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.

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

A pipette (also called a pipet, pipettor or chemical dropper) is a laboratory instrument used to transport a measured volume of liquid.

Use and variations

Pipettes are commonly used in chemistry and molecular biology research as well as medical tests. Pipettes come in several designs for various purposes with differing levels of accuracy and precision, from single piece glass pipettes to more complex adjustable or electronic pipettes. A pipette works by creating a vacuum above the liquid-holding chamber and selectively releasing this vacuum to draw up and dispense liquid.

Pipettes that dispense between 1 and 1000 ?l are termed micropipettes, while macropipettes dispense a greater volume of liquid.

Glass pipettes

The original pipette is made of glass. It is more commonly used in chemistry, with aqueous solutions. There are two types. One type, the volumetric pipette, has a large bulb, and is calibrated for a single volume. Typical volumes are 10, 25, and 50 mL. Alternatively, Mohr pipettes are straight-walled, and graduated for different volumes such as 5 mL in 0.5 mL increments. The single volume pipette is usually more accurate, with an error of ± 0.1 or 0.2 mL.

The pipette is filled by dipping the tip in the volume to be measured, and drawing up the liquid with a pipette filler past the inscribed mark. The volume is then set by releasing the vacuum using the pipette filler or a damp finger. While moving the pipette to the receiving vessel, care must be taken not to shake the pipette because the column of fluid may “bounce”.

Piston-driven air displacement pipettes

Biohit Pipette

These are the most accurate and precise pipettes. They are more commonly used in biology, though they are commonly used by chemists as well. The plastic pipette tips are designed for aqueous solutions, and are not recommended for use with organic solvents which may dissolve the plastic.

These pipettes operate by piston-driven air displacement. A vacuum is generated by the vertical travel of a metal or ceramic piston within an airtight sleeve. As the piston moves upward, driven by the depression of the plunger, a vacuum is created in the space left vacant by the piston. Air from the tip rises to fill the space left vacant, and the tip air is then replaced by the liquid, which is drawn up into the tip and thus available for transport and dispensing elsewhere.

Sterile technique prevents liquid from coming into contact with the pipette. Instead, the liquid is drawn into and dispensed from a disposable pipette tip which is changed between transfers. Depressing the tip ejector button removes the tip, which is cast off without being handled by the operator and disposed of safely in an appropriate container.

The plunger is depressed to both draw up and dispense the liquid. Normal operation consists of depressing the plunger button to the first stop while the pipette is held in the air. The tip is then submerged in the liquid to be transported and the plunger is released in a slow and even manner. This draws the liquid up into the tip. The instrument is then moved to the desired dispensing location. The plunger is again depressed to the first stop, and then to the second stop, or ‘blowout’, position. This action will fully evacuate the tip and dispense the liquid. In an adjustable pipette, the volume of liquid contained in the tip is variable; it can be changed via a dial or other mechanism, depending on the model. Some pipettes include a small window which displays the currently selected volume.

Certain considerations should be observed to ensure maximum accuracy and repeatability:

