Electrophoresis of Nucleic Acids

In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally-occurring negative charge carried by their sugar-phosphate backbone. Double-stranded DNA fragments naturally behave as long rods, so their migration through the gel is relative to their radius of gyration, or, for non-cyclic fragments, size. Single-stranded DNA or RNA tend to fold up into molecules with complex shapes and migrate through the gel in a complicated manner based on their tertiary structure. Therefore, agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again.

Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis.

DNA electrophoresis is an analytical technique used to separate DNA fragments by size. An electric field forces the fragments to migrate through a gel. DNA molecules normally migrate from negative to positive potential due to the net negative charge of the phosphate backbone of the DNA chain. At the scale of the length of DNA molecules, the gel looks much like a random, intricate network. Longer molecules migrate more slowly because they are more easily 'trapped' in the network.

After the separation is completed, the fractions of DNA fragments of different length are often visualizing a fluorescent dye specific for DNA, such as ethidium bromide. The gel shows bands corresponding to different DNA molecules populations with different molecular weight. Fragment size is usually reported in "nucleotides", "base pairs" or "kb" (for 1000's of base pairs) depending upon whether single- or double-stranded DNA has been separated. Fragment size determination is typically done by comparison to commercially available DNA ladders containing linear DNA fragments of known length.

The types of gel most commonly used for DNA electrophoresis are agarose (for relatively long DNA molecules) and polyacrylamide (for high resolution of short DNA molecules, for example in DNA sequencing). Gels have conventionally been run in a "slab" format such as that shown in the figure, but capillary electrophoresis has become important for applications such as high-throughput DNA sequencing. Electrophoresis techniques used in the assessment of DNA damage include alkaline gel electrophoresis and pulsed field gel electrophoresis. The measurement and analysis are mostly done with a specialized gel analysis software. Capillary electrophoresis results are typically displayed in a trace view called an electropherogram.

The DNA strand is cut into smaller fragments using DNA endonuclease, then samples of the DNA solution (DNA sample and buffer) are placed in the wells of the gel and allowed to run for some time (the less the voltage of the electophoresis, the longer time for the DNA sample to run through the gel, and this results in a more accurate separation).

Agarose gel electrophoresis

Background

Agarose gel electrophoresis is the easiest and commonest way of separating and analyzing DNA. The purpose of the gel might be to look at the DNA, to quantify it or to isolate a particular band. The DNA is visualised in the gel by addition of ethidium bromide. This binds strongly to DNA by intercalating between the bases and is fluorescent meaning that it absorbs invisible UV light and transmits the energy as visible orange light.

What percentage gel?

Most agarose gels are made between 0.7% and 2%. A 0.7% gel will show good separation (resolution) of large DNA fragments (5–10kb) and a 2% gel will show good resolution for small fragments (0.2–1kb). Some people go as high as 3% for separating very tiny fragments but a vertical polyacrylamide gel is more appropriate in this case. Low percentage gels are very weak and may break when you try to lift them. High percentage gels are often brittle and do not set evenly. I usually make 1% gels.

Which gel tank?

Small 8x10cm gels (minigels) are very popular and give good photographs. Larger gels are used for applications such as Southern and Northern blotting. The volume of agarose required for a minigel is around 30–50mL, for a larger gel it may be 250mL. This method assumes you are making a mini-gel.

How much DNA should I load?

The big question. You may be preparing an analytical gel to just look at your DNA. Alternatively, you may be preparing a preparative gel to separate a DNA fragment before cutting it out of the gel for further treatment. Either way you want to be able to see the DNA bands under UV light in an ethidium-bromide-stained gel. Typically, a band is easily visible if it contains about 20ng of DNA.

Now consider an example. Suppose you are digesting a plasmid that comprises 3kb of vector and 2kb of insert. You are using EcoRI (a common restriction enzyme) and you expect to see three bands: the linearised vector (3kb), the 5' end of the insert (0.5kb) and the 3' end of the insert (1.5kb). In order to see the smallest band (0.5kb) you want it to contain at least 20ng of DNA. The smallest band is 1/10th the size of the uncut plasmid. Therefore you need to cut 10x20ng that is 200ng of DNA (0.2µg). Then your three bands will contain 120ng, 20ng and 60ng of DNA respectively. All three bands will be clearly visible on the gel and the biggest band will be six times brighter than the smallest band.

Now imagine cutting the same plasmid with BamHI (another popular restriction enzyme) and that BamHI only cuts the plasmid once, to linearise it. If you digest 200ng of DNA in this case then the band will contain 200ng of DNA and will be very bright and will probably be overloaded.

Too much DNA loaded onto a gel is a bad thing. The band appears to run fast (implying that it is smaller than it really is) and in extreme cases can mess up the electrical field for the other bands, making them appear the wrong size also.

Too little DNA is only a problem in that you will not be able to see the smallest bands because they are too faint.

Having said all that, I usually digest and load 2–4µL of the 50µL obtained from a kit miniprep. But you see how it depends on the number and size of the bands expected.
For PCR reactions, it depends on the PCR but in routine applications 10–20µL should be plenty to see the product on the gel.

Which comb?

