The Basic Principles Of Immunohistochemistry

Published Date: 02 Nov 2017

Chapter 17

The protein is the end product of the genome. DNA is transcribed to RNA which is then translated into protein. The gene is constant; however, the protein structure can vary depending on the way the RNA is translated and post translational modifications.

The term ‘proteome’ was first used during the mid 1990s to describe the complete set of proteins related to the genome in a living organism or a living cell. The term ‘proteomics’ indicates large-scale characterization of the entire protein sequence and structure of a cell type, tissue, or organism. The complete characterization of all proteins has been the goal of proteomics since its inception almost 25 years ago. The features on the basis of which a protein can be described are multiple; these include localisation, interaction, domain structure, modification and activity of the protein in question. Since the proteome depends on splice isoforms and post translational modifications, the protein structure can vary considerably even if the gene coding for the protein is constant. Therefore, the understanding of the proteome is considerably more complex and difficult than understanding the genome. Unlike genomics, proteomics represents a more complex body of information to gather and analyse. Even for a single cell, the proteome changes in response to different stimuli.

It is important to understand proteomics because ultimately it is the protein sequence that dictates how the cell and therefore, how the tissue behaves. Study of the genome can provide at the best a limited understanding of functionality; whereas, study of the proteome can provide more complete understanding.

Initially, proteomics involved the separation of proteins and peptides by two dimensional gel-electrophoresis (2DGE) and their detection by matrix assisted laser desorption ionization mass spectrometry (MALDI MS). This meant a simple analysis of the proteins but over the years, proteomics has evolved to encompass profiling of proteins, deciphering their structure and their functional activity. Coinciding with the availability of completed genome sequences, there has been a shift to more comprehensive proteome analyses.

Essentially, proteomics involves protein profiling for a diseased and a normal tissue. Functional proteomics is the study of protein function. This includes study of post translational modifications (PTMs) of proteins and the interaction of proteins with substrates and small molecules. The concept of post translational modifications has already been dealt with in chapter 6.

Study of post-translational modifications is the key to understanding the functions and roles of proteins in a living organism. Since the proteins interact with other molecules, knowing the reaction pathways in which they are involved helps to understand the ‘wiring diagram’ of a cell.

<H1>WHY PROTEOMICS?

All information cannot be obtained from the study of the genes alone. It is impossible to elucidate mechanisms of disease, aging, and effects of the environment solely by studying the genome. Proteins, not genes, are responsible for the phenotypes of cells. Only through the study of proteins can protein modifications be characterized and the targets of drugs identified.

<H2>Annotation of the genome

Proteomics is essential in identifying the total number of genes in a given genome. This "functional annotation" of a genome is necessary because it is still difficult to accurately interpret the genomic data. Therefore, genomic information has to be integrated with the data obtained from protein studies to confirm the existence of a particular gene (Fig 17.1).

Protein

Amino Acid Sequence

Retrospective knowledge of the nucleotide sequence (Hence a knowledge of the mRNA sequence)

Confirmation/ discovery of the gene

Fig. 17.1: - Diagrammatic representation of gene discovery working retrospectively from the knowledge of the protein.

<H2>Protein expression studies

In recent years, the analysis of mRNA expression by various methods has become increasingly popular. M RNA is an unstable substrate. Further, , the analysis of mRNA is not a direct reflection of the protein content in the cell. mRNA is subject to posttranscriptional control in the form of alternative splicing, polyadenylation, and mRNA editing. It is possible to have many different protein isoforms are generated from a single gene in this step. mRNA is also subjected to regulation at the level of protein translation. Proteins, having been formed, are then subjected to posttranslational modification. It is estimated that up to 200 different types of posttranslational protein modification exist. The concept of ‘one gene, one protein’ is an oversimplification and therefore, cannot be used to define proteins accurately. Thus, protein analysis always scores over mRNA analysis in understanding the functioning of the human body.

<H2>Protein function

According to one study, no function can be assigned to about one-third of the sequences in organisms for which the genomes have been sequenced. The complete identification of all proteins in a genome would aid the structural genomics, where the ultimate goal is to obtain a 3-D structure for all the proteins in a proteome. This is necessary because the functions of many proteins can only be inferred by examination of their 3-D structure.

<H2>Protein modifications

One of the most important applications of proteomics is the characterization of posttranslational protein modifications. Proteins undergo posttranslational modification in response to a variety of intracellular and extracellular signals. By using the proteomics approach, these modifications can be analyzed simultaneously for many proteins.

<H2>Protein localization and compartmentalization

One of the most important regulatory mechanisms currently known is protein localization. A wrong localization of proteins has profound effects on cellular function (e.g., cystic fibrosis). Proteomics aims to identify the subcellular location of each protein. This information can be used to create a 3-D protein map of the cell, providing novel information about protein regulation.

<H2>Protein-protein interactions

Of fundamental importance in biology is the understanding of protein-protein interactions. Proteomics aims to develop a complete 3-D map of all protein interactions in the cell.

Nuggets

The protein is the end product of the genome. The proteome is the complete set of proteins in a living organism or a living cell, related to the genome expressing these proteins.

Proteomics involves profiling of proteins, deciphering their structure and their functional activity.

