RAJALAKSHMI ENGINEERING COLLEGE

Rajalakshmi Engineering College

Department of Biotechnology

Faculty Name : Mr.M.Sankar (Lecturer)

Staff code: BT33 Semester :VII SEC A&B

IMMUNOTECHNOLOGY BT2046

UNIT I ANTIGENS

An antigen is a substance/molecule that when introduced into the body triggers the production of an antibody by the immune system which will then kill or neutralize the antigen that is recognized as a foreign and potentially harmful invader. These invaders can be molecules such as pollen or cells such as bacteria. Originally the term came from antibody generator and was a molecule that binds specifically to an antibody, but the term now also refers to any molecule or molecular fragment that can be bound by a major histocompatibility complex (MHC) and presented to a T-cell receptor"Self" antigens are usually tolerated by the immune system; whereas "Non-self" antigens are identified as intruders and attacked by the immune system. Autoimmune disorders arise from the immune system reacting to its own antigens.

Antigen

Each antibody binds to a specific antigen; an interaction similar to a lock and key.

Similarly, an immunogen is a specific type of antigen. An immunogen is defined as a substance that is able to provoke an adaptive immune response if injected on its own.Said another way, an immunogen is able to induce an immune response, while an antigen is able to combine with the products of an immune response once they are made. The overlapping concepts of immunogenicity and antigenicity are thereby subtly different. According to a current text book:

Immunogenicity is the ability to induce a humoral and/or cell-mediated immune response

Antigenicity is the ability to combine specifically with the final products of the [immune response] (i.e. secreted antibodies and/or surface receptors on T-cells). Although all molecules that have the property of immunogenicity also have the property of antigenicity, the reverse is not true."

At the molecular level, an antigen is characterized by its ability to be "bound" at the antigen-binding site of an antibody. Note also that antibodies tend to discriminate between the specific molecular structures presented on the surface of the antigen (as illustrated in the Figure). Antigens are usually proteins or polysaccharides. This includes parts (coats, capsules, cell walls, flagella, fimbrae, and toxins) of bacteria, viruses, and other microorganisms. Lipids and nucleic acids are antigenic only when combined with proteins and polysaccharides. Non-microbial exogenous (non-self) antigens can include pollen, egg white, and proteins from transplanted tissues and organs or on the surface of transfused blood cells. Vaccines are examples of immunogenic antigens intentionally administered to induce acquired immunity in the recipient.

Cells present their immunogenic-antigens to the immune system via a histocompatibility molecule. Depending on the antigen presented and the type of the histocompatibility molecule, several types of immune cells can become activated.

Related concepts

Epitope - The distinct molecular surface features of an antigen capable of being bound by an antibody (a.k.a. antigenic determinant). Antigenic molecules, normally being "large" biological polymers, usually present several surface features that can act as points of interaction for specific antibodies. Any such distinct molecular feature constitutes an epitope. Most antigens therefore have the potential to be bound by several distinct antibodies, each of which is specific to a particular epitope. Using the "lock and key" metaphor, the antigen itself can be seen as a string of keys - any epitope being a "key" - each of which can match a different lock. Different antibody idiotypes, each having distinctly formed complementarity determining regions, correspond to the various "locks" that can match "the keys" (epitopes) presented on the antigen molecule.

Allergen - A substance capable of causing an allergic reaction. The (detrimental) reaction may result after exposure via ingestion, inhalation, injection, or contact with skin.

Superantigen - A class of antigens which cause non-specific activation of T-cells resulting in polyclonal T cell activation and massive cytokine release.

Tolerogen - A substance that invokes a specific immune non-responsiveness due to its molecular form. If its molecular form is changed, a tolerogen can become an immunogen.

Immunoglobulin binding protein - These proteins are capable of binding to antibodies at positions outside of the antigen-binding site. That is, whereas antigens are the "target" of antibodies, immunoglobulin binding proteins "attack" antibodies. Protein A, protein G and protein L are examples of proteins that strongly bind to various antibody isotypes.

Origin of the term antigen

In 1899 Ladislas Deutsch (Laszlo Detre) (18741939) named the hypothetical substances halfway between bacterial constituents and antibodies "substances immunogenes ou antigenes". He originally believed those substances to be precursors of antibodies, just like zymogen is a precursor of zymase. But by 1903 he understood that an antigen induces the production of immune bodies (antibodies) and wrote that the word antigen was a contraction of "Antisomatogen = Immunkrperbildner". The Oxford English Dictionary indicates that the logical construction should be "anti(body)-gen"[6].

Classification of antigens

Antigens can be classified in order of their class.

Exogenous antigens

Exogenous antigens are antigens that have entered the body from the outside, for example by inhalation, ingestion, or injection. The immune system's response to exogenous antigens is often subclinical. By endocytosis or phagocytosis, exogenous antigens are taken into the antigen-presenting cells (APCs) and processed into fragments. APCs then present the fragments to T helper cells (CD4+) by the use of class II histocompatibility molecules on their surface. Some T cells are specific for the peptide:MHC complex. They become activated and start to secrete cytokines. Cytokines are substances that can activate cytotoxic T lymphocytes (CTL), antibody-secreting B cells, macrophages, and other particles.

Some antigens start out as exogenous antigens, and later become endogenous (for example, intracellular viruses). Intracellular antigens can be released back into circulation upon the destruction of the infected cell, again.

Endogenous antigens

Endogenous antigens are antigens that have been generated within previously normal cells as a result of normal cell metabolism, or because of viral or intracellular bacterial infection. The fragments are then presented on the cell surface in the complex with MHC class I molecules. If activated cytotoxic CD8+ T cells recognize them, the T cells begin to secrete various toxins that cause the lysis or apoptosis of the infected cell. In order to keep the cytotoxic cells from killing cells just for presenting self-proteins, self-reactive T cells are deleted from the repertoire as a result of tolerance (also known as negative selection). Endogenous antigens include xenogenic (heterologous), autologous and idiotypic or allogenic (homologous) antigens.

Autoantigens

An autoantigen is usually a normal protein or complex of proteins (and sometimes DNA or RNA) that is recognized by the immune system of patients suffering from a specific autoimmune disease. These antigens should, under normal conditions, not be the target of the immune system, but, due to mainly genetic and environmental factors, the normal immunological tolerance for such an antigen has been lost in these patients.

Tumor antigens

Tumor antigens or neoantigens are[citation needed] those antigens that are presented by MHC I or MHC II molecules on the surface of tumor cells. These antigens can sometimes be presented by tumor cells and never by the normal ones. In this case, they are called tumor-specific antigens (TSAs) and, in general, result from a tumor-specific mutation. More common are antigens that are presented by tumor cells and normal cells, and they are called tumor-associated antigens (TAAs). Cytotoxic T lymphocytes that recognize these antigens may be able to destroy the tumor cells before they proliferate or metastasize.

Tumor antigens can also be on the surface of the tumor in the form of, for example, a mutated receptor, in which case they will be recognized by B cells.

Nativity

A native antigen is an antigen that is not yet processed by an APC to smaller parts. T cells cannot bind native antigens, but require that they be processed by APCs, whereas B cells can be activated by native ones.

Antigenic specificity

Antigen(ic) specificity is the ability of the host cells to recognize an antigen specifically as a unique molecular entity and distinguish it from another with exquisite precision. Antigen specificity is due primarily to the side-chain conformations of the antigen. It is a measurement, although the degree of specificity may not be easy to measure, and need not be linear or of the nature of a rate-limited step or equation.

Structure of antigen

Preparation Of Antigens For Raising Antibodies

Polyclonal antibodies (or antisera) are antibodies that are obtained from different B cell resources. They are a combination of immunoglobulin molecules secreted against a specific antigen, each identifying a different epitope.

Production

These antibodies are typically produced by immunization of a suitable mammal, such as a mouse, rabbit or goat. Larger mammals are often preferred as the amount of serum that can be collected is greater. An antigen is injected into the mammal. This induces the B-lymphocytes to produce IgG immunoglobulins specific for the antigen. This polyclonal IgG is purified from the mammals serum.By contrast, monoclonal antibodies are derived from a single cell line.

Many methodologies exist for polyclonal antibody production in laboratory animals. Institutional guidelines governing animal use and procedures relating to these methodologies are generally oriented around humane considerations and appropriate conduct for adjuvant (agents which modify the effect of other agents while having few if any direct effects when given by themselves) use. This includes adjuvant selection, routes and sites of administration, injection volumes per site and number of sites per animal. Institutional policies generally include allowable volumes of blood per collection and safety precautions including appropriate restraint and sedation or anesthesia of animals for injury prevention to animals or personnel.

