DNA Repair

DNA repair refers to a collection of processes by which a cell identifies and corrects damage to the DNA molecules that encode its genome. In human cells, both normal metabolic activities and environmental factors such as UV light can cause DNA damage, resulting in as many as 1 million individual molecular lesions per cell per day.Many of these lesions cause structural damage to the DNA molecule and can alter or eliminate the cell's ability to transcribe the gene that the affected DNA encodes. Other lesions induce potentially harmful mutations in the cell's genome, which affect the survival of its daughter cells after it undergoes mitosis. Consequently, the DNA repair process is constantly active as it responds to damage in the DNA structure.

The rate of DNA repair is dependent on many factors, including the cell type, the age of the cell, and the extracellular environment. A cell that has accumulated a large amount of DNA damage, or one that no longer effectively repairs damage incurred to its DNA, can enter one of three possible states:

1. an irreversible state of dormancy, known as senescence
2. cell suicide, also known as apoptosis or programmed cell death
3. unregulated cell division, which can lead to the formation of a tumor that is cancerous

The DNA repair ability of a cell is vital to the integrity of its genome and thus to its normal functioning and that of the organism. Many genes that were initially shown to influence lifespan have turned out to be involved in DNA damage repair and protection. Failure to correct molecular lesions in cells that form gametes can introduce mutations into the genomes of the offspring and thus influence the rate of evolution.

DNA damage

DNA damage, due to environmental factors and normal metabolic processes inside the cell, occurs at a rate of 1,000 to 1,000,000 molecular lesions per cell per day. While this constitutes only 0.000165% of the human genome's approximately 6 billion bases (3 billion base pairs), unrepaired lesions in critical genes (such as tumor suppressor genes) can impede a cell's ability to carry out its function and appreciably increase the likelihood of tumor formation.

The vast majority of DNA damage affects the primary structure of the double helix; that is, the bases themselves are chemically modified. These modifications can in turn disrupt the molecules' regular helical structure by introducing non-native chemical bonds or bulky adducts that do not fit in the standard double helix. Unlike proteins and RNA, DNA usually lacks tertiary structure and therefore damage or disturbance does not occur at that level. DNA is, however, supercoiled and wound around "packaging" proteins called histones (in eukaryotes), and both superstructures are vulnerable to the effects of DNA damage.

DNA repair mechanisms

Cells cannot function if DNA damage corrupts the integrity and accessibility of essential information in the genome (but cells remain superficially functional when so-called "non-essential" genes are missing or damaged). Depending on the type of damage inflicted on the DNA's double helical structure, a variety of repair strategies restore lost information. If possible, cells use the unmodified complementary strand of the DNA or the sister chromatid as a template to losslessly recover the original information. Without access to a template, cells use an error-prone recovery mechanism known as translesion synthesis as a last resort.

Damage to DNA alters the spatial configuration of the helix and such alterations can be detected by the cell. Once damage is localized, specific DNA repair molecules bind at or near the site of damage, inducing other molecules to bind and form a complex that enables the actual repair to take place. The types of molecules involved and the mechanism of repair that is mobilized depend on the type of damage that has occurred and the phase of the cell cycle that the cell is in.

Direct reversal

Cells are known to eliminate three types of damage to their DNA by chemically reversing it. These mechanisms do not require a template, since the types of damage they counteract can only occur in one of the four bases. Such direct reversal mechanisms are specific to the type of damage incurred and do not involve breakage of the phosphodiester backbone. The formation of thymine dimers (a common type of cyclobutyl dimer) upon irradiation with UV light results in an abnormal covalent bond between adjacent thymidine bases. The photoreactivation process directly reverses this damage by the action of the enzyme photolyase, whose activation is obligately dependent on energy absorbed from blue/UV light (300–500nm wavelength) to promote catalysis. Another type of damage, methylation of guanine bases, is directly reversed by the protein methyl guanine methyl transferase (MGMT), the bacterial equivalent of which is called as ogt. This is an expensive process because each MGMT molecule can only be used once; that is, the reaction is stoichiometric rather than catalytic.A generalized response to methylating agents in bacteria is known as the adaptive response and confers a level of resistance to alkylating agents upon sustained exposure by upregulation of alkylation repair enzymes. The third type of DNA damage reversed by cells is certain methylation of the bases cytosine and adenine.

When only one of the two strands of a double helix has a defect, the other strand can be used as a template to guide the correction of the damaged strand. In order to repair damage to one of the two paired molecules of DNA, there exist a number of excision repair mechanisms that remove the damaged nucleotide and replace it with an undamaged nucleotide complementary to that found in the undamaged DNA strand.