This depends on the volume of DNA you are loading and the number of samples. Combs with many tiny teeth may hold 10µL. This is no good if you want to load 20µL of restriction digest plus 5µL of loading buffer. When deciding whether a comb has enough teeth, remember that you need to load at least one marker lane, preferably two.

Making the gel (for a 1% gel, 50mL volume)

Weigh out 0.5g of agarose into a 250mL conical flask. Add 50mL of 0.5xTBE , swirl to mix.

It is good to use a large container, as long as it fits in the microwave, because the agarose boils over easily.

Microwave for about 1 minute to dissolve the agarose.

The agarose solution can boil over very easily so keep checking it. It is good to stop it after 45 seconds and give it a swirl. It can become superheated and NOT boil until you take it out whereupon it boils out all over you hands. So wear gloves and hold it at arms length. You can use a bunsen burner instead of a microwave - just remember to keep watching it.

Leave it to cool on the bench for 5 minutes down to about 60°C (just too hot to keep holding in bare hands).

If you had to boil it for a long time to dissolve the agarose then you may have lost some water to water-vapour. You can weigh the flask before and after heating and add in a little distilled water to make up this lost volume. While the agarose is cooling, prepare the gel tank ready, on a level surface.

Add 1µL of ethidium bromide (10mg/mL) and swirl to mix

The reason for allowing the agarose to cool a little before this step is to minimise production of ethidium bromide vapour. Ethidium Bromide is mutagenic and should be handled with extreme caution. Dispose of the contaminated tip into a dedicated ethidium bromide waste container. 10mg/mL ethidium bromide solution is made up using tablets (to avoid weighing out powder) and is stored at 4°C in the dark with TOXIC labels on it.

Pour the gel slowly into the tank. Push any bubbles away to the side using a disposable tip. Insert the comb and double check that it is correctly positioned.

The benefit of pouring slowly is that most bubbles stay up in the flask. Rinse out the flask immediately.

Leave to set for at least 30 minutes, preferably 1 hour, with the lid on if possible.

The gel may look set much sooner but running DNA into a gel too soon can give terrible-looking results with smeary diffuse bands.

Pour 0.5x TBE buffer into the gel tank to submerge the gel to 2–5mm depth. This is the running buffer.

You must use the same buffer at this stage as you used to make the gel. ie. If you used 0.6xTBE in the gel then use 0.6xTBE for the running buffer. Remember to remove the metal gel-formers if your gel tank uses them.

Preparing the samples

Transfer an appropriate amount of each sample to a fresh microfuge tube.

It may be 10µL of a 50µL PCR reaction or 5µL of a 20µL restriction enzyme digestion. If you are loading the entire 20µL of a 20µL PCR reaction or enzyme digestion (as I often do) then there is no need to use fresh tubes, just add in the loading buffer into the PCR tubes. Write in your lab-book the physical order of the tubes so you can identify the lanes on the gel photograph.

Add an appropriate amount of loading buffer into each tube and leave the tip in the tube.

Add 0.2 volumes of loading buffer, eg. 2µL into a 10µL sample. The tip will be used again to load the gel.

Load the first well with marker.

I store my markers ready-mixed with loading buffer at 4°C. I know to load 2µL and how much DNA is in each band. See below for more on this.
Avoid using the end wells if possible. For example, If you have 12 samples and 2 markers then you will use 14 lanes in total. If your comb formed 18 wells then you will not be using 4 wells. It is best to not use the outer wells because they are the most likely to run aberrantly.

Continue loading the samples and finish of with a final lane of marker

I load gels from right to left with the wells facing me. This is because gels are published, by convention, as if the wells were at the top and the DNA had run down the page. If this seems confusing then you can load left to right with the wells facing away from you.

Close the gel tank, switch on the power-source and run the gel at 5V/cm.

For example, if the electrodes are 10cm apart then run the gel at 50V. It is fine to run the gel slower than this but do not run any faster. Above 5V/cm the agarose may heat up and begin to melt with disastrous effects on your gel's resolution. Some people run the gel slowly at first (eg. 2V/cm for 10 minutes) to allow the DNA to move into the gel slowly and evenly, and then speed up the gel later. This may give better resolution. It is OK to run gels overnight at very low voltages, eg. 0.25–0.5V/cm, if you want to go home at 11 O'clock already.

Check that a current is flowing

You can check this on the power-source, the milliamps should be in the same ball-park as the voltage, but the the best way is to look at the electrodes and check that they are evolving gas (ie. bubbles). If not then check the connections, that the power-source is plugged in etc.etc. This has been known to happen if people use water instead of running buffer.

Monitor the progress of the gel by reference to the marker dye.

Stop the gel when the bromophenol blue has run 3/4 the length of the gel.

Switch off and unplug the gel tank and carry the gel (in its holder if possible) to the dark-room to look at on the UV light-box.

Some gel holders are not UV transparent so you have to carefully place the gel onto the glass surface of the light-box. UV is carcinogenic and must not be allowed to shine on naked skin or eyes. So wear face protection, gloves and long sleeves.