Proteomics is important for annotation of the genome, protein expression studies, protein function, protein modifications, protein localisation and compartmentalization and to study protein–proteion interactions.

<H1>TYPES OF PROTEOMICS

Protein expression proteomics. The quantitative comparison of protein expression in samples that differ in some variable is known as expression proteomics. The expression of the entire proteome or the subproteomes in different samples can be compared. It is then possible to identify novel proteins in signal transduction. It also enables identification of disease-specific proteins. An example of this approach is application of proteomics for cancer where a novel new protein maybe expressed in the cancerous cell but not in the normal cell.

Structural proteomics. Studies concerned with mapping of either the structure of protein complexes or the protein itself in a specific cellular organelle are known as structural proteomics. It identifies all proteins in a protein complex or organelle. It also determines where the proteins are located and characterises the protein–protein interactions. This information helps to piece together the overall architecture of cells and explain how expression of certain proteins gives a cell its unique characteristics.

Functional proteomics. In functional proteomics, a selected group of proteins is studied and characterised. It can provide information about protein signalling, disease mechanisms and protein-drug interactions.

<H1>TOOLS OF PROTEOMICS

The tools used in proteomics are many and include Western blotting, immunohistochemistry, mammalian two-hybrid, peptide and protein microarrays, x-ray crystallography, NMR spectroscopy, fluorescence microscopy, mass spectrometry and flow cytometry. Mass spectrometry is the tool which is most commonly used and sometimes proteomics is used synonymously with mass spectrometry. Western blotting has already been covered in chapter 12.. Therefore, techniques other than Western blotting would be described here with special emphasis on mass spectrometry.

<H2>MAMMALIAN TWO HYBRID

Interactomes are dynamic protein networks. Specific protein–protein interactions are studied in the mammalian two hybrid. Important factors in these interactions include protein modifications and/or structural alterations in the interacting proteins which can occur only under specific cellular conditions. Yeast and mammalian two-hybrids allow the assayed proteins to undergo proper modifications in their native cellular context. Thus, it allows protein interactions to be tracked as a function of time, space and physiological context.

<H2>ONE- AND TWO-DIMENSIONAL GEL ELECTROPHORESIS

For many proteomics applications, 1 Dimensional Electrophoresis(1 DE) is the method of choice to resolve protein mixtures. 1 – DE uses the molecular mass as a basis of separation of proteins. . Proteins are solubilized in sodium dodecyl sulfate (SDS). 1-DE is simple to perform, reproducible, and can be used to resolve proteins with molecular masses of 10 to 300 kDa. The most common application of 1-DE is the characterization of proteins after some form of protein purification. This is because of the limited resolving power of a 1-D gel.

If there is a more complex protein mixture such as a crude cell lysate, then 2-DE can be used. In 2-DE, proteins are separated by two properties. They are resolved according to their net charge in the first dimension and according to their molecular mass in the second dimension. Therefore, it uses charge as the basis of initial separation. The second phase is similar to 1-DE electrophoresis. The combination of these two techniques produces resolution far exceeding than that obtained in 1-DE.

One of the greatest strengths of 2-DE is the ability to resolve proteins that have undergone some form of posttranslational modification. This resolution is possible in 2-DE because many types of protein modifications confer a difference in charge as well as a change in mass on the protein.

The primary application of 2-DE is in protein expression profiling. In this approach, the protein expression of any two samples can be qualitatively and quantitatively compared. The appearance or disappearance of spots can provide information about differential protein expression, while the intensity of those spots provides quantitative information about protein expression levels.

A single 2-DE gel can resolve thousands of proteins; it remains a powerful tool for preparing a catalogue of proteins. Many 2-DE databases have been constructed and are available online..

Despite so many advantages with 2-DE electrophoresis, a number of problems still persist. It is a labour intensive and time consuming process. The number and the type of proteins that can be resolved is limited. It is difficult to detect proteins which are present in low concentrations (low copy proteins). Because of these limitations, the largest application of 2-DE in the future will probably involve the analysis of protein complexes or subproteomes as opposed to whole proteomes.

Fig. 17.2: Image of 2D gel electrophoresis. As can be seen in the sample, the initial separation is in the vertical direction with the higher molecular weight proteins above and the lower molecular weight proteins below. Then the separation is based on charge. This allows even post translational proteins to be separated since post translational modifications make changes in the mass and the charge of the protein.

<H2>PROTEIN X-RAY CRYSTALLOGRAPHY

Protein X-ray crystallography and NMR spectroscopy are currently two methods which can analyse tertiary protein structures. These methods dominate the field of structural biology.

X-rays are electromagnetic waves with a very short wavelength of around one Angstrom. X rays can be generated in several ways. They can be of very high intensity and may be highly focussed. The protein of interest is placed in the path of a monochromatic single wavelength X ray beam. When the X ray hits the crystal, X-ray diffraction takes place. Diffraction is seen whenever wave of any nature encounters an obstacle which can be any material object. This results in bending of wave around that object. This is also called ‘scattering of the wave.’

