Lung tissue stained with the H&E technique. Nuclei are darkly stained in this image.

H&E stain, HE stain or hematoxylin and eosin stain, is a popular staining method in histology. It is the most widely used stain in medical diagnosis; for example when a pathologist looks at a biopsy of a suspected cancer, the histological section is likely to be stained with H&E and termed H&E section, H+E section, or HE section.

The staining method involves application of hemalum, which is a complex formed from aluminium ions and oxidized hematoxylin. This colors nuclei of cells (and a few other objects, such as keratohyalin granules) blue. Materials colored blue by hemalum are often said to be basophilic, but this is an incorrect use of the word. The nuclear staining is folowed by counterstaining with an aqueous or alcoholic solution of eosin Y, which colors eosinophilic other structures in various shades of red, pink and orange.

Haematoxylin Solutions

Haematoxylin stains are commonly employed for histologic studies, often employed to color the nuclei of cells (and a few other objects, such as keratohyalin granules) blue. The mordants used to demonstrate nuclear and cytoplasmic structures are alum and iron, forming lakes or colored complexes (dye-mordant-tissue complexes), the color of which will depend on the salt used. Aluminium salt lakes are usually colored blue white while ferric salt lakes are colored blue-black.

The three main alum haematoxylin solutions employed are Ehrlich’s haematoxylin, Harris’s haematoxylin and Mayer’s haematoxylin. The name haemalum is preferable to “haematoxylin” for these solutions because haematein, a product of oxidation of haematoxylin, is the compound that combines with aluminium ions to form the active dye-metal complex. Alum haematoxylin solutions impart to the nuclei of cells a light transparent red stain which rapidly turns blue on exposure to any neutral or alkaline liquid.

Alum or potassium aluminium sulfate used as the mordant usually dissociates in an alkaline solution, combining with OH^? of water to form insoluble aluminium hydroxide. In the presence of excess acid, aluminium hydroxide cannot be formed thus failure of aluminium haematoxylin dye-lake to form, due to lack of OH^? ions. Hence, acid solutions of alum haematoxylin become red. During staining alum haematoxylin stained sections are usually passed on to a neutral or alkaline solution (e.g. hard tap water or 1% ammonium hydroxide) in order to neutralize the acid and form an insoluble blue aluminium haematin complex. This procedure is known as blueing.

When tap water is not sufficiently alkaline, or is even acid and is unsatisfactory for blueing haematoxylin, a tap water substitute consisting of 3.5 g NaHCO₃ and 20 g MgSO₄.7H₂O in one liter of water with thymol (to inhibit formation of moulds), is used to accelerate blueing of thin paraffin sections. Addition of a trace of any alkali to tap or distilled water also provides an effective blueing solution; a few drops of strong ammonium hydroxide or of saturated aqueous lithium carbonate, added immediately before use, are sufficient for a 400 ml staining dish full of water. Use of very cold water slows down the blueing process, whereas warming accelerates it. In fact, the use of water below 10°C for blueing sections may even produce pink artifact discolorations in the tissue.

The staining of nuclei by hemalum does not require the presence of DNA and is probably due to binding of the dye-metal complex to arginine-rich basic nucleoproteins such as histones. The mechanism is different from that of nuclear staining by basic (cationic) dyes such as thionine or toluidine blue. Staining by basic dyes is prevented by chemical or enzymatic extraction of nucleic acids. Such extractions do not prevent staining of nuclei by hemalum.

Eosin Solutions

Eosin is a fluorescent red dye resulting from the action of bromine on fluorescein. It can be used to stain cytoplasm, collagen and muscle fibers for examination under the microscope. Structures that stain readily with eosin are termed eosinophilic.Eosin is most often used as a counterstain to haematoxylin in H&E (haematoxylin and eosin) staining.  Eosin stains red blood cells intensely red. Eosin is an acidic dye and shows up in the basic parts of the cell, ie the cytoplasm. For staining, eosin Y is typically used in concentrations of 1 to 5 percent weight by volume, dissolved in water or ethanol. For prevention of mold growth in aqueous solutions, thymol is sometimes added. A small concentration (0.5 percent) of acetic acid usually gives a deeper red stain to the tissue.