Loading buffers

The loading buffer gives colour and density to the sample to make it easy to load into the wells. Also, the dyes are negatively charged in neutral buffers and thus move in the same direction as the DNA during electrophoresis. This allows you to monitor the progress of the gel. The most common dyes are bromophenol blue (Sigma B8026) and xylene cyanol (Sigma X4126). Density is provided by glycerol or sucrose.

Typical recipe

• 25mg bromophenol blue or xylene cyanol
• 4g sucrose
• H₂O to 10mL

The exact amount of dye is not important
Store at 4°C to avoid mould growing in the sucrose. 10mL of loading buffer will last for years.

Bromophenol blue migrates at a rate equivalent to 200–400bp DNA. If you want to see fragments anywhere near this size (ie. anything smaller than 600bp) then use the other dye because the bromophenol blue will obscure the visibility of the small fragments.
Xylene cyanol migrates at approximately 4kb equivalence. So do not use this if you want to visualise fragments of 4kb.

Size markers

There are lots of different kinds of DNA size markers. In the old days the cheapest defined DNA was from bacteriophage so alot of markers are phage DNA cut with restriction enzymes. Many of these are still very popular eg, lambda HindIII, lambda PstI, PhiX174 HaeIII. These give bands with known sizes but the sizes are arbitrary. Choose a marker with good resolution for the fragment size you expect to see in you sample lanes. For example, for tiny PCR products you might choose PhiX174 HaeIII but for 6kb fragments you would choose lambda HindIII. More recently, companies have started producing ladder markers with bands at defined intervals, eg. 0.5, 1, 1.5, 2, 2.5kb and so on up to 10kb. If you know the total amount of DNA loaded into a marker lane, and you know the sizes of all the bands, you can calculate the amount of DNA in each band visible on the gel. This can be very useful for quantifying the amount of DNA in your sample bands by comparison with the marker bands. It is good to load two markers lanes, flanking the samples. Lots of companies sell DNA size markers. It pays to shop around for the cheapest. Often the local kitchen-sink biotech company sells excellent markers.

TBE

TBE stands for Tris Borate EDTA.

People also use TAE (Tris Acetate EDTA). Make up a 10x stock using cheap reagents. Do not use expensive 'analytical grade' reageants. Cheap Tris base and boric acid can be bought in bulk.

Recipe for 2L of 10xTBE

• 218g Tris base
• 110g Boric acid
• 9.3g EDTA

Dissolve the ingredients in 1.9L of distilled water. pH to about 8.3 using NaOH and make up to 2L.

No responsibility is assumed by methodbook.net for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. It is the users responsibility to ensure that all procedures are carried out according to appropriate Health and Safety requirements.

Capillary electrophoresis

Capillary electrophoresis (CE), also known as capillary zone electrophoresis (CZE), can be used to separate ionic species by their charge and frictional forces. In traditional electrophoresis, electrically charged analytes move in a conductive liquid medium under the influence of an electric field. Introduced in the 1960s, the technique of capillary electrophoresis (CE) was designed to separate species based on their size to charge ratio in the interior of a small capillary filled with an electrolyte. While its use has been sporadic, CE offers unparalleled resolution and selectivity allowing for separation of analytes with very little physical difference. Efficiencies of millions of plates are routinely reported. Once thought impossible, separation of large proteins differing in only one amino acid (ie. D-Lysine substituted for L-Lysine) and even an isotopic separation of ¹⁴N and ¹⁵N ammonium hydroxide have been reported. No other technique has shown such powerful selectivity with the ability for extremely high sensitivity. As few as 6 molecules of a substance have been separated and detected with the help of laser-induced fluorescence (LIF).

Instrumentation

The instrumentation needed to perform capillary electrophoresis is relatively simple. A basic schematic of a capillary electrophoresis system is shown in figure 1. The system's main components are a sample vial, source and destination vials, a capillary, electrodes, a high-voltage power supply, a detector, and a data output and handling device. The source vial, destination vial and capillary are filled with an electrolyte such as an aqueous buffer solution. To introduce the sample, the capillary inlet is placed into a vial containing the sample and then returned to the source vial (sample is introduced into the capillary via capillary action, pressure, or siphoning). The migration of the analytes is then initiated by an electric field that is applied between the source and destination vials and is supplied to the electrodes by the high-voltage power supply. It is important to note that all ions, positive or negative, are pulled through the capillary in the same direction by electroosmotic flow, as will be explained. The analytes separate as they migrate due to their electrophoretic mobility, as will be explained, and are detected near the outlet end of the capillary. The output of the detector is sent to a data output and handling device such as an integrator or computer. The data is then displayed as an electropherogram, which reports detector response as a function of time. Separated chemical compounds appear as peaks with different retention times in an electropherogram.

Detection

Separation by capillary electrophoresis can be detected by several detection devices. The majority of commercial systems use UV or UV-Vis absorbance as their primary mode of detection. In these systems, a section of the capillary itself is used as the detection cell. The use of on-tube detection enables detection of separated analytes with no loss of resolution. In general, capillaries used in capillary electrophoresis are coated with a polymer for increased stability. The portion of the capillary used for UV detection, however, must be optically transparent. Bare capillaries can break relatively easily and, as a result, capillaries with transparent coatings are available to increase the stability of the cell window. The path length of the detection cell in capillary electrophoresis (~ 50 micrometers) is far less than that of a traditional UV cell (~ 1 cm). According to the Beer-Lambert law, the sensitivity of the detector is proportional to the path length of the cell. To improve the sensitivity, the path length can be increased, though this results in a loss of resolution. The capillary tube itself can be expanded at the detection point, creating a "bubble cell" with a longer path length or additional tubing can be added at the detection point as shown in figure 2. Both of these methods, however, will decrease the resolution of the separation.