The phenomenon of scattering can also occur when a wave encounters a small opening or a slit which causes spreading of the wave in all directions. In both cases, the new scattered waves interact with each other and may either get stronger or cancel each other. Those waves which get stronger are detected by an X ray detector. Thousands of different diffraction spots are then collected. Depending on the type of protein, the data is processed in order to produce a three dimensional image of the protein.

The protein is then rotated and again exposed to the X-rays. The data is reanalysed. The complete structure of the protein is obtained using several computer programs like Mosflm, XDS and HKL 2000. The intensities are used to calculate an electron density map into which a model of the protein is built. The electron density map shows the positions of the amino acids, the main chain and the side chains. These can be used to trace the whole polypeptide chain of the protein in question.

Fig. 17.3: X Ray crystallography. A protein crystal is bombarded by X rays. The X rays diffract and produce different diffraction patterns. The protein are rotated in different axes and different diffraction maps are produced. Based on these diffraction patterns, an electron dense map is made. Extrapolating these electron density maps leads to the final structure of the protein.

<H2>EDMAN SEQUENCING

One of the earliest methods used for protein identification was microsequencing by Edman Chemistry to obtain N-terminal amino acid sequences. With the advent of the mass spectrometer, the use of Edman’s sequencing has declined; however, it still continues to be used when mass spectrometer is not available.

Essentially, the protein is electrophoresed on a polyacrylamide gel and blotted on an inert membrane. The proteins of interest are excised and fragmented chemically. On an average, three to five peptides are generated and the membrane is placed in an automated Edman’s sequencer without any further manipulation. Six to twelve cycles are run and the data is fed into algorithms which compare the data against protein. The FASTF or TFASTF algorithms are usually used.

<H2>MASS SPECTROMETRY

Mass spectrometry remains the method of choice for detection of new proteins. It can be used to identify proteomic changes that occur in diseases and to gain insight into their pathogenesis. Such proteomic changes include alterations in the amount of proteins, their post-translational modifications and subcellular localization.

Essentially the study of proteomics is done using mass spectrometry. Several mass spectrometry platforms are commercially available. One commonly used method is the surface enhanced laser desorption/ionisation technology. This allows matrix assisted laser desorption and time of flight analysis. The variations in time of flight based on mass and electric charge of different protein components are utilised to separate the different protein components. With the aid of computer analyses, results are displayed as peaks representing the different peptide components of a given sample.

<H3>Methodology

A mass spectrometer converts individual molecules to ions so that they can move about and be manipulated by external electric and magnetic fields. Modern mass spectrometers allow identification of peptides and proteins at subfemtomolar levels, thus, potentially enabling the study of low-abundant proteins, such as most proteins involved in signal-transduction processes. Proteins are usually separated either by 1 D or 2D gel electrophoresis or by a chromatographic technique. After separation of the proteins, the proteins are digested. This is done because measurement of masses of fragmented peptides is more accurate than measuring masses of intact proteins. Following protein digestion, mass spectrometry is performed which measures masses of peptides and reveals their primary structures.

Fig 17.4 – Flowchart showing the sequence of events which are followed in Mass Spectrophotometry.

The three essential functions of a mass spectrometer, and their associated components, are as follows:

A small sample of compound is ionized, usually to cations. This occurs by loss of an electron and is done by the ion source.

The ions are sorted and separated according to their mass and charge. This is done using the mass analyzer.

The separated ions are then detected and tallied, and the results are displayed on a chart. This is done using the detector.

A diagrammatic representation of mass spectrophotometry is shown in Fig 17.4.

As ions are very reactive and short-lived, their formation and manipulation must be conducted in a vacuum. The pressure under which ions should be handled is roughly less than a billionth of an atmosphere.

Ionization is effected by the high energy beam of electrons. Ion separation is achieved by accelerating and focusing the ions in a beam, which is bent by an external magnetic field. The ions are then detected electronically and the resulting information is stored and analyzed in a computer.

A mass spectrum is usually presented as a vertical bar graph. Each bar in the graph represents an ion having a specific mass-to-charge ratio (m/z) and the length of the bar indicates the relative abundance of the ion. The most intense ion is assigned an abundance of 100, and it is referred to as the base peak. Most of the ions formed in a mass spectrometer have a single charge, so the m/z value is equivalent to mass itself. Modern mass spectrometers easily distinguish ions differing by only a single atomic mass unit (amu), and thus provide completely accurate values for the molecular mass of a compound.

<H3>Sample ionization

For biological samples to be analyzed by MS, the molecules should be charged and dry. This is accomplished by converting them to desolvated ions. The two most common methods for this are electrospray ionization (ESI) and matrix-assisted laser desorption/ionization (MALDI). In both methods, peptides are converted to ions by the addition or loss of one or more protons. In both the methods, there is no significant loss of sample integrity.

Electrospray ionization. In this method, the liquid sample flows from a microcapillary tube into the orifice of the mass spectrometer. Because of this flow, a potential difference between the capillary and the inlet to the mass spectrometer is generated. This results in the formation of a fine mist of charged droplets. The solvent then evaporates and the size of the droplets decreases. This then results in the formation of desolvated ions.