The primary goal of antibody production in laboratory animals is to obtain high titer, high affinity antisera for use in experimentation or diagnostic tests. Adjuvants are used to improve or enhance an immune response to antigens. Most adjuvants provide for an injection site, antigen depot which allows for a slow release of antigen into draining lymph nodes.

Many adjuvants also contain or act directly as:

1. surfactants which promote concentration of protein antigens molecules over a large surface area, and

2. immunostimulatory molecules or properties. Adjuvants are generally used with soluble protein antigens to increase antibody titers and induce a prolonged response with accompanying memory.

Such antigens by themselves are generally poor immunogens. Most complex protein antigens induce multiple B-cell clones during the immune response, thus, the response is polyclonal. Immune responses to non-protein antigens are generally poorly or enhanced by adjuvants and there is no system memory.

Antibodies are currently also being produced from isolation of human B-lymphocytes to produce specific recombinant polyclonal antibodies. The biotechnology company, Symphogen, produces this type of antibody for therapeutic applications. They are the first research company to develop recombinant polyclonal antibody drugs to reach phase two trials. This production prevents viral and prion transmission.

Animal selection

Animals frequently used for polyclonal antibody production include chickens, goats, guinea pigs, hamsters, horses, mice, rats, and sheep. However, the rabbit is the most commonly used laboratory animal for this purpose. Animal selection should be based upon:

1. the amount of antibody needed,

2. the relationship between the donor of the antigen and the recipient antibody producer (generally the more distant the phylogenetic relationship, the greater the potential for high titer antibody response) and

3. the necessary characteristics [e.g., class, subclass (isotype), complement fixing nature] of the antibodies to be made. Immunization and phlebotomies are stress associated and, at least when using rabbits and rodents, specific pathogen free (SPF) animals are preferred. Use of such animals can dramatically reduce morbidity and mortality due to pathogenic organisms, especially Pasteurella multocida in rabbits.

Goats or horses are generally used when large quantities of antisera are required. Many investigators favor chickens because of their phylogenetic distance from mammals. Chickens transfer high quantities of IgY (IgG) into the egg yolk and harvesting antibodies from eggs eliminates the need for the invasive bleeding procedure. One weeks eggs can contain 10 times more antibodies than the volume of rabbit blood obtained from one weekly bleeding. However, there are some disadvantages when using certain chicken derived antibodies in immunoassays. Chicken IgY does not fix mammalian complement component C1 and it does not perform as a precipitating antibody using standard solutions.

Although mice are used most frequently for monoclonal antibody production, their small size usually prevents their use for sufficient quantities of polyclonal, serum antibodies. However, polyclonal antibodies in mice can be collected from ascites fluid using any one of a number of ascites producing methodologies.

When using rabbits, young adult animals (2.53.0kg or 5.5-6.5lbs) should be used for primary immunization because of the vigorous antibody response. Immune function peaks at puberty and primary responses to new antigens decline with age. Female rabbits are generally preferred because they are more docile and are reported to mount a more vigorous immune response than males. At least two animals per antigen should be used when using outbred animals. This principle reduces potential total failure resulting from non-responsiveness to antigens of individual animals.

Antigen preparation

The size, extent of aggregation and relative nativity of protein antigens can all dramatically affect the quality and quantity of antibody produced. Small polypeptides (80%) is obtained. However, this method may be problematic for antibodies that are easily damaged, as harsh conditions are generally used. A low pH can break the bonds to remove the antibody from the column. In addition to possibly affecting the product, low pH can cause Protein A/G itself to leak off the column and appear in the eluted sample. Gentle elution buffer systems that employ high salt concentrations are also available to avoid exposing sensitive antibodies to low pH. Cost is also an important consideration with this method because immobilized Protein A/G is a more expensive resin.

To achieve maximum purity in a single step, affinity purification can be performed, using the antigen to provide exquisite specificity for the antibody. In this method, the antigen used to generate the antibody is covalently attached to an agarose support. If the antigen is a peptide, it is commonly synthesized with a terminal cysteine, which allows selective attachment to a carrier protein, such as KLH during development and to the support for purification. The antibody-containing media is then incubated with the immobilized antigen, either in batch or as the antibody is passed through a column, where it selectively binds and can be retained while impurities are washed away. An elution with a low pH buffer or a more gentle, high salt elution buffer is then used to recover purified antibody from the support.

To further select for antibodies, the antibodies can be precipitated out using sodium sulfate or ammonium sulfate. Antibodies precipitate at low concentrations of the salt, while most other proteins precipitate at higher concentrations. The appropriate level of salt is added in order to achieve the best separation. Excess salt must then be removed by a desalting method such as dialysis.

The final purity can be analyzed using a chromatogram. Any impurities will produce peaks, and the volume under the peak indicates the amount of the impurity. Alternatively, gel electrophoresis and capillary electrophoresis can be carried out. Impurities will produce bands of varying intensity, depending on how much of the impurity is present.

Recombinant

The production of recombinant monoclonal antibodies involves technologies, referred to as repertoire cloning or phage display/yeast display. Recombinant antibody engineering involves the use of viruses or yeast to create antibodies, rather than mice. These techniques rely on rapid cloning of immunoglobulin gene segments to create libraries of antibodies with slightly different amino acid sequences from which antibodies with desired specificities can be selected. The phage antibody libraries are a variant of the phage antigen libraries first invented by George Pieczenik These techniques can be used to enhance the specificity with which antibodies recognize antigens, their stability in various environmental conditions, their therapeutic efficacy, and their detectability in diagnostic applications. Fermentation chambers have been used to produce these antibodies on a large scale.

Chimeric antibodies

Early on, a major problem for the therapeutic use of monoclonal antibodies in medicine was that initial methods used to produce them yielded mouse, not human antibodies. While structurally similardifferences between the two sufficient to invoke an immune response occurred when murine monoclonal antibodies were injected into humans and resulted in their rapid removal from the blood, systemic inflammatory effects, and the production of human anti-mouse antibodies (HAMA).

In an effort to overcome this obstacle, approaches using recombinant DNA have been explored since the late 1980s. In one approach, mouse DNA encoding the binding portion of a monoclonal antibody was merged with human antibody-producing DNA in living cells, and the expression of this chimeric DNA through cell culture yielded half-mouse, half-human monoclonal antibody. For this product, the descriptive terms "chimeric" and "humanised" monoclonal antibody have been used to reflect the amount of human DNA used in the recombinant process.

'Fully' human monoclonal antibodies

Ever since the discovery that monoclonal antibodies could be generated in-vitro, scientists have targeted the creation of 'fully' human antibodies to avoid some of the side effects of humanised and chimeric antibodies. Two successful approaches were identified - phage display-generated antibodies and mice genetically engineered to produce more human-like antibodies.

One of the most successful commercial organisations behind therapeutic monoclonal antibodies was Cambridge Antibody Technology (CAT). Scientists at CAT demonstrated that phage display could be used such that variable antibody domains could be expressed on filamentous phage antibodies. This was reported in a key Nature publication.

CAT developed their display technologies further into several, patented antibody discovery/functional genomics tools, which were named ProximolTM and ProAbTM. ProAb was announced in December 1997 and involved highthroughput screening of antibody libraries against diseased and non-diseased tissue, whilst Proximol used a free radical enzymatic reaction to label molecules in proximity to a given protein

Genetically engineered mice, so called transgenic mice, can be modified to produce human antibodies, and this has been exploited by a number of commercial organisations:

Medarex - who market their UltiMab platform

Abgenix - who marketed their Xenomouse technology. Abgenix were acquired in April 2006 by Amgen.

Regeneron's VelocImmune technology.

Monoclonal antibodies have been generated and approved to treat: cancer, cardiovascular disease, inflammatory diseases, macular degeneration, transplant rejection, multiple sclerosis, and viral infection (see monoclonal antibody therapy).

In August 2006 the Pharmaceutical Research and Manufacturers of America reported that U.S. companies had 160 different monoclonal antibodies in clinical trials or awaiting approval by the Food and Drug Administration.

Applications

Diagnostic tests

Once monoclonal antibodies for a given substance have been produced, they can be used to detect the presence of this substance. The Western blot test and immuno dot blot tests detect the protein on a membrane. They are also very useful in immunohistochemistry, which detect antigen in fixed tissue sections and immunofluorescence test, which detect the substance in a frozen tissue section or in live cells.

