Optimizing Gel Electrophoresis: Ampere, Volts and Buffer concentrations

Optimizing Gel Electrophoresis: Ampere, Volts and Buffer concentrations

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I am a master student in biochemistry, and I have used gel electrophoresis many times before. What I want to know is how one should adjust the mA (mAmpere) compared to the voltage and the buffer one uses.

Normally I used 1xTAE buffer (run at 90V) or 1x sodium borate buffer (run at 110V). I know that if I adjust the voltage up, the buffer heats up faster, the samples travel faster, but the "picture" of the gel becomes more blurry. This is however all I know about adjust the parameter.

So my questions can be summarized as the following:

  1. How should I adjust the ampere, is there a range I should be in compared to the buffer or sample I am running?

  2. Does the ampere effect the travel speed if adjusted up or down?

  3. Can I adjust the concentration of the buffer to run sample at a higher voltage, what effects will this have (to increase or decrease concentration)?

  4. Is there a relationship between the ampere and the voltage which I can manipulate, in this case: how and why?

  1. As far as I know you can either adjust the ampere or the voltage as both is dependent of the resistance from the buffer. Which means that I would suggest you to let the voltage as it is.

  2. As the ampere = voltage / resistance and voltage = resistance * ampere and as you can see that the voltage has to get higher if the ampere is increased and the ampere has to get higher if the voltage is increased (at least if I understand it correctly) the travel speed should be increased the higher the ampere is set.

  3. I would neither decrease nor increase the concentration of the buffer. Maybe easily using another kind of buffer would help you. But this depends on the different samples you are running. Maybe that could help you: Brody, J.R., and Kern, S.E. (2004). History and principles of conductive media for standard DNA electrophoresis. Analytical Biochemistry 333, 1-13.

  4. The resistance is more or less given. Therefore as far as I know: No.

Agarose and Metaphor Gels

1.Add agarose to 1X TBE (or TAE) buffer. For gel size 20 x 24 cm, use 300-400 ml buffer and 0.7 to 1.0% agarose.
2. Melt agarose in 500 ml flask in microwave oven, mixing several times during heating. Let cool to 55 C (until flask can be held).
3. Tape the ends of gel tray and place on a level bench.
4. Add ethidium bromide: 2.5 ul of 10 mg/ml stock per 100 ml. (gel cam also be stained in ethidium bromide bath after electrophoresis (see point 9).

NOTE: Ethidium bromide is mutagenic - wear gloves when handling, and use extra caution. Change gloves when contaminated and dispose in separate waste for ethidium bromide.

5. Pour agarose into tray and insert combs. Remove bubbles with a pipette tip. Allow to solidify.
6. Remove tape and place tray in gel boxes. Pour enough 1X TBE (or TAE) buffer into the gel box to cover the gel by at least 0.5 cm. Remove combs when ready to load samples.
7. Load 1 ug Lambda digested with Hind III (5 ul of 200 ng/ul stock) as molecular weight marker, then load samples.
8. Run at 15-25 V for 12-24 hours.
9. If no ethidium bromide was added to the agarose: stain gel in 1 ug/ml ethidium bromide (100 ul of 10 mg/ml ethidium bromide in 1000 ml dd H2O) for 20 min. 10. Rinse gel for 20 min in 1000 ml ddH2O.
11. Slide gel onto UV transilluminator and take photo.
Photographing tip:
Place small piece of paper with writing or transparent ruler on the gel to help focus.

MetaPhor® Agarose
High resolution agarose
MetaPhor agarose is a high resolution agarose that challenges polyacrylamide. MetaPhor agarose is an intermediate melting temperature (75° C) agarose with twice the resolution capabilities of the finest-sieving agarose products. Using submarine gel electrophoresis, you can resolve PCR products and small DNA fragments that differ in size by 2%.

Analytical Specifications
Gelling temperature (3%) = 35° C
Melting temperature (3%) = 75° C
Gel strength (3%) = 300 g/cm²
• High resolution separation of 20-800 bp DNA fragments
• Recovery of fragments under 800 bp
• Fine analysis of PCR? products
• AMPFLP, STR and tri- and tetranucleotide repeat analysis

Suggested Agarose Concentrations

Migration of double-stranded DNA in relation to Bromophenol Blue (BPB) and Xylene Cyanol (XC) in MetaPhor agarose gels.