Matrix-assisted laser desorption/ionization. In MALDI, the sample is incorporated into matrix molecules and then subjected to irradiation by a laser. The laser promotes the formation of molecular ions. In MALDI, the entire system can be automated. This is the single biggest advantage of MALDI. Another advantage of MALDI over ESI is that samples can often be used directly without any purification after in-gel digestion.

A diagrammatic representation of EIS and MALDI is shown in Fig 17.5

Fig 17.5 – A diagrammatic representation of Electrospray Ionisation and Matrix Assisted Laser Desorption/ Ionisation

<H3>Mass analysis

Mass analysis follows the conversion of proteins or peptides to molecular ions. This is accomplished by mass analyzers in a mass spectrometer, which resolve the molecular ions on the basis of their mass and charge in vacuum. There are three types of mass analyzers:

Quadrupole mass analyzers. It is the most common mass analyzer that is being used. Here, ions are transmitted through an electric field created by an array of four parallel metal rods called the quadrupole. A quadrupole may either transmit all ions or may serve as a mass filter to allow the transmission of ions of a certain mass-to-charge (m/z) ratio.

Time of flight mass analyzers. A time-of-flight (TOF) instrument is the simplest mass analyzer that is being used. It measures the m/z ratio of an ion by determining the time required for it to traverse the length of a flight tube. Some TOF mass analyzers have an ion mirror at the end of the flight tube, which reflects ions back through the flight tube to a detector. This causes a doubling of the length the ion has to traverse. This improves the sensitivity of the analysis. .

Ion trap mass analyzers. These function to trap molecular ions in a 3-D electric field. They thus ‘store’ ions which are then selectively ejected.

<H3>Types of Mass Spectrometers

Most mass spectrometers consist of four basic elements:

An ionization source,

One or more mass analyzers,

An ion mirror, and

A detector.

The analysis of proteins or peptides by MS can be divided into two general categories:

Peptide mass analysis

Amino acid sequencing.

In peptide mass analysis or peptide mass fingerprinting, the masses of individual peptides in a mixture are measured and used to create a mass spectrum. In amino acid sequencing, a procedure known as tandem mass spectrometry, or MS/MS, is used to fragment a specific peptide into smaller peptides, which can then be used to deduce the amino acid sequence.

With this background in mind, we move on to the main types of instruments.

Triple quadrupole. Triple-quadrupole mass spectrometers are most commonly used to obtain amino acid sequences.

Quadrupole-TOF. It is a form of a hybrid spectrometer in which the quadrupole collision cell (q) of a triple-quadrupole machine is combined with a time-of-flight analyzer (TOF). The main applications of a QqTOF mass spectrometer are protein identification by amino acid sequencing and characterization of protein modifications.

MALDI-TOF. The principal application of a MALDI TOF mass spectrometer is peptide mass fingerprinting because it can be completely automated, making it the method of choice for large-scale proteomics work.

MALDI-QqTOF. The MALDI-QqTOF mass spectrometer was developed to permit both peptide mass fingerprinting and amino acid sequencing. It was formed by the combination of a MALDI ion source with a QqTOF mass analyzer.

FT- ICR. A Fourier transform ion cyclotron resonance (FT-ICR) mass spectrometer is an ion-trapping instrument that can achieve higher mass resolution and mass accuracy than any other type of mass spectrometer. It has a high resolution and can be used in the analysis of complex peptide mixtures.

Nuggets

Types of proteomics include protein expression proteomics, structural proteomics and functional proteomics.

The mammalian two-hybrid system allows one to study protein-protein interactions.

2-D electrophoresis allows us to separate proteins based on charge and mass. It allows resolution of proteins that have undergone some form of posttranslational modification.

Protein X ray crystallography and NMR spectroscopy are currently two methods which can analyse tertiary protein structures. It is based on the principle of scattering of X rays by proteins.

Mass spectrometry remains the method of choice for the detection of new proteins.

A mass spectrometer converts individual molecules to ions so that they can be moved about and manipulated by external electric and magnetic fields. It allows for identification of peptides and proteins at subfemtomolar levels.

Mass spectrometry measures masses of peptides and reveals their primary structures.

Sample ionisation can be done by electrospray ionisation or matrix-assisted laser desorption/ionisation.

Mass analysis can be done by quadrupole mass analysers, time-of-flight mass analysers and ion trap mass analyzers.

<H1>MICRODISSECTION TECHNIQUES

Very often, it is necessary to study the proteomics of a specific cell type in a mixed cell population; for example, it maybe necessary to study breast cancer cells in a specimen of a modified radical mastectomy. The cancer cells may comprise only 10% of the total cells. One way to reduce contamination is to culture the cells from that tissue. However, cultured cells may not accurately reflect the molecular changes taking place in that tissue. The best way would be to remove those particular cells and analyse them separately. This can be done by laser tissue microdissection. A diagrammatic representation of laser microdissection is shown in Fig 17.6.

In microdissection, microscopic homogenous cellular population is extracted from complex tissue. This population can then be compared with the adjacent non-involved tissue. Laser capture microdissection (LCM) has been developed to provide scientists with a fast and dependable method of capturing cells from tissues under direct microscopic visualisation. The development of LCM has allowed scientists to determine specific protein expression profiles. It is also possible to correlate the pattern of expressed genes and post translationally modified proteins with histopathology, and study the interactions between cellular subtypes in an organ or tissue microenvironment.