Fluorescence detection can also be used in capillary electrophoresis for samples that naturally fluoresce or are chemically modified to contain fluorescent tags. This mode of detection offers high sensitivity and improved selectivity for these samples, but cannot be utilized for samples that do not fluoresce. The set-up for fluorescence detection in a capillary electrophoresis system can be complicated. The method requires that the light beam be focused on the capillary, which can be difficult for many light sources. Laser-induced fluorescence has been used in CE systems with detection limits as low as 10^-18 to 10^-21 mol. The sensitivity of the technique is attributed to the high intensity of the incident light and the ability to accurately focus the light on the capillary.

In order to obtain the identity of sample components, capillary electrophoresis can be directly coupled with mass spectrometers or Surface Enhanced Raman Spectroscopy (SERS). In most systems, the capillary outlet is introduced into an ion source that utilizes electrospray ionization (ESI). The resulting ions are then analyzed by the mass spectrometer. This set-up requires volatile buffer solutions, which will affect the range of separation modes that can be employed and the degree of resolution that can be achieved. The measurement and analysis are mostly done with a specialized gel analysis software.

For CE-SERS, capillary electrophoresis eluants can be deposited onto a SERS-active substrate. Analyte retention times can be translated into spatial distance by moving the SERS-active substrate at a constant rate during capillary electrophoresis. This allows the subsequent spectroscopic technique to be applied to specific eluants for identification with high sensitivity. SERS-active substrates can be chosen that do not interfere with the spectrum of the analytes.

Modes of separation

The separation of compounds by capillary electrophoresis is dependent on the differential migration of analytes in an applied electric field. The electrophoretic migration velocity (u_p) of an analyte toward the electrode of opposite charge is:

where μ_p is the electrophoretic mobility and E is the electric field strength. The electrophoretic mobility is proportional to the ionic charge of a sample and inversely proportional to any frictional forces present in the buffer. When two species in a sample have different charges or experience different frictional forces, they will separate from one another as they migrate through a buffer solution. The frictional forces experienced by an analyte ion depend on the viscosity (η) of the medium and the size and shape of the ion. Accordingly, the electrophoretic mobility of an analyte at a given pH is given by:

where z is the net charge of the analyte and r is the Stokes radius of the analyte. The Stokes radius is given by:

where k_B is the Boltzmann constant, and T is the temperature, D is the diffusion coefficient. These equations indicate that the electrophoretic mobility of the analyte is proportional to the charge of the analyte and inversely proportional to its radius. The electrophoretic mobility can be determined experimentally from the migration time and the field strength:

where L is the distance from the inlet to the detection point, t_r is the time required for the analyte to reach the detection point (migration time), V is the applied voltage (field strength), and L_t is the total length of the capillary. Since only charged ions are affected by the electric field, neutral analytes are poorly separated by capillary electrophoresis.

The velocity of migration of an analyte in capillary electrophoresis will also depend upon the rate of electroosmotic flow (EOF) of the buffer solution. In a typical system, the electroosmotic flow is directed toward the negatively charged cathode so that the buffer flows through the capillary from the source vial to the destination vial. Separated by differing electrophoretic mobilities, analytes migrate toward the electrode of opposite charge. As a result, negatively charged analytes are attracted to the positively charged anode, counter to the EOF, while positively charged analytes are attracted to the cathode, in agreement with the EOF as depicted in figure 3.

Figure 3: Diagram of the separation of charged and neutral analytes (A) according to their respective electrophoretic and electroosmotic flow mobilities

The velocity of the electroosmotic flow, u_o can be written as:

u_o = μ_oE

where μ_o is the electroosmotic mobility, which is defined as:

where ζ is the zeta potential of the capillary wall, and ε is the relative permittivity of the buffer solution. Experimentally, the electroosmotic mobility can be determined by measuring the retention time of a neutral analyte. The velocity (u) of an analyte in an electric field can then be defined as:

u_p + u_o = (μ_p + μ_o)E

Since the electroosmotic flow of the buffer solution is generally greater than that of the electrophoretic flow of the analytes, all analytes are carried along with the buffer solution toward the cathode. Even small, triply charged anions can be redirected to the cathode by the relatively powerful EOF of the buffer solution. Negatively charged analytes are retained longer in the capilliary due to their conflicting electrophoretic mobilities. The order of migration seen by the detector is shown in figure 3: small multiply charged cations migrate quickly and small multiply charged anions are retained strongly.