Therapeutic treatment

Monoclonal antibody therapy for Cancer treatment

One possible treatment for cancer involves monoclonal antibodies that bind only to cancer cell-specific antigens and induce an immunological response against the target cancer cell. Such mAb could also be modified for delivery of a toxin, radioisotope, cytokine or other active conjugate; it is also possible to design bispecific antibodies that can bind with their Fab regions both to target antigen and to a conjugate or effector cell. In fact, every intact antibody can bind to cell receptors or other proteins with its Fc region.

Monoclonal antibodies for cancer. ADEPT, antibody directed enzyme prodrug therapy; ADCC, antibody dependent cell-mediated cytotoxicity; CDC, complement dependent cytotoxicity; MAb, monoclonal antibody; scFv, single-chain Fv fragment.

The illustration below shows all these possibilities:

MAbs approved by the FDA include

Bevacizumab

Cetuximab

Panitumumab

Traztuzumab

Pertuzumab

Autoimmune diseases

Monoclonal antibodies used for autoimmune diseases include infliximab and adalimumab, which are effective in rheumatoid arthritis, Crohn's disease and ulcerative Colitis by their ability to bind to and inhibit TNF-.[23] Basiliximab and daclizumab inhibit IL-2 on activated T cells and thereby help preventing acute rejection of kidney transplants.[23] Omalizumab inhibits human immunoglobulin E (IgE) and is useful in moderate-to-severe allergic asthma.

Examples

Below are examples of clinically important monoclonal antibodies.

Main category

Type

Application

Mechanism/Target

Mode

Anti-inflammatory

infliximab

rheumatoid arthritis

Crohn's disease

Ulcerative Colitis

inhibits TNF-

chimeric

adalimumab

rheumatoid arthritis

Crohn's disease

Ulcerative Colitis

inhibits TNF-

human

etanercept

rheumatoid arthritis

Contains decoy TNF receptor

fusion protein

basiliximab

Acute rejection of kidney transplants

inhibits IL-2 on activated T cells

chimeric

daclizumab

Acute rejection of kidney transplants

inhibits IL-2 on activated T cells

humanized

omalizumab

moderate-to-severe allergic asthma

inhibits human immunoglobulin E (IgE)

humanized

Anti-cancer

gemtuzumab

relapsed acute myeloid leukaemia

targets myeloid cell surface antigen CD33 on leukemia cells

humanized

alemtuzumab

B cell leukemia

targets an antigen CD52 on T- and B-lymphocytes

humanized

rituximab

non-Hodgkin's lymphoma

targets phosphoprotein CD20 on B lymphocytes

chimeric

trastuzumab

breast cancer with HER2/neu overexpression

targets the HER2/neu (erbB2) receptor

humanized

nimotuzumab

Approved in squamous cell carcinomas, Glioma

Clinical trials for other indications underway

EGFR inhibitor

Humanized

cetuximab

Approved in squamous cell carcinomas, colorectal carcinoma

EGFR inhibitor

Chimeric

bevacizumab

Anti-angiogenic cancer therapy

inhibits VEGF

humanized

Other

palivizumab[

RSV infections in children

inhibits an RSV fusion (F) protein

humanized

abciximab

Prevent coagulation in coronary angioplasty

inhibits the receptor GpIIb/IIIa on platelets

chimeric

Rapid Characterization of Monoclonal Antibodies using the Piezoelectric Immunosensor

Monoclonal antibodies with specificity against the Francisella tularensis outer lipopolysaccharide (LPS) membrane were prepared and characterized using the piezoelectric

immunosensor with immobilized LPS antigen from F. tularensis. Signals obtained by the

immunosensor were compared with ELISA and similar sensitivity was noticed. Signal of

negative controls obtained using the biosensor was below 0.5% of the signal obtained for the

selected specific antibody clone 4H3B9D3. Testing of cross reactivity based on the sensors

with immobilized LPS from Escherichia coli and Bacillus subtilis confirmed selectivity of

this antibody. Furthermore, the 4H3B9D3 antibody was successfully isotypized as IgM using

the piezoelectric sensors with secondary antibodies. Kinetics parameters of antibody were

evaluated in the flow-through arrangement. The kinetic rate constants for the antibody

4H3B9D3 were ka = (2.31 0.20)105 l mol-1s-1 (association) and kd = (0.0010 0.00062) s-1(dissociation) indicating very good affinity to the LPS antigen.

Characterization of Monoclonal Antibody Products

Characterization tests

Provide detailed information on the molecule/product

Requirement for Reference Standard characterization

Required for comparability studies

Often technically challenging for routine use

Characterization tests for Mab products

Primary Structure

LC/MS Peptide Maps

N-terminal Sequencing

Verification of C-terminus

Disulfide Bond Determination

Glycan Map

Intact Mass Determination

Secondary and Tertiary Structure

FTIR

Far UV CD

Fluorescence

Near UV CD

Others

Non-reduced CE-SDS

CEX-HPLC, low pH

DSC

AUC

SE-MALS

Extinction Coefficient

Excipients

Process impurities

Functional characterization

Antigen binding

Additional cell-based assays

Epitope mapping

Fc RI, RIII binding

ADCC

CDC

FcRn binding

Production Polyclonal antibodies Seen in detail under the topic of antigen preparation Unit I

Characterisation of Polyclonal antibodies

Basis of polyclonality

Responses are polyclonal in nature as each clone somewhat specializes in producing antibodies against a given epitope, and because, each antigen contains multiple epitopes, each of which in turn can be recognized by more than one clone of B cells. But, to be able to react to innumerable antigens, as well as, multiple constituent epitopes, the immune system requires the ability to recognize a very great number of epitopes in all, i.e., there should be a great diversity of B cell clones.

Clonality of B cells

Memory and nave B cells normally exist in relatively small numbers. As the body needs to be able to respond to a large number of potential pathogens, it maintains a pool of B cells with a wide range of specificities. Consequently, while there is almost always at least one B (naive or memory) cell capable of responding to any given epitope (of all that the immune system can react against), there are very few exact duplicates. However, when a single B cell encounters an antigen to which it can bind, it can proliferate very rapidly. Such a group of cells with identical specificity towards the epitope is known as a clone, and is derived from a common "mother" cell. All the "daughter" B cells match the original "mother" cell in their epitope specificity, and they secrete antibodies with identical paratopes. So, in this context, a polyclonal response is one in which multiple clones of B cells react to the same antigen.

Single antigen contains multiple overlapping epitopes

Blind Monks Examining an Elephant: An allegory for the polyclonal response: Each clone or antibody recognizes different parts of a single, larger antigen

A single antigen can be thought of as a sequence of multiple overlapping epitopes. Many unique B cell clones may be able to bind to the individual epitopes. This imparts even greater multiplicity to the overall response. All of these B cells can become activated and produce large colonies of plasma cell clones, each of which can secrete up to 1000 antibody molecules against each epitope per second.

Multiple clones recognize single epitope

In addition to different B cells reacting to different epitopes on the same antigen, B cells belonging to different clones may also be able to react to the same epitope. An epitope that can be attacked by many different B cells is said to be highly immunogenic. In these cases, the binding affinities for respective epitope-paratope pairs vary, with some B cell clones producing antibodies that bind strongly to the epitope, and others producing antibodies that bind weakly.

Clonal selection

For more details on lymph nodes, germinal centers of lymph nodes and clonal selection of B cells, see Lymph node, Germinal center, Clonal selection.

The clones that bind to a particular epitope with greater strength are more likely to be selected for further proliferation in the germinal centers of the follicles in various lymphoid tissues like the lymph nodes. This is not unlike natural selection: clones are selected for their fitness to attack the epitopes (strength of binding) on the encountered pathogen. What makes the analogy even stronger is that the B lymphocytes have to compete with each other for signals that promote their survival in the germinal centers.

Diversity of B cell clones

Although there are many diverse pathogens, many of which are constantly mutating, it is a surprise that a majority of individuals remain free of infections. Thus, maintenance of health requires the body to recognize all pathogens (antigens they present or produce) likely to exist. This is achieved by maintaining a pool of immensely large (about 109) clones of B cells, each of which reacts against a specific epitope by recognizing and producing antibodies against it. However, at any given time very few clones actually remain receptive to their specific epitope. Thus, approximately 107 different epitopes can be recognized by all the B cell clones combined. Moreover, in a lifetime, an individual usually requires the generation of antibodies against very few antigens in comparison with the number that the body can recognize and respond against.

Significance of the phenomenonIncreased probability of recognizing any antigen

If an antigen can be recognized by more than one component of its structure, it is less likely to be "missed" by the immune system. Mutation of pathogenic organisms can result in modification of antigenand, hence, epitopestructure. If the immune system "remembers" what the other epitopes look like, the antigen, and the organism, will still be recognized and subjected to the body's immune response. Thus, the polyclonal response widens the range of pathogens that can be recognized.