Other colors, e.g. yellow and brown, can be present in the sample; they are caused by intrinsic pigments, e.g. melanin.

Some structures do not stain well. Basal laminae need to be stained by PAS stain or some silver stains in order to exhibit appropriate contrast. Reticular fibers also require silver stain. Hydrophobic structures also tend to remain clear; these are usually rich in fats, eg. adipocytes, myelin around neuron axons, and Golgi apparatus membranes.

Protocols

There are a large number of H&E protocols available for the histotechnologist. For most tissues, these approaches can be used interchangably, and selection of a particular protocol will be based upon the particular needs of the investigator. Primary differences are dye composition, staining protocol, and intensity of blue dye. Staining contrast for a particular tissue will differ depending upon the approach that is used.

Mayer’s Hematoxylin Protocol

Solutions

Mayer’s Hematoxylin

1. Dissolve 50 g aluminum potassium sulfate (alum) in 1000 ml distilled water.
2. When alum is completely dissolved, add 1 gm hematoxylin.
3. When hematoxylin is completely dissolved, add 0.2 gm sodium iodate and 20 ml acetic acid.
4. Bring solution to boil and cool, and filter

Staining Method

Staining times will vary based upon depth of stain requiredFor slide-mounted immunohistochemistry, counterstain tissue for 30 seconds. For H&E staining, counterstain tissue for 5 minutes.

In order to blue the stain, put slides through 4 changes of tap water, 5 minutes each.

Results

This recipe should create sharp blue nucleus staining with little background.

Harris’ Hematoxylin and Eosin (H&E) Staining Protocol

Solutions and Reagents

Staining Procedure

1. Deparaffinize sections, 2 changes of xylene, 10 minutes each.
2. Re-hydrate in 2 changes of absolute alcohol, 5 minutes each.
3. 95% alcohol for 2 minutes and 70% alcohol for 2 miuntes.
4. Wash briefly in distilled water.
5. Stain in Harris hematoxylin solution for 8 minutes.
6. Wash in running tap water for 5 minutes.
7. Differentiate in 1% acid alcohol for 30 seconds.
8. Wash running tap water for 1 minute.
9. Bluing in 0.2% ammonia water or saturated lithium carbonate solution for 30 seconds to 1 minute.
10. Wash in running tap water for 5 minutes.
11. Rinse in 95% alcohol, 10 dips.
12. Counterstain in eosin-phloxine solution for 30 seconds to 1 minute.
13. Dehydrate through 95% alcohol, 2 changes of absolute alcohol, 5 minutes each.
14. Clear in 2 changes of xylene, 5 minutes each.
15. Mount with xylene based mounting medium.

Results

Nuclei should be blue, cytoplasm pink to red.

References

Kiernan JA (2008) Histological and Histochemical Methods: Theory and Practice. 4th ed. Bloxham, UK: Scion.

Lillie RD, Pizzolato P, Donaldson PT (1976) Nuclear stains with soluble metachrome mordant lake dyes. The effect of chemical endgroup blocking reactions and the artificial introduction of acid groups into tissues. Histochemistry 49: 23-35.

Llewellyn BD (2009) Nuclear staining with alum-hematoxylin. Biotech. Histochem. 84: 159-177.

Puchtler H, Meloan SN, Waldrop FS (1986) Application of current chemical concepts to metal-haematein and -brazilein stains. Histochemistry 85: 353-364.

Yarn drying after being dyed in the early American tradition, at Conner Prairie living history museum.

A mordant is a substance used to set dyes on fabrics or tissue sections by forming a coordination complex with the dye which then attaches to the fabric or tissue.^[1] It may be used for dyeing fabrics, or for intensifying stains in cell or tissue preparations. A mordant is always a polyvalent metal ion.^[2] The resulting coordination complex of dye and ion is colloidal and can be either acidic or alkaline.

