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

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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) (1874–1939) 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 = Immunkörperbildner". 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

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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 mammal’s 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 week’s 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 (2.5–3.0 kg 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.

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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-α.[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- |infliximab |rheumatoid arthritis |inhibits TNF-α |chimeric |

|inflammatory | |Crohn's disease | | |

| | |Ulcerative Colitis | | |

| |adalimumab |rheumatoid arthritis |inhibits TNF-α |human |

| | |Crohn's disease | | |

| | |Ulcerative Colitis | | |

| |etanercept |rheumatoid arthritis |Contains decoy TNF receptor |fusion protein |

| |basiliximab |Acute rejection of kidney |inhibits IL-2 on activated T cells |chimeric |

| | |transplants | | |

| |daclizumab |Acute rejection of kidney |inhibits IL-2 on activated T cells |humanized |

| | |transplants | | |

| |omalizumab |moderate-to-severe allergic asthma |inhibits human immunoglobulin E |humanized |

| | | |(IgE) | |

|Anti-cancer |gemtuzumab |relapsed acute myeloid leukaemia |targets myeloid cell surface antigen|humanized |

| | | |CD33 on leukemia cells | |

| |alemtuzumab |B cell leukemia |targets an antigen CD52 on T- and |humanized |

| | | |B-lymphocytes | |

| |rituximab |non-Hodgkin's lymphoma |targets phosphoprotein CD20 on B |chimeric |

| | | |lymphocytes | |

| |trastuzumab |breast cancer with HER2/neu |targets the HER2/neu (erbB2) |humanized |

| | |overexpression |receptor | |

| |nimotuzumab |Approved in squamous cell |EGFR inhibitor |Humanized |

| | |carcinomas, Glioma | | |

| | |Clinical trials for other | | |

| | |indications underway | | |

| |cetuximab |Approved in squamous cell |EGFR inhibitor |Chimeric |

| | |carcinomas, colorectal carcinoma | | |

| |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 |inhibits the receptor GpIIb/IIIa on |chimeric |

| | |angioplasty |platelets | |

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

– Fcγ 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 naïve 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 phenomenon

Increased 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 antigen—and, hence, epitope—structure. 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][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 therapies—particularly 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

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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.

Procedure

Tissue 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 non–disulfide–linked 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.

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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.

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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.

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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]

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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.

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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 19 μm 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

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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 50 ng 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][20]

SDS gradient gel electrophoresis of proteins

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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

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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 blot

Tissue 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.

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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.

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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.

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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.

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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.

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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.[pic]

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 Bøg-Hansen.

Agarose as 1 % gel slabs of about 1 mm 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 280 nm, 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 spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate towards the bottom of the tube faster than lighter material. During centrifugation in the absence of sucrose, as particles move farther and farther from the center of rotation, they experience more and more centrifugal force (the further they move, the faster they move). The problem with this is that the useful separation range of within the vessel is restricted to a small observable window. Spinning a sample twice as long doesn't mean the particle of interest will go twice as far, in fact, it will go significantly further. However, when the proteins are moving through a sucrose gradient, they encounter liquid of increasing density and viscosity. A properly designed sucrose gradient will counteract the increasing centrifugal force so the particles move in close proportion to the time they have been in the centrifugal field. Samples separated by these gradients are referred to as "rate zonal" centrifugations. After separating the protein/particles, the gradient is then fractionated and collected.

4. Chromatographic methods

Usually a protein purification protocol contains one or more chromatographic steps. The basic procedure in chromatography is to flow the solution containing the protein through a column packed with various materials. Different proteins interact differently with the column material, and can thus be separated by the time required to pass the column, or the conditions required to elute the protein from the column. Usually proteins are detected as they are coming off the column by their absorbance at 280 nm. Many different chromatographic methods exist:

a. Size exclusion chromatography

Gel permeation chromatography

Chromatography can be used to separate protein in solution or denaturing conditions by using porous gels. This technique is known as size exclusion chromatography. The principle is that smaller molecules have to traverse a larger volume in a porous matrix. Consequentially, proteins of a certain range in size will require a variable volume of eluent (solvent) before being collected at the other end of the column of gel.

In the context of protein purification, the eluant is usually pooled in different test tubes. All test tubes containing no measurable trace of the protein to purify are discarded. The remaining solution is thus made of the protein to purify and any other similarly-sized proteins.

b. Separation based on charge or hydrophobicity

Hydrophobic Interaction Chromatography

c. Ion exchange chromatography

Ion exchange chromatography separates compounds according to the nature and degree of their ionic charge. The column to be used is selected according to its type and strength of charge. Anion exchange resins have a positive charge and are used to retain and separate negatively charged compounds, while cation exchange resins have a negative charge and are used to separate positively charged molecules.

Before the separation begins a buffer is pumped through the column to equilibrate the opposing charged ions. Upon injection of the sample, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge. The most weakly charged compounds will elute first, followed by those with successively stronger charges. Becauses of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation.

Ion exchange chromatography is a very powerful tool for use in protein purification and is frequently used in both analytical and preparative separations.

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Nickel-affinity column. The resin is blue since it has bound nickel.

d. Affinity chromatography

Affinity Chromatography is a separation technique based upon molecular conformation, which frequently utilizes application specific resins. These resins have ligands attached to their surfaces which are specific for the compounds to be separated. Most frequently, these ligands function in a fashion similar to that of antibody-antigen interactions. This "lock and key" fit between the ligand and its target compound makes it highly specific, frequently generating a single peak, while all else in the sample is unretained.