Fig 17.6 – A diagrammatic representation of a microdissection technique.

<H1>SERUM PROTEOMICS

By the time cancer is diagnosed, the cancer cell has already completed a major portion of its life cycle. Most patients with cancer of the breast, lung and colon have metastatic colonies at the time of diagnosis. At this stage, therapeutic modalities are few and may not significantly influence the course of the disease. It would be invaluable if cancer could be detected early so that treatment can be initiated early.

A clinically useful biomarker should be measurable in a readily accessible fluid such as serum, urine or saliva. Clinical proteomics is capable of detecting such cells. Serum or plasma is the preferred medium for discovery because it contains cells dislodged by the constant movement of blood throughout the body. As always, the problem in the detection of a metastatic clone of cells is the heterogeneity of the cancer cell population. Therefore, it maybe possible to use a protein ‘A’ to detect 40% of the metastatic clones, marker ‘B’ to detect 50% of the metastatic clones and marker ‘C’ to detect 30 % of the metastatic clones. However, if all the three markers are used, it would be possible to detect 100% of the metastatic clone in all patients.

The low molecular weight range of the serum proteome (indicating peptides less than 50,000 Daltons) is called a peptidome due to an abundance of protein fragments and peptides. Some investigators dismiss this range of proteome as biological trash and say that the peptides are too small to be biologically relevant. Other investigators have proposed that the peptidome may contain a large number of peptides of biological significance. Tissue proteins that are too small to pass through the endothelium into the circulation may still be seen as fragments of the parent molecule. Metastatic cells often exhibit this pattern. This peptidome can then be analysed by using tools like mass spectrometry to identify molecules which can be used to predict the presence of metastatic cell clones.

Nuggets

Microdissection techniques allow us to separate the protein of interest from a tissue for further analysis.

Serum proteomics allows us to pick up cancer cells early from the blood so that early treatment can be initiated.

<H1>PROTEINOPEDIA

Clinical proteomics deals with the application of proteomic technologies to help decipher the changes that occur in cells, tissues, and organs in diseased conditions. There is an increasing knowledge about the different kinds of proteins seen in normal and diseased tissues. It is essential that this knowledge be integrated and kept on a platform so that the data is available to all the researchers. The integration of this data also means that researchers at a later date can question the data and refine their knowledge.

In future, it may even be possible to diagnose a particular disease condition from organ-specific proteomic signatures present in the serum. For this, we must first systematically obtain proteomic data from individual organs. Such data can be archived, and meta-analysis can be carried out to decipher the signatures.

<H2>Genomic Versus Proteomic Data

In the case of genomic data, the International Nucleotide Sequence Consortium has already established a working principle according to which any sequence data that is submitted to any one of the 3 members, Gen Bank, European Molecular Biology Laboratory (EMBL), and DNA Data Bank of Japan (DDBJ), would automatically be reflected in the other data bases. However, unlike genomic data, proteomic data is diverse with a multitude of experimental platforms and data types with the result that there are no general working principles for data submission that apply to all types of proteomic data. Proteomic data is, therefore, more difficult to handle and process as compared to genomic data.

Very often, proteomic data is published as supplementary information. This has several disadvantages; the principal disadvantage being that the supplementary data is not easily available. On the contrary, data contributed to centralized repositories can be downloaded freely and is more searchable. It is thus possible for information from diverse research papers to be integrated and submitted for use in the form of a human proteinopedia.

<H2>Standardization and Vocabulary Issues in Proteomic Data

Gene nomenclature is regulated by the Human Genome Organization, whereas naming of the proteins is largely left to individual investigators. Therefore, retrieval of information for proteins in a comprehensive fashion becomes difficult. Some standardised vocabularies are emerging like the ‘eVOC’ for describing tissue expression, 'gene ontology for cellular component, molecular function, and biological process, while ‘RESID’ and ‘Proteomics Standard Initiative Molecular Interaction’ vocabularies are available for post-translational modifications and protein-protein interactions, respectively. However, these vocabularies are still not standardised completely.

<H2>Concept of a Human Proteinopedia

It is important to remember that in the context of sharing human protein data, there are two major issues. Firstly, all the data should be shared regardless of the size of the data base. Secondly, there should be a central portal where the available data is compiled and displayed in the context of a gene/protein. The latter feature would permit users to construct complex queries such as "what are the post-translational modifications on one’s protein of interest, what are its interacting proteins, its subcellular localization, and if it is overexpressed in cancers".

Such queries cannot be answered in any of the existing proteomic repositories although some provide links to other data bases for certain data types.

The human Proteinpedia was conceived with these aims in mind. It is a community portal for sharing human proteomic data that is developed with the active participation of more than 70 laboratories around the world. It allows researchers to share their human proteomic data in a manner that is somewhat similar to that of Wikipedia. However, experimental evidence is mandatory for inclusion of data in human Proteinpedia and the contributions are always linked to the investigator and the laboratory. Annotations pertaining to post-translational modifications, expression in cell lines or tissues, protein-protein interactions, enzyme substrate, and subcellular localization can be submitted. Human Proteinpedia includes data from diseases such as cancers, thereby allowing the biomedical community to take a system’s view of the disease proteome. Moreover, it can accommodate data from multiple experimental platforms such as yeast two-hybrid screens, peptide/protein arrays, immunohistochemistry, Western blots, mass spectrometry, co-immunoprecipitation, and fluorescence microscopy. Thus, Human Proteinpedia represents an early attempt to unify human proteomic data under a single resource.