Common dye mordants

Mordants include tannic acid, alum, urine, chrome alum, sodium chloride, and certain salts of aluminium, chromium, copper, iron, iodine, potassium, sodium, and tin.

Iodine is often referred to as a mordant in Gram stains but is in fact a trapping agent. ^[3]

Dyeing methods

The three methods used for mordanting are:

• Pre-mordanting (onchrome): The substrate is treated with the mordant and then dyed.
• Meta-mordanting (metachrome): The mordant is added in the dye bath itself.
• Post-mordanting (afterchrome): The dyed material is treated with a mordant.

The type of mordant used changes the shade obtained after dyeing and also affects the fastness property of the dye. The application of mordant, either pre-, meta- or post-mordant methods, is influenced by:

• The action of the mordant on the substrate: if the mordant and dye methods are harsh (e.g. an acidic mordant with an acidic dye), pre- or post- mordanting limits the potential for damage to the substrate.
• The stability of the mordant and/or dye lake: the formation of a stable dye lake means that the mordant can be added in the dye without risk of losing the dye properties (meta-mordanting).

Dye results can also rely on the mordant chosen as the introduction of the mordant into the dye will have a marked effect on the final colour.

The dye lake

The dye lake is formed when the complex of dye and mordant are combined, which then attaches to the substrate.^[2]

The term “lake” is derived from the term lac, the secretions of the Indian wood insect Laccifer lacca (formerly known as the Coccus lacca.^[4] The type of mordant used can change the colour of both the dye-plus-mordant solution and influence the shade of the final product.

Cotton

Since metallic mordants are soluble in water and are loosely held by the cotton fibres, these mordants have to be precipitated on the fabric by converting them into insoluble form, or by first treating the fibres with oil or tannic acid and then impregnating treated fabric with solution of mordant, whereby the metallic mordants are held on to cotton via oil or tannic acid.

Wool

Unlike cotton, wool is highly receptive toward mordants. Due to its amphoteric nature wool can absorb acids and bases equally effectively. When wool is treated with a metallic salt it hydrolyses the salt into an acidic and basic component. The basic component is absorbed at –COOH group and the acidic component is removed during washing. Wool also has a tendency to absorb fine precipitates from solutions; these cling to the surface of fibres and dye particles attached to these contaminants result in poor rubbing fastness.

Silk

Like wool, silk is also amphoteric and can absorb both acids as well as bases. However, wool has thio groups (-SH) from the cystine amino acid, which act as reducing agent and can reduce hexavalent chromium of potassium dichromate to trivalent form. The trivalent chromium forms the complex with the fibre and dye. Therefore potassium dichromate cannot be used as mordant effectively.

Animal and Plant Tissues

In Histology, mordants are indispensable in adhering dyes to tissues for microscopic examination.

Methods for mordant application depend on the desired stain and tissues under study; pre-, meta- and post-mordanting techniques are used as required.

The most commonly used stain used in diagnostic histology of animal tissues is Harris’ haematoxylin as part of a haematoxylin and eosin (H&E) stain.

References

1. International Union of Pure and Applied Chemistry (1993). “mordant”. Compendium of Chemical Terminology Internet edition.
2. ^{a b} Llewellyn, Bryan D. (May, 2005). “Stain Theory – How stains work”. http://stainsfile.info/stainsfile/theory/mordant.htm/. Retrieved 2009-09-20.
3. Llewellyn, Bryan D. (May, 2005). “Stain Theory – Trapping agents”. http://stainsfile.info/StainsFile/theory/trapping.htm/. Retrieved 2009-09-20.
4. Llewellyn, Bryan D. (May, 2005). “Stain Theory – Lac”. http://stainsfile.info/StainsFile/dyes/75450.htm/. Retrieved 2009-09-20

In biochemistry, PMSF (phenylmethanesulfonylfluoride or phenylmethylsulfonyl fluoride) is a serine protease inhibitor commonly used in the preparation of cell lysates. PMSF does not inhibit all serine proteases. It is rapidly degraded in water and stock solutions are usually made up in anhydrous ethanol, isopropanol, corn oil, or DMSO. Proteolytic inhibition occurs when a concentration between 0.1 – 1 mM PMSF is used. The half-life is short in aqueous solutions (110 min at pH=7 and 35 min at pH=8).