• Operator consistency is paramount to repeatable operation. The necessity of operator practice and development of good pipetting practices and habits is absolute. Light guided pipetting aides are used to help reduce errors and speed up liquid handling protocols.
• When drawing up liquid the tip should be dipped 3 to 5 mm below the surface of the liquid, always at a 90 degree angle.
• When dispensing the pipette should be held at a 45 degree angle, and the tip placed against the side of the receiving vessel. Glass vessels are preferred; the surface tension of the glass provides additional torsion that results in complete evacuation of the tip.
• The tip must never be wiped off or blotted in any way, even from the exterior, while liquid is in the tip. These actions tend to attract and thus bleed off some of the liquid, resulting in decreased accuracy and repeatability.
• A dry tip should always be pre-wetted by drawing up and dispensing the chosen volume a minimum of three times. This action reduces the surface tension on the inside walls of the tip and also provides the proper level of inter-tip humidity, which reduces evaporation of the sample liquid.
• Most pipettes are calibrated “to deliver” (TD) and not “to contain” (TC). If they are TD pipettes they should not be rinsed after they have delivered their contents. If the pipette were calibrated TC it should be rinsed to obtain the correct amount of material. If the fluid to be measured is quite viscous or sticky (such as glycerol solutions) the pipette must be calibrated and in this case the outside of the tip must be carefully wiped with a lint free tissue to remove the adhering liquid – while being careful to not touch the opening of the pipette tip, which may require some practice. Accuracy in delivering liquids with high or low viscosity may require a “positive displacement” pipettor, which is quite distinct from an air displacement pipettor.
• For maximum accuracy, and especially necessary when calibrating the pipette, relative humidity in the ambient environment should be maintained between 50% and 75%, and in no case should the humidity be allowed to dip below 50%. This limits the rate of sample evaporation which can cause significant errors, especially at lower volumes.

The importance of operator skill cannot be overstated. A high-quality, well-calibrated pipette in the hand of an uninterested or untrained operator is an unreliable instrument. Additionally, there are four factors that can reduce the accuracy and repeatability of even highly-skilled operators, and these factors must be counteracted if optimal accuracy is to be achieved:

• Heat from the operator’s hand is absorbed through the handle of the instrument and transferred to the metallic components inside. If the pipette is operated continuously for a prolonged period of time this heat buildup becomes significant, causing the internal components to expand and changing the interplay between components. This reduces the consistency, accuracy, and repeatability of the instrument. The volume dispensed is dependent on the sizes of the piston and the springs that cause its travel. As these change in size the volume dispensed changes also. This effect is more pronounced in low-volume instruments. Additionally, the expansion of a metallic component that interacts with a non-metallic one that does not expand as readily in the presence of heat may cause the instrument to seem to stick, hang up, or react more slowly. Pipettes with thin handles are particularly susceptible to this phenomenon. Plumper handles are both more ergonomic and less likely to suffer from heat transfer problems. The best technique for maximum accuracy is to employ multiple pipettes and rotate them often, storing them between uses in a stand that holds them vertically.

• Operator fatigue is an often-overlooked but crucial component when seeking maximal accuracy and repeatability. Human beings are not robots, and repetitive motions cause stress in human joints and muscles. Even a well-trained and experienced operator will see a decrease in accuracy and repeatability as length of time on the job increases. It is for this reason that pipette calibration service providers that are dedicated to excellence limit the number of pipettes that can be calibrated by an individual technician to a maximum daily number. Each pipette, and each customer, deserves a high level of care in the treatment of the instrument. Additionally, some dedicated professionals train themselves to pipette ambidextrously, allowing them to reduce arm and finger strain by alternating hands. Another solution is choosing an electronic pipettor which significantly reduces hand fatigue. Once the operating button is touched the pipettor operates always the same way producing user independent accuracy and precision.

• Long-term pipette operation can lead to repetitive strain injuries (RSI), such as carpal tunnel syndrome. These disorders may cause significant reductions in accuracy and repeatability by altering the proper pipetting techniques that are crucial to achieving optimal accuracy. Preventive measures include learning to pipette with both hands and alternating their usage, taking frequent breaks while pipetting, and choosing the most ergonomic pipette available. Instruments with plumper handles are generally superior in this regard. On the other hand, electronic pipettors which operate with a light touch reduce RSI significantly.

• Letting the pipette “rest” for at least one minute after a volume change is made. This does not apply to single-volume instruments, also called set volume or fixed volume pipettes. A change in the dispensed volume of an adjustable pipette involves modifying the internal tensioning of a spring that governs the piston’s travel distance. Springs subjected to changing tensioning behave more smoothly and consistently when they are allowed to enjoy an interval of rest to settle into their new configuration. A pipette that is left idle for at least one minute after a volume adjustment will perform more accurately than one that is pressed into service prematurely. This is especially important when calibrating a pipette.