Limitation of immune system against rapidly mutating viruses

The clone 1 that got stimulated by first antigen gets stimulated by second antigen, too, which best binds with naive cell of clone 2. However, antibodies produced by plasma cells of clone 1 inhibit the proliferation of clone 2.

Many viruses undergo frequent mutations that result in changes in amino acid composition of their important proteins. Epitopes located on the protein may also undergo alterations in the process. Such an altered epitope binds less strongly with the antibodies specific to the unaltered epitope that would have stimulated the immune system. This is unfortunate because somatic hypermutation does give rise to clones capable of producing soluble antibodies that would have bound the altered epitope avidly enough to neutralize it. But these clones would consist of naive cells which are not allowed to proliferate by the weakly binding antibodies produced by the priorly stimulated clone. This doctrine is known as the original antigenic sin. This phenomenon comes into play particularly in immune responses against influenza, dengue and HIV viruses. This limitation, however, is not imposed by the phenomenon of polyclonal response, but rather, against it by an immune response that is biased in favor of experienced memory cells against the "novice" naive cells.

Increased chances of autoimmune reactions

In autoimmunity the immune system wrongly recognizes certain native molecules in the body as foreign (self-antigen), and mounts an immune response against them. Since these native molecules, as normal parts of the body, will naturally always exist in the body, the attacks against them can get stronger over time (akin to secondary immune response). Moreover, many organisms exhibit molecular mimicry, which involves showing those antigens on their surface that are antigenically similar to the host proteins. This has two possible consequences: first, either the organism will be spared as a self antigen; or secondly, that the antibodies produced against it will also bind to the mimicked native proteins. The antibodies will attack the self-antigens and the tissues harboring them by activating various mechanisms like the complement activation and antibody-dependent cell-mediated cytotoxicity. Hence, wider the range of antibody-specificities, greater the chance that one or the other will react against self-antigens (native molecules of the body).[26]

HYPERLINK "http://en.wikipedia.org/wiki/Polyclonal_response" \l "cite_note-36#cite_note-36" [27]

Difficulty in producing monoclonal antibodies

Monoclonal antibodies are structurally identical immunoglobulin molecules with identical epitope-specificity (all of them bind with the same epitope with same affinity) as against their polyclonal counterparts which have varying affinities for the same epitope. They are usually not produced in a natural immune response, but only in diseased states like multiple myeloma, or through specialized laboratory techniques. Because of their specificity, monoclonal antibodies are used in certain applications to quantify or detect the presence of substances (which act as antigen for the monoclonal antibodies), and for targeting individual cells (e.g. cancer cells). Monoclonal antibodies find use in various diagnostic modalities (see: western blot and immunofluorescence) and therapiesparticularly of cancer and diseases with autoimmune component. But, since virtually all responses in nature are polyclonal, it makes production of immensely useful monoclonal antibodies less straightforward.

SDS-PAGE

Picture of an SDS-PAGE. The molecular marker is in the left lane

SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis, is a technique widely used in biochemistry, forensics, genetics and molecular biology to separate proteins according to their electrophoretic mobility (a function of length of polypeptide chain or molecular weight). SDS gel electrophoresis of samples have identical charge per unit mass due to binding of SDS results in fractionation by size.

ProcedureTissue preparation

Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonicator. Cells may also be broken open by one of the above mechanical methods. However, it should be noted that bacteria, virus or environmental samples can be the source of protein and thus Western blotting is not restricted to cellular studies only.

A combination of biochemical and mechanical techniques including various types of filtration and centrifugation can be used to separate different cell compartments and organelles.

The solution of proteins to be analyzed is mixed with SDS, an anionic detergent which denatures secondary and nondisulfidelinked tertiary structures, and applies a negative charge to each protein in proportion to its mass. Heating the samples to at least 60 degrees C shakes up the molecules, helping SDS to bind.

A tracking dye may be added to the protein solution (of a size smaller than protein) to allow the experimenter to track the progress of the protein solution through the gel during the electrophoretic run.

Preparing acrylamide gels

The gels generally consist of acrylamide, bisacrylamide, SDS, and a Tris-Cl buffer with adjusted pH. The solution is degassed under a vacuum to prevent air bubbles during polymerization. Ammonium persulfate and TEMED are added when the gel is ready to be polymerized. The separating or resolving gel is usually more basic and has a higher polyacrylamide content than the loading gel.

Gels are polymerized in a gel caster. First the separating gel is poured and allowed to polymerize. Next a thin layer of isopropanol is added. Next the loading gel is poured and a comb is placed to create the wells. After the loading gel is polymerized the comb can be removed and the gel is ready for electrophoresis.

Electrophoresis

First the anode and cathode buffers are prepared. The anode buffer usually contains Tris-Cl, distilled deionized water and is adjusted to a higher pH than the cathode buffer. The cathode buffer contains SDS, Tris, Tricine, and distilled deionized water.[7] [8]

The electrophoresis apparatus is set up with cathode buffer covering the gel in the negative electrode chamber, and anode buffer in the lower positive electrode chamber. Next, the denatured sample proteins are added to the wells one end of the gel with a syringe or pipette. Finally, the apparatus is hooked up to a power source under appropriate running conditions to separate the protein bands.

An electric field is applied across the gel, causing the negatively-charged proteins to migrate across the gel towards the positive (+) electrode (anode). Depending on their size, each protein will move differently through the gel matrix: short proteins will more easily fit through the pores in the gel, while larger ones will have more difficulty (they encounter more resistance). After a set amount of time (usually a few hours- though this depends on the voltage applied across the gel; higher voltages run faster but tend to produce somewhat poorer resolution), the proteins will have differentially migrated based on their size; smaller proteins will have traveled farther down the gel, while larger ones will have remained closer to the point of origin. Therefore, proteins may be separated roughly according to size (and therefore, molecular weight), certain glycoproteins behave anomalously on SDS gels.

Staining

Two SDS-PAGE-gels after a completed run

Following electrophoresis, the gel may be stained (most commonly with Coomassie Brilliant Blue R-250 or silver stain), allowing visualization of the separated proteins, or processed further (e.g. Western blot). After staining, different proteins will appear as distinct bands within the gel. It is common to run molecular weight size markers of known molecular weight in a separate lane in the gel, in order to calibrate the gel and determine the weight of unknown proteins by comparing the distance traveled relative to the marker. The gel is actually formed because the acrylamide solution contains a small amount, generally about 1 part in 35 of bisacrylamide, which can form cross-links between two polyacrylamide molecules. The ratio of acrylamide to bisacrylamide can be varied for special purposes. The acrylamide concentration of the gel can also be varied, generally in the range from 5% to 25%. Lower percentage gels are better for resolving very high molecular weight proteins, while much higher percentages are needed to resolve smaller proteins. Determining how much of the various solutions to mix together to make gels of particular acrylamide concentration can be done on line

Gel electrophoresis is usually the first choice as an assay of protein purity due to its reliability and ease. The presence of SDS and the denaturing step causes proteins to be separated solely based on size. False negatives and positives are possible. A comigrating contaminant can appear as the same band as the desired protein. This comigration could also cause a protein to run at a different position or to not be able to penetrate the gel. This is why it is important to stain the entire gel including the stacking section. Coomassie Brilliant Blue will also bind with less affinity to glycoproteins and fibrous proteins, which interferes with quantification.

Chemical ingredients and their roles

Polyacrylamide gel (PAG) had been known as a potential embedding medium for sectioning tissues as early as 1964. Two independent groups, Davis and Raymond, employed PAG in electrophoresis in 1959. It possesses several electrophoretically desirable features that make it a versatile medium. PAGE separates protein molecules according to both size and charge. It is a synthetic gel, thermo-stable, transparent, strong, relatively chemically inert, can be prepared with a wide range of average pore sizes. The pore size of a gel is determined by two factors, the total amount of acrylamide present (%T) (T = Total acrylamide-bisacrylamide monomer concentration) and the amount of cross-linker (%C) (C = Crosslinker concentration). Pore size decreases with increasing%T; with cross-linking, 5%C gives the smallest pore size. Any increase or decrease in%C increases the pore size, as pore size with respect to%C is a parabolic function with vertex as 5%C. This appears to be because of nonhomogeneous bundling of strands in the gel.

This gel material can also withstand high voltage gradients, feasible for various staining and destaining procedures, and can be digested to extract separated fractions or dried for autoradiography and permanent recording. DISC electrophoresis utilizes gels of different pore sizes. The name DISC was derived from the discontinuities in the electrophoretic matrix and coincidentally from the discoid shape of the separated zones of ions. There are two layers of gel, namely stacking or spacer gel, and resolving or separating gel.

