Biochemistry | Turkish Chemistry
May 11

Gel electrophoresis

electrophoresis is a technique used for the separation of deoxyribonucleic acid (), ribonucleic acid (), or molecules using an electric current applied to a matrix.[1] It is usually performed for analytical purposes, but may be used as a preparative technique prior to use of other methods such as mass spectrometry, RFLP, PCR, cloning, sequencing, or Southern blotting for further characterization.

The term “” in this instance refers to the matrix used to contain, then separate the target molecules. In most cases, the is a crosslinked polymer whose composition and porosity is chosen based on the specific weight and composition of the target to be analyzed. When separating proteins or small nucleic acids (, , or oligonucleotides) the is usually composed of different concentrations of acrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids (greater than a few hundred bases), the preferred matrix is purified agarose. In both cases, the forms a solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is a neurotoxin and must be handled using appropriate safety precautions to avoid poisoning. Another matrix is agarose, long unbranched chains of uncharged carbohydrate without cross links giving a with large pores allowing separation of macromolecules and macromolecular complexes.

“Electrophoresis” refers to the electromotive force (EMF) that is used to move the molecules through the matrix. By placing the molecules in wells in the and applying an electric current, the molecules will move through the matrix at different rates, determined largely by their mass when the charge to mass ratio (Z) of all species is uniform, toward the anode if negatively charged or toward the cathode if positively charged

After the electrophoresis is complete, the molecules in the can be stained to make them visible. Ethidium bromide, silver, or coomassie blue dye may be used for this process. Other methods may also be used to visualize the separation of the mixture’s components on the . If the analyte molecules fluoresce under ultraviolet light, a photograph can be taken of the under ultraviolet lighting conditions. If the molecules to be separated contain radioactivity added for visibility, an autoradiogram can be recorded of the .

If several mixtures have initially been injected next to each other, they will run parallel in individual lanes. Depending on the number of different molecules, each lane shows separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or to indistinguishable smears representing multiple unresolved components.

Bands in different lanes that end up at the same distance from the top contain molecules that passed through the with the same speed, which usually means they are approximately the same size. There are molecular weight size markers available that contain a mixture of molecules of known sizes. If such a marker was run on one lane in the parallel to the unknown samples, the bands observed can be compared to those of the unknown in order to determine their size. The distance a band travels is approximately inversely proportional to the logarithm of the size of the molecule.

Applications

electrophoresis is used in forensics, molecular biology, genetics, microbiology and . The results can be analyzed quantitatively by visualizing the with UV light and a imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same . The measurement and analysis are mostly done with specialized software.

Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of electrophoresis, providing a wide range of field-specific applications.

Nucleic acids

In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally-occurring negative charge carried by their sugar-phosphate backbone.[3]

Double-stranded fragments naturally behave as long rods, so their migration through the is relative to their radius of gyration, or, for non-cyclic fragments, size. Single-stranded or tend to fold up into molecules with complex shapes and migrate through the in a complicated manner based on their tertiary structure. Therefore, agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again.[4]

electrophoresis of large or is usually done by agarose electrophoresis. See the “Chain termination method” page for an example of a polyacrylamide sequencing . Characterization through ligand interaction of nucleic acids or fragments may be performed by mobility shift affinity electrophoresis.

Proteins

SDS-PAGE autoradiography – The indicated proteins are present in different concentrations in the two samples.

Proteins, unlike nucleic acids, can have varying charges and complex shapes, therefore they may not migrate into the polyacryl amide at similar rates, or at all, when placing a negative to positive EMF on the sample. Proteins therefore, are usually denatured in the presence of a detergent such as sodium dodecyl sulfate/sodium dodecyl phosphate (SDS/SDP) that coats the proteins with a negative charge.[1] Generally, the amount of SDS bound is relative to the size of the (usually 1.4g SDS per gram of ), so that the resulting denatured proteins have an overall negative charge, and all the proteins have a similar charge to mass ratio. Since denatured proteins act like long rods instead of having a complex tertiary shape, the rate at which the resulting SDS coated proteins migrate in the is relative only to its size and not its charge or shape.[1]

Proteins are usually analyzed by sodium dodecyl sulfate polyacrylamide electrophoresis (SDS-PAGE), by native electrophoresis, by quantitative preparative native continuous polyacrylamide electrophoresis (QPNC-PAGE), or by 2-D electrophoresis.

Characterization through ligand interaction may be performed by electroblotting or by affinity electrophoresis in agarose or by capillary electrophoresis as for estimation of binding constants and determination of structural features like glycan content through lectin binding.