Many membrane proteins are glycoproteins and can be purified by lectin affinity chromatography. Detergent-solubilized proteins can be allowed to bind to a chromatography resin that has been modified to have a covalently attached lectin. Proteins that do not bind to the lectin are washed away and then specifically bound glycoproteins can be eluted by adding a high concentration of a sugar that competes with the bound glycoproteins at the lectin binding site. Some lectins have high affinity binding to oligosaccharides of glycoproteins that is hard to compete with sugars, and bound glycoproteins need to be released by denaturing the lectin.

e. Metal binding

A common technique involves engineering a sequence of 6 to 8 histidines into the N- or C-terminal of the protein. The polyhistidine binds strongly to divalent metal ions such as nickel and cobalt. The protein can be passed through a column containing immobilized nickel ions, which binds the polyhistidine tag. All untagged proteins pass through the column. The protein can be eluted with imidazole, which competes with the polyhistidine tag for binding to the column, or by a decrease in pH (typically to 4.5), which decreases the affinity of the tag for the resin. While this procedure is generally used for the purification of recombinant proteins with an engineered affinity tag (such as a 6xHis tag or Clontech's HAT tag), it can also be used for natural proteins with an inherent affinity for divalent cations.

f. Immunoaffinity chromatography

Immunoaffinity chromatography uses the specific binding of an antibody to the target protein to selectively purify the protein. The procedure involves immobilizing an antibody to a column material, which then selectively binds the protein, while everything else flows through. The protein can be eluted by changing the pH or the salinity. Because this method does not involve engineering in a tag, it can be used for proteins from natural sources.

5. Purification of a tagged protein

Adding a tag to the protein such as RuBPS gives the protein a binding affinity it would not otherwise have. Usually the recombinant protein is the only protein in the mixture with this affinity, which aids in separation. The most common tag is the Histidine-tag (His-tag), that has affinity towards nickel or cobalt ions. Thus by immobilizing nickel or cobalt ions on a resin, an affinity support that specifically binds to histidine-tagged proteins can be created. Since the protein is the only component with a His-tag, all other proteins will pass through the column, and leave the His-tagged protein bound to the resin. The protein is released from the column in a process called elution, which in this case involves adding imidazole, to compete with the His-tags for nickel binding, as it has a ring structure similar to histidine. The protein of interest is now the major protein component in the eluted mixture, and can easily be separated from any minor unwanted contaminants by a second step of purification, such as size exclusion chromatography or RP-HPLC.

Another way to tag proteins is to engineer an antigen peptide tag onto the protein, and then purify the protein on a column or by incubating with a loose resin that is coated with an immobilized antibody. This particular procedure is known as immunoprecipitation. Immunoprecipitation is quite capable of generating an extremely specific interaction which usually results in binding only the desired protein. The purified tagged proteins can then easily be separated from the other proteins in solution and later eluted back into clean solution.

When the tags are not needed anymore, they can be cleaved off by a protease. This often involves engineering a protease cleavage site between the tag and the protein.

Concentration of the purified protein

A selectively permeable membrane can be mounted in a centrifuge tube. The buffer is forced through the membrane by centrifugation, leaving the protein in the upper chamber.

At the end of a protein purification, the protein often has to be concentrated. Different methods exist.

Lyophilization

If the solution doesn't contain any other soluble component than the protein in question the protein can be lyophilized (dried). This is commonly done after an HPLC run. This simply removes all volatile component leaving the proteins behind.

Ultrafiltration

Ultrafiltration concentrates a protein solution using selective permeable membranes. The function of the membrane is to let the water and small molecules pass through while retaining the protein. The solution is forced against the membrane by mechanical pump or gas pressure or centrifugation.

Analytical

Denaturing-Condition Electrophoresis

Gel electrophoresis is a common laboratory technique that can be used both as preparative and analytical method. The principle of electrophoresis relies on the movement of a charged ion in an electric field. In practice, the proteins are denatured in a solution containing a detergent (SDS). In these conditions, the proteins are unfolded and coated with negatively charged detergent molecules. The proteins in SDS-PAGE are separated on the sole basis of their size.

In analytical methods, the protein migrate as bands based on size. Each band can be detected using stains such as Coomassie blue dye or silver stain. Preparative methods to purify large amounts of protein, require the extraction of the protein from the electrophoretic gel. This extraction may involve excision of the gel containing a band, or eluting the band directly off the gel as it runs off the end of the gel.

In the context of a purification strategy, denaturing condition electrophoresis provides an improved resolution over size exclusion chromatography, but does not scale to large quantity of proteins in a sample as well as the late chromatography columns.

Non-Denaturing-Condition Electrophoresis

An important non-denaturing electrophoretic procedure for isolating bioactive metalloproteins in complex protein mixtures is termed 'quantitative native continuous polyacrylamide gel electrophoresis (QPNC-PAGE).

Method for concentration and purification of antigens and antibodies

In carrying out a purification and/or concentration step of antigens or antibodies, a substrate is immersed in a first aqueous medium containing a specifically reacting antigen to coat said substrate with a monomolecular layer of said specifically reacting antigen. The resulting coated substrate is then immersed in a second aqueous medium containing immunologically reactive antibody specific to the antigen in the first aqueous medium to complex said immunologically reactive antibody with said specifically reacting antigen. The resulting substrate is then immersed in a reagent capable of cleaving the immunological bond between said immunologically reactive antibody and said specifically reactive antigen and forming a solution of said immunologically reactive antibody in the immunological bond-cleaving solution and leaving said specifically reacting antigen coated on said substrate. The method can be reversed for preparing a purified concentration of an immunologically reactive antigen whereby a specifically reacting antibody is substituted for the antigen and the corresponding immunologically reactive antigen is substituted for the antibody.

Synthesis of Antigens Through Cloned Genes -  Hundreds of genes in eukaryotes have been cloned either from genomic DNA or from cDNA. These cloned genes included a number of genes for specific antigens, and in some cases have been used for the synthesis of antigens leading to the preparation of vaccines.

Following two examples can be used to illustrate the use of cloned genes for vaccine preparation

(i) Cloning of Hepatitis B virus (HBY) genome. The HBV genome has been cloned in the plasmid pBR322 and propagated in E. coli. From this clone, antigen could be produced in good quantity, which reacted with hepatitis B core antibody (HBAb). This has therefore been used to produce hepatitis B vaccine, which was later approved for mass vaccination in several countries.

(ii) Cloning of human malarial gene. Despite the great menace and threat to human health due to malarial parasite Plasmodium falciparum, no anti-malaria vaccine, could be developed so far. Recently with cloning of a gene coding for surface protein of the sporozoite of P. falciparum, there is a hope for developing a vaccine.

In human host, malarial parasite passes. through several antigenically distinct phases, namely;

(i) sporozoite: the form in which the parasite is injected with mosquito bite; sporozoites enter the liver and multiply and develop into

(ii) merozoites, which in turn invade and multiply in red blood cells; small fraction of these merozoites in red blood cells form

(iii) gametocytes, which may be picked up by a mosquito to start another cycle.