Always wear eye protection when dissolving agarose and guard yourself and others against scalding solutions.

Microwave Instructions for Agarose Preparation
1. Choose a beaker that is 2-4 times the volume of the solution.
2. Add chilled 1X or 0.5X electrophoresis buffer and a stir bar to the beaker.
3. Slowly sprinkle in the agarose powder while the solution is rapidly stirred.
4. Remove the stir bar if not Teflon® coated.
5. Soak the agarose in the buffer for 15 minutes before heating. This reduces the tendency of the agarose solution to foam during heating.
6. Weigh the beaker and solution before heating.
7. Cover the beaker with plastic wrap.
8. Pierce a small hole in the plastic wrap for ventilation.

For agarose concentrations > 4%, the following additional steps will further help prevent the agarose solution from foaming during melting/dissolution:
a. Heat the beaker in the microwave oven on Medium power for 1 minute.
b. Remove the solution from the microwave.
c. Allow the solution to sit on the bench for 15 minutes.

9. Heat the beaker in the microwave oven on Medium power for 2 minutes.
10. Remove the beaker from the microwave oven. Caution: Any microwaved
solution may become superheated and foam over when agitated.
11. GENTLY swirl the beaker to resuspend any settled powder and gel pieces.
12. Reheat the beaker on HIGH power until the solution comes to a boil.
13. Hold at boiling point for 1 minute or until all of the particles are dissolved.
14. Remove the beaker from the microwave oven.
15. GENTLY swirl the beaker to thoroughly mix the agarose solution.
16. After dissolution, add sufficient hot distilled water to obtain the initial weight.
17. Mix thoroughly.
18. Cool the solution to 50-60°C prior to casting. Once the gel is cast, allow the molten agarose to cool and gel at room temperature. The gel must then be placed at 4° C for 20 minutes to obtain optimal resolution and gel handling characteristics.

Hot Plate Instructions for Agarose Preparation
1 . Choose a beaker that is 2-4 times the volume of the solution.
2. Add chilled electrophoresis buffer and a stir bar to the beaker.
3. Slowly sprinkle the agarose powder while the solution is rapidly stirred.
4. Weigh the beaker and solution before heating.
5. Cover the beaker with plastic wrap.
6. Pierce a small hole in the plastic wrap for ventilation.
7. Bring the solution to a boil while stirring.
8. Maintain gentle boiling until all the agarose is dissolved (approximately 10 minutes).
9. Add sufficient hot distilled water to obtain the initial weight.
10. Mix thoroughly.
11 . Cool the solution to 50-60°C prior to casting. Once the gel is cast, allow the molten agarose to cool and gel at room temperature. The gel must then be placed at 4° C for 20 minutes to obtain optimal resolution and gel handling characteristics.

How Does PFGE Work?

PFGE stemmed from the observation that DNA molecules elongate upon application of an electric field and return to an unelongated state upon removal of the electric field this relaxation rate is dependent on the size of the DNA.

When the orientation of the electric field is changed during electrophoresis, the DNA molecules must return to their elongated form prior to reorientation, thus affecting the migration rate. This effect can be used to greatly extend the size range over which electrophoretic DNA separations are possible.

When the electrical field is applied to the gel, the DNA molecules elongate in the direction of the electrical field. The first electrical field is then switched to the second field according to the run specifications. The DNA must change conformation and reorient before it can migrate in the direction of this field.

As long as the alternating fields are equal with respect to the voltage and pulse duration, the DNA will migrate in a straight path down the gel (see below).

Time-lapse representation of DNA molecules undergoing PFGE.


A number of factors can affect the migration of nucleic acids: the dimension of the gel pores, the voltage used, the ionic strength of the buffer, and the concentration intercalating dye such as ethidium bromide if used during electrophoresis. [2]

Size of DNA Edit

The gel sieves the DNA by the size of the DNA molecule whereby smaller molecules travel faster. Double-stranded DNA moves at a rate that is approximately inversely proportional to the logarithm of the number of base pairs. This relationship however breaks down with very large DNA fragments and it is not possible to separate them using standard agarose gel electrophoresis. The limit of resolution depends on gel composition and field strength. [3] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [4] Separation of very large DNA fragments requires pulse field gel electrophoresis (PFGE). In field inversion gel electrophoresis (FIGE, a kind of PFGE), it is possible to have "band inversion" - where large molecules may move faster than small molecules.