Calibration

For sustained accuracy and consistent and repeatable operation, pipettes should be calibrated at periodic intervals. These intervals vary depending on several factors:

• The skill and training of the operators. Skilled operators tend to operate the instrument more correctly and make fewer accuracy-robbing mistakes.
• The liquid dispensed by the pipette. Corrosive and volatile liquids tend to emit vapors which ascend into the pipette shaft even under proper operating conditions and may corrode the metal piston and springs, or the seals and o-rings that provide an air-tight seal between the piston and the surrounding sleeve.
• Proper and careful handling. Pipettes that are frequently dropped, are subjected to careless handling or horseplay, or that are not properly stored in a vertical position, will tend to degrade in accuracy over time.
• The accuracy required by the instrument. Applications requiring maximum accuracy also demand more frequent calibration. Instruments used for purely research applications or in educational settings generally require less frequent calibration.

Under average conditions, most pipettes can be calibrated semi-annually (every six months) and provide satisfactory performance. Institutions that are regulated by the Food and Drug Administration‘s GMP/GLP regulations generally benefit from quarterly calibration, or every three months. Critical applications may require monthly service, while research and educational institutions may need only annual service. These are general guidelines and any decision on the appropriate calibration interval should be made carefully and include considerations of the pipette in question (some are more reliable than others), the conditions under which the pipette is used, and the operators who use it.

Calibration is generally accomplished through means of gravimetric analysis. This entails dispensing samples of distilled water into a receiving vessel perched atop a precision analytical balance. The density of water is a well-known constant, and thus the mass of the dispensed sample provides an accurate indication of the volume dispensed. Relative humidity, ambient temperature, and barometric pressure are factors in the accuracy of the measurement, and are usually combined in a complex formula and computed as the Z-factor. This Z-factor is then used to modify the raw mass data output of the balance and provide an adjusted and more accurate measurement.

The colormetric method uses precise concentrations of colored water to affect the measurement and determine the volume dispensed. A spectrophotomer is used to measure the color difference before and after aspiration of the sample, providing a very accurate reading. This method is more expensive than the more common gravimetric method, given the cost of the colored reagents, and is recommended when optimal accuracy is required. It is also recommended for extremely low-volume pipette calibration, in the 2 microliter range, because the inherent uncertainties of the gravimetic method, performed with standard laboratory balances, becomes excessive. Properly calibrated microbalances, capable of reading in the range of micrograms (10^-6 g) can also be used effectively for gravimetric analysis of low-volume micropipettes.

Other pipette types

• Pasteur pipettes, also known as droppers are used to transfer small amounts of liquids, but are not graduated. Pasteur pipettes are made of plastic or glass.
• Transfer pipettes, are similar to Pasteur pipettes. However, they are made exclusively from plastic and their bulb can serve as the liquid-holding chamber.
• Serological pipettes are measuring pipettes that have graduations extending all the way to the tip.
• Mohr pipettes are measuring pipettes that resemble serological pipettes, with the primary difference that the graduations do not extend all the way to the tip.
• Dispensable pipettes are often made of plastic and intended to be used to administer medicine into the eye or ear of a patient (see image).

Pipette accessories

• Pipette fillers are used to fill the pipette easily, avoiding the need for mouth pipetting.
• Pipette helpers are battery-operated and are designed to be used with disposable pipette tubes. These pipettes cannot be calibrated and their accuracy is determined by that of the printed graduations on the disposable tubes.
• Light-guided pipetting systems are pipetting accessories which are computer based. They utilize flat screen LCD monitors or LED arrays to light up source and destination wells in microplates or vials for accurate well to well pipetting. Some of these systems use text to speech to alert the operator during plate or volume changes when pipetting lab protocols.
• Pipette tips. The pipettors and injection molded plastic disposable tips form together a reliable pipetting system. It is recommended to use original manufacturers tips to guarantee the precision and accuracy of the pipettes. The precision-made pipettor tips provide excellent reproducibility and accuracy. Pipettor tips are available in autoclavable boxes, refills and bulk packaging. Non-sterile, pre-sterilized and filtered tips are usually available in single trays as RNase, DNase and endotoxin certified free.

