Coomassie-stained Polyacrylamide gel

Coomassie dyes (also known as Coomassie Brilliant Dyes) are a family of dyes commonly used to stain proteins in sodium dodecyl sulfate and blue native polyacrylamide gel electrophoresis (SDS-PAGE and BN-PAGE, respectively) gels. The gels are soaked in dye and excess stain is then eluted with a solvent (“destaining”). This treatment allows the visualization of protein bands. The gel usually contains a set of molecular weight marker (proteins of pre-determined weight) so that protein molecular weight can be estimated in an unknown solution during the visualization.

Alternatively, Coomassie may be added to undenatured protein in PAGE in place of SDS, in a technique called BN-PAGE; both SDS and CBB have the effect of imparting a net negative charge upon the proteins. Without the Coomassie, the technique is known as CN-PAGE (colorless native), and will only separate negatively-charged proteins.

Variations

The original Coomassie dye was developed as a wool dye and named to commemorate the 1896 British occupation of Coommassie (now Kumasi) in Ghana. The first of the Coomassie series was Coommassie Blue R-250 (“R” standing for “reddish” and “250″ being the dye strength indicator). Coommassie Blue G-250 (“G” for “greenish”) and Coomassie Violet R-150 later followed. The most commonly used dyes in the laboratory for staining PAGE gels are Coomassie Blue R-250 and G-250. Although G-250 is more sensitive, R-250 affords better resolution, and is often used instead.^[1]

Medical Uses

Brilliant Blue G (BBG) has recently been used in scientific experiments to treat spinal injuries in laboratory rats.^[2] It acts by reducing the body’s natural swelling response, which can cause neurons in the area to die of metabolic stress. Testing is still in progress to determine if this treatment can be used effectively in humans. The recent tests have administered the dye within 15 minutes of injury, but to be effective in a real-life setting, where it may take time for a patient to reach the emergency room, the treatment should be effective even when administered up to two hours after injury. The only reported side effect was that the rats turned blue.^[3][4]

Coomassie Brilliant Blue G-250

Laboratory usage

The Coomassie dyes (R-250 and G-250) are used for quantification of protein, and work by binding  to proteins through Van der Waals attractions and through ionic interactions between dye sulfonic acid groups and positive protein amine groups . Coomassie R-250, is the most commonly used variant for detection of protein, allowing for detection of as little as 0.1 ug protein. The coomassie G-250 dyes is less sensitive, with a lower limit of around 0.5 µg for most proteins, but this is somewhat offset by the speed of destaining offered by this dye compared to R-250. Although R-250 is typically color invariant, Coomassie G-250 undergoes a color shift  where the dye changes from dark blue black at pH 7 to a light tan  below pH 2. The leuco form recovers its blue color upon binding to protein, apparently due to a neutral pH environment surrounding the the protein molecule. Consequently, a gel placed in an acidified solution of Coomassie G-250 will manifest blue protein bands on a light amber background. The bands develop rapidly and there is no need to destain, for the background color is so light as to be essentially clear. Consequently, the loss in sensitivity of G-250 can offset by the speed and convenience of the protocol, which saves up to 11 hours versus the most sensitive R-250 procedures.

In addition to their use in gels, Coomassie dyes are an integral component of the Bradford Method for determining protein concentration in a solution.  When Coomassie Brilliant Blue G-250 binds to proteins in acid solution, it has an absorbance shift from 465 nm to 595 nm. The absorbance data can then be used in Beer’s law to determine protein concentration and ultimately the actual amount of protein in a given solution.

Protocols for Staining Gels with Coomassie Blue R-250 or Coomassie Blue G-250

RAPID PROTOCOL – COOMASSIE BLUE R-250

RAPID PROTOCOL – COOMASSIE BLUE G-250

References

1. Merril CR (1990) “Gel-staining techniques” In Deutscher MP (Ed) Guide to protein purification. Methods in Enzymology volume 182. Academic press Inc.
2. Peng, Weiguo; Maria L. Cotrina, Xiaoning Han, Hongmei Yu, Lane Bekar, Livnat Blum, Takahiro Takano, Guo-Feng Tian, Steven A. Goldman, Maiken Nedergaard (2009-07-28). “Systemic administration of an antagonist of the ATP-sensitive receptor P2X7 improves recovery after spinal cord injury”. Proceedings of the National Academy of Sciences 106 (30): 12489–12493. doi:10.1073/pnas.0902531106. http://www.pnas.org/content/106/30/12489.abstract. Retrieved 2009-08-02.
3. Published: 7:00AM BST 28 Jul 2009 (2009-07-28). “Blue M&Ms ‘mend spinal injuries’”. Telegraph. http://www.telegraph.co.uk/science/science-news/5921266/Blue-MandMs-mend-spinal-injuries.html. Retrieved 2010-01-19.
4. “Blue Food Dye Treats Spine Injury in Rats | Wired Science”. Wired.com. 2009-07-27. http://www.wired.com/wiredscience/2009/07/bluerats/. Retrieved 2010-01-19.

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