Conformation of DNA Edit

The conformation of the DNA molecule can significantly affect the movement of the DNA, for example, supercoiled DNA usually moves faster than relaxed DNA because it is tightly coiled and hence more compact. In a normal plasmid DNA preparation, multiple forms of DNA may be present, [5] and gel from the electrophoresis of the plasmids would normally show a main band which would be the negatively supercoiled form, while other forms of DNA may appear as minor fainter bands. These minor bands may be nicked DNA (open circular form) and the relaxed closed circular form which normally run slower than supercoiled DNA, and the single-stranded form (which can sometimes appear depending on the preparation methods) may move ahead of the supercoiled DNA. The rate at which the various forms move however can change using different electrophoresis conditions, for example linear DNA may run faster or slower than supercoiled DNA depending on conditions, [6] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [4] Unless supercoiled DNA markers are used, the size of a circular DNA like plasmid therefore may be more accurately gauged after it has been linearized by restriction digest.

DNA damage due to increased cross-linking will also reduce electrophoretic DNA migration in a dose-dependent way. [7] [8]

Concentration of ethidium bromide Edit

Circular DNA are more strongly affected by ethidium bromide concentration than linear DNA if ethidium bromide is present in the gel during electrophoresis. All naturally occurring DNA circles are underwound, but ethidium bromide which intercalates into circular DNA can change the charge, length, as well as the superhelicity of the DNA molecule, therefore its presence during electrophoresis can affect its movement in gel. Increasing ethidium bromide intercalated into the DNA can change it from a negatively supercoiled molecule into a fully relaxed form, then to positively coiled superhelix at maximum intercalation. [9] Agarose gel electrophoresis can be used to resolve circular DNA with different supercoiling topology.

Gel concentration Edit

The concentration of the gel determines the pore size of the gel which affect the migration of DNA. The resolution of the DNA changes with the percentage concentration of the gel. Increasing the agarose concentration of a gel reduces the migration speed and improves separation of smaller DNA molecules, while lowering gel concentration permits large DNA molecules to be separated. For a standard agarose gel electrophoresis, a 0.7% gives good separation or resolution of large 5–10kb DNA fragments, while 2% gel gives good resolution for small 0.2–1kb fragments. Up to 3% can be used for separating very tiny fragments but a vertical polyacrylamide gel would be more appropriate for resolving small fragments. High concentrations gel however requires longer run times (sometimes days) and high percentage gels are often brittle and may not set evenly. High percentage agarose gels should be run with PFGE or FIGE. Low percentage gels (0.1−0.2%) are fragile and may break. 1% gels are common for many applications. [10]

Applied field Edit

At low voltages, the rate of migration of the DNA is proportional to the voltage applied, i.e. the higher the voltage, the faster the DNA moves. However, in increasing electric field strength, the mobility of high-molecular-weight DNA fragments increases differentially, and the effective range of separation decreases and resolution therefore is lower at high voltage. For optimal resolution of DNA greater than 2kb in size in standard gel electrophoresis, 5 to 8 V/cm is recommended. [6] Voltage is also limited by the fact that it heats the gel and may cause the gel to melt if a gel is run at high voltage for a prolonged period, particularly for low-melting point agarose gel.

The mobility of DNA however may change in an unsteady field. In a field that is periodically reversed, the mobility of DNA of a particular size may drop significantly at a particular cycling frequency. [11] This phenomenon can result in band inversion whereby larger DNA fragments move faster than smaller ones in PFGE.

The negative charge of its phosphate backbone moves the DNA towards the positively charged anode during electrophoresis. However, the migration of DNA molecules in solution, in the absence of a gel matrix, is independent of molecular weight during electrophoresis, i.e. there is no separation by size without a gel matrix. [12] Hydrodynamic interaction between different parts of the DNA are cut off by streaming counterions moving in the opposite direction, so no mechanism exists to generate a dependence of velocity on length on a scale larger than screening length of about 10 nm. [11] This makes it different from other processes such as sedimentation or diffusion where long-ranged hydrodynamic interaction are important.