PMSF binds specifically to the active site serine residue in a serine protease, but does not bind to any other serine residues in the protein. Since PMSF binds covalently to the enzyme at the active serine residue, the complex can be viewed by X-ray crystallography; it can therefore be used as a chemical label to identify an essential active site SER in an enzyme.

Enzyme(active)Ser-O-H + F-SO₂CH₂C₆H₅ ? EnzymeSer-O-SO₂CH₂C₆H₅ + HF

Serine protease + PMSF ? Irreversible enzyme-PMS complex + HF

PMSF is a cytotoxic chemical that should only be handled inside a fume hood; the LD₅₀ for this compound is less than 500 mg/kg.

Preparation of PMSF (10 mM), 10 ml

1. Weigh out  17.4g PMSF
2. Add isopropanol to 10ml and dissolve.
3. No need to filter sterilize.
4. Aliquot and store at –20°C.

PMSF is extremely unstable in aqueous solutions with a half-life of approximately 30 minutes. Add solution immediately before use.

Recommended Suppliers

PMSF: Sigma (P-7626)

The three dimensional structure of ampicillin.

Ampicillin is a beta-lactam antibiotic that has been used extensively to treat bacterial infections since 1961. It is often used as a selective agent in molecular biology to select for and to confirm the uptake of genes (e.g., of plasmids) by bacteria (e.g., E. coli). A gene that is to be inserted into a bacterium is coupled to a gene coding for an ampicillin resistance (in E. coli, usually the bla (TEM-1) gene, coding for ?-lactamase). The treated bacteria are then grown in a medium containing ampicillin (typically 50–100 mg/L). Only the bacteria that successfully take up the desired genes become ampicillin resistant, and therefore contain the other desired gene as well. It can be used with Cloaxicillin as well. As a powder ampicillin is white with slight yellow cast and is soluble in water (150 mg/ml).

Preparation  of Ampicillin (100mg/ml), 50ml

1. Weight out 5g Ampicillin
2. Add double distilled H2O to 50ml.
3. Sterilize using a 0.22 µm filter.
4. Aliquot and store at –20°C. (Use at 100µg/ml).

Recommended Suppliers

Ampicillin: Sigma-Aldrich, A-9518

30% acrylamide solution is used for the creation of polyacrylamide gels in gel electrophoresis techniques, such as western blotting. Acrylamide needs to be handled using best laboratory practice (such as wearing appropriate gloves, lab coat etc. and having safe systems of work) to avoid poisonous exposure since it is a neurotoxin.

Preparation of acrylamide solution (30%), 500 ml

1. Weigh out 29 grams 2X Acrylamide.
2. Weigh out 1 gram of N,N’-methylenebisacrylamide.
3. Add acrylamide and N,N’-methylenebisacrylamide to 300ml double-distilled H2O.
4. Heat to 37°C to dissolve chemicals.
5. Adjust the final volume to 500ml with double-distilled H2O.

Recommended suppliers

Acrylamide: Serva, 10675

N,N’-methylenebisacrylamide: Serva, 29195

Two-dimensional gel electrophoresis, abbreviated as 2-DE or 2-D electrophoresis, is a form of gel electrophoresis commonly used to analyze proteins. Mixtures of proteins are separated by two properties in two dimensions on 2D gels.

Basis for separation

2-D electrophoresis begins with 1-D electrophoresis but then separates the molecules by a second property in a direction 90 degrees from the first. In 1-D electrophoresis, proteins (or other molecules) are separated in one dimension, so that all the proteins/molecules will lie along a lane but be separated from each other by a property (e.g. isoelectric point). The result is that the molecules are spread out across a 2-D gel. Because it is unlikely that two molecules will be similar in two distinct properties, molecules are more effectively separated in 2-D electrophoresis than in 1-D electrophoresis.