Stacking gel

The stacking gel is a large pore PAG (4%T). This gel is prepared with Tris/HCl buffer pH 6.8 of about 2 pH units lower than that of electrophoresis buffer (Tris/Glycine). These conditions provide an environment for Kohlrausch reactions determining molar conductivity, as a result, SDS-coated proteins are concentrated to several fold and a thin starting zone of the order of 19m is achieved in a few minutes. This gel is cast over the resolving gel. The height of the stacking gel region is always maintained more than double the height and the volume of the sample to be applied. This is based on isotachophoresis.

Chemical ingredients

Tris (tris (hydroxy methyl) aminomethane) (C4H11NO3; mW: 121.14). It has been used as a buffer because it is an innocuous substance to most proteins. Its pKa is 8.3 at 20 C, making it a very satisfactory buffer in the pH range from roughly 7 to 9.

Glycine (Amino Acetic Acid) (C2H5NO2; mW: 75.07). Glycine has been used as the source of trailing ion or slow ion because its pKa is 9.69 and mobility of glycinate are such that the effective mobility can be set at a value below that of the slowest known proteins of net negative charge in the pH range. The minimum pH of this range is approximately 8.0.

Acrylamide (C3H5NO; mW: 71.08). It is a white crystalline powder. While dissolving in water, autopolymerization of acrylamide takes place. It is a slow spontaneous process by which acrylamide molecules join together by head on tail fashion. But in presence of free radicals generating system, acrylamide monomers are activated into a free-radical state. These activated monomers polymerise quickly and form long chain polymers. This kind of reaction is known as Vinyl addition polymerisation. A solution of these polymer chains becomes viscous but does not form a gel, because the chains simply slide over one another. Gel formation requires hooking various chains together. Acrylamide is a neurotoxin. It is also essential to store acrylamide in a cool dark and dry place to reduce autopolymerisation and hydrolysis.

Bisacrylamide (N,N'-Methylenebisacrylamide) (C7H10N2O2; mW: 154.17). Bisacrylamide is the most frequently used cross linking agent for poly acrylamide gels. Chemically it is thought of having two-acrylamide molecules coupled head to head at their non-reactive ends.

Sodium Dodecyl Sulfate (SDS) (C12H25NaO4S; mW: 288.38). SDS is the most common dissociating agent used to denature native proteins to individual polypeptides. When a protein mixture is heated to 100 C in presence of SDS, the detergent wraps around the polypeptide backbone. It binds to polypeptides in a constant weight ratio of 1.4 g/g of polypeptide. In this process, the intrinsic charges of polypeptides becomes negligible when compared to the negative charges contributed by SDS. Thus polypeptides after treatment becomes a rod like structure possessing a uniform charge density, that is same net negative charge per unit length. Mobilities of these proteins will be a linear function of the logarithms of their molecular weights.

Without SDS, different proteins with similar molecular weights would migrate differently due to differences in mass charge ratio, as each protein has an isoelectric point and molecular weight particular to its primary structure. This is known as Native PAGE. Adding SDS solves this problem, as it binds to and unfolds the protein, giving a near uniform negative charge along the length of the polypeptide.

Ammonium persulfate (APS) (N2H8S2O8; mW: 228.2). APS is an initiator for gel formation.

TEMED (N, N, N', N'-tetramethylethylenediamine) (C6H16N2; mW: 116.21). Chemical polymerisation of acrylamide gel is used for SDS-PAGE. It can be initiated by ammonium persulfate and the quaternary amine, N,N,N',N'-tetramethylethylenediamine (TEMED). The rate of polymerisation and the properties of the resulting gel depends on the concentration of APS and TEMED. Increasing the amount of APS and TEMED results in a decrease in the average polymer chain length, an increase in gel turbidity and a decrease in gel elasticity. Decreasing the amount of initiators shows the reverse effect. The lowest catalytic concentrations that will allow polymerisation in the optimal period of time should be used. APS and TEMED are used, approximately in equimolar concentrations in the range of 1 to 10 mM.

Chemicals for processing and visualization

The following chemicals are used for processing of the gel and the protein samples visualized in it:

Bromophenol blue (BPB) (3',3",5',5" tetrabromophenolsulfonphthalein) (C19H10Br4O5S; mW: 669.99). BPB is the universal marker dye. Proteins and nucleic acids are mostly colourless. When they are subjected to electrophoresis, it is important to stop the run before they run off the gel. BPB is the most commonly employed tracking dye, because it is viable in alkali and neutral pH, it is a small molecule, it is ionisable and it is negatively charged above pH 4.6 and hence moves towards the anode. Being a small molecule it moves ahead of most proteins and nucleic acids. As it reaches the anodic end of the electrophoresis medium electrophoresis is stopped. It can bind with proteins weakly and give blue colour.

Glycerol (C3H8O3; mW: 92.09). It is a preservative and a weighing agent. Addition of glycerol (20-30 or 50%) is often recommended for the storage of enzymes. Glycerol maintains the protein solution at very low temperature, without freezing. It also helps to weigh down the sample into the wells without being spread while loading.

Coomassie Brilliant Blue R-250 (CBB)(C45H44N3NaO7S2; mW: 825.97). CBB is the most popular protein stain. It is an anionic dye, which binds with proteins non-specifically. The structure of CBB is predominantly non-polar. So is usually used (0.025%) in methanolic solution (40%) and acetic acid (7%). Proteins in the gel are fixed by acetic acid and simultaneously stained. The excess dye incorporated in the gel can be removed by destaining with the same solution but without the dye. The proteins are detected as blue bands on a clear background. As SDS is also anionic, it may interfere with staining process. Therefore, large volume of staining solution is recommended, approximately ten times the volume of the gel.

n-Butanol (C4H10O; mW: 74.12). Water saturated butanol is used as an overlay solution on the resolving gel.

Dithiothreitol (DTT; C4H10O2S2; mW: 154.25). DTT is a reducing agent used to disrupt disulfide bonds to ensure the protein is fully denatured before loading on the gel; ensuring the protein runs uniformly. Traditionally the toxic and less potent 2-mercaptoethanol was used.

Reducing SDS-PAGE

Besides the addition of SDS, proteins may optionally be briefly heated to near boiling in the presence of a reducing agent, such as dithiothreitol (DTT) or traditionally 2-mercaptoethanol (beta-mercaptoethanol/BME), which further denatures the proteins by reducing disulfide linkages, thus overcoming some forms of tertiary protein folding, and breaking up quaternary protein structure (oligomeric subunits). This is known as reducing SDS-PAGE, and is most commonly used. Non-reducing SDS-PAGE (no boiling and no reducing agent) may be used when native structure is important in further analysis (e.g. enzyme activity, shown by the use of zymograms). For example, quantitative preparative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE) is a new method for separating native metalloproteins in complex biological matrices.

Silver staining

Silver stained SDS Polyacrylamide gels

In the 14th century the silver staining technique was developed for colouring the surface of glass. It has been used extensively for this purpose since the 16th century. The colour produced by the early silver stains ranged between light yellow and an orange-red. Camillo Golgi perfected the silver staining for the study of the nervous system. Golgi's method stains a limited number of cells at random in their entirety.[14] The exact chemical mechanism by which this happens is still largely unknown.[15] Silver staining was introduced by Kerenyi and Gallyas as a sensitive procedure to detect trace amounts of proteins in gels.[16] The technique has been extended to the study of other biological macromolecules that have been separated in a variety of supports.[17] Classical Coomassie Brilliant Blue staining can usually detect a 50ng protein band, Silver staining increases the sensitivity typically 50 times. Many variables can influence the colour intensity and every protein has its own staining characteristics; clean glassware, pure reagents and water of highest purity are the key points to successful staining.[18]

Buffer systems

Most protein separations are performed using a "discontinuous" buffer system that significantly enhances the sharpness of the bands within the gel. During electrophoresis in a discontinuous gel system, an ion gradient is formed in the early stage of electrophoresis that causes all of the proteins to focus into a single sharp band. This occurs in a region of the gel that has larger pores so that the gel matrix does not retard the migration during the focusing or "stacking" event. Negative ions from the buffer in the tank then "outrun" the SDS-covered protein "stack" and eliminate the ion gradient so that the proteins subsequently separate by the sieving action in the lower, "resolving" region of the gel.