The gel matrix is therefore responsible for the separation of DNA by size during electrophoresis, however the precise mechanism responsible the separation is not entirely clear. A number of models exists for the mechanism of separation of biomolecules in gel matrix, a widely accepted one is the Ogston model which treats the polymer matrix as a sieve consisting of randomly distributed network of inter-connected pores. [13] A globular protein or a random coil DNA moves through the connected pores large enough to accommodate its passage, and the movement of larger molecules is more likely to be impeded and slowed down by collisions with the gel matrix, and the molecules of different sizes can therefore be separated in this process of sieving. [11]

The Ogston model however breaks down for large molecules whereby the pores are significantly smaller than size of the molecule. For DNA molecules of size greater than 1 kb, a reptation model (or its variants) is most commonly used. This model assumes that the DNA can crawl in a "snake-like" fashion (hence "reptation") through the pores as an elongated molecule. At higher electric field strength, this turned into a biased reptation model, whereby the leading end of the molecule become strongly biased in the forward direction, and this leading edge pulls the rest of the molecule along. In the fully biased mode, the mobility reached a saturation point and DNA beyond a certain size cannot be separated. [13] Perfect parallel alignment of the chain with the field however is not observed in practice as that would mean the same mobility for long and short molecules. [11] Further refinement of the biased reptation model takes into account of the internal fluctuations of the chain. [14]

The biased reptation model has also been used to explain the mobility of DNA in PFGE. The orientation of the DNA is progressively built up by reptation after the onset of a field, and the time it reached the steady state velocity is dependent on the size of the molecule. When the field is changed, larger molecules take longer to reorientate, it is therefore possible to discriminate between the long chains that cannot reach its steady state velocity from the short ones that travel most of the time in steady velocity. [14] Other models, however, also exist.

Real-time fluorescence microscopy of stained molecules showed more subtle dynamics during electrophoresis, with the DNA showing considerable elasticity as it alternately stretching in the direction of the applied field and then contracting into a ball, or becoming hooked into a U-shape when it gets caught on the polymer fibres. [15] [16] This observation may be termed the "caterpillar" model. [17] Other model proposes that the DNA gets entangled with the polymer matrix, and the larger the molecule, the more likely it is to become entangled and its movement impeded. [18]

The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide, usually abbreviated as EtBr. It fluoresces under UV light when intercalated into the major groove of DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than

20 ng DNA becomes distinctly visible. EtBr is a known mutagen, [19] and safer alternatives are available, such as GelRed, produced by Biotium, which binds to the minor groove. [20]

SYBR Green I is another dsDNA stain, produced by Invitrogen. It is more expensive, but 25 times more sensitive, and possibly safer than EtBr, though there is no data addressing its mutagenicity or toxicity in humans. [21]

SYBR Safe is a variant of SYBR Green that has been shown to have low enough levels of mutagenicity and toxicity to be deemed nonhazardous waste under U.S. Federal regulations. [22] It has similar sensitivity levels to EtBr, [22] but, like SYBR Green, is significantly more expensive. In countries where safe disposal of hazardous waste is mandatory, the costs of EtBr disposal can easily outstrip the initial price difference, however.

Since EtBr stained DNA is not visible in natural light, scientists mix DNA with negatively charged loading buffers before adding the mixture to the gel. Loading buffers are useful because they are visible in natural light (as opposed to UV light for EtBr stained DNA), and they co-sediment with DNA (meaning they move at the same speed as DNA of a certain length). Xylene cyanol and Bromophenol blue are common dyes found in loading buffers they run about the same speed as DNA fragments that are 5000 bp and 300 bp in length respectively, but the precise position varies with percentage of the gel. Other less frequently used progress markers are Cresol Red and Orange G which run at about 125 bp and 50 bp, respectively.

Visualization can also be achieved by transferring DNA after SDS-PAGE to a nitrocellulose membrane followed by exposure to a hybridization probe. This process is termed Southern blotting.