1. Base excision repair (BER), which repairs damage to a single nucleotide caused by oxidation, alkylation, hydrolysis, or deamination. The base is removed with glycosylase and ultimately replaced by repair synthesis with DNA ligase.
2. Nucleotide excision repair (NER), which repairs damage affecting longer strands of 2–30 bases. This process recognizes bulky, helix-distorting changes such as thymine dimers as well as single-strand breaks (repaired with enzymes such UvrABC endonuclease). A specialized form of NER known as Transcription-Coupled Repair (TCR) deploys high-priority NER repair enzymes to genes that are being actively transcribed.
3. Mismatch repair (MMR), which corrects errors of DNA replication and recombination that result in mispaired (but normal, that is non- damaged) nucleotides following DNA replication.
4. 
   

Double-strand breaks

Double-strand breaks (DSBs), in which both strands in the double helix are severed, are particularly hazardous to the cell because they can lead to genome rearrangements. Two mechanisms exist to repair DSBs: non-homologous end joining (NHEJ) and recombinational repair

, a specialized DNA Ligase that forms a complex with the cofactor XRCC4, directly joins the two ends. To guide accurate repair, NHEJ relies on short homologous sequences called microhomologies present on the single-stranded tails of the DNA ends to be joined. If these overhangs are compatible, repair is usually accurate. NHEJ can also introduce mutations during repair. Loss of damaged nucleotides at the break site can lead to deletions, and joining of nonmatching termini forms translocations. NHEJ is especially important before the cell has replicated its DNA, since there is no template available for repair by homologous recombination. There are "backup" NHEJ pathways in higher eukaryotes. Besides its role as a genome caretaker, NHEJ is required for joining hairpin-capped double-strand breaks induced during V(D)J recombination, the process that generates diversity in B-cell and T-cell receptors in the vertebrate immune system.

Recombinational repair requires the presence of an identical or nearly identical sequence to be used as a template for repair of the break. The enzymatic machinery responsible for this repair process is nearly identical to the machinery responsible for chromosomal crossover during meiosis. This pathway allows a damaged chromosome to be repaired using a sister chromatid (available in G2 after DNA replication) or a homologous chromosome as a template. DSBs caused by the replication machinery attempting to synthesize across a single-strand break or unrepaired lesion cause collapse of the replication fork and are typically repaired by recombination.

Topoisomerases introduce both single- and double-strand breaks in the course of changing the DNA's state of supercoiling, which is especially common in regions near an open replication fork. Such breaks are not considered DNA damage because they are a natural intermediate in the topoisomerase biochemical mechanism and are immediately repaired by the enzymes that created them.

A team of French researchers bombarded Deinococcus radiodurans to study the mechanism of double-strand break DNA repair in that organism. At least two copies of the genome, with random DNA breaks, can form DNA fragments through annealing. Partially overlapping fragments are then used for synthesis of homologous regions through a moving D-loop that can continue extension until they find complementary partner strands. In the final step there is crossover by means of RecA-dependent homologous recombination.

Translesion synthesis

Translesion synthesis is a DNA damage tolerance process that allows the DNA replication machinery to replicate past DNA lesions such as thymine dimers or AP sites. It involves the switching out of regular DNA polymerases for specialized translesion polymerases, often with larger active sites that can facilitate the insertion of bases opposite damaged nucleotides. The polymerase switching is thought to be mediated by, among other factors, the post-translational modification of the replication processivity factor PCNA. Translesion synthesis polymerases often have low fidelity (high propensity to insert wrong bases) relative to regular polymerases. However, many are extremely efficient at inserting correct bases opposite specific types of damage. For example, Pol η mediates error-free bypass of lesions induced by UV irradiation, whereas Pol ζ introduces mutations at these sites. From a cellular perspective, risking the introduction of point mutations during translesion synthesis may be preferable to resorting to more drastic mechanisms of DNA repair, which may cause gross chromosomal aberrations or cell death.

Taq Polymerase Enzyme

 

Taq polymerase is a thermostable DNA-dependent DNA polymerase that catalyzes the template-directed polymerization of dNTPs at high temperatures. Taq Polymerase was first isolated in 1976 from Thermus aquaticus strain YT-1.