Many people continue to use a tris-glycine or "Laemmli" buffering system that stacks at a pH of 6.8 and resolves at a pH of ~8.3-9.0. These pHs promote disulfide bond formation between cysteine residues in the proteins, especially when they are present at high concentrations because the pKa of cysteine ranges from 8-9 and because reducing agent present in the loading buffer doesn't co-migrate with the proteins. Recent advances in buffering technology alleviate this problem by resolving the proteins at a pH well below the pKa of cysteine (e.g., bis-tris, pH 6.5) and include reducing agents (e.g. sodium bisulfite) that move into the gel ahead of the proteins to maintain a reducing environment. An additional benefit of using buffers with lower pHs is that the acrylamide gel is more stable so the gels can be stored for long periods of time before use.[19]

HYPERLINK "http://en.wikipedia.org/wiki/SDS-PAGE" \l "cite_note-19" [20]

SDS gradient gel electrophoresis of proteins

Migration of proteins in SDS gels of varying acrylamide concentrations (%T). The migration of nine proteins ranging from 94 kDa to 14.4 kDa is shown. Stacking and unstacking occurs continuously in the gel, for every protein at a different gel concentration. The dotted line indicates the discontinuity at the Gly/Cl moving boundary. Proteins between the fast leading electrolyte and the slow trailing electrolyte are not diluted by diffusion.

As voltage is applied, the anions (and negatively charged sample molecules) migrate toward the positive electrode (anode) in the lower chamber, the leading ion is Cl ( high mobility and high concentration); glycinate is the trailing ion (low mobility and low concentration). SDS-protein particles do not migrate freely at the border between the Cl of the gel buffer and the Gly of the cathode buffer. Friedrich Kohlrausch found that Ohm's law also applies to dissolved electrolytes. Because of the voltage drop between the Cl- and Glycine-buffers, proteins are compressed (stacked) into micrometer thin layers. The boundary moves through a pore gradient and the protein stack gradually disperses due to an frictional resistance increase of the gel matrix. Stacking and unstacking occurs continuously in the gradient gel, for every protein at a different position. For a complete protein unstacking the polyacrylamide-gel concentration must exceed 16% T. The two-gel system of "Laemmli" is a simple gradient gel. The pH discontinuity of the buffers is of no significance for the separation quality, and a "stacking-gel" with a different pH is not needed.

Western blot

Western blot analysis of proteins separated by SDS-PAGE.

The Western blot (alternatively, protein immunoblot) is an extremely useful analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein.

There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against tens of thousands of different proteins. Commercial antibodies can be expensive, although the unbound antibody can be reused between experiments. This method is used in the fields of molecular biology, biochemistry, immunogenetics and other molecular biology disciplines.

Other related techniques include using antibodies to detect proteins in tissues and cells by immunostaining and enzyme-linked immunosorbent assay (ELISA).

The method originated from the laboratory of George Stark at Stanford. The name Western blot was given to the technique by W. Neal Burnette and is a play on the name Southern blot, a technique for DNA detection developed earlier by Edwin Southern. Detection of RNA is termed northern blotting and the detection of post-translational modification of protein is termed eastern blotting.

Steps in a Western blotTissue preparation

Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are first broken down mechanically using a blender (for larger sample volumes), using a homogenizer (smaller volumes), or by sonication. Cells may also be broken open by one of the above mechanical methods. However, it should be noted that bacteria, virus or environmental samples can be the source of protein and thus Western blotting is not restricted to cellular studies only.

Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to solubilize proteins. Protease and phosphatase inhibitors are often added to prevent the digestion of the sample by its own enzymes. Tissue preparation is often done at cold temperatures to avoid protein denaturing and degradation.

A combination of biochemical and mechanical techniques including various types of filtration and centrifugation can be used to separate different cell compartments and organelles.

Gel electrophoresis

The proteins of the sample are separated using gel electrophoresis. Separation of proteins may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these factors. The nature of the separation depends on the treatment of the sample and the nature of the gel. This is a very useful way to determine a protein.

By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers loaded with sodium dodecyl sulfate (SDS). SDS-PAGE (SDS polyacrylamide gel electrophoresis) maintains polypeptides in a denatured state once they have been treated with strong reducing agents to remove secondary and tertiary structure (e.g. disulfide bonds [S-S] to sulfhydryl groups [SH and SH]) and thus allows separation of proteins by their molecular weight. Sampled proteins become covered in the negatively charged SDS and move to the positively charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster through this mesh and the proteins are thus separated according to size (usually measured in kilodaltons, kDa). The concentration of acrylamide determines the resolution of the gel - the greater the acrylamide concentration the better the resolution of lower molecular weight proteins. The lower the acrylamide concentration the better the resolution of higher molecular weight proteins. Proteins travel only in one dimension along the gel for most blots.

Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder, a commercially available mixture of proteins having defined molecular weights, typically stained so as to form visible, coloured bands. When voltage is applied along the gel, proteins migrate into it at different speeds. These different rates of advancement (different electrophoretic mobilities) separate into bands within each lane.

It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single sample out in two dimensions. Proteins are separated according to isoelectric point (pH at which they have neutral net charge) in the first dimension, and according to their molecular weight in the second dimension.

Transfer

In order to make the proteins accessible to antibody detection, they are moved from within the gel onto a membrane made of nitrocellulose or polyvinylidene difluoride (PVDF). The membrane is placed on top of the gel, and a stack of filter papers placed on top of that. The entire stack is placed in a buffer solution which moves up the paper by capillary action, bringing the proteins with it. Another method for transferring the proteins is called electroblotting and uses an electric current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The protein move from within the gel onto the membrane while maintaining the organization they had within the gel. As a result of this "blotting" process, the proteins are exposed on a thin surface layer for detection (see below). Both varieties of membrane are chosen for their non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is based upon hydrophobic interactions, as well as charged interactions between the membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more fragile and do not stand up well to repeated probings.

The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie Brilliant Blue or Ponceau S dyes. Ponceau S is the more common of the two, due to Ponceau S's higher sensitivity and its water solubility makes it easier to subsequently destain and probe the membrane as described below.

Blocking

Since the membrane has been chosen for its ability to bind protein and as both antibodies and the target are proteins, steps must be taken to prevent interactions between the membrane and the antibody used for detection of the target protein. Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein - typically 3-5% Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive) in Tris-Buffered Saline (TBS), with a minute percentage of detergent such as Tween 20 or Triton X-100. The protein in the dilute solution attaches to the membrane in all places where the target proteins have not attached. Thus, when the antibody is added, there is no room on the membrane for it to attach other than on the binding sites of the specific target protein. This reduces "noise" in the final product of the Western blot, leading to clearer results, and eliminates false positives.

Detection

During the detection process the membrane is "probed" for the protein of interest with a modified antibody which is linked to a reporter enzyme, which when exposed to an appropriate substrate drives a colourimetric reaction and produces a colour. For a variety of reasons, this traditionally takes place in a two-step process, although there are now one-step detection methods available for certain applications.

Two steps

Primary antibody

Antibodies are generated when a host species or immune cell culture is exposed to the protein of interest (or a part thereof). Normally, this is part of the immune response, whereas here they are harvested and used as sensitive and specific detection tools that bind the protein directly.

After blocking, a dilute solution of primary antibody (generally between 0.5 and 5 micrograms/mL) is incubated with the membrane under gentle agitation. Typically, the solution is composed of buffered saline solution with a small percentage of detergent, and sometimes with powdered milk or BSA. The antibody solution and the membrane can be sealed and incubated together for anywhere from 30 minutes to overnight. It can also be incubated at different temperatures, with warmer temperatures being associated with more binding, both specific (to the target protein, the "signal") and non-specific ("noise").

Secondary antibody

After rinsing the membrane to remove unbound primary antibody, the membrane is exposed to another antibody, directed at a species-specific portion of the primary antibody. Antibodies come from animal sources (or animal sourced hybridoma cultures); an anti-mouse secondary will bind to almost any mouse-sourced primary antibody, which allows some cost savings by allowing an entire lab to share a single source of mass-produced antibody, and provides far more consistent results. This is known as a secondary antibody, and due to its targeting properties, tends to be referred to as "anti-mouse," "anti-goat," etc. The secondary antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or horseradish peroxidase. This means that several secondary antibodies will bind to one primary antibody and enhance the signal.

Most commonly, a horseradish peroxidase-linked secondary is used to cleave a chemiluminescent agent, and the reaction product produces luminescence in proportion to the amount of protein. A sensitive sheet of photographic film is placed against the membrane, and exposure to the light from the reaction creates an image of the antibodies bound to the blot. A cheaper but less sensitive approach utilizes a 4-chloronaphthol stain with 1% hydrogen peroxide; reaction of peroxide radicals with 4-chloronaphthol produces a dark brown stain that can be photographed without using specialized photographic film.