The smallest pipette

A zeptoliter pipette has been developed at Brookhaven National Laboratory. The pipette is made of a carbon shell, within which is an alloy of gold-germanium alloy. The pipette was used to learn about how crystallization takes place.^[1]

References

Description

In molecular biology, c-Fos is a cellular proto-oncogene belonging to the immediate early gene family of transcription factors. c-Fos has a leucine-zipper DNA binding domain, and a transactivation domain at the C-terminus. Transcription of c-Fos is upregulated in response to many extracellular signals, e.g. growth factors. Additionally, phosphorylation by MAPK, PKA, PKC or cdc2 alters the activity and stability of c-Fos. Members of the Fos family dimerise with Jun to form the AP-1 transcription factor, which upregulates transcription of a diverse range of genes involved in everything from proliferation and differentiation to defense against invasion and cell damage.

The AP-1 complex has been implicated in transformation and progression of cancer, and both Fos and Jun were first discovered in rat fibroblasts.

The viral homologue of c-Fos, v-Fos, is found in the retrovirus Finkel-Biskis-Jinkins murine osteogenic sarcoma virus. In neuroscience research, neuroscientists measure expression of c-fos as an indirect marker of neuronal activity because c-fos is often expressed when neurons fire action potentials.

Staining procedure

1. This is a free-floating staining procedure for formalin-fixed brain tissue. Sections should be cut between 15-30 µm.
2. Transfer sections in 6-well plates loaded with PBS 0.1 M (one brain per well).
3. Rinse sections twice, 10 minutes each rinse, with PBS 0.1 M on a shaker.
4. Incubate sections with fresh 0.3% H₂O₂ in PBS 0.1 M for 30 minutes at room temperature on a shaker.
5. Rinse sections 3 x 10 minutes with PBS 0.1 M on a shaker.
6. Incubate sections with blocking solution  for 60 min at room temperature on a shaker.
7. Incubate sections with primary antibody diluted in blocking solution overnight at room temperature on a shaker.  With certain antibodies, to reduce background staining, consider an incubation for 2-3 days at 4°C.
8. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
9. Incubate sections with biotinylated secondary antibody, diluted in blocking solution for 2 hours at room temperature on a shaker.
10. Rinse sections 4 x 10 minutes in PBS 0.1 M on a shaker.
11. Prepare ABC solution  at least 30 minutes prior to incubation to allow for ABC complex to form. Add 2 drops of solution A and 2 drops of solution B per 10 ml of blocking solution. Solutions A and B can also be added to plain PBS 0.1 M.
12. Incubate sections in ABC solution for 1-2 hours at room temperature on a shaker.
13. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
14. Incubate sections in DAB solution for 8 minutes at room temperature on a shaker. DAB solution is highly toxic and carcinogen. Wear gloves and handle with care.
15. Add three drops of 0.3%  H₂O₂ (~125 ul) to each well to reveal staining. When background is satisfactory (after 1 to 5 min), halt the reaction by adding PBS 0.1 M.
16. Rinse sections 4 x 10 minutes with PBS 0.1 M on a shaker.
17. Transfer sections to slides using a brush, allow to air dry. It is best to transfer sections as soon as possible but well plates can be stored for a few days in the fridge at 4°C.
18. Dehydrate slides twice in ethanol 100% for 5 minutes each.
19. Incubate slides twice in toluene or xylene for 5 minutes each.
20. Add mounting medium to slides while still wet. Place coverslips to slides and allow to dry. Examine staining by microscopy.