Unlike genomic data, however, proteomic data is diverse with a multitude of experimental platforms and data types. The aim of researchers should be to unify all this data under a single portal. This has been done partially using the Human Proteinpedia. In this platform, all the data is integrated and stored and is available for researchers when they need it. However, complex integrations between proteins need to be evaluated. After that, it is hoped that our understanding of the human proteome would reach a stage when it can influence diagnostics and therapeutics.

Box 15.4

It is essential that the knowledge of proteins be integrated and kept on a single platform so that the data is available to all the researchers. This is the concept of a proteinopedia.

Unlike genomic data, proteomic data is diverse with a multitude of experimental platforms and data types, and therefore, it is not standardised. Proteomic data is, therefore, more difficult to handle and process as compared to genomic data.

The concept of a Human Proteinopedia is to share all data. . The data should be shared regardless of the size of the data base. There should be a central portal where the available data is compiled and displayed in the context of a gene/ protein.

<H1>BASIC PRINCIPLES OF IMMUNOHISTOCHEMISTRY

Immunohistochemistry (IHC) or Immunocytochemistry is a method of localising specific antigens in a cell or tissue based on antigen–antibody recognition. Essentially, in an IHC, the antigen antibody reaction occurs and it is visualised at a light microscopic level. An example of a section stained using immunohistochemistry is shown in Fig 17.7

Fig 17.7 - An example of the use of immunohistochemistry in clinical practice. The picture shows a section of the skin with prominent dermal blood vessels. The vessels have been stained using CD 34, a marker for endothelial cells.

The initial pioneer of this technique was Coons who used immunofluorscence to detect specific antigens in frozen sections. This was not very useful to the histopathologist since it was a very restricted technique; not all tissues could be processed in frozen section and immunofluorscence is a relatively difficult technique. However, since the early 1990s, the method began to be used in general pathology. Detection systems have now become sensitive and antibodies specific. The development of horse radish peroxidase as a detection technique has considerably enhanced the sensitivity of detection of antigens.

The development of the ‘hybridoma’ technique facilitated the development of IHC and the manufacture of abundant, highly specific antibodies. After frozen section, IHC was extended to include paraffin sections. Soon afterwards, IHC became a routine application for all paraffin embedded tissues. The concept of unmasking antigens was developed by Huang et al. This was followed by the concept of antigen retrieval. With all these techniques, IHC has now become a standard procedure used in most pathological laboratories.

<H2>BASIC PRINCIPLES

The standard haematoxylin and eosin stain is capable of providing a lot of information to the pathologists but very often, problems persist during diagnosis. Initially, histochemical stains were used. Several improvements in diagnostics were made by intelligently using the Periodic Acid Schiffs and the reticulin stain. These stains were capable of highlighting specific cellular or histological features and therefore, could improve the diagnosis. However, they could still not provide a specific diagnosis which was left to IHC.

The basic principle of IHC is the localisation of target components in a cell or tissue. The aim is to amplify the signal while reducing the non specific background staining. It is now possible to perform IHC routinely on tissues in several situations.

<H2>ANTIBODIES AS STAINING AGENTS

An antibody is capable of combining with an antigen. The recognition of an antigen is based on the recognition of its three dimensional structure which is a critical issue in IHC. It must be remembered that all fixatives distort the tissues. The most commonly used fixative is formalin which causes protein–protein links and therefore, distorts the structure of the protein. Since the proteins are folded, it maybe impossible for the antibody to combine with the antigen until the epitopes are exposed. This would be covered later under the heading of antigen retrieval.

An epitope is an antigenic determinant, the determinant being the exact site of the molecule with which the antibody combines. The epitope is a cluster of amino acids residues. It is a functional unit and not a fixed structure of the protein. Epitopes can be classified as continuous and discontinuous. A continuous epitope is one where there is a continuous chain of residues in a polypeptide chain, whereas, discontinuous is where residues from different parts of a polypeptide chain are brought together because of protein folding. A diagrammatic representation of continuous and discontinuous epitopes is shown in Fig 17.8.

Fig 17.8 – A diagrammatic representation of a protein showing the continuous and discontinuous epitopes.

Antibodies are both sensitive and specific for protein antigens. Initially, it was difficult to manufacture antibodies on a large scale but with the development of the hybridoma technique, it became possible to produce endless amounts of antibodies. Essentially what is done in this technique is that myeloma cells are fused with the cells expressing the protein of choice. Polyclonal antibodies show a reaction to a variety of antigens but a monoclonal antibody shows a reaction to only one or a very few select antigens. A polyclonal antibody is more sensitive but a monoclonal antibody is naturally more specific.

Most commercially available antibodies are highly reliable for IHC but this needs to be checked by evaluating the specificity of the staining pattern and the expected type of staining to be seen. It is always useful to compare the staining pattern with the literature and see if the staining pattern obtained is the same as what has been quoted in other studies.