Recent advances in the research of artificial antigen have shown that artificial antigens can

be valuable approach for the treatment of some diseases as well as the detection of pesticide residues.By directly/indirectly coupling hapten to an appropriated carrier (macromolecule), artificial antigen can induce animals to produce hapten-specific antibody. Based on this principle, various vaccines have been developed. More impotently, new analytical method, immunological analysis has also been established. Comparing the conventional technologies, such as chromatographic methods, this promising method offers an alternative with high specificity, sensitivity, simplicity and suitability for the analysis of a large number of samples in a short period of time.

ELISA

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A 96-well microtiter plate being used for ELISA.

Enzyme-linked immunosorbent assay (ELISA), also known as an enzyme immunoassay (EIA), is a biochemical technique used mainly in immunology to detect the presence of an antibody or an antigen in a sample. The ELISA has been used as a diagnostic tool in medicine and plant pathology, as well as a quality control check in various industries. In simple terms, in ELISA, an unknown amount of antigen is affixed to a surface, and then a specific antibody is applied over the surface so that it can bind to the antigen. This antibody is linked to an enzyme, and in the final step a substance is added that the enzyme can convert to some detectable signal, most commonly a colour change in a chemical substrate.

Performing an ELISA involves at least one antibody with specificity for a particular antigen. The sample with an unknown amount of antigen is immobilized on a solid support (usually a polystyrene microtiter plate) either non-specifically (via adsorption to the surface) or specifically (via capture by another antibody specific to the same antigen, in a "sandwich" ELISA). After the antigen is immobilized the detection antibody is added, forming a complex with the antigen. The detection antibody can be covalently linked to an enzyme, or can itself be detected by a secondary antibody which is linked to an enzyme through bioconjugation. Between each step the plate is typically washed with a mild detergent solution to remove any proteins or antibodies that are not specifically bound. After the final wash step the plate is developed by adding an enzymatic substrate to produce a visible signal, which indicates the quantity of antigen in the sample.

Traditional ELISA typically involves chromogenic reporters and substrates which produce some kind of observable color change to indicate the presence of antigen or analyte. Newer ELISA-like techniques utilize fluorogenic, electrochemiluminescent, and real-time PCR reporters to create quantifiable signals. These new reporters can have various advantages including higher sensitivities and multiplexing. Technically, newer assays of this type are not strictly ELISAs as they are not "enzyme-linked" but are instead linked to some non-enzymatic reporter. However, given that the general principles in these assays are largely similar, they are often grouped in the same category as ELISAs.

Applications

ELISA results using S-OIV A neuraminidase antibody at 1 μg/ml to probe the immunogenic and the corresponding seasonal influenza A neuraminidase peptides at 50, 10, 2 and 0 ng/ml.

Because the ELISA can be performed to evaluate either the presence of antigen or the presence of antibody in a sample, it is a useful tool for determining serum antibody concentrations (such as with the HIV test[3] or West Nile Virus). It has also found applications in the food industry in detecting potential food allergens such as milk, peanuts, walnuts, almonds, and eggs. ELISA can also be used in toxicology as a rapid presumptive screen for certain classes of drugs.

The ELISA was the first screening test widely used for HIV because of its high sensitivity. In an ELISA, a person's serum is diluted 400-fold and applied to a plate to which HIV antigens are attached. If antibodies to HIV are present in the serum, they may bind to these HIV antigens. The plate is then washed to remove all other components of the serum. A specially prepared "secondary antibody" — an antibody that binds to other antibodies — is then applied to the plate, followed by another wash. This secondary antibody is chemically linked in advance to an enzyme. Thus, the plate will contain enzyme in proportion to the amount of secondary antibody bound to the plate. A substrate for the enzyme is applied, and catalysis by the enzyme leads to a change in color or fluorescence. ELISA results are reported as a number; the most controversial aspect of this test is determining the "cut-off" point between a positive and negative result.

A cut-off point may be determined by comparing it with a known standard. If an ELISA test is used for drug screening at workplace, a cut-off concentration, 50 ng/mL, for example, is established, and a sample will be prepared which contains the standard concentration of analyte. Unknowns that generate a signal that is stronger than the known sample are "positive". Those that generate weaker signal are "negative."

History

Before the development of the ELISA, the only option for conducting an immunoassay was radioimmunoassay, a technique using radioactively-labeled antigens or antibodies. In radioimmunoassay, the radioactivity provides the signal which indicates whether a specific antigen or antibody is present in the sample. Radioimmunoassay was first described in a paper by Rosalyn Sussman Yalow and Solomon Berson published in 1960.

Because radioactivity poses a potential health threat, a safer alternative was sought. A suitable alternative to radioimmunoassay would substitute a non-radioactive signal in place of the radioactive signal. When enzymes (such as peroxidase) react with appropriate substrates (such as ABTS or 3,3’,5,5’-Tetramethylbenzidine), this causes a change in color, which is used as a signal. However, the signal has to be associated with the presence of antibody or antigen, which is why the enzyme has to be linked to an appropriate antibody. This linking process was independently developed by Stratis Avrameas and G.B. Pierce. Since it is necessary to remove any unbound antibody or antigen by washing, the antibody or antigen has to be fixed to the surface of the container, i.e. the immunosorbent has to be prepared. A technique to accomplish this was published by Wide and Jerker Porath in 1966.

In 1971, Peter Perlmann and Eva Engvall at Stockholm University in Sweden, and Anton Schuurs and Bauke van Weemen in The Netherlands, independently published papers which synthesized this knowledge into methods to perform EIA/ELISA.

Types

"Indirect" ELISA

The steps of "indirect" ELISA follows the mechanism below:-

• A buffered solution of the antigen to be tested for is added to each well of a microtiter plate, where it is given time to adhere to the plastic through charge interactions.

• A solution of non-reacting protein, such as bovine serum albumin, or casein is added to block any plastic surface in the well that remains uncoated by the antigen.

• Next the primary antibody, generally in the form of serum is added, which contains a mixture of the serum donor's antibodies, of unknown concentration, some of which may bind specifically to the test antigen that is coating the well.