Reagents

• Sodium phosphate, monobasic anhydrous NaH₂PO₄ (FW 120.0). Sigma,  S-0751, 1Kg
• Sodium phosphate, dibasic anhydrous, Na₂HPO₄ (FW 142.0). Sigma, S-0876, 1Kg
• Hydrogen peroxide, H2O2  30% (w/w) solution. Sigma, H-1009, 100 ml
• Albumin Bovine fraction V, min 96%, electrophoresis. Sigma, A-9647, 50g
• 3,3′-diaminobenzidine tablets (DAB). Sigma, D-5905, 50 tablets
• Goat serum. BioWest, Cat# S2000, 100ml or similar
• Vectastain ABC Kit, Elite standard. Vector, PK-6100
• Triton X-100 (t-Octylphenoxypolyethoxyethanol). Sigma, T-9284, 100 ml
• Toluene or xylene from VWR or Fisher
• Ethanol 100%

Antibodies

Titrate new batches of antibodies for appropriate concentration before using in experiments as effective concentrations may vary across batches of antibody?.

Primary

Rabbit anti-Fos polyclonal IgG, Oncogene Research Products (Ab-5, Cat.# PC38). Recommended dilution, 1:20 000.

Secondary

Biotin-SP-conjugated affiniPure Goat anti-rabbit IgG (H+L) (minimal cross reaction to Human , Mouse and rat serum proteins). Made in goat. Jackson Immunoresearch, Cat.# 111-065-144. Recommended dilution: 1:2000.

Solutions

Phosphate buffer solution, 0.2 M,  pH 7.4

1. Collect 1000 ml of distilled water in a graduated cylinder. Pour about 400 ml of water in a beaker and stir.
2. Weigh 4.8 g of Sodium Phosphate monobasic NaH₂PO₄ and 22.72 g of sodium phosphate dibasic Na₂HPO₄ .
3. Add to the 400 ml of water. When dissolved, add the rest of the water and continue stirring for 5 min. Take pH which should be around 7.4.

Phosphate buffer solution, 0.1 M,  pH 7.4

Make phosphate buffer 0.2M solution as described above and add 1000 ml of distilled water to bring it to 0.1 M, total volume 2 liters. pH should be around 7.4. Solution can be kept at room temperature or at 4°C.

Blocking solution (PBS 0.1 M; 0.1 % BSA; 0.2% Triton X-100; 2% serum)

Collect about 800 ml of phosphate buffer 0.1 M in a graduated cylinder. Add 20 ml of serum, 2 ml of Triton X-100 and 1 g of BSA. Stir for 10 min. Add more PBS 0.1 M to reach 1000 ml. Stir another 5 min. Store blocking solution in 50-ml aliquots (50-ml Falcon tubes) at -20°C

DAB solution, 0.05% (w/v)

Add 1 tablet (10 mg) of DAB in 20 ml of PBS 0.1 M in a 50-ml Falcon tube. Vortex vigorously until dissolved. Solution should be used fresh, or may be frozen in single-use aliquots and stored at -20C until use. Wear gloves and inactivate solution using a 10% bleach solution (dilute DAB with an equal volume of bleach) when finished and dispose in appropriate biohazard container. DAB is highly toxic and carcinogen; do not dump solution down the drain without treatment.

Neutralization of DAB

Although chlorine bleach is commonly employed in many laboratories as a neutralization procedure, it is not effective in removing the mutagenic properties of DAB. A potassium permanganate-sulfuric acid procedure must be used.

1. Take up bulk quantities of diaminobenzidine tetrahydrochloride dehydrate in water and bulk quantities of the free base in 0.1 M hydrochloric acid so that the concentration of DAB does not exceed 0.9 mg/ml.  Dilute solutions with the same buffer, if necessary, so that the concentration does not exceed 0.9 mg/ml.
2. For each 10 ml of solution, add 5 ml of 0.2 M potassium permanganate solution and 5 ml of 2 M sulfuric acid solution.
3. Allow the mixture to stand overnight, decolorize by the addition of sodium ascorbate, neutralize and dispose solution down the drain with copious amounts of water.