The two dimensions that proteins are separated into using this technique can be isoelectric point, protein complex mass in the native state, and protein mass.

To separate the proteins by isoelectric point is called isoelectric focusing (IEF). Thereby, a gradient of pH is applied to a gel and an electric potential is applied across the gel, making one end more positive than the other. At all pHs other than their isoelectric point, proteins will be charged. If they are positively charged, they will be pulled towards the more negative end of the gel and if they are negatively charged they will be pulled to the more positive end of the gel. The proteins applied in the first dimension will move along the gel and will accumulate at their isoelectric point; that is, the point at which the overall charge on the protein is 0 (a neutral charge).

For the analysis of the functioning of proteins in a cell, the knowledge of their cooperation is essential. Most often proteins act together in complexes to be fully functional. The analysis of this sub organelle organisation of the cell requires techniques conserving the native state of the protein complexes. In native polyacrylamide gel electrophoresis (native PAGE), proteins remain in their native state and are separated in the electric field following their mass and the mass of their complexes respectively. To obtain a separation by size and not by net charge, as in IEF, an additional charge is transferred to the proteins by the use of coomassie or lithium dodecyl sulfate (LDS). After completion of the first dimension the complexes are destroyed by applying the denaturing SDS-PAGE in the second dimension, where the proteins of which the complexes are composed of are separated by their mass.

Before separating the proteins by mass, they are treated with sodium dodecyl sulfate (SDS) along with other reagents (SDS-PAGE in 1-D). This denatures the proteins (that is, it unfolds them into long, straight molecules) and binds a number of SDS molecules roughly proportional to the protein’s length. Because a protein’s length (when unfolded) is roughly proportional to its mass, this is equivalent to saying that it attaches a number of SDS molecules roughly proportional to the protein’s mass. Since the SDS molecules are negatively charged, the result of this is that all of the proteins will have approximately the same mass-to-charge ratio as each other. In addition, proteins will not migrate when they have no charge (a result of the isoelectric focusing step) therefore the coating of the protein in SDS (negatively charged) allows migration of the proteins in the second dimension (NB SDS is not compatible for use in the first dimension as it is charged and a nonionic or zwitterionic detergent needs to be used). In the second dimension, an electric potential is again applied, but at a 90 degree angle from the first field. The proteins will be attracted to the more positive side of the gel proportionally to their mass-to-charge ratio. As previously explained, this ratio will be nearly the same for all proteins. The proteins’ progress will be slowed by frictional forces. The gel therefore acts like a molecular sieve when the current is applied, separating the proteins on the basis of their molecular weight with larger proteins being retained higher in the gel and smaller proteins being able to pass through the sieve and reach lower regions of the gel.

The result of this is a gel with proteins spread out on its surface. These proteins can then be detected by a variety of means, but the most commonly used stains are silver and coomassie staining. In this case, a silver colloid is applied to the gel. The silver binds to cysteine groups within the protein. The silver is darkened by exposure to ultra-violet light. The darkness of the silver can be related to the amount of silver and therefore the amount of protein at a given location on the gel. This measurement can only give approximate amounts, but is adequate for most purposes.

Molecules other than proteins can be separated by 2D electrophoresis. In supercoiling assays, coiled DNA is separated in the first dimension and denatured by a DNA intercalator (such as ethidium bromide or the less carcinogenic chloroquine) in the second. This is comparable to the combination of native PAGE /SDS-PAGE in protein separation.

In summary 2D provides resolution according to two traits, whereof one is most often molecular charge. The investigated molecule needs not be protein.

2D Gel Analysis Software

Warping: Images of two 2D electrophoresis gels, overlaid with Delta2D. First image is colored in orange, second one colored in blue. Due to running differences, corresponding spots do not overlap.