• Afterwards, a secondary antibody is added, which will bind any antibody produced by a member of the donor's species (for example, an antibody produced in a mouse that will bind any rabbit antibody). This secondary antibody often has an enzyme attached to it, which has a negligible effect on the binding properties of the antibody.

• A substrate for this enzyme is then added. Often, this substrate changes color upon reaction with the enzyme. The color change shows that secondary antibody has bound to primary antibody, which strongly implies that the donor has had an immune reaction to the test antigen. This can be helpful in a clinical setting, and in R&D.

• The higher the concentration of the primary antibody that was present in the serum, the stronger the color change. Often a spectrometer is used to give quantitative values for color strength.

The enzyme acts as an amplifier; even if only few enzyme-linked antibodies remain bound, the enzyme molecules will produce many signal molecules. Within common-sense limitations the enzyme can go on producing color indefinitely, but the more primary antibody is present in the donor serum, the more secondary antibody + enzyme will bind, and the faster color will develop. A major disadvantage of the indirect ELISA is that the method of antigen immobilization is non-specific; when serum is used as the source of test antigen, all proteins in the sample may stick to the microtiter plate well, so small concentrations of analyte in serum must compete with other serum proteins when binding to the well surface. The sandwich or direct ELISA provides a solution to this problem, by using a "capture" antibody specific for the test antigen to pull it out of the serum's molecular mixture.

ELISA may be run in a qualitative or quantitative format. Qualitative results provide a simple positive or negative result (yes or no) for a sample. The cutoff between positive and negative is determined by the analyst and may be statistical. Two or three times the standard deviation (error inherent in a test) is often used to distinguish positive from negative samples. In quantitative ELISA, the optical density (OD) of the sample is compared to a standard curve, which is typically a serial dilution of a known-concentration solution of the target molecule. For example if your test sample returns an OD of 1.0, the point on your standard curve that gave OD = 1.0 must be of the same analyte concentration as your sample.

Sandwich ELISA

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A sandwich ELISA. (1) Plate is coated with a capture antibody; (2) sample is added, and any antigen present binds to capture antibody; (3) detecting antibody is added, and binds to antigen; (4) enzyme-linked secondary antibody is added, and binds to detecting antibody; (5) substrate is added, and is converted by enzyme to detectable form.

A less-common variant of this technique, called "sandwich" ELISA, is used to detect sample antigen. The steps are as follows:

1. Prepare a surface to which a known quantity of capture antibody is bound.

2. Block any non specific binding sites on the surface.

3. Apply the antigen-containing sample to the plate.

4. Wash the plate, so that unbound antigen is removed.

5. Apply enzyme linked primary antibodies as detection antibodies which also bind specifically to the antigen.

6. Wash the plate, so that the unbound antibody-enzyme conjugates are removed.

7. Apply a chemical which is converted by the enzyme into a color or fluorescent or electrochemical signal.

8. Measure the absorbency or fluorescence or electrochemical signal (e.g., current) of the plate wells to determine the presence and quantity of antigen.

The image to the right includes the use of a secondary antibody conjugated to an enzyme, though technically this is not necessary if the primary antibody is conjugated to an enzyme. However, use of a secondary-antibody conjugate avoids the expensive process of creating enzyme-linked antibodies for every antigen one might want to detect. By using an enzyme-linked antibody that binds the Fc region of other antibodies, this same enzyme-linked antibody can be used in a variety of situations. Without the first layer of "capture" antibody, any proteins in the sample (including serum proteins) may competitively adsorb to the plate surface, lowering the quantity of antigen immobilized.Use of the purified specific antibody to attach the antigen to the plastic eliminates a need to purify the antigen from complicated mixtures before the measurement, simplifying the assay, and increasing the specificity and the sensitivity of the assay.

Application of sandwich ELISA to home pregnancy testing using hCG hormone antibody.

Competitive ELISA

A third use of ELISA is through competitive binding. The steps for this ELISA are somewhat different than the first two examples:

1. Unlabeled antibody is incubated in the presence of its antigen (Sample)

2. These bound antibody/antigen complexes are then added to an antigen coated well.

3. The plate is washed, so that unbound antibody is removed. (The more antigen in the sample, the less antibody will be able to bind to the antigen in the well, hence "competition.")

4. The secondary antibody, specific to the primary antibody is added. This second antibody is coupled to the enzyme.

5. A substrate is added, and remaining enzymes elicit a chromogenic or fluorescent signal.

For competitive ELISA, the higher the sample antigen concentration, the weaker the eventual signal. The major advantage of a competitive ELISA is the ability to use crude or impure samples and still selectively bind any antigen that may be present.

(Note that some competitive ELISA kits include enzyme-linked antigen rather than enzyme-linked antibody. The labeled antigen competes for primary antibody binding sites with your sample antigen (unlabeled). The more antigen in the sample, the less labeled antigen is retained in the well and the weaker the signal).Commonly the antigen is not first positioned in the well.

Multiple and Portable ELISA (M&P ELISA)

A new technique (EP 1 499 894 B1 in EPO Bulletin 25.02.209 N. 2009/09; USPTO 7510687 in USPTO Bulletin 31.03.2009; ZL 03810029.0 in SIPO PRC Bulletin 08.04.2009) uses a solid phase made up of an immunosorbent polystyrene rod with 8-12 protruding ogives. The entire device is immersed in a test tube containing the collected sample and the following steps (washing, incubation in conjugate and incubation in chromogenous) are carried out by dipping the ogives in microwells of standard microplates pre-filled with reagents.

The advantages of this technique are as follows:

1. The ogives can each be sensitized to a different reagent, allowing the simultaneous detection of different antibodies and / or different antigens for multi-target assays;

2. The sample volume can be increased to improve the test sensitivity in clinical (saliva, urine), food (bulk milk, pooled eggs) and environmental (water) samples;

3. One ogive is left unsensitized to measure the non-specific reactions of the sample;

4. The use of laboratory supplies for dispensing sample aliquots, washing solution and reagents in microwells is not required, facilitating the deveopment of ready-to-use lab-kits and on-site kits.

Radioimmunoassay

Radioimmunoassay (RIA) is an in vitro nuclear medicine very sensitive technique used to measure concentrations of antigens (for example, hormone levels in the blood) without the need to use a bioassay.