<H2>BLOCKING BACKGROUND STAINING

There are two possibilities which may contribute to background staining; the first is that there maybe a non specific antibody binding and the second is that there maybe endogenous enzyme activity.

The problem of non specific binding can easily be resolved by using highly specific monoclonal antibodies. Polyclonal antibodies should be avoided as explained previously. It is also a good idea to incubate the tissue with normal human serum prior to staining with the antibody. This occupies unwanted antigenic sites and thus allows the primary antibody to have a more specific binding. Diluting the antibody also helps; an optimum dilution of the antibody prevents non specific binding. Antibodies are highly charged molecules and may bind non specifically to collagen and other tissue components. This can also be prevented by incubation with serum.

The problem of endogenous enzyme activity is more complex. Some enzymes like peroxidase are preserved in tissues even after fixation. These are capable of breaking peroxide used in the final stage of staining and cause a non specific colour reaction. The removal of the peroxidase activity is a must in immunohistochemical staining.

Usually endogenous peroxidase activity is removed by incubation of sections in a mixture of H2O2 and methanol. Some workers use a mixture of 0.075% hydrochloric acid and ethanol. If it is not possible to block endogenous peroxidase activity by these methods, other methods such as the immunogold or the glucose oxidase method should be used for the development of colour.

<H2>DETECTION SYSTEMS

Detection systems are used to tag the antibody so that it can be visualised by light microscopy. A variety of flags have been used including fluorescent compounds and active enzymes which are capable of generating a coloured reaction from a suitable substrate.

Direct conjugate labelled antibody method –

This is the simplest method to use. A label is attached to the antibody and the antibody is then directly applied to the tissues. The aim is to attach a maximum number of labels to the antibody so that it can be optimally visualised in the tissues. The labelling process should not inactivate either the antibody or the label. There should be no free label that may possibly bind to the tissues and create a non specific antibody binding. Lastly, the binding antibody should be monoclonal, otherwise again non specific reactions may occur. All these conditions are difficult to achieve in a pathology laboratory and hence this method is not generally used. However, the direct conjugate labelled antibody method would be remembered as a simple method which had the advantage of rapidity and ease of performance.

Indirect of sandwich procedure –

In this procedure, the conjugated antibody is separated from the primary antibody. The conjugation process is applied only to the secondary antibody. This allows the reaction to be far more flexible as compared to the direct conjugate method. The other advantages are that the primary antibody can often be used at a higher working dilution. All labelled antibody methods performed by the indirect procedure have the same principle.

Biotin Avidin method –

This method exploits the high affinity between biotin and avidin. Biotin can be linked to the primary antibody and therefore, it localises the site of the antigen. Avidin conjugated to horse radish peroxidase is then added. It binds to the biotinylated antibody and thus localises the peroxide molecule at the site of the antigen. The antigen can then be detected by using a substrate for peroxidase. Some tissues contain endogenous biotin and this may cause problems in the reaction.

The problems associated with the avidin biotin system can be circumvented by substituting streptavidin for avidin. The streptavidin can be directly conjugated to the enzyme molecule. It is capable of binding biotin with high affinity and thus provides specific detection and amplification system for antigen–antibody binding. In addition, streptavidin does not contain carbohydrates (which bind non specifically to lectins). This offers significant advantage. The reagent is highly stable and can be stored without any problems for long periods.

Polyvalent systems –

In the polyvalent system, the secondary reagent contains multiple antibodies raised against immunoglobulins of different species. This allows one secondary reagent to be used for both polyclonal and monoclonal antibodies.

Protein A methods –

Protein A derived from Staphylococcus has the capacity to bind to the Fc portion of the immunoglobulin molecule from several different species. The primary antibody binds with protein A and not the secondary antibody. Thus, this method does not have the sensitivity of the PAP or the biotin avidin technique.

Other methods –

The enzyme labelled antigen methods, polymeric labelling two stage method and the tyramine signal amplification methods are other methods used for antigen detection.

<H2>ANTIGEN RETRIEVAL

In the current literature, the term ‘antigen retrieval’ is defined as a high-temperature heating method to recover the antigenicity of tissue sections that had been masked by formalin fixation. The term ‘AR’ was also applied for non-heating methods, including the enzymatic method. The AR technique has widespread application in pathology and other fields of morphology. Antigen retrieval has shown a pronounced enhancement of IHC staining on archival formalin-fixed, paraffin-embedded tissue sections for a variety of antibodies. The improvements in staining has occurred in several ways including a reduction of the detection thresholds of immunostaining (increasing sensitivity) and retrieval of some antigens, such as Ki- 67 and androgen receptor which would otherwise have been negative in IHC. The entire exercise is performed in order to improve the threshold of antigen detection, it is therefore an important aspect of the pre detection phase. .

Antigen retrieval has also achieved satisfactory results in IHC for tissues embedded in plastic embedding media used for immunoelectron microscopy (IEM), celloidin-embedded tissues, as well as in cell smear preparations fixed in non-cross linking fixatives. The heat-induced retrieval approach has also been applied to in situ hybridization (ISH), TUNEL [terminal deoxynucleotidyl transferase (TdT)-mediated dUTP-biotin nick end-labeling], and FISH.