As with the ELISPOT and ELISA procedures, the enzyme can be provided with a substrate molecule that will be converted by the enzyme to a colored reaction product that will be visible on the membrane (see the figure below with blue bands).

Another method of secondary antibody detection utilizes a near-infrared (NIR) fluorophore-linked antibody. Light produced from the excitation of a fluorescent dye is static, making fluorescent detection a more precise and accurate measure of the difference in signal produced by labeled antibodies bound to proteins on a Western blot. Proteins can be accurately quantified because the signal generated by the different amounts of proteins on the membranes is measured in a static state, as compared to chemiluminescence, in which light is measured in a dynamic state.

A third alternative is to use a radioactive label rather than an enzyme coupled to the secondary antibody, such as labeling an antibody-binding protein like Staphylococcus Protein A or Streptavidin with a radioactive isotope of iodine. Since other methods are safer, quicker, and cheaper, this method is now rarely used; however, an advantage of this approach is the sensitivity of auto-radiography based imaging, which enables highly accurate protein quantification when combined with optical software (e.g. Optiquant).

One step

Historically, the probing process was performed in two steps because of the relative ease of producing primary and secondary antibodies in separate processes. This gives researchers and corporations huge advantages in terms of flexibility, and adds an amplification step to the detection process. Given the advent of high-throughput protein analysis and lower limits of detection, however, there has been interest in developing one-step probing systems that would allow the process to occur faster and with less consumables. This requires a probe antibody which both recognizes the protein of interest and contains a detectable label, probes which are often available for known protein tags. The primary probe is incubated with the membrane in a manner similar to that for the primary antibody in a two-step process, and then is ready for direct detection after a series of wash steps.

Western blot using radioactive detection system

Analysis

After the unbound probes are washed away, the Western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all Westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is repeated for a structural protein, such as actin or tubulin, that should not change between samples. The amount of target protein is indexed to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.

Colorimetric detection

The colorimetric detection method depends on incubation of the Western blot with a substrate that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary antibody. This converts the soluble dye into an insoluble form of a different color that precipitates next to the enzyme and thereby stains the membrane. Development of the blot is then stopped by washing away the soluble dye. Protein levels are evaluated through densitometry (how intense the stain is) or spectrophotometry.

Chemiluminescent detection

Chemiluminescent detection methods depend on incubation of the Western blot with a substrate that will luminesce when exposed to the reporter on the secondary antibody. The light is then detected by photographic film, and more recently by CCD cameras which capture a digital image of the Western blot. The image is analysed by densitometry, which evaluates the relative amount of protein staining and quantifies the results in terms of optical density. Newer software allows further data analysis such as molecular weight analysis if appropriate standards are used.

Radioactive detection

Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the Western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is declining, because it is very expensive, health and safety risks are high, and ECL (enhanced chemiluminescence) provides a useful alternative.

Fluorescent detection

The fluorescently labeled probe is excited by light and the emission of the excitation is then detected by a photosensor such as CCD camera equipped with appropriate emission filters which captures a digital image of the Western blot and allows further data analysis such as molecular weight analysis and a quantitative Western blot analysis. Fluorescence is considered to be among the most sensitive detection methods for blotting analysis.

Secondary probing

One major difference between nitrocellulose and PVDF membranes relates to the ability of each to support "stripping" antibodies off and reusing the membrane for subsequent antibody probes. While there are well-established protocols available for stripping nitrocellulose membranes, the sturdier PVDF allows for easier stripping, and for more reuse before background noise limits experiments. Another difference is that, unlike nitrocellulose, PVDF must be soaked in 95% ethanol, isopropanol or methanol before use. PVDF membranes also tend to be thicker and more resistant to damage during use.

2-D gel electrophoresis

2-dimensional SDS-PAGE uses the principles and techniques outlined above. 2-D SDS-PAGE, as the name suggests, involves the migration of polypeptides in 2 dimensions. For example, in the first dimension polypeptides are separated according to isoelectric point, while in the second dimension polypeptides are separated according to their molecular weight. The isoelectric point of a given protein is determined by the relative number of positively (e.g. lysine and arginine) and negatively (e.g. glutamate and aspartate) charged amino acids, with negatively charged amino acids contributing to a high isoelectric point and positively charged amino acids contributing to a low isoelectric point. Samples could also be separated first under nonreducing conditions using SDS-PAGE and under reducing conditions in the second dimension, which breaks apart disulfide bonds that hold subunits together. SDS-PAGE might also be coupled with urea-PAGE for a 2-dimensional gel.

In principle, this method allows for the separation of all cellular proteins on a single large gel. A major advantage of this method is that it often distinguishes between different isoforms of a particular protein - e.g. a protein that has been phosphorylated (by addition of a negatively charged group). Proteins that have been separated can be cut out of the gel and then analysed by mass spectrometry, which identifies the protein.

Please refer to reference articles for examples of the application of 2-D SDS PAGE.

Medical diagnostic applications

The confirmatory HIV test employs a Western blot to detect anti-HIV antibody in a human serum sample. Proteins from known HIV-infected cells are separated and blotted on a membrane as above. Then, the serum to be tested is applied in the primary antibody incubation step; free antibody is washed away, and a secondary anti-human antibody linked to an enzyme signal is added. The stained bands then indicate the proteins to which the patient's serum contains antibody.

A Western blot is also used as the definitive test for Bovine spongiform encephalopathy (BSE, commonly referred to as 'mad cow disease').

Some forms of Lyme disease testing employ Western blotting.

Western blot can also be used as a confirmatory test for Hepatitis B infection.

In veterinary medicine, Western blot is sometimes used to confirm FIV+ status in cats.

Immunoelectrophoresis

Immunoelectrophoresis is a general name for a number of biochemical methods for separation and characterization of proteins based on electrophoresis and reaction with antibodies. All variants of immunoelectrophoresis require immunoglobulins, also known as antibodies reacting with the proteins to be separated or characterized. The methods were developed and used extensively during the second half of the 20th century. In somewhat chronological order: Immunoelectrophoretic analysis (one-dimensional immunoelectrophoresis ad modum Grabar), crossed immunoelectrophoresis (two-dimensional quantitative immunoelectrophoresis ad modum Clarke and Freeman or ad modum Laurell), rocket-immunoelectrophoresis (one-dimensional quantitative immunoelectrophoresis ad modum Laurell), fused rocket immunoelectrophoresis ad modum Svendsen and Harboe, affinity immunoelectrophoresis ad modum Bg-Hansen.

Agarose as 1% gel slabs of about 1mm thickness buffered at high pH (around 8.6) is traditionally preferred for the electrophoresis as well as the reaction with antibodies. The agarose was chosen as the gel matrix because it has large pores allowing free passage and separation of proteins, but provides an anchor for the immunoprecipitates of protein and specific antibodies. The high pH was chosen because antibodies are practically immobile at high pH.

Immunoprecipitates may be seen in the wet agarose gel, but are stained with protein stains like Coomassie Brilliant Blue in the dried gel. In contrast to SDS-gel electrophoresis, the electrophoresis in agarose allows native conditions, preserving the native structure and activities of the proteins under investigation, therefore immunoelectrophoresis allows characterization of enzyme activities and ligand binding etc in addition to electrophoretic separation.

The immunoelectrophoretic analysis ad modum Grabar is the classical method of immunoelectrophoresis. Proteins are separated by electrophoresis, then antibodies are applied in a trough next to the separated proteins and immunoprecipitates are formed after a period of diffusion of the separated proteins and antibodies against each other. The introduction of the immunoelectrophoretic analysis gave a great boost to protein chemistry, some of the very first results were the resolution of proteins in biological fluids and biological extracts. Among the important observations made were the great number of different proteins in serum, the existence of several immunoglobulin classes and their electrophoretic heterogeneity.

Crossed immunoelectrophoresis is also called two-dimensional quantitative immunoelectrophoresis ad modum Clarke and Freeman or ad modum Laurell. In this method the proteins are first separated during the first dimension electrophoresis, then instead of the diffusion towards the antibodies, the proteins are electrophoresed into an antibody-containing gel in the second dimension. Immunoprecipitation will take place during the second dimension electrophorsis and the immunoprecipitates have a characteristic bell-shape, each precipitate representing one antigen, the position of the precipitate being dependent on the amount of protein as well as the amount of specific antibody in the gel, so relative quantification can be performed. The sensitivity and resolving power of crossed immunoelectrophoresis is than that of the classical immunoelectrophoretic analysis and there are multiple variations of the technique useful for various purposes. Crossed immunoelectrophoresis has been used for studies of proteins in biological fluids, particularly human serum, and biological extracts.