0.3% (v/v) H₂O₂ solution

Add 0.5 ml of H₂O₂ 30% solution to 50 ml of PBS 0.1 M in a 50-ml Falcon tube. Vortex. Use fresh.

Equipment

• Microscope
• 2D Shaker
• 6-well plates
• Gelatin-coated slides or precleaned superfrost plus slides (25 x 75 x 1 mm). VWR, Cat.# 48311-703
• Coverlips (micro cover glasses) 24 x 60 mm, No. 1. VWR, Cat.# 48404 454.
• Mounting medium (Eukit or Cytoseal 280 from Richard-Allan Scientific (8311-4) or similar)

Phosphate buffered saline (abbreviated as PBS) is a buffer solution commonly used in biological research. It is a salty solution containing sodium chloride, sodium phosphate, and (in some formulations) potassium chloride and potassium phosphate. The buffer helps to maintain a constant pH. The osmolarity and ion concentrations of the solution usually match those of the human body (isotonic).

Applications

PBS has many uses because it is isotonic and non-toxic to cells. It can be used to dilute substances. It is used to rinse containers containing cells. PBS can be used as a diluent in methods to dry biomolecules, as water molecules within it will be structured around the substance (protein, for example) to be ‘dried’ and immobilized to a solid surface. The thin film of water that binds to the substance prevents denaturation or other conformational changes. Carbonate buffers may be used for the same purpose but with less effectiveness. PBS can be used to take a reference spectrum when measuring the protein adsorption in ellipsometry.

Additives can be used to add function. For example, PBS with EDTA is also used to disengage attached and clumped cells. Divalent metals such as zinc, however, cannot be added as this will result in precipitation. For these types of applications, Good’s buffers are recommended.

Preparation

There are many different ways to prepare PBS. Some formulations do not contain potassium, while others contain calcium or magnesium^[1]. One of the most common preparations is described below.

A 10 liter stock of 10x PBS can be prepared by dissolving 800 g NaCl, 20 g KCl, 144 g Na₂HPO₄ · 2H₂O and 24 g KH₂PO₄ in 8 L of distilled water, and topping up to 10 L. The pH is ~6.8, but when diluted to 1x PBS it should change to 7.4. When making buffer solutions, it is good practice to always measure the pH directly using a pH meter. If necessary, pH can be adjusted using hydrochloric acid or sodium hydroxide.

On dilution, the resultant 1x PBS should have a final concentration of 137 mM NaCl, 10 mM Phosphate, 2.7 mM KCl, and a pH of 7.4.
Another preparation is described in Molecular Cloning by Sambrook, Fritsch and Maniatis, Apendix B.12^[2] as follows:

For 1 litre of 1X PBS, prepare as follows:

1. Start with 800 ml of distilled water:
2. Add 8 g of NaCl.
3. Add 0.2 g of KCl.
4. Add 1.44 g of Na₂HPO₄.
5. Add 0.24 g of KH₂PO₄.
6. Adjust the pH to 7.4 with HCl.
7. Add distilled water to a total volume of 1 liter.

Dispense the solution into aliquots and sterilize them by autoclaving (20 min, 121°C, liquid cycle). Store at room temperature.

References

1. Dulbecco, R. et al. (1954): Plaque formation and isolation of pure lines with poliomyelitis viruses. In: J. Exp. Med. vol. 99 (2), pp. 167-182. PMID 13130792
2. Sambrook, Fritsch, and Maniatis (1989) Molecular Cloning: A Laboratory Manual, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, volume 3, apendix B.12
3. Portions of this article are from “Phosphate buffered saline. In Wikipedia, the free encyclopedia. Retrieved September 17, 2008, from http://en.wikipedia.org/wiki/Phosphate_buffered_saline.” This article has been reviewed for scientific accuracy and is used in accordance with Wikipedia’s GNU Free Documentation License (GFDL).