Friday, August 27, 2010

DNA–protein interactions

DNA–protein interactions are fundamental to the existence of life forms,providing the key to the genetic plan as well as mechanisms for its maintenanceand evolution. The study of these interactions is therefore fundamental to our understanding of growth, development, differentiation, evolution, and disease. The manipulation of DNA–protein interactions is also becoming increasingly important to the biotechnology industry, permitting among other things the reprogramming of gene expression. The success of the first edition of DNA– Protein Interactions; Principles and Protocols was the result of Dr. G. Geoff Kneale's efforts in bringing together a broad range of relevant techniques. In producing the second edition of this book, I have tried to further increase this diversity while presenting the reader with alternative approaches to obtaining
the same information. A major barrier to the study of interactions between biological macromolecules has always been detection and hence the need to obtain sufficient material. The development of molecular cloning and subsequently of protein overexpression systems has essentially breached this barrier. However, in the case of DNA–protein interactions, the problem of quantity and hence of detection is often offset by the high degree of selectivity and stability of DNA– protein interactions. DNA–protein binding reactions will often go to near completion at very low component concentrations even within crude protein extracts. Thus, although many techniques described in this volume were initially developed to study interactions between highly purified components, these same techniques are often just as applicable to the identification of novel DNA–protein interactions within systems as undefined as a whole cell extract. In general, these techniques use a DNA rather than a protein detection system because the former is more sensitive. Radiolabeled DNA fragments are easily produced by a range of techniques commonly available to molecular biologists. DNA–protein complexes may be studied at three distinct levels—at the level of the DNA, of the protein, and of the complex. At the level of the DNA, the DNA binding site may be delimited and exact base sequence requirements defined. The DNA conformation can be studied and the exact bases contacted by the protein identified. At the protein level, the protein species binding a given DNA sequence can be identified. The amino acids contacting DNA and the protein surface facing the DNA may be defined and the amino acids essential to the recognition process can be identified. Furthermore, the protein’s tertiary structure and its conformational changes on complex formation can be studied. Finally, global parameters of a DNA–protein complex such as stoichiometry, the kinetics of its formation and dissociation, its stability, and the energy of interaction can be measured. Filter binding, electrophoretic mobility shift assay (EMSA/gel shift), DNaseI footprinting, and Southwestern blotting have been the most commonly used techniques to identify potentially interesting DNA target sites and to define the proteins that bind them. For example, gel shift or footprinting of a cloned gene regulation sequence by proteins in a crude cell extract may define binding activities for a given DNA sequence that correlates with gene expression or silencing. These techniques can be used as an assay during subsequent isolation of the protein(s) responsible. Interference assays, SELEX, and more refined footprinting techniques, such as hydroxy radical footprinting and DNA bending assays, can then be used to study the DNA component of the DNA–protein complex, whereas the protein binding surface can be probed by amino acid side chain modification, DNA–protein crosslinking, and of course by the production of protein mutants. Genetic approaches have also opened the way to engineer proteins recognizing chosen DNA targets. DNA–protein crosslinking has in recent years become a very important approach to investigate the relative positions of proteins in multicomponent protein–DNA complexes such as the transcription initiation complex. Here, crosslinkable groups are incorporated at specific DNA sequences and these are used to map out the “positions” of different protein components along the DNA. Extension of this technique can also allow the mapping of the crosslink within the protein sequence. Similar data can be obtained by incorporating crosslinking groups at known sites within the protein and then identifying the nucleotides targeted.
Once the basic parameters of a DNA–protein interaction have been defined, it is inevitable that a deeper understanding of the driving forces behind the DNA–protein interaction and the biological consequences of its formation will require physical and physicochemical approaches. These can be either static or dynamic measurements, but most techniques have been developed to deal with steady-state situations. Equilibrium constants can be obtained by surface plasmon resonance, by spectroscopic assays that differentiate
complexed and uncomplexed components, and, for more stable products, by footprinting and gel shift. Spectroscopy can also give specific answers about the conformation of proteins and any conformational changes they undergo on interacting with DNA as well as providing a rapid quantitative measure of mcomplex formation. Microcalorimetry gives a global estimation of the forces stabilizing a given complex. Static pictures of protein–DNA interactions can be obtained by several techniques. At atomic resolution, X-ray crystallography,
and nuclear magnetic resonance (NMR) studies require large amounts of highly homogeneous material. Lower resolution images can be obtained by electron and, more recently, by atomic force microscopies. Large multiprotein complexes are generally beyond the scope of NMR or even of X-ray crystallography. These are therefore more often studied using the electron microscope, either in a direct imaging mode or via the analysis of data obtained from 2D pseudocrystalline arrays.
Dynamic measurements of complex formation or dissociation can be obtained by biochemical techniques when the DNA–protein complexes have half-lives of several minutes to several hours. For footprinting and crosslinking,
a general rule is that the complexes should be stable for a time well in excess of the proposed period of the enzymatic or chemical reaction. For gel shift, the complex half-life should at least approach that of the time of gel migration, although the cage effect may tend to stabilize the complex within the gel matrix,
extending the applicability of this technique. More rapid assembly kinetics, multistep assembly processes, and short-lived DNA–protein complexes require much more rapid techniques such as UV laser-induced crosslinking,
surface plasmon resonance, and spectroscopic assays. UV-laser induced DNA– protein crosslinking is a promising development because it potentially permits the kinetics of complex assembly to be followed both in vitro and in vivo.