The Properties of Taq Polymerase Enzyme

Taq Polymerase catalyzes the DNA-dependent polymerization of dNTPs, one unit of the enzyme is defined as the amount of enzyme that will incorporate 10 nmol of radioactively labeled dTTP into acid insoluble material at 80^oC in 30 minutes. The enzyme could be purified to a specific activity of 200000 U/mg.
The activity of the enzyme is dependent on bivalent cations, such as Mg²⁺. The optimum concentration of₂ is 2 mM. Monovalent cations also have an effect on the activity of Taq Polymerase. The monovalent cation is K⁺ in a form of KCl when it is using with the optimum concentration 50 mM, when KCl concentration more than 75 mM it can inhibit the activity of taq polymerase. Another monovalent cations, such as NaCl, NH₄Cl and NH₄acetate, cannot substitute KCl without a decrease in specific activity. MgCl
Maximum polymerization rates of Taq Polymerase are obtained with 0.7-0.8 mM dNTPs. While at dNTP concentrations of 4-6 mM, substrate inhibition is observed. Denaturing agents, detergents, and organic solvents in low concentration are tolerated by Taq Polymerase, while at higher concentrations, the inhibition of enzyme activity is observed.
The major distinguishing feature of Taq Polymerase is its extreme thermal stability. The enzyme can withstand temperatures in excess of 95^oC for prolonged periods, and in fact, its optimum for reaction is 75^oC. The rate of reaction is reduced to 50% at 60°C, and to 10% at 37^oC.

Taq Polymerase and PCR Technique

The major application of Taq polymerase at present is in the polymerase chain reaction (PCR). This technique is a simple method of amplifying minute quantities of DNA for a variety of subsequent procedures, including cloning, sequencing, hybridization, and genome mapping. The application of taq polymerase to the PCR was the basis for the success of the technique. It is caused by these following factors:

• Taq polymerase enzyme is stable up to 95^oC, so it is not necessary to replenish the enzyme after each PCR cycle.
• The activity of the enzyme is maximum at the temperature between 70 and 75^oC, which minimizes secondary structures of the template, resulting in high polymerization yield.
• The annealing can be chosen from 30-70^oC, allowing an optimal adaptation of cycle parameters to appropriate annealing temperatures of the primers; therefore, by products are hardly generated.

The enzyme from Thermus thermophilus (when used in a manganese-containing buffer) has an additional reverse transcriptase activity, which extends the use of PCR directly to cDNA synthesis. Another use of Taq polymerase 1s directly in DNA sequencing, where the high temperatures employed help reduce problems caused by secondary structure in the template and allow an increase in the stringency of primers used.

Protein Hydrolysis: Acid And Alkaline Method

There are three general methods to hydrolyze protein into its composition, amino acids. Those methods are acid hydrolysis, alkaline hydrolysis, and enzymatic hydrolysis. Strong acid is ordinarily the method of choice, and constant boiling hydrochloric acid, 6 M, is most frequently used. The reaction is usually carried out in evacuated sealed tubes or under N2 (Nitrogen) at110 Celcius degree for 18 to 96 hours. Under these conditions, peptide bonds are quantitatively hydrolyzed (although relatively long periods are required for the complete hydrolysis of bonds to valine, leucine, and isoleucine).


While the complete alkaline hydrolysis of proteins, is achieved with 2 to 4 M sodium hydroxide at 100 Celcius degree for 4 to 8 hours. This is of limited application for routine analysis, because cysteine, serine, threonine, and arginine are destroyed in the process, and partial destruction by deamination of other amino acids occurs. The complete enzymatic hydrolysis of proteins is difficult, because most enzymes attack only specific peptide bonds rapidly.

In this particular I only provide two methods of protein hydrolysis, acid hydrolysis and alkaline hydrolysis. Here are the methods:

Materials that you need:

3M p-toluenesulfonic acid.

0.2% tryptamine 3-[2-Aminoethyl] indole.

3M mercaptoethanesulfonic acid (Pierce).

1M sodium hydroxide.

Acid Hydrolysis of Protein:

1. 1 mL of 3M p-toluenesulphonic acid, containing 0.2% tryptamine (0.2% 3-[2-aminoethyl] indole) is added to the protein dried in a Pyrex glass tube (1.2 x 6 cm or similar, in which a constriction has been made by heating in an oxygen/gas flame).
2. The solution is sealed under vacuum and heated in an oven for 24 to 72 hours at 110 Celcius degree.
3. Altematively, you can use 3M mercaptoethanesulfonic acid as p-toluenesulphonic acid replacing, The sample is hydrolyzed for a similar time and temperature.
4. The tube is allowed to cool and cracked open with a heated glass rod held against a horizontal scratch made in the side of the tube.
5. The acid is taken to near neutrality by carefully adding 2 mililiters of 1M sodium hydroxide.
6. After this hydrolysis you can continue carrying out to quantitatively analyze certain amino acids, such as tryptophan.