Warping: Images of two 2D electrophoresis gels after warping. First image is colored in orange, second one colored in blue. Corresponding spots overlap after warping. Common spots are colored black, orange spots are only present (or much stronger) on the first image, blue spots are only present (or much stronger) on the second image.

In quantitative proteomics, these tools primarily analyze bio-markers by quantifying individual proteins, and showing the separation between one or more protein “spots” on a scanned image of a 2-DE gel. Additionally, these tools match spots between gels of similar samples to show, for example, proteomic differences between early and advanced stages of an illness. Software packages include Delta2D, ImageMaster, Melanie, PDQuest, Progenesis and REDFIN – among others. While this technology is widely utilized, the intelligence has not been perfected. For example, while PDQuest and Progenesis tend to agree on the quantification and analysis of well-defined well-separated protein spots, they deliver different results and analysis tendencies with less-defined less-separated spots.^[1]

Challenges for automatic software-based analysis include:

• incompletely separated (overlapping) spots (less-defined and/or separated)
• weak spots / noise (e.g., “ghost spots”)
• running differences between gels (e.g., protein migrates to different positions on different gels)
• unmatched/undetected spots, leading to missing values^[2]
• mismatched spots
• errors in quantification (several distinct spots may be erroneously detected as a single spot by the software and/or parts of a spot may be excluded from quantification)
• differences in software algorithms and therefore analysis tendencies

Generated picking lists can be used for the automated in-gel digestion of protein spots, and subsequent identification of the proteins by mass spectrometry.

For an overview of the current approach for software analysis of 2DE gel images see ^[3] or ^[4].

External links

• A 2-D electrophoresis tutorial on the web site of the Parasitology Group at Aberystwyth University
• JVirGel Create virtual 2-D Gels from sequence data.
• fixingproteomics.org Protocols for preparing samples and running 2-D Gels.nnn
• Gel IQ A freely downloadable software tool for assessing the quality of 2D gel image analysis data.

References

1. Arora PS, Yamagiwa H, Srivastava A, Bolander ME, Sarkar G (2005). “Comparative evaluation of two two-dimensional gel electrophoresis image analysis software applications using synovial fluids from patients with joint disease”. J Orthop Sci 10 (2): 160–6. doi:10.1007/s00776-004-0878-0. PMID 15815863. http://www.springerlink.com/openurl.asp?genre=article&doi=10.1007/s00776-004-0878-0.
2. Pedreschi R, Hertog ML, Carpentier SC, et al (April 2008). “Treatment of missing values for multivariate statistical analysis of gel-based proteomics data”. Proteomics 8 (7): 1371–83. doi:10.1002/pmic.200700975. PMID 18383008.
3. Berth M, Moser FM, Kolbe M, Bernhardt J (October 2007). “The state of the art in the analysis of two-dimensional gel electrophoresis images”. Appl. Microbiol. Biotechnol. 76 (6): 1223–43. doi:10.1007/s00253-007-1128-0. PMID 17713763.
4. Bandow JE, Baker JD, Berth M, et al (August 2008). “Improved image analysis workflow for 2-D gels enables large-scale 2-D gel-based proteomics studies–COPD biomarker discovery study”. Proteomics 8 (15): 3030–41. doi:10.1002/pmic.200701184. PMID 18618493. http://www3.interscience.wiley.com/journal/120749796.

Introduction

The western blot (alternatively, protein immunoblot) is a semi-quantitative analytical technique used to detect specific proteins in a given sample of tissue homogenate or extract, often used to determine the effect of an experimental treatment on protein expression in cells or tissue. The approach 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.^[2][3]

There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against tens of thousands of different proteins^[4]. 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^[5] 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.

Western blotting is used in a wide variety of medical diagnostic applications such,

• 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.

Steps involved 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.