Electroosmotic flow is observed when an electric field is applied to a solution in a capillary that has fixed charges on its interior wall. Charge is accumulated on the inner surface of a capillary when a buffer solution is placed inside the capillary. In a fused-silica capillary, silanol (Si-OH) groups attached to the interior wall of the capillary are ionized to negatively charged silanoate (Si-O^-) groups at pH values greater than three. The ionization of the capillary wall can be enhanced by first running a basic solution, such as NaOH or KOH through the capillary prior to introducing the buffer solution. Attracted to the negatively charged silanoate groups, the positively charged cations of the buffer solution will form two inner layers of cations (called the diffuse double layer or the electrical double layer) on the capillary wall as shown in figure 4. The first layer is referred to as the fixed layer because it is held tightly to the silanoate groups. The outer layer, called the mobile layer, is farther from the silanoate groups. The mobile cation layer is pulled in the direction of the negatively charged cathode when an electric field is applied. Since these cations are solvated, the bulk buffer solution migrates with the mobile layer, causing the electroosmotic flow of the buffer solution. Other capillaries including Teflon capillaries also exhibit electroosmotic flow. The EOF of these capillaries is probably the result of adsorption of the electrically charged ions of the buffer onto the capillary walls. The rate of EOF is dependent on the field strength and the charge density of the capillary wall. The wall's charge density is proportional to the pH of the buffer solution. The electroosmotic flow will increase with pH until all of the available silanols lining the wall of the capillary are fully ionized.

Figure 4: Depiction of the interior of a fused-silica gel capillary in the presence of a buffer solution.

Efficiency and resolution

The number of theoretical plates, or separation efficiency, in capillary electrophoresis is given by:

where N is the number of theoretical plates, μ is the apparent mobility in the separation medium and D_m is the diffusion coefficient of the analyte. According to this equation, the efficiency of separation is only limited by diffusion and is proportional to the strength of the electric field. The efficiency of capillary electrophoresis separations is typically much higher than the efficiency of other separation techniques like HPLC. Unlike HPLC, in capillary electrophoresis there is no mass transfer between phases.^[3] In addition, the flow profile in EOF-driven systems is flat, rather than the rounded laminar flow profile characteristic of the pressure-driven flow in chromatography columns as shown in figure 5. As a result, EOF does not significantly contribute to band broadening as in pressure-driven chromatography. Capillary electrophoresis separations can have several hundred thousand theoretical plates.

The resolution (R_s) of capillary electrophoresis separations can be written as:

According to this equation, maximum resolution is reached when the electrophoretic and electroosmotic mobilities are similar in magnitude and opposite in sign. In addition, it can be seen that high resolution requires lower velocity and, correspondingly, increased analysis time.

Related techniques

As discussed above, separations in a capillary electrophoresis system are typically dependent on the analytes having different electrophoretic mobilities. However, some classes of analyte cannot be separated by this effect because they are neutral (uncharged) or because they may not differ significantly in electrophoretic mobility. However, there are several techniques that can help separate such analytes with a capillary electrophoresis system. Adding a surfactant to the electrolyte can facilitate the separation of neutral compounds by micellar electrokinetic chromatography. Charged polymers such as DNA can be separated by filling the capillary with a gel matrix that retards longer strands more than shorter strands. This is called capillary gel electrophoresis. This is a high-resolution alternative to slab gel electrophoresis. Some capillary electrophoresis systems can also be used for microscale liquid chromatography or capillary electrochromatography. A capillary electrophoresis system can also be used for isotachophoresis and isoelectric focusing.

NOTE: Figures are not shown here

Electrophoresis

Active fractions isolated by a preparative fractionation procedure may be subjected to a number of analytical fractionation procedures to determine their purity. Analytical fractionations are distinguished from preparative fractionations by the criteria shown in Table 1.
Table 1. The difference between preparative and analytical fractionations.
Parameters: Analytical / Preparative
Scale: Small / Large
Fate of sample: Destroyed / Preserved
Product Information: Active / fraction

In an analytical fractionation, therefore, a small amount of sample is sacrificed in order to gain information about the state of purity of the material being analysed. Of the many physico-chemical techniques which have contributed to our knowledge of proteins (and nucleic acids), electrophoretic techniques occupy a position of primary importance. Electrophoresis finds its greatest usefulness in the analysis of mixtures and in the determination of purity, although certain forms of electrophoresis may be applied on a preparative scale.
Electrophoresis is a technique used to separate and sometimes purify macromolecules - especially proteins and nucleic acids - that differ in size, charge or conformation. As such, it is one of the most widely-used techniques in biochemistry and molecular biology.
Principles of electrophoresis

Electrophoresis may be defined as the migration of charged ions in an electric field. In metal conductors, electric current flows between electrodes and is carried by ions. The negative electrode - the cathode - donates electrons and the positive electrode - the anode - takes up electrons to complete the circuit. The ions that result from the take up of electrons from the cathode will be negatively charged and will thus migrate towards the positive anode.
Because of their anodic migration, negative ions are called “anions”. Electric current is carried by the movement of electrons, largely along the surface of the metal. In solutions, the Ions which result from the donation of an electron to the electron deficient (i.e. positively charged) anode will themselves be electron deficient, and thus positively charged. These will migrate to the cathode and are thus called cations.

