste" DNA, and it produces a molecule of recombinant DNA, or DNA assembled out of fragments from multiple sources. Next, the recombinant plasmid is introduced into bacteria. Bacteria carrying the plasmid are selected and grown up. As they reproduce, they replicate the plasmid and pass it on to their offspring, making copies of the DNA it contains. What is the point of making many copies of a DNA sequence in a plasmid? In some cases, we need lots of DNA copies to conduct experiments or build new plasmids. In other cases, the piece of DNA encodes a useful protein, and the bacteria are used as "factories" to make the protein. For instance, the human insulin gene is expressed in E. coli bacteria to make insulin used by diabetics.

Dye-terminator sequencing

utilizes fluorescent labeling of the chain terminator ddNTPs, which permits sequencing in a single reaction, rather than four reactions as in the labeled-primer method. Laser and detector "read" the fragments. Dye-terminator sequencing utilizes labelling of the chain terminator ddNTPs, which permits sequencing in a single reaction, rather than four reactions as in the labelled-primer method. In dye-terminator sequencing, each of the four dideoxynucleotide chain terminators is labelled with fluorescent dyes, each of which emit light at different wavelengths. Owing to its greater expediency and speed, dye-terminator sequencing is now the mainstay in automated sequencing. Its limitations include dye effects due to differences in the incorporation of the dye-labelled chain terminators into the DNA fragment, resulting in unequal peak heights and shapes in the electronic DNA sequence trace chromatogram after capillary electrophoresis (see figure to the left). This problem has been addressed with the use of modified DNA polymerase enzyme systems and dyes that minimize incorporation variability, as well as methods for eliminating "dye blobs". The dye-terminator sequencing method, along with automated high-throughput DNA sequence analyzers, was used for the vast majority of sequencing projects until the introduction of Next Generation Sequencing.