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

Gel electrophoresis

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

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

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

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

Transfer

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

The uniformity and overall effectiveness of transfer of protein from the gel to the membrane can be checked by staining the membrane with Coomassie 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.^[6] Coomassie dyes are generally incompatible with western b

Blocking

Since the membrane has been chosen for its ability to bind protein, and 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 (since the antibody is a protein itself). Blocking of non-specific binding is achieved by placing the membrane in a dilute solution of protein – typically Bovine serum albumin (BSA) or non-fat dry milk (both are inexpensive), with a minute percentage of detergent such as Tween 20. 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 step

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

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

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

Another method of secondary antibody detection utilizes a near-infrared (NIR) fluorophore-linked antibody. Light produced from the excitation of a fluorescent dye is static, making fluorescent detection a more precise and accurate measure of the difference in signal produced by labeled antibodies bound to proteins on a Western blot. Proteins can be accurately quantified because the signal generated by the different amounts of proteins on the membranes is measured in a static state, as compared to chemiluminescence, in which light is measured in a dynamic state. ^[7]

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 with a radioactive isotope of iodine. Since other methods are safer, quicker and cheaper this method is now rarely used.

One step

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

Western blot using radioactive detection system

Detection Methods

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, also refered to as a loading control (follow link for examples of appropriate loading controls) that are not expected to change between samples. The amount of target protein is indexed to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.

Colorimetric detection

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

Chemiluminescent detection

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

Radioactive detection

Radioactive labels do not require enzyme substrates, but rather allow the placement of medical X-ray film directly against the western blot which develops as it is exposed to the label and creates dark regions which correspond to the protein bands of interest (see image to the right). The importance of radioactive detections methods is declining^{[citation needed]}, because it is very expensive, health and safety risks are high and ECL 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, and we refer you to the expanded article for examples of the application of 2-D SDS PAGE. 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.

Medical diagnostic applications

References

1. Hempelmann E, Schirmer RH, Fritsch G, Hundt E, Gröschel-Stewart U (1987). “Studies on glutathione reductase and methemoglobin from human erythrocytes parasitized with Plasmodium falciparum.”. Mol Biochem Parasitol. 23 (1): 19–24. doi:10.1016/0166-6851(87)90182-4. PMID 3553936.
2. Towbin H, Staehelin T, Gordon J. (1979). “Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications.”. Proc Natl Acad Sci U S A. 76 (9): 4350–4354. doi:10.1073/pnas.76.9.4350. PMID 388439.
3. Renart J, Reiser J, Stark GR (1979). “Transfer of proteins from gels to diazobenzyloxymethyl-paper and detection with antisera: a method for studying antibody specificity and antigen structure.”. Proc Natl Acad Sci U S A. 76 (7): 3116–3120. doi:10.1073/pnas.76.7.3116. PMID 91164.
4. “Western blot antibody”. exactantigen.com. http://www.exactantigen.com/review/western-blot-antibody.html. Retrieved 2009-01-29.
5. W. Neil Burnette (April 1981). “‘Western blotting’: electrophoretic transfer of proteins from sodium dodecyl sulfate — polyacrylamide gels to unmodified nitrocellulose and radiographic detection with antibody and radioiodinated protein A“. Analytical Biochemistry (United States: Academic Press) 112 (2): 195–203. doi:10.1016/0003-2697(81)90281-5. ISSN 0003-2697. PMID 6266278. . Retrieved 2008-04-03.
6. Source.

Good’s buffers are twelve buffering agents selected and described by Norman Good and colleagues in 1966. Good selected the buffers based on a number of criteria which make them candidates for use in biochemistry and biological research. Many remain staples in modern biology laboratories.

Selection criteria

Good sought to identify buffering compounds which met several criteria likely to be of value in biological research.