• The force that an ion of charge q will experience when placed in an electric field E is called the electric force (Fef) and is defined as:
Fef = q x E

• This force is opposed by the frictional force,( Ff):
Ff = f V

Where f = frictional coefficient
f = 6hpr for a sphere of radius r (Stoke’s equation)

• In a constant electric field Fef = Ff thus the velocity of a particle moving in an applied electric field, V will also be constant:
V (cm/s) = (q x E) / f = q x E / 6hpr

• Electrophoretic mobility, m, is defined as the velocity/unit electric field
m= V/ E
m= q/f (m=q/6hpr)

The electrophoretic mobility is thus a function of the charge on the protein ion and the medium through which it is traveling. Electrophoretic techniques exploit the fact that different ions have different mobilities in an electric field and so can be separated by electrophoresis.
The flow of electricity in electrophoresis is subject to the same physical laws as other forms of electricity. For example, Ohm's law applies:
I=V/R
Where I = current (amps)
V = potential difference (volts)
R = resistance (ohms).
The unit of electrical charge is the coulomb and the unit of current [the ampere (amp)] may be defined as coulombs sec-1.

The flow- of electricity involves work, which generates heat, and the work (W, in joules) done in transferring a charge of q coulombs between a potential difference of V volts is:-
W=q.V=I.tV
W= I2R.t (Jouleís law of heating)

Which means that I2R.t joules of heat will be developed in the conductor.
The power (in watts) (defined as the rate of work) gives the rate of heating (joules sec-1).

Watts= I2R.t/t= I2R
Heating of the electrophoretic medium has the following effects
1. An increased rate of sample diffusion.
2. Mixing of separated samples due to convection current.
3. Thermal instability, protein denaturation and loss of enzyme activity.
4. Decrease in buffer viscosity and reduction in the resistivity of the medium.

Support Media

GEL ELECTROPHORESIS
Electrophoresis of macromolecules can be carried out in solution. However, the ability to separate molecules is compromised by their diffusion. Greater resolution is achieved if electrophoresis is carried out on semi-solid supports such as polyacrylamide or agarose gels. Gels are formed by cross-linking polymers in aqueous medium. This will form a 3-dimensional meshwork which the molecules must pass through. Polyacrylamide is a common gel for protein electrophoresis whereas agarose is more commonly used for nucleic acids. Agarose gels have a larger pore size than acrylamide gels and are better suited for larger macromolecules. However, either type of gel can be applied to either nucleic acids or proteins depending on the application.

Gels are formed from long polymers in a cross-linked lattice. The space between the polymers are the pores. Higher concentrations of the polymer will result in smaller average pore sizes. Polyacrylamide gels are formed by covalently cross-linking acrylamide monomers with bis-acrylamide with a free radical like persulfate (SO4•). The cross-linking of the acrylamide polymers results in 'pores' of a defined size. The total acrylamide concentration and the ratio of bis-acrylamide to acrylamide will determine the average pore size. The polyacrylamide solution is poured into a mold and polymerized. This mold can be a cylindrical tube, but is usually a 'slab' poured between two glass plates

Since the gel is solid with respect to the mold, all molecules are forced through the gel. Smaller molecules will be able to pass through this lattice more easily resulting in larger molecules having a lower mobility than smaller molecules. In other words, the gel acts like a molecular sieve and retains the larger molecules while letting the smaller ones pass through. (This is opposite of gel filtration where the larger molecules have a higher mobility because they to not enter the gel.) Therefore, the frictional coefficient is related to how easily a protein passes through the pores of the gel and size will be the major determinant of the mobility of molecules in a gel matrix. Protein shape and other factors will still affect mobility, but to a lesser extent. Substituting size for the frictional coefficient results in:
Agarose is a polysaccharide extracted from seaweed. It is typically used at concentrations of 0.5 to 2%. The higher the agarose concentration the "stiffer" the gel. Agarose gels are extremely easy to prepare: you simply mix agarose powder with buffer solution, melt it by heating, and pour the gel. It is also non-toxic.
Agarose gels have a large range of separation, but relatively low resolving power. By varying the concentration of agarose, fragments of DNA from about 200 to 50,000 bp can be separated using standard electrophoretic techniques.
The length of the polymer chains is dictated by the concentration of acrylamide used, which is typically between 3.5 and 20%. Polyacrylamide gels are significantly more annoying to prepare than agarose gels. Because oxygen inhibits the polymerization process, they must be poured between glass plates (or cylinders).
Acrylamide is a potent neurotoxin and should be handled with care! Wear disposable gloves when handling solutions of acrylamide, and a mask when weighing out powder. Polyacrylamide is considered to be non-toxic, but polyacrylamide gels should also be handled with gloves due to the possible presence of free acrylamide.
Polyacrylamide gels have a rather small range of separation, but very high resolving power. In the case of DNA, polyacrylamide is used for separating fragments of less than about 500 bp. However, under appropriate conditions, fragments of DNA differing is length by a single base pair are easily resolved. In contrast to agarose, polyacrylamide gels are used extensively for separating and characterizing mixtures of proteins.
PAGE