May 11

Restriction enzyme

A restriction enzyme (or restriction endonuclease) is an enzyme that cuts double-stranded or single stranded DNA at specific recognition nucleotide sequences known as restriction sites.[1][2][3] Such enzymes, found in bacteria and archaea, are thought to have evolved to provide a defense mechanism against invading viruses.[4][5] Inside a bacterial host, the restriction enzymes selectively cut up foreign DNA in a process called restriction; host DNA is methylated by a modification enzyme (a methylase) to protect it from the restriction enzyme’s activity. Collectively, these two processes form the restriction modification system.[6] To cut the DNA, a restriction enzyme makes two incisions, once through each sugar-phosphate backbone (i.e. each strand) of the DNA double helix.

After isolating the first restriction enzyme, HindII, in 1970[7], and the subsequent discovery and characterization of numerous restriction endonucleases,[8] the Nobel Prize in Medicine was awarded, in 1978, to Daniel Nathans, Werner Arber, and Hamilton Smith.[9] Their discovery led to the development of recombinant DNA technology that allowed, for example, the large scale production of human insulin for diabetics using E. coli bacteria.[10] Over 3000 restriction enzymes have been studied in detail, and more than 600 of these are available commercially[11] and are routinely used for DNA modification and manipulation in laboratories

Restriction enzymes recognize a specific sequence of nucleotides[2] and produce a double-stranded cut in the DNA. While recognition sequences vary widely, with lengths between 4 and 8 nucleotides, many of them are palindromic, which correspond to nitrogenous base sequences that read the same backwards and forwards.[15] In theory, there are two types of palindromic sequences that can be possible in DNA. The mirror-like palindrome is similar to those found in ordinary text, in which a sequence reads the same forward and backwards on the same DNA strand (i.e., single stranded) as in GTAATG. The inverted repeat palindrome is also a sequence that reads the same forward and backwards, but the forward and backward sequences are found in complementary DNA strands (i.e., double stranded) as in GTATAC (Notice that GTATAC is complementary to CATATG)[16]. The inverted repeat is more common and has major biological importance than the mirror-like.

 

EcoRI digestion produces “sticky” ends, whereas SmaI restriction enzyme cleavage produces “blunt” ends

Recognition sequences in DNA differ for each restriction enzyme, producing differences in the length, sequence and strand orientation (5′ end or the 3′ end) of a sticky-end “overhang” of an enzyme restriction.[17]

Different restriction enzymes that recognize the same sequence are known as neoschizomers. These often cleave in a different locales of the sequence; however, different enzymes which recognize and cleave in the same location are known as an isoschizomer.

Bacteria prevent their own DNA from being cut by modifying their nucleotides via DNA methylation

Restriction enzymes as tools

See the main article on restriction digests.

Isolated restriction enzymes are used to manipulate DNA for different scientific applications.

They are used to assist insertion of genes into plasmid vectors during gene cloning and protein expression experiments. For optimal use, plasmids that are commonly used for gene cloning are modified to include a short polylinker sequence (called the multiple cloning site, or MCS) rich in restriction enzyme recognition sequences. This allows flexibility when inserting gene fragments into the plasmid vector; restriction sites contained naturally within genes influence the choice of endonuclease for digesting the DNA since it is necessary to avoid restriction of wanted DNA while intentionally cutting the ends of the DNA. To clone a gene fragment into a vector, both plasmid DNA and gene insert are typically cut with the same restriction enzymes, and then glued together with the assistance of an enzyme known as a DNA ligase.[27][28]

Restriction enzymes can also be used to distinguish gene alleles by specifically recognizing single base changes in DNA known as single nucleotide polymorphisms (SNPs).[29][30] This is only possible if a SNP alters the restriction site present in the allele. In this method, the restriction enzyme can be used to genotype a DNA sample without the need for expensive gene sequencing. The sample is first digested with the restriction enzyme to generate DNA fragments, and then the different sized fragments separated by gel . In general, alleles with correct restriction sites will generate two visible bands of DNA on the gel, and those with altered restriction sites will not be cut and will generate only a single band. The number of bands reveals the sample subject’s genotype, an example of restriction mapping.

In a similar manner, restriction enzymes are used to digest genomic DNA for gene analysis by Southern blot. This technique allows researchers to identify how many copies (or paralogues) of a gene are present in the genome of one individual, or how many gene mutations (polymorphisms) have occurred within a population. The latter example is called restriction fragment length polymorphism

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Rutin Biyokimya Analitleri-5

Rutin Biyokimya -5
Total Bilirubin, Direkt Bilirubin, , Ãœre, , , ,

Prealbumin, Haptoglobulin, Transferrin, IgA, IgM, IgG, IgE, CA 15-3, CA 19-9,

 Total PSA, Serbest PSA, CEA

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