Although the RIA technique is extremely sensitive and extremely specific, it requires specialized equipment, but remains the least expensive method to perform such tests. It requires special precautions and licensing, since radioactive substances are used. Today it has been supplanted by the ELISA method, where the antigen-antibody reaction is measured using colorimetric signals instead of a radioactive signal. However, because of its robustness, consistent results and low price per test , RIA methods are again becoming popular.

The RAST test (radioallergosorbent test) is an example of radioimmunoassay. It is used to detect the causative allergen for an allergy.

Method

To perform a radioimmunoassay, a known quantity of an antigen is made radioactive, frequently by labeling it with gamma-radioactive isotopes of iodine attached to tyrosine. This radiolabeled antigen is then mixed with a known amount of antibody for that antigen, and as a result, the two chemically bind to one another. Then, a sample of serum from a patient containing an unknown quantity of that same antigen is added. This causes the unlabeled (or "cold") antigen from the serum to compete with the radiolabeled antigen ("hot") for antibody binding sites. As the concentration of "cold" antigen is increased, more of it binds to the antibody, displacing the radiolabeled variant, and reducing the ratio of antibody-bound radiolabeled antigen to free radiolabeled antigen. The bound antigens are then separated from the unbound ones, and the radioactivity of the free antigen remaining in the supernatant is measured using a gamma counter. Using known standards, a binding curve can then be generated which allows the amount of antigen in the patient's serum to be derived.

History

It was developed by Rosalyn Yalow and Solomon Aaron Berson in the 1950s. In 1977, Rosalyn Sussman Yalow received the Nobel Prize in Medicine for the development of the RIA for insulin: the precise measurement of minute amounts of such a hormone was considered a breakthrough in endocrinology.

With this technique, separating bound from unbound antigen is crucial. Initially, the method of separation employed was the use of a second "anti-antibody" directed against the first for precipitation and centrifugation. The use of charcoal suspension for precipitation was extended but replaced later by Drs. Werner and Acebedo at Columbia University for RIA of T3 and T4. An ultramicro RIA for human TSH was published in BBRC (1975) by Drs. Acebedo, Hayek et al.[3]

Applications of Radioimmunoassays

Endocrinology

14 Insulin, HCG, Vasopressin

15 Detects Endocrine Disorders

16 Physiology of Endocrine Function

Pharmacology

18 Morphine

19 Detect Drug Abuse or Drug Poisoning

20 Study Drug Kinetics

Non isotopic methods of detection of antigens

Sensitive non-isotopic DNA hybridisation assay or immediate-early antigen detection for rapid identification of human cytomegalovirus in urine.

A sensitive non-radioactive DNA hybridisation assay employing digoxigenin-labelled probes was compared with immediate-early antigen detection and conventional virus isolation for the identification of human cytomegalovirus (HCMV) in 249 urine samples. Of 44 specimens yielding HCMV by virus isolation, more were positive by DNA hybridisation (32; 73%) than by immediate-early antigen detection (25; 52%) (P = 0.05). The specificity of the hybridisation assay in 45 apparently falsely positive specimens was supported by detection of HCMV DNA in 40 of these specimens using the polymerase chain reaction. Many urine specimens may thus contain large amounts of non-viable virus or free viral DNA. Evaluation of various protocols for the extraction and denaturation of virus DNA prior to hybridisation showed that proteinase K digestion with phenol/chloroform extraction was the most sensitive and reliable procedure. We conclude that the non-radioactive DNA hybridisation assay described is a potentially valuable routine diagnostic test.

Cytomegalovirus detection by nonisotopic in situ DNA hybridization and viral antigen immunostaining using a two-color technique.

Rapid methods of specific viral diagnosis in formalin fixed, paraffin embedded tissues include identification of viral incusions in routinely stained histologic sections, immunologic staining of viral antigens, and in situ nucleic acid hybridization. To correlate in situ hybridization with immunologic detection methods, sequential two-color staining was used on tissues from 12 patients, each containing characteristic cytomegalovirus (CMV) inclusions, using a biotinylated CMV DNA probe in an avidin-alkaline phosphatase-linked reaction followed by avidin-biotin complex immunoperoxidase staining of CMV antigen. CMV genetic material was seen in all 17 tissues. CMV antigen was detected in 11 of 17 tissues (65%). The DNA hybridization technique provided more intense staining, detected greater numbers of inclusions, and had less background staining than the immunoperoxidase technique. The alkaline phosphatase reaction product was stable through subsequent immunostaining steps, and immunologic reactivity of CMV antigen was not significantly reduced by prior hybridization steps. CMV DNA probe was localized predominantly within cell nuclei, while CMV antigen immunostaining was predominantly cytoplasmic. It was concluded that sequential in situ hybridization and immunocytochemistry can be performed on standard histologic sections. Furthermore, it is likely that the majority of CMV nucleic acid detected by this tissue hybridization technique is unencapsidated, intranuclear viral DNA and not DNA contained within complete CMV nucleocapsids.

Chemiluminescence

Chemiluminescence (sometimes "chemoluminescence") is the emission of light with limited emission of heat (luminescence), as the result of a chemical reaction. Given reactants A and B, with an excited intermediate â—Š,

[A] + [B] → [◊] → [Products] + light

For example, if [A] is luminol and [B] is hydrogen peroxide in the presence of a suitable catalyst we have:

luminol + H2O2 → 3-APA[◊] → 3-APA + light

where:

• where 3-APA is 3-aminophthalate

• 3-APA[◊] is the excited state fluorescing as it decays to a lower energy level.

The decay of the excited state[â—Š] to a lower energy level causes the emission of light. In theory, one photon of light should be given off for each molecule of reactant, or Avogadro's number of photons per mole. In actual practice, non-enzymatic reactions seldom exceed 1% QC, quantum efficiency.

In a chemical reaction, reactants collide to form a transition state, the enthalpic maximum in a reaction coordinate diagram, which proceeds to the product. Normally, reactants form products of lesser chemical energy. The difference in energy between reactants and products, represented as ΔHrxn, is turned into heat, physically realized as excitations in the vibrational state of the normal modes of the product. Since vibrational energy is generally much greater than the thermal agitation, it is rapidly dispersed into the solvent through solvent molecules' rotation and translation. This is how exothermic reactions make their solutions hotter. In a chemiluminescent reaction, the direct product of a reaction is delivered in an excited electronic state, which then decays into an electronic ground state through either fluorescence or phosphorescence, depending on the spin state of the electronic excited state formed. This is possible because chemical bond formation can occur on a timescale faster than electronic transitions, and therefore can result in discrete products in excited electronic states.