Alkaline Hydrolysis Protein:

1. 0.5 mL of 3M sodium hydroxide is added to the protein dried in a Pyrex glass tube.
2. The solution is sealed under vacuum and heated in an oven for 4 to 8 hours at 100 Celcius degree.
3. After cooling and cracking open, the alkali is neutralized carefully with an equivalent amount of 1M HCl.

Epigenome

The "epigenome" is a parallel to the word "genome", and refers to the overall epigenetic state of a cell. The phrase "genetic code" has also been adapted—the "epigenetic code" has been used to describe the set of epigenetic features that create different phenotypes in different cells. Taken to its extreme, the "epigenetic code" could represent the total state of the cell, with the position of each molecule accounted for in an epigenomic map, a diagrammatic representation of the gene expression, DNA methylation and histone modification status of a particular genomic region. More typically, the term is used in reference to systematic efforts to measure specific, relevant forms of epigenetic information such as the histone code or DNA methylation patterns.

 

Quantitative Estimation of DNA Concentrations

DNA, RNA, and protein strongly absorb ultraviolet light in the 260 to 280 nm range. UV spectroscopy can be used as a quantitative technique to measure nucleic acid concentration and protein contamination. Nucleic acids strongly absorb at 260 nm and less strongly at 280 nm while proteins do the opposite. The general rules for determining the concentrations of nucleic acids at 260 nm are:

1. 1 Optical Density (OD) unit of double-stranded DNA is 50 micrograms/ml.
2. 1 OD unit of single-stranded DNA is 33 micrograms/ml.
3. 1 OD unit of single-stranded RNA is 40 micrograms/ml.

Proteins absorb strongly at 280 nm where 1 OD unit is 1 mg/ml. When using UV spectroscopy for estimating DNA concentrations, it is very important to remove all protein and RNA from the DNA solution. Good estimations can only be made on clean preparations.

An estimate of the purity of a DNA preparation can be made by measuring the absorbance at both 260 nm and 280 nm. Pure solutions of nucleic acid will absorb approximately twice as much at 260 nm as at 280 nm. Experimentally, the ratio of 260 nm/280 nm of a pure DNA solution is between 1.8 to 2.0. As protein contamination increases, the ratio decreases. Additionally, the presence of contaminating oranic solvents, such as phenol, can affect estimations of concentration and purity.

Materials you need are:

UV Spectrophotometer

Quartz or UV compatible cuvettes

TE buffer

DNA template

Method:

1. Fill the cuvette with water or TE buffer. Zero the spectrophotometer at 260 nm with this blank.
2. DNA from plasmid and genomic preparations is typically at a concentration exceeding 1 micrograms/microliter. Consequently, DNA is usually diluted before measuring its absorbance. An unfortunate result of this measurement is that the DNA is expended as a result of the dilution. Be sure these is adequate DNA to waste. Start by diluting the DNA sample 1 microliter : 999 microliters of TE buffer (the dilution can be done directly in the cuvette). Mix the dilution thoroughly.
3. Measure the optical density (OD). Multiply the resulting OD by 50 micrograms/ml. For a 1:1000 dilution, the mass of DNA is equal to micrograms/microliter.
4. Similarly, the same sample can be measured at 280 nm. A ratio of the OD-260nm/OD-280nm is an indicator of DNA purity. A ratio of 1.8 or higher indicates minimal protein contamination.

Isolation Of Genomic DNA Rrom Yeast

Isolating genomic DNA from yeast involves culturing the microbe, harvesting the cell, enzymatically removing the cell wall, lysing the protoplast, and finally separating the DNA from the other cell debris.
Materials that you need are:

Yeast culture-prepared previously

Spectrophotometer with cuvettes

50 mM EDTA, pH 8-ice cold

50 mM Tris, pH 9.5, 2% 2-mercaptoethanol

1.2 M sorbitol, 50 mM Tris, pH 7.5

Lyticase solution-500 U/ml in 50 mM Tris, pH 7.5

10% Sodium Dodecyl Sulfate (SDS)-used for checking protoplast formation

Lysis buffer-100 mM Tris, pH 7.5, 100 mM EDTA, 150 mMNaCl, 50 micrograms/ml RNase A

Lysis buffer with 2% SDS

95% Ethanol-stored at minus 20 degree Celcius

TE buffer-10 M Tris, pH 8, 1 mM EDTA

3 M potassium acetate, pH 5.5

Here are the step by step methods:

1. The yeast can be cultured for as long as 48 hours at 30 degree Celcius. The optical density of a 1:10 dilution of the culture in water can be as high as 1.0 at 520 nm.
2. Harvest 5 ml of cells by centrifugation (5 minutes at 5000 rpm). Resuspend the yeast in 1 ml of cold 50 mM EDTA, pH 8, and transfer to a 1.5 ml microfuge tube. Centrifuge for 1 minute, decant, and resuspend again in 50 mM EDTA.
3. Pellet the cells as before and suspend the cells in 1 ml of 50 mM Tris, pH 9.5, 2% 2-mercaptoethanol. Incubate for 10 min at room temperature. Centrifuge and decant.
4. Resuspend the cells in 800 micro liter of 1.2 M sorbitol, 50 mM Tris, pH 7.5. The sorbitol act as an osmotic support and prevents rupture of the cells as the wall is removed. As the yeast cell walls degrade, membranes can easily overextend and rupture.
5. Add 200 micro liter of Lyticase (500 U/ml in 50 mM Tris, pH 7.5). Place the cells on a rocker and incubate at 37 degree Celcius for one hour. Lyticase is a yeast cell wall degrading enzyme isolated from the bacteria Arthrobacter luteus.
6. Examine the suspension under a microscope to ensure protoplast formation. As the yeast wall is degraded, the cell membrane can ooze out of the sack. Viewed with phase contrast microscopy, yeast protoplasts are characteristically refractile (or bright) spheres, and yeast cell wall shells appear as gray ghosts (cell walls without membrane and cytosol). Combine 10 micro liter of 10% SDS with 10 micro liter of yeast protoplasts. Examine the cells under the microscope. The absence of refractile yeast indicates the protoplasts were lysed by the SDS.
7. Pellet the protoplasts by centrifuging at 10000 rpm for five minutes. Resuspend the cells in 1 ml of 100 mM Tris, pH 7.5, 100 mM EDTA, 150 mM NaCl (lysis buffer). Transfer the cells to a 5 ml polypropylene tube. Add 1 ml of lysis buffer with 2% SDS. Mix and incubate at 30 Celcius egree for 30 minutes. Check the cells under a microscope for lysis.
8. Centrifuge the lysate at 5000 rpm for 15 min to pellet cellular debris. Decant the upper phase containing the DNA.
9. Using a pipet, determine the volume of the DNA solution. Add 1/10th volume (e.g., 100 micro liter for every ml) of 3 M potassium acetate to the solution. In the presence of ions Na and K, DNA precipitates if mixed with either ethanol or isopropanol. Incubate the DNA at -20 Celcius degree for 30 min (or overnight if possible. Centrifuge the solution at 7000 rpm for 20 minutes. The DNA appears as white pellet. Decant and remove as much moisture as possible, but do not allow the pellet to dry. Once genomic DNA drys, it can be very difficult to resuspend.
10. Resuspend the DNA in 100 micro liter of TE buffer and freeze.

Gold Nanoparticles In Cancer Cell Detection

“Gold nanoparticles are very good at scattering and absorbing light,” said Mostafa El-Sayed, director of the Laser Dyanamics Laboratory and chemistry professor at Georgia Tech. “We wanted to see if we could harness that scattering property in a living cell to make cancer detection easier. So far, the results are extremely promising.”

Many cancer cells have a protein, known as Epidermal Growth Factor Receptor (EFGR), all over their surface, while healthy cells typically do not express the protein as strongly. By conjugating, or binding, the gold nanoparticles to an antibody for EFGR, suitably named anti-EFGR, researchers were able to get the nanoparticles to attach themselves to the cancer cells.

“If you add this conjugated nanoparticle solution to healthy cells and cancerous cells and you look at the image, you can tell with a simple microscope that the whole cancer cell is shining,” said El-Sayed. “The healthy cell doesn’t bind to the nanoparticles specifically, so you don’t see where the cells are. With this technique, if you see a well defined cell glowing, that’s cancer.”

In the study, researchers found that the gold nanoparticles have 600 percent greater affinity for cancer cells than for noncancerous cells. The particles that worked the best were 35 nanometers in size. Researchers tested their technique using cell cultures of two different types of oral cancer and one nonmalignant cell line. The shape of the strong absorption spectrum of the gold nanoparticles are also found to distinguish between cancer cells and noncancerous cells.

What makes this technique so promising, said El-Sayed, is that it doesn’t require expensive high-powered microscopes or lasers to view the results, as other techniques require. All it takes is a simple, inexpensive microscope and white light.

Another benefit is that the results are instantaneous. “If you take cells from a cancer stricken tissue and spray them with these gold nanoparticles that have this antibody you can see the results immediately. The scattering is so strong that you can detect a single particle,” said El-Sayed.

Finally, the technique isn’t toxic to human cells. A similar technique using artificial atoms known as Quantum Dots uses semiconductor crystals to mark cancer cells, but the semiconductor material is potentially toxic to the cells and humans.