1. NONDENATURING (Native) PAGE
Nondenaturing system is used to separate intact proteins, especially oligomeric proteins, by a nondestructive means for later assessment of biological activity. On nondenaturing system, protein separation depends on a combination of differences in molecular size and shape as well as charge. Separation by size is accomplished by varying the pore size of the acrylamide polymer as a function of both the concentration of the acrylamide (range about 3-30%, w/v) and the amount of cross-linker used. In general, the higher the acrylamide concentration, the smaller the protein that remains in gel: this can be counteracted by decreasing the amount of cross-linker used, which in turn increases the degree of gel swelling during standard staining and washing procedures. Separation by charge in nondenaturing gel systems is permitted because the protein separation can be performed at any pH between 3 and 11 to allow for maximal charge differences neighboring protein species. These include the molecular weight of the stability in the range of pH 4 to 9. Once these parameters are known optimal resolution can be obtained by varying certain components within a single gel system. For example the initial choice of the gel system for separation of acidic protein of molecular weight 20,000 might be a system operating at an alkaline pH i.e.9 with an acylamide concentration of 12-15 %.
It should be noted that biologically active proteins such as enzymes often require specials electrophoresis conditions in order to retain activity after separation. For example the heat lability of the some enzymes requires that electrophoresis be conducted most conveniently in a refrigerated room or if necessary using circulating water of 0-4C in the cooling jacket of the slab gel apparatus. In addition ammonium persulphate an agent commonly used in gel polymerization reaction often interferes with enzymes activity after elution from the gel. For this reason the photo activated polymerizing agent riboflavin can also be used instead of ammonium persulphate in the preparation of the stacking gel. For the various systems if ammonium persulphate affects enzyme activity then preelectrophoresis can be used. Dithioithreitol (40-100 mM) or 2-mercaptoethanol (upto 1 M) may be included in the sample buffer to reduce some disulfide linkages, although detergent denaturing is often necessary for complete disulfide reduction.

2. SDS- PAGE
Acrylamide mixed with bisacrylamide forms a cross-linked polymer network when the polymerizing agent, ammonium persulfate, is added (Figure 1). The ammonium persulfate produces free radicals faster in the presence of TEMED (N, N, N, N’-tetramethylenediamine). The size of the pores created in the gel is inversely related to the amount of acrylamide used. Gels with a low percentage of acrylamide are typically used to resolve large proteins and high percentage gels are used to resolve small proteins.

Figure 2. Polymerization and cross-linking of acrylamide.
However, polyacrylamide of sufficient stability will always be so dense, that friction influences the migration of proteins in the gel. The electrophoretic mobility of a protein is determined mostly by its net charge per unit mass at the given pH, but is inversely proportional to its frictional coefficient in the gel, determined by the proteins size and shape. In the denaturing, reductive variant of PAGE, SDS-PAGE all differences between the proteins in charge per unit mass has been eliminated by the SDS (sodium dodecylsulphate) and the proteins migrate solely according to subunit size. The charged SDS molecules bind all along the polypetide chain, giving the chain equal charge per unit length. Thus, the denatured polypeptide chains are separated electrophoretically only according to their subunit length. The polyacrylamide gels may be cast as i fixed concentration gel (e.g. 10%) or as a gradient gel (e.g. 7-15 %). There is also a choice between continuous and discontinuous buffer systems. If the same buffer ions are present throughout sample, gel and electrophoresis buffer the electrophoresis is referred to as continuous buffer PAGE. In discontinuous buffer PAGE the separation is usually preceded by a stacking of the sample to a narrow band in a low-porosity stacking gel.
The high pore size stacking gel is to concentrate the protein sample into a sharp band before it inters to the main separating gel. This is achieved by utilizing differences in the ionic strength and pH between the electrophoresis buffer and the stacking gel. The band sharpening effect relies on the fact that negatively charged glycine ions (in running buffer) have a lower electrophoretic mobility than do the protein-SDS complex which in turn has lower mobility than the chloride ion of the loaded buffer and the stacking gel. When current is switched on, all the ionic species have to migrate at the same speed otherwise there would be a break in the electrical current. The glycinate ions can only move at the same speed as Cl- ion if they are in the region of the higher field strength. Field strength is inversely proportional to conductivity which is proportional to conc. The result is that three species of interest adjust their concentrations so that [Cl-] > [protein-SDS] > [glycinate ions]. There is only a small quantity of protein-SDS complexes so they concentrate in a very tight bond between glycinate and Cl- boundaries. Once the glycinate reaches the separating gel it becomes more fully ionized in higher pH environment and its mobility increases. Thus the interface between the glycinate and Cl- leaves behind the protein-SDS complexes which are left to electrophoresis at their own rates. The negatively charged protein SDS complexes now continue to move towards the anode and because they have the same charge per unit length they travel into the separating gel under the applied electric field with the same mobility .However as they pass through the separating gel the protein separate owing to the molecular sieving properties of the gel. Detection of the separated protein bands may be done in several ways. Staining for protein with Coomassie Brilliant Blue G-250 is a standard procedure detecting 0.1 to 1µg per band (depending on band dimensions). Coomassie Brilliant Blue G- 250 can also be used and by using dilute staining solution, destaining can be omitted. An increase in detection sensitivity of approximately a factor 100 can be obtained with silver staining.
Determination of Molecular Weight
This is done by SDS-PAGE of proteins known molecular weight along with the proteins to be characterized. A linear relationship exists between the logarithm of the molecular weight of an SDS-denatured polypeptide and its Rf value. The Rf is calculated as the ratio of the distance migrated by the molecule to that migrated by a marker dye-front. A simple way of determining relative molecular weight by electrophoresis (Mr) is to plot a standard curve of distance migrated vs. log10MW for known samples, and read off the logMr of the sample after measuring distance migrated on the same gel.





