Chemiluminescence differs from fluorescence in that the electronic excited state is derived from the product of a chemical reaction rather than the more typical way of creating electronic excited states, namely absorption. It is the antithesis of a photochemical reaction, in which light is used to drive an endothermic chemical reaction. Here, light is generated from a chemically exothermic reaction.

A standard example of chemiluminescence in the laboratory setting is found in the luminol test, where evidence of blood is taken when the sample glows upon contact with iron. When chemiluminescence takes place in living organisms, the phenomenon is called bioluminescence. A lightstick emits light by chemiluminescence.

Liquid-phase reactions

• Luminol in an alkaline solution with hydrogen peroxide in the presence of iron or copper, or an auxiliary oxidant, produces chemiluminescence. The luminol reaction is

luminol + H2O2 → 3-APA[◊] → 3-APA + light

[pic]

The quantum efficiency, QC is 1%. For the laboratory experiment see references.

• Cyalume, as used in a lightstick, emits light by chemiluminescence of a fluorescent dye (also called fluorescor) activated by cyalume reacting with hydrogen peroxide in the presence of a catalyst,such as sodium salicylate. It is the most efficient chemiluminescent reaction known. up to 15% quantum efficiency.

cyalume + H2O2 + dye → phenol + 2CO2 + dye[◊]

When the activated fluorescent dye decays to a lower energy level, light is given off. The color depends upon the dye.

|Color |Sensitiser |

|Blue |9,10-Diphenylanthracene |

|Green |9,10-Bis(phenylethynyl)anthracene |

|Yellow-green |Tetracene |

|Yellow |1-Chloro-9,10-bis(phenylethynyl)anthracene |

|Orange |5,12-Bis(phenylethynyl)naphthacene, Rubrene, Rhodamine 6G |

|Red |Rhodamine B |

• Oxalyl chloride (C2O2Cl2) produces light when oxidized - but only in the presence of a sensitiser, similar to the above examples. If oxalyl chloride is treated with H2O2 in non-aqueous media (e.g. CH2Cl2) in the presence of a sensitiser, emission of light is obtained. The colour, intensity and duration of light emission depend on the sensitiser used. Rodamin 6 G gives bright orange light with moderate duration of emission.

• Ru(bipy)32+ is a ruthenium(II) complex which undergoes oxidation to ruthenium(III) if certain oxidizing agents are introduced. If ruthenium(III) complex is then reduced in alkaline medium, emission of light occurs. First, there is a reaction:

2Ru(bipy)32+ + PbO2 + 4H+ → 2Ru(bipy)33+ + Pb2+ + 2H2O

Here, Ru(III) is obtained. Further reaction includes use of solution of sodium tetrahydroborate(III), NaBH4 in alkaline medium. When the solution is added, Ru(III) is reduced to Ru(II) and orange light is emitted.

• TMAE (tetrakis(dimethylamino)ethylene) emits clear blue-green light upon oxidation by air.

• Pyrogallol (1,2,3-trihydroxibenzene) is also capable of light emission. If an aqueous solution of pyrogallol, NaOH and K2CO3 is mixed with formaldehyde, short-lived red emission occurs.

• Singlet oxygen (O2) can also emit light. If solutions of 30% hydrogen peroxide and 5% alkaline sodium hypochlorite (NaClO) are mixed, red light is emitted. It is barely visible, though - for this reason a sensitiser is often included to boost light emission in terms of brightness and intensity. Again, both colour and intensity of light depend on the sensitiser used.

• Lucigenin oxidation is also very well known among chemiluminescence reactions. If an aqueous lucigenin solution is mixed with highly alkaline aqueous solution containing ethanol or acetone and hydrogen peroxide, very bright green emission is produced that decays to greenish blue and finally blue emission. The duration of the emission can be up to a couple of minutes under the right circumstances.

• Solutions containing ions of manganese (VII, IV, III) show chemiluminescence (690 nm) when reduced by sodium borohydride at low pH to Mn(II)

• Other liquid phase chemilumiscence reagents:

o peroxyoxalates

o Aryl oxalates

o Acridiniumesters

o dioxetanes

Gas-phase reactions

A green and blue glowstick.

• One of the oldest known chemoluminescent reactions is that of elemental white phosphorus oxidizing in moist air, producing a green glow. This is a gas-phase reaction of phosphorus vapor, above the solid, with oxygen producing the excited states (PO)2 and HPO.

• Another gas phase reaction is the basis of nitric oxide detection in commercial analytic instruments applied to environmental air quality testing. Ozone is combined with nitric oxide to form nitrogen dioxide in an activated state.

NO+O3 → NO2[◊]+ O2

The activated NO2[â—Š] luminesces broadband visible to infrared light as it reverts to a lower energy state. A photomultiplier and associated electronics counts the photons which are proportional to the amount of NO present. To determine the amount of nitrogen dioxide, NO2, in a sample (containing no NO) it must first be converted to nitric oxide, NO, by passing the sample through a converter before the above ozone activation reaction is applied. The ozone reaction produces a photon count proportional to NO which is proportional to NO2 before it was converted to NO. In the case of a mixed sample containing both NO and NO2, the above reaction yields the amount of NO and NO2 combined in the air sample, assuming that the sample is passed through the converter. If the mixed sample is not passed through the converter, the ozone reaction produces activated NO2[â—Š] only in proportion to the NO in the sample. The NO2 in the sample is not activated by the ozone reaction. Though unactivated NO2 is present with the activated NO2[â—Š], photons are only emitted by the activated species which is proportional to original NO. Final step, subtract NO from (NO + NO2) to yield NO2

Enhanced chemiluminescence

Enhanced chemiluminescence is a common technique for a variety of detection assays in biology. A horseradish peroxidase enzyme (HRP) is tethered to the molecule of interest (usually through labeling an immunoglobulin that specifically recognizes the molecule). This enzyme complex, then catalyzes the conversion of the enhanced chemiluminescent substrate into a sensitized reagent in the vicinity of the molecule of interest, which on further oxidation by hydrogen peroxide, produces a triplet (excited) carbonyl which emits light when it decays to the singlet carbonyl. Enhanced chemiluminescence allows detection of minute quantities of a biomolecule. Proteins can be detected down to femtomole quantities, well below the detection limit for most assay systems.