Rocket immunoelectrophoresis is one-dimensional quantitative immunoelectrophoresis. The methods has been used for quantitation of human serum proteins before automated methods became available.

Fused rocket immunoelectrophoresis is a modification of one-dimensional quantitative immunoelectrophorsis used for detailed measurement of proteins in fractions from protein separation experiments.

Affinity immunoelectrophoresis is based on changes in the electrophoretic pattern of proteins through biospecific interaction or complex formation with other macromolecules or ligands. Affinity immunoelectrophoresis has been used for estimation of binding constants, as for instance with lectins or for characterization of proteins with specific features like glycan content or ligand binding. Some variants of affinity immunoelectrophoresis are similar to affinity chromatography by use of immobilized ligands.

The open structure of the immunoprecipitate in the agarose gel will allow additional binding of radioactively labeled antibodies to reveal specific proteins. This variation has been used for identification of allergens through reaction with IgE.

Two factors determine that immunoelectrophoretic methods are not widely used. First they are rather work intensive and require some manual expertise. Second they require rather large amounts of polyclonal antibodies. Today gel electrophoresis followed by electroblotting is the preferred method for protein characterization because its ease of operation, its high sensitivity, and its low requirement for specific antibodies. In addition proteins are separated by gel electrophoresis on the basis of their apparent molecular weight, which is not accomplished by immunoelectrophoresis, but nevertheless immunoelectrophoretic methods are still useful when non-reducing conditions are needed.

Protein purification

Protein purification is a series of processes intended to isolate a single type of protein from a complex mixture. Protein purification is vital for the characterisation of the function, structure and interactions of the protein of interest. The starting material is usually a biological tissue or a microbial culture. The various steps in the purification process may free the protein from a matrix that confines it, separate the protein and non-protein parts of the mixture, and finally separate the desired protein from all other proteins. Separation of one protein from all others is typically the most laborious aspect of protein purification. Separation steps exploit differences in protein size, physico-chemical properties and binding affinity.

Purpose

Purification may be preparative or analytical. Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products such as enzymes (e.g. lactase), nutritional proteins (e.g. soy protein isolate), and certain biopharmaceuticals (e.g. insulin). Analytical purification produces a relatively small amount of a protein for a variety of research or analytical purposes, including identification, quantification, and studies of the protein's structure, post-translational modifications and function. Among the first purified proteins were urease and Concanavalin A.

Strategies

Choice of a starting material is key to the design of a purification process. In a plant or animal, a particular protein usually isn't distributed homogeneously throughout the body; different organs or tissues have higher or lower concentrations of the protein. Use of only the tissues or organs with the highest concentration decreases the volumes needed to produce a given amount of purified protein. If the protein is present in low abundance, or if it has a high value, scientists may use recombinant DNA technology to develop cells that will produce large quantities of the desired protein (this is known as an expression system). Recombinant expression allows the protein to be tagged, e.g. by a His-tag, to facilitate purification, which means that the purification can be done in fewer steps. In addition, recombinant expression usually starts with a higher fraction of the desired protein than is present in a natural source.

An analytical purification generally utilizes three properties to separate proteins. First, proteins may be purified according to their isoelectric points by running them through a pH graded gel or an ion exchange column. Second, proteins can be separated according to their size or molecular weight via size exclusion chromatography or by SDS-PAGE (sodium dodecyl sulfate-polyacrylamide gel electrophoresis) analysis. Proteins are often purified by using 2D-PAGE and are then analysed by peptide mass fingerprinting to establish the protein identity. This is very useful for scientific purposes and the detection limits for protein are nowadays very low and nanogram amounts of protein are sufficient for their analysis. Thirdly, proteins may be separated by polarity/hydrophobicity via high performance liquid chromatography or reversed-phase chromatography.

Evaluating purification yield

The most general method to monitor the purification process is by running a SDS-PAGE of the different steps. This method only gives a rough measure of the amounts of different proteins in the mixture, and it is not able to distinguish between proteins with similar apparent molecular weight.

If the protein has a distinguishing spectroscopic feature or an enzymatic activity, this property can be used to detect and quantify the specific protein, and thus to select the fractions of the separation, that contains the protein. If antibodies against the protein are available then western blotting and ELISA can specifically detect and quantify the amount of desired protein. Some proteins function as receptors and can be detected during purification steps by a ligand binding assay, often using a radioactive ligand.

In order to evaluate the process of multistep purification, the amount of the specific protein has to be compared to the amount of total protein. The latter can be determined by the Bradford total protein assay or by absorbance of light at 280 nm, however some reagents used during the purification process may interfere with the quantification. For example, imidazole (commonly used for purification of polyhistidine-tagged recombinant proteins) is an amino acid analogue and at low concentrations will interfere with the bicinchoninic acid (BCA) assay for total protein quantification. Impurities in low-grade imidazole will also absorb at 280nm, resulting in an inaccurate reading of protein concentration from UV absorbance.

Another method to be considered is Surface Plasmon Resonance (SPR). SPR can detect binding of label free molecules on the surface of a chip. If the desired protein is an antibody, binding can be translated to directly to the activity of the protein. One can express the active concentration of the protein as the percent of the total protein. SPR can be a powerful method for quickly determining protein activity and overall yield. It is a powerful technology that requires an instrument to perform.

Methods of protein purification

The methods used in protein purification can roughly be divided into analytical and preparative methods. The distinction is not exact, but the deciding factor is the amount of protein that can practically be purified with that method. Analytical methods aim to detect and identify a protein in a mixture, whereas preparative methods aim to produce large quantities of the protein for other purposes, such as structural biology or industrial use. In general, the preparative methods can be used in analytical applications, but not the other way around.

1. Extraction

Depending on the source, the protein has to be brought into solution by breaking the tissue or cells containing it. There are several methods to achieve this: Repeated freezing and thawing, sonication, homogenization by high pressure, filtration (either via cellulose-based depth filters or cross-flow filtration[1]), or permeabilization by organic solvents. The method of choice depends on how fragile the protein is and how sturdy the cells are. After this extraction process soluble proteins will be in the solvent, and can be separated from cell membranes, DNA etc. by centrifugation. The extraction process also extracts proteases, which will start digesting the proteins in the solution. If the protein is sensitive to proteolysis, it is usually desirable to proceed quickly, and keep the extract cooled, to slow down proteolysis.

2. Precipitation and differential solubilization

In bulk protein purification, a common first step to isolate proteins is precipitation with ammonium sulfate (NH4)2SO4. This is performed by adding increasing amounts of ammonium sulfate and collecting the different fractions of precipitate protein. One advantage of this method is that it can be performed inexpensively with very large volumes.

The first proteins to be purified are water-soluble proteins. Purification of integral membrane proteins requires disruption of the cell membrane in order to isolate any one particular protein from others that are in the same membrane compartment. Sometimes a particular membrane fraction can be isolated first, such as isolating mitochondria from cells before purifying a protein located in a mitochondrial membrane. A detergent such as sodium dodecyl sulfate (SDS) can be used to dissolve cell membranes and keep membrane proteins in solution during purification; however, because SDS causes denaturation, milder detergents such as Triton X-100 or CHAPS can be used to retain the protein's native conformation during complete purification.

3. Ultracentrifugation

Centrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid. When a vessel (typically a tube or bottle) containing a mixture of proteins or other particulate matter, such as bacterial cells, is rotated at high speeds, the angular momentum yields an outward force to each particle that is proportional to its mass. The tendency of a given particle to move through the liquid because of this force is offset by the resistance the liquid exerts on the particle. The net effect of "spinning" the sample in a centrifuge is that massive, small, and dense particles move outward faster than less massive particles or particles with more "drag" in the liquid. When suspensions of particles are "spun" in a centrifuge, a "pellet" may form at the bottom of the vessel that is enriched for the most massive particles with low drag in the liquid. Non-compacted particles still remaining mostly in the liquid are called the "supernatant" and can be removed from the vessel to separate the supernatant from the pellet. The rate of centrifugation is specified by the angular acceleration applied to the sample, typically measured in comparison to the g. If samples are centrifuged long enough, the particles in the vessel will reach equilibrium wherein the particles accumulate specifically at a point in the vessel where their buoyant density is balanced with centrifugal force. Such an "equilibrium" centrifugation can allow extensive purification of a given particle.

Sucrose gradient centrifugation a linear concentration gradient of sugar (typically sucrose, glycerol, or a silica based density gradient media, like Percoll) is generated in a tube such that the highest concentration is on the bottom and lowest on top. Percoll is a trademark owned by GE Healthcare companies. A protein sample is then layered on top of the gradient and spu