Isoelectric focusing (IEF)
IEF, also known as electrofocusing, is a technique for separating different molecules by their electric charge differences. It is a type of zone electrophoresis, usually performed in a gel, that takes advantage of the fact that a molecule's charge changes with the pH of its surroundings. A protein which is in a pH region below its pI will be positively charged and so will migrate towards the cathode. However, as it migrates, the charge will decrease until the protein reaches a pH which is its pI. At this point it has no net charge and so migration ceases. Should it overshoot this point, it will enter a region of pH above its pI and so become negatively charged. It will then reverse its direction of migration and now migrate towards the anode. Therefore proteins become focused into sharp stationary bands with each protein positioned at a point in the pH gradient corresponding to its pI. The technique is capable of extremely high resolution with proteins differing by a single charge being fractionated into separate bands.

Molecules to be focused are distributed over a medium that has a pH gradient (usually created by aliphatic ampholytes). An electric current is passed through the medium, creating a "positive" anode and "negative" cathode end. Negatively charged molecules migrate through the pH gradient in the medium toward the "positive" end while positively charged molecules move toward the "negative" end. As a particle moves towards the pole opposite of its charge it moves through the changing pH gradient until it reaches a point in which the pH of that molecules isoelectric point is reached. At this point the molecule no longer has a net electric charge (due to the protonation or deprotonation of the associated functional groups) and as such will not proceed any further within the gel. The gradient is initially established before adding the particles of interest by first subjecting a solution of small molecules such as polyampholytes with varying pI values to electrophoresis.
The method is applied particularly often in the study of proteins, which separate based on their relative content of acidic and basic residues, whose value is represented by the pI. Proteins are introduced into a Immobilized pH gradient gel composed of polyacrylamide, starch, or agarose where a pH gradient has been established. Gels with large pores are usually used in this process to eliminate any "sieving" effects, or artifacts in the pI caused by differing migration rates for proteins of differing sizes. Isoelectric focusing can resolve proteins that differ in pI value by as little as 0.01. Isoelectric focusing is the first step in two-dimensional gel electrophoresis, in which proteins are first separated by their pI and then further separated by molecular weight through SDS-PAGE.

Two-dimensional gel electrophoresis
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.
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, PDQuest and Progenesis - 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.
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]
• differences in software algorithms and therefore analysis tendencies
New approaches taken by software packages, such as Delta2D and Progenesis, go a long way in compensating for running differences between gels and solving the missing values problem to allow for robust statistical analysis of 2-DE gels. Generated picking lists can be used for the automated in-gel digestion of protein spots, and subsequent identification of the proteins by mass spectrometry. Although 2-DE automated image analysis technology has not been perfected, manual visual analysis is no longer realistic for objective and statistically valid experiment designs.

Western blot
The western blot (alternately, immunoblot) is a method of detecting specific proteins in a given sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D structure of the protein (native/ non-denaturing conditions). The proteins are then transferred to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using antibodies specific to the target protein. There are now many reagent companies that specialize in providing antibodies (both monoclonal and polyclonal antibodies) against many thousands of different proteins. Commercial antibodies can be expensive, though 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[1] 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.
Steps in a Western blot
1. 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.
A combination of biochemical and mechanical techniques – including various types of filtration and centrifugation – can be used to separate different cell compartments and organelles.
2. 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.
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. S-S disulfide bonds to 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 kilo Daltons, kD). 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. An example of a ladder is the GE Full Range Molecular weight ladder (Figure 1). 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.
3. 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 fluoride (PVDF). The membrane is placed on top of the gel, and a stack of tissue 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. Coomassie is the more sensitive of the two, although Ponceau S's water solubility makes it easier to subsequently destain and probe the membrane as described below.
4. 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. 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.
5. 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.
5.1 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 just about 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 enhances the signal.
Most commonly, a horseradish peroxidase-linked secondary is used in conjunction with 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.
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).
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.
5.2 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.
6. Analysis
After the unbound probes are washed away, the Western blot is ready for detection of the probes that are labeled and bound to the protein of interest. In practical terms, not all Westerns reveal protein only at one band in a membrane. Size approximations are taken by comparing the stained bands to that of the marker or ladder loaded during electrophoresis. The process is repeated for a structural protein, such as actin or tubulin, that should not change between samples. The amount of target protein is indexed to the structural protein to control between groups. This practice ensures correction for the amount of total protein on the membrane in case of errors or incomplete transfers.
6.1 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.

6.2 Chemiluminescence
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 captures 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. So-called "enhanced chemiluminescent" (ECL) detection is considered to be among the most sensitive detection methods for blotting analysis.
6.3 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.
6.4 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.
6.5 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.