There are several important issues involved in antigen retrieval. . First, not all antigen structures modified by formalin can be restored using conventional AR protocols. Therefore, standardisation of antigen retrieval techniques is very important. Optimum AR protocols need to be developed for certain antigens under investigation. For a few proteins, higher temperature (boiling) may induce a negative result of IHC staining. In this case, a lower-temperature heating treatment or a combining retrieval protocol (heat and enzyme digestion) may provide better results.

Knowledge of the exact localization of a certain protein (antigen) in tissues is critical to interpret not only the accuracy of IHC staining results but also the reliability of AR treatment. Control groups, including a tissue section that is not treated by AR, are required to rule out false-positive results and altered immunostaining pattern.

Although the AR technique is a simple method, it is necessary to understand the factors that influence the effectiveness of IHC staining, particularly two major factors, the heating conditions and the pH value of the AR solution. Antigens can be broadly grouped under three categories with respect to the importance of pH on AR. These are as follows:

Most antigens showed no significant variation using AR solutions with a pH value ranging between 1.0 to 10.0

Some antigens like nuclear antigens showed optimum results at low pH

Some antigens like HMB 45 showed good results at high pH.

Antigen retrieval can be done by heating the tissue in a microwave or pressure cooker. It is generally accepted that heating in a pressure cooker gives better results. It is believed that pressure cooking creates uniform heat which helps in better antigen retrieval.

Antigen retrieval can also be performed using enzymes like proteinase K, trypsin and pepsin. The principle is as follows:

Formalin forms protein cross-links that mask the antigenic sites in tissue specimens and prevent immunohistochemical detection of certain proteins. The enzymes are designed to break the protein cross-links and therefore unmask the antigens and epitopes in formalin-fixed and paraffin embedded tissue sections. This enhances the staining intensity of antibodies.

However, since the procedure is difficult to standardize, it is rarely performed.

<H2>FIXATION

In performing their protective role, fixatives denature proteins by coagulation or by forming additive compounds or by a combination of the two. Thus, conformational changes in the structure of proteins occurs causing inactivation of enzymes. The resulting complexes differ from the non denatured proteins in both chemical and antigenic profiles. These changes can adversely affect immunohistochemical analysis.

By far, the largest proportion of samples used for immunostaining is embedded in paraffin, and the usual fixative used is formalin. There may be shrinkage or distortion during fixation or subsequent paraffin-embedding, but generally formalin-based fixatives are excellent for most immunostains. Formaldehyde fixes not by coagulation, but by reacting primarily with basic amino acids to form cross-linking "methylene bridges." This means that there is relatively low permeability to macromolecules and that the structures of intracytoplasmic proteins are not significantly altered. It is the great variation in time and conditions for fixation that cause the majority of problems in immunochemistry. Although some antigens are not well demonstrated after fixation in formaldehyde-based fixatives, many can be demonstrated after the use of appropriate pre-treatment methods. If monoclonal antibodies are to be utilized on formalin-fixed, paraffin-embedded tissue sections, there are three considerations, which should be kept in mind:

Does formaldehyde react with the epitope under investigation?

Does it react with adjacent amino acids causing conformational changes?

Does paraffin processing destroy the epitope under investigation?

Among the other fixatives, mercuric fixatives show better antibody penetration, resulting in a more intense immunostain. However, loss of immunoreactivity occurs through blockage of specific epitopes. This is particularly evident with monoclonal antibodies. Zenkers solution and B5 show a good cytoplasmic immunostain but poor surface antigenic stains. Alcohol is very good for immunohistochemistry since it permits good antibody penetration and does not block immunoreactive determinants. Alcohol precipitates carbohydrates and therefore particularly useful for surface membrane antigens. Proteolytic digestion or antigen retrieval is of no use following alcohol fixation and results in destruction of the tissue section or smear. Acetone is also an excellent preservative of immunoreactive sites, leaving most sites intact, but it is a very poor penetrator. For this reason, it is used only for smears and cryostat sections.

<H1>CONCLUSION - It is generally being realised that genomic data is an inadequate tool when it comes to understanding disease. For understanding disease in its entirety, the understanding of proteins is essential. Large gaps in knowledge remain as far as proteomics is concerned, especially in terms of the structure of proteins, post translational modifications and protein protein interactions. With better understanding of proteins, our understanding of human disease and potential therapeutic interventions is bound to improve.

Box 15.5

Immunohistochemistry uses antibodies as staining agents. It localises target proteins in an organ or a tissue.

Antibodies localise onto epitopes. Epitopes are three dimensional and maybe continuous or discontinuous.

There are two possibilities which may contribute to background staining; the first is that there maybe a non specific antibody binding and the second is that there maybe endogenous enzyme activity.

Detection can be by the direct conjugate labelled antibody method or by the indirect sandwich procedure.

Antigen retrieval is essential for IHC. It can be done using either heat or by enzymatic digestion.

Fixation adversely affects immunohistochemistry. Formalin is the most commonly used fixative and some of the effects have to be reversed before IHC.