1. pK_a. Because most biological reactions take place at near-neutral pH between 6 and 8, ideal buffers would have pK_a values in this region to provide maximum buffering capacity there.
2. Solubility. For ease in handling and because biological systems are in aqueous systems, good solubility in water was required. Low solubility in nonpolar solvents (fats, oils, and organic solvents) was also considered beneficial, as this would tend to prevent the buffer compound from accumulating in nonpolar compartments in biological systems: cell membranes and other cell compartments.
3. Membrane impermeability. Ideally, a buffer will not readily pass through cell membranes, this will also reduce the accumulation of buffer compound within cells.
4. Minimal salt effects. Highly ionic buffers may cause problems or complications in some biological systems.
5. Well-behaved cation interactions. If the buffers form complexes with cationic ligands, the complexes formed should remain soluble. Ideally, at least some of the buffering compounds will not form complexes.
6. Stability. The buffers should be chemically stable, resisting enzymatic and non-enzymatic degradation.
7. Optical absorbance. Buffers should not absorb visible or ultraviolet light at wavelengths longer than 230 nm so as not to interfere with commonly-used spectrophotometric assays.
8. Ease of preparation. Buffers should be easily prepared and purified from inexpensive materials.

The twelve buffers selected by Good are tabulated below.

Description

A semi-permanent mounting media for immunofluorescence microscopy.

Method

1. Add the following reagents to a 250 ml flask or beaker:

• 24 g analytical grade glycerol (Sigma #G-6279)
• 9.6 g Mowiol 4-88 (Fluka, #81381, can be purchased through Sigma-Aldrich)
• 24 ml distilled water
• 48 ml 0.2M Tris buffer, pH 8.5

2. Stir with a clean stir bar on a hot plate on warm (not boiling)at least 4-5 hours until the majority of the Mowiol powder goes into solution.
3. Aliquot into 50 ml centrifuge tubes, weigh and balance
4. Centrifuge at 5000g for 15 minutes. Carefully remove the supernatant without disturbing the pellet at the bottom of the flask.
5. Aliquot into 15 ml conical tubes – add only 10 mls to each tube to allow for expansion with freezing.
6. Aliquots may be stored at -20C for 12 months. Store at room temperature no more than one month.
7. To use, warm solution to room temperature to eliminate bubble formation. use approximately 10 ul of mounting media for an 18 mm coverslip.
8. Allow slides to dry overnight at room temperature in a light-tight box.

Description

Weil’s stain is a modification for paraffin sections of the Weigert-Pal-Kulschitsky technique. The underlying principle of these methods involves the reduction of chrome salt to chromium dioxide by myelin. The chromium subsequently acts as a mordant for the haematoxylin, intensifying the stain.

Procedure

This procedure is generally conducted on sections from formalin-fixed, paraffin-embedded tissue that are cut between 8-15 µm. Spinal cord tissue is rich in myelinated axons and can be used as a positive control.

1. Dewax and hydrate sections to distilled water.
2. Put slides in freshly prepared Staining Solution at 56-60C for 30 minutes.
3. Wash slides well in water.
4. Partially differentiate in iron alum differentiating solution until myelin sheaths stand out blueish-black on a pale grey background, approximately 5 minutes. If you are unsure, check your sections under the microscope at1 minute intervals).
5. Wash slides in tap water for 10 minutes.
6. Complete differentiation in Weigert’s differentiator, 1 to 2 minutes. Control this differentiation step carefully checking under the microscope, until the myelin is an intense deep blue  against a creamy or clear background.
7. Wash well in tap water.
8. Dehydrate through a series of graded ethanol baths, clear in xylene, and mount.

RESULTS

Myelin-containing structures will be stained black, red blood cells will be black, nuclei will be blue, and the background should be clear or yellow.

Solutions

Haematoxylin Solution, working strength

• 10g Haematoxylin
• 100 ml absolute ethanol.

Allow solution to “ripen” naturally for six (6) weeks, forming the basis of a stock solution. Just prior to staining, create a working strength solution by diluting the stock 1:4 with distilled water

Iron Alum Solution

• 4% (w/v) aqueous ferric ammonium sulphate.

Staining Solution

Mix equal volumes of preheated (56-60C) working strength Haematoxylin and Iron Alum solutions just prior to use.

Weigert’s Differentiator, 200 ml

• Borax (sodium tetraborate), 2g
• Potassium ferricyanide, 2.5g
• Distilled water, 200ml