Applications

• gas analysis: for determining small amounts of impurities or poisons in air. Other compounds can also be determined by this method (ozone, N-oxides, S-compounds). A typical example is NO determination with detection limits down to 1 ppb

• analysis of inorganic species in liquid phase

• analysis of organic species: useful with enzymes, where the substrate isn't directly involved in chemiluminescence reaction, but the product is

• detection and assay of biomolecules in systems such as ELISA and Western blots

• DNA sequencing using pyrosequencing

• Lighting objects. Chemiluminescence kites, emergency lighting, glow sticks (party decorations).

• Combustion analysis: certain radical species (such as CH* and OH*) give off radiation at specific wavelengths. The heat release rate is calculated by measuring the amount of light radiated from a flame at those wavelengths.

• children's toys

Electrochemiluminescence

Electrochemiluminescence or electrogenerated chemiluminescence (ECL) is a kind of luminescence produced during electrochemical reactions in solutions. In electrogenerated chemiluminescence, electrochemically generated intermediates undergo a highly exergonic reaction to produce an electronically excited state that then emits light. ECL excitation is caused by energetic electron transfer (redox) reactions of electrogenerated species. Such luminescence excitation is a form of chemiluminescence where one/all reactants are produced electrochemically on the electrodes.

ECL is usually observed during application of potential (several volts) to electrodes of electrochemical cell that contains solution of luminescent species (polycyclic aromatic hydrocarbons, metal complexes) in aprotic organic solvent (ECL composition).

Application

ECL proved to be very useful in analytical applications as a highly sensitive and selective method. It combines analytical advantages of chemiluminescent analysis (absence of background optical signal) with ease of reaction control by applying electrode potential. Enhanced selectivity of ECL analysis is reached by variation of electrode potential thus controlling species that are oxidized/reduced at the electrode and take part in ECL reaction.

It generally uses Ruthenium complexes, esp [Ru (Bpy)3]2+ (which releases a photon at ~620 nm) regenerating with TPA (Tripropylamine) in liquid phase or liquid-solid interface. It can be used as monolayer immobilized on an electrode surface (made eg of nafion, or special thin films made by Langmuir-Blogett technique or self-assembly technique) or as a coreactant or more commonly as a tag and used in HPLC, Ru tagged antibody based immunoassays, Ru Tagged DNA probes for PCR etc, NADH or H2O2 generation based biosensors, oxalate and organic amine detection and many other applications and can be detected from picomolar sensitivity to dynamic range of more than six orders of magnitude. Photon detection is done with photomultiplier tubes (PMT) or silicon photodiode or gold coated fiber-optic sensors. ECL is heavily used commercially for many clinical lab applications.

UNIT III ASSESSMENT OF CELL MEDIATED IMMUNITY

Identification of lymphocytes and their subsets in blood

To clarify the immune mechanism in myocarditis, immunofluorescence techniques with laser flow cytometry were used to examine serial changes in lymphocyte subsets in the heart, spleen, and peripheral blood of DBA/2 and BALB/c mice inoculated with encephalomyocarditis virus (Experiment I). B cells were identified by staining with fluorescein isothiocyanate-labelled rabbit anti-mouse immunoglobulin. T- cell subsets were identified with rat anti-Thy 1.2, and nonpolymorphic Lyt 1 and Lyt 2 monoclonal antibodies plus fluorescein isothiocyanate- labelled anti-mouse immunoglobulin. On days 7 and 14 postinfection, the percentage of Thy 1.2+ (pan T) cells in both strains had decreased in the peripheral blood; B cells showed no significant changes throughout the entire period. On the other hand, Thy 1.2+ (pan T) and Lyt 1+, 23+ (precursor and immature) T cells appeared to occupy the major portion of the myocardium on days 7 and 14 when congestive heart failure developed. To confirm this, serial immunohistologic studies (immunoperoxidase staining) of the hearts of DBA/2 and BALB/c mice with encephalomyocarditis virus-induced myocarditis were performed (Experiment II). In Experiment II, most of the stained cells in the hearts of both strains were Thy 1.2 positive and Lyt 1 and Lyt 2 positive on days 7 and 14. Thus, Experiments I and II demonstrated that lymphocytes at the site of inflammation in acute viral myocarditis carried antigenic markers that differed from those of peripheral lymphocytes and suggested that Thy 1.2+ (pan T) cells, especially the Lyt 1+, 23+ subset (immature T cells and T-cell subset precursors) were involved in the development of myocarditis in these animals.

T cell activation parameters

1. T lymphocyte activation gene identification by coregulated expression on DNA m croarrays.

High-capacity methods for assessing gene function have become increasingly important because of the increasing number of newly identified genes emerging from large-scale genome sequencing and cDNA cloning efforts. We investigated the use of DNA microarrays to identify uncharacterized genes specifically involved in human T cell activation. Activation of human peripheral blood T lymphocytes induced significant changes in hundreds of transcripts, but most of these were not unique to T cell activation. Variation of experimental parameters and analysis techniques allowed better enrichment for gene expression changes unique to T cell activation. Best results were achieved by identification of genes that were most highly coregulated with the T-cell-specific transcript interleukin 2 (IL2) in a "compendium" of experiments involving both T cells and other cell types. Among the genes most highly coregulated with IL2 were many genes known to function during T cell activation, together with ESTs of unknown function. Four of these ESTs were extended to novel full-length clones encoding T-cell-regulated proteins with predicted functions in GTP metabolism, cell organization, and signal transduction.