SDS-PAGE

SDS-PAGE (Sodium Dodecyl Sulfate- Polyacrylamide Gel Electrophoresis) is a powerful technique which is used for the separation of proteins and nucleic acids. Electrophoresis is the migration of charged molecules in a media upon application of an electric field. The rate of migration depends on the charge on the molecule, its molecular mass, size and the strength of the electric field. Usually, this technique is routinely used for the analysis of proteins. The most commonly used matrix is agarose or polyacrylamide. These matrix forms a porous support and the size of the pores can be varied by changing the concentration of the matrix. Agarose is used mostly for separation of larger macromolecules, including nucleic acids, proteins and their complexes. On the other hand, polyacrylamide is used for the separation of proteins and small oligonucleotides. The charge on a protein is determined by the pH of the medium and the amino acid composition of the protein. Each protein has an isoelectric point which is the pH at which the protein has no net charge. Thus, at a pH below the isoelectric point, the protein will be net positive charge and migrate towards cathode, but at higher pH, it will be negatively charged and move towards anode. Thus, the movement of protein will not only depend on the mass, but also on the charge. Nucleic acids, however, remain negative at any pH due to the presence of the phosphate group of each nucleotide. Electrophoretic separation of nucleic acids is therefore strictly according to size. Sodium dodecyl sulphate (SDS) is an anionic detergent which denatures proteins by binding to the polypeptide backbone. This makes the protein molecule negatively charged. This negative charge is proportionately distributed throughout the molecule, yielding the same charge density per unit length. In order to remove the disulphide bridges in proteins before they adopt the random-coil configuration necessary for separation by size, the proteins are reduced either by 2-mercaptoethanol or dithiothreito. Thus, in denaturing SDS-PAGE separations, migration is determined not by intrinsic electrical charge of the polypeptide, but by molecular weight. To increase the resolution of protein separation, a discontinuous buffer system is often used. The stacking gel contains a low pH, range of 6.8. At this pH, the major ion species, glycine, from the buffer is less ionized and hence moves very slowly. This leads to a trapping effect of the protein molecules, thereby concentrating them in the form of a band. As the protein enters the smaller pore sized separating gel and a higher pH, glycine is ionized, the voltage gradient is dissipated and the protein is separated based on size.

Protein Precipitation Mehod

(1) Clarify the protein solution (in most cases the lysates) by centrifugation.
(2) Transfer the supernatant into an ice cold beaker with a magnetic bead.
(3) Note the exact amount of the supernatant.
(4) Keep the beaker chilled by placing it in an ice tray.
(5) Transfer the beaker with the ice tray onto a magnetic stirrer

(6) Weigh the amount of ammonium sulfate to be added. The amount depends on the volume of the solution and the percentage saturation of the salt needed. Refer to the precipitation chart. In case of protein purification, a step precipitation is carried out.
(7) Slowly add the ammonium sulphate with stirring. One needs to be careful as the addition of the salt should be very slow. Add a small amount at a time and then allow it to dissolve before further addition.
(8) Keep it on the stirrer for 1hr precipitation to occur in ice.
(9) Centrifuge at 10,000g for 15min at 4oC.
(10) The pellet contains the precipitated protein which could be dissolved in a suitable buffer for further analysis and purification.
(11) For a second round of precipitation of a different protein, the supernatant is again used and the above same steps are followed.

Protein Precipitation

Many cytosolic proteins are water soluble and their solubility is a function of the ionic strength and pH of the solution. The commonly used salt for this purpose is Ammonium Sulphate, due to its high solubility even at lower temperatures. Proteins in aqueous solutions are heavily hydrated, and with the addition of salt, the water molecules become more attracted to the salt than to the protein due to the higher charge. This competition for hydration is usually more favorable towards the salt, which leads to interaction between the proteins, resulting in aggregation and finally precipitation. The precipitate can then be collected by centrifugation and the protein pellet is re-dissolved in a low salt buffer. Since different proteins have distinct characteristics, it is often the case that they precipitate (or ‘salt out’) at a particular concentration of salt.

Protein expression

Protein expression is tightly regulated for normal functioning of a cell or organism. To understand protein structure and function in detail, they often need to be separated from other cellular components (lipids, nucleic acids, sugars, etc.) and isolated to homogeneity. After recovering a protein to near homogeneity, it should retain all its native biological characteristics of structure and activity. To achieve this objective, one needs to take into account the physical and chemical property of proteins (size, charge, solubility, hydrophobicity, precipitation, etc.). These common characteristics of the protein can be exploited to separate it from other components of the cell. With the introduction of recombinant DNA technology, protein purification technique has been enhanced and also simplified. Purification protocols vary, depending on the precise nature of the protein. General steps include (i) chromatography, (ii) precipitation and/or (iii) extraction.

Animal cell




The cell is the functional basic unit of life. It was discovered by Robert Hooke and is the functional unit of all known living organisms. It is the smallest unit of life that is classified as a living thing, and is often called the building block of life

Online Mendelian Inheritance in Man

OMIM is a comprehensive, authoritative, and timely compendium of human genes and genetic phenotypes. The full-text, referenced overviews in OMIM contain information on all known mendelian disorders and over 12,000 genes. OMIM focuses on the relationship between phenotype and genotype. It is updated daily, and the entries contain copious links to other genetics resources.
This database was initiated in the early 1960s by Dr. Victor A. McKusick as a catalog of mendelian traits and disorders, entitled Mendelian Inheritance in Man (MIM). Twelve book editions of MIM were published between 1966 and 1998. The online version, OMIM, was created in 1985 by a collaboration between the National Library of Medicine and the William H. Welch Medical Library at Johns Hopkins. It was made generally available on the internet starting in 1987. In 1995, OMIM was developed for the World Wide Web by NCBI, the National Center for Biotechnology Information.