2. Determination of soluble CD21 as a parameter of B cell activation.

In this study we established a novel solid-phase immunoassay for CD21 using the time-resolved fluorescence of lanthanide chelates. The capture assay was able to detect concentrations of as low as 100 pg of CD21 antigen per millilitre of sample and was used for quantitative determination of CD21 in lysates of different cell lines as well as in patient serum specimens. CD21 was measured in lysates of tonsils and cell lines of B, T cell and myelomonocyte lineage, and appeared to consist of monomeric antigen under the detergent conditions used. Elevated levels of soluble CD21 were observed in serum of patients with Epstein-Barr virus (EBV) infection, a disease known to be associated with polyclonal B cell activation, and in infection with the lymphotropic rubella virus. Significantly increased levels were also found in malignancies which are associated with EBV. In patients with nasopharyngeal carcinoma (NPC), a correlation with the titre of EBV-specific IgA was observed, thus supporting a possible role of soluble CD21 as a marker for disease activity in certain malignancies. Our data suggest that measurement of soluble CD21 could serve as a marker for activation of the immune system and diseases involving the B cell lymphoid system. Possible mechanisms and functions of soluble CD21 are discussed.

3. Multiple CD4 and CD8 T-cell activation parameters predict vaccine efficacy in vivo mediated by individual DC-activating agonists

A systematic comparison of the immunostimulatory capacity of TLR 2, 3, 4, 5, 7 and 9 agonists and an agonistic CD40-specific antibody was performed in a single long peptide vaccination model. All adjuvants activated DC in vitro but not all induced a strong functional T-cell response in vivo. Optimal clonal CD8+ T-cell expansion depended on the capacity of agonists to mature pro-inflammatory DC and the duration of their in vivo stimulatory effect. Strong agonists promoted the induction of both antigen-specific IFNγ-producing CD4+ T-helper cells and high numbers of IFNγ producing CD8+ effector T-cells that killed target cells in vivo. Importantly, the capacity of an agonist to function as an adjuvant depended on the vaccine strategy used. Collectively, the multi-parameter system presented here can be used as a general road map to develop therapeutic vaccines.

4. Anti-Qa-2-induced T cell activation. The parameters of activation, the definition of mitogenic and nonmitogenic antibodies, and the differential effects on CD4+ vs CD8+ T cells

The MHC Ag Qa-2 is a glycolipid anchored class I molecule expressed at high levels on all peripheral T lymphocytes. In this study we found that anti-Qa-2 antibodies could stimulate the proliferation of murine T cells in vitro. Anti-Qa-2-induced proliferation required secondary cross-linking with anti-Ig antibody and the presence of PMA. Only Qa-2+ strains could be induced to proliferate by anti-Qa-2 antibody, but under the conditions employed, anti-CD3 could induce proliferation in Qa-2+ and Qa-2-strains. Interestingly, only anti-Qa-2 reagents directed against the alpha 3 domain of the Qa-2 class I molecule were effective in inducing proliferation. Furthermore, unlike purified CD4+ cells, purified CD8+ cells were unable to be stimulated by the anti-Qa-2 antibodies. These results lead to the inclusion of Qa-2 in a group of physiologically relevant, glycolipid-anchored, cell-surface molecules, mobilization of which can generate signals that initiate the proliferation of T cells. Such molecules may play a secondary role in cellular activation after the primary engagement of the TCR.

5. Resistance to macrophage-mediated killing as a factor influencing the pathogenesis of chronic cutaneous leishmaniasis

Cutaneous leishmaniasis can be either a spontaneously healing or chronic disease, depending upon the strain of parasite and the immunological status of the host. We have investigated parasite factors responsible for the variable pathogenesis observed in leishmanial infections by testing the sensitivity of several leishmanial strains to intracellular killing in lymphokine (LK) activated mouse macrophages. Significant microbicidal activity against Leishmania tropica, a strain which heals in C57BL/6 (B6) mice, was found. In contrast, a strain (Maria) which has previously been shown to induce chronic nonhealing cutaneous lesions in B6 mice was resistant to killing in activated macrophages. This resistance to killing was observed in macrophages activated by LK obtained from either Bacille Calmette-Guerin-, L. tropica, or the Maria strain infected mice. The inability of LK activated macrophages to kill the Maria strain was shown not to be due to parasite induced inhibition of killing mechanisms, since Maria strain infected, LK treated macrophages exhibited tumoricidial activity similar to uninfected macrophages. Furthermore, LK activated macrophages simultaneously infected with the Maria strain and another intracellular pathogen, Toxoplasma gondii, killed Toxoplasma, but not the Maria strain. Temperature was also found to significantly influence the multiplication and killing of Leishmania parasites. As would be expected from their cutaneous nature, L. tropica and Maria strain parasites multiplied better at 35 degrees C than at 37 degrees C. Also consistent with the failure of cutaneous strains to visceralize in immunocompetent mice was the observation that the killing of leishmanial parasites was enhanced at the higher temperature. Thus, the temperature dependent growth capacity and sensitivity to killing of a given leishmanial strain in macrophages may be important factors influencing the pathogenesis of cutaneous leishmaniasis.

6. Effect of patulin on microbicidal activity of mouse peritoneal macrophages

Patulin, a fungal metabolite shown previously to exert immunosuppressive effects on the cellular and humoral immune systems, was examined for its in vitro effects on some functions of murine peritoneal macrophages. The cells were pre-incubated for 2hr with mycotoxin concentrations of 0.01–2 μg/ml. Phagocytosis and phagosome-lysosome fusion were diminished above 0.1 μg patulin/ml and lysosomal nzymes and microbicidal activity above 0.5 μg/ml,whereas O2− production was inhibited only above 2 μg/ml. This indicated that the killing mechanism did not depend on products of the oxidative burst. The concentrations used did not decrease the cell viability. Under natural circumstances, patulin may constitute a health risk for animals.

CYTOKINES

Cytokines are believed to be involved in the pathogenesis and enhanced expression in patients with Hodgkin’s and Non-Hodgkins lymphoma. Based on this phenomenon, a multicentric study was carried out in various lymphoma cases. The diagnosis of lymphoma was made on tissue biopsies and fine needle aspiration cytology (FNAC). Out of a total of 72 cases studied, 45 were of Hodgkin’s lymphoma (62.5%) and 27 cases were of Non-Hodgkin’s lymphoma (37.5%). Maximum cases of Hodgkin’s disease occurred in the age group of 30-40 years and males outnumbered females. Hodgkin’s lymphoma cases were predominantly of mixed cellularity histologic type (46.66%) whereas majority cases of Non-Hodgkin’s lymphoma were of high grade histologic type (48.14%) with predominance in the age group 51-60 years. In both these type of lymphomas, the IL-2R and IL-6 levels were found to be increased more than four fold (as compared to healthy controls) (p ................
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