For fluorescent dyes, after electrophoresis the gel is illuminated with an ultraviolet lamp (usually by placing it on a light box, while using protective gear to limit exposure to ultraviolet radiation). The illuminator apparatus mostly also contains imaging apparatus that takes an image of the gel, after illumination with UV radiation. The ethidium bromide fluoresces reddish-orange in the presence of DNA, since it has intercalated with the DNA. The DNA band can also be cut out of the gel, and can then be dissolved to retrieve the purified DNA. The gel can then be photographed usually with a digital or polaroid camera. Although the stained nucleic acid fluoresces reddish-orange, images are usually shown in black and white (see figures). UV damage to the DNA sample can reduce the efficiency of subsequent manipulation of the sample, such as ligation and cloning. Shorter wavelength UV radiations (302 or 312 nm) cause greater damage, for example exposure for as little as 45 seconds can significantly reduce transformation efficiency. Therefore if the DNA is to be use for downstream procedures, exposure to a shorter wavelength UV radiations should be limited, instead higher-wavelength UV radiation (365 nm) which cause less damage should be used. Higher wavelength radiations however produces weaker fluorescence, therefore if it is necessary to capture the gel image, a shorter wavelength UV light can be used a short time. Addition of Cytidine or guanosine to the electrophoresis buffer at 1 mM concentration may protect the DNA from damage. [23] Alternatively, a blue light excitation source with a blue-excitable stain such as SYBR Green or GelGreen may be used.

Gel electrophoresis research often takes advantage of software-based image analysis tools, such as ImageJ.

Fundamentals Of Electrophoresis

Electrophoresis refers to the migration of charged molecules through a liquid or gel medium under the influence of an electric field. Zone electrophoresis, or the migration of macromol-ecules through a porous support medium, or gel, is almost universally used in molecular biology laboratories today. Electrophoresis (all discussion in this chapter will involve zone electrophoresis but will be referred to as electrophoresis for brevity) is a powerful separation tool, being able to detect the differential migration of macromolecules with only subtly different structures. The two formats that are used are slab gel electrophoresis, in which the electrophoresis takes place in support medium, agarose, or polyacrylamide, and capillary electrophoresis (CE), an instrumental technique in which the electrophoresis takes place in a capillary tube.

The rate of migration of a macromolecule through a gel matrix is dependent on several factors, including (1) the net charge on the molecule at the pH at which the assay is conducted, (2) the size and shape of the molecule, (3) the electric field strength or voltage drop, (4) the pore size of the gel, and (5) temperature. The forces acting on the analyte to drive it through the gel are the charge on the molecule and the electric field strength. Equation

(1) describes the electrophoretic driving force (Table 1).

The forces acting to retard the movement of the molecule are the frictional forces, determined by the velocity of the ana-lyte, the pore size of the gel, and the size and shape of the molecule. The opposition of the acceleration of the analyte by the frictional forces is described by Stokes' Law as shown in Eq.

(2) (Table 1). When the electrophoretic driving force equals the frictional force (F = F), the result is a constant velocity of the analyte molecule through the gel matrix [Eq. (2), Table 1]. The term viX describes the velocity of the analyte through the gel matrix (cmis) per unit field strength (Vicm) under constant conditions (i.e., the same buffer conditions and the same gel viscosity). This term (given by the symbol ||) is defined as the electrophoretic mobility of the analyte (expressed as cm2iV-s). From the units of the electrophoretic mobility, it can be seen that if the gel is run under constant-voltage conditions (Vicm constant) for a given period of time (s), then (cm2iV-s)(V-sicm) is distance of migration (in cm). Thus, if gels are run such that the product of the voltage and the time are constant, the same analyte will migrate the same distance into the gel. For convenience, this is typically expressed as volt-hours. For example, if one gel is run at 50 V for 10 h (500 V-h) and an identical gel (constant length, viscosity, and buffer concentration) is run at 100 V for 5 h (500 V-h), the same analyte will appear at the same position on both gels. This ability to reproduce gel profiles is one of the principal reasons that constant-voltage run conditions are preferred for DNA analysis. An exception to this is the preferred conditions for DNA sequencing. As will be discussed later, sequencing gels are run at elevated temperature to ensure the adequate denaturation of the single-stranded DNA molecules in this case, running the gel at constant watts is helpful in maintaining an even heating of the gel.

In protein analysis, the pH of the electrophoresis buffer can be a powerful tool in optimizing specific separations. This is because the type (acidic or basic) and number of ionizable groups found on proteins are variable. However, for DNA analysis, it is the charge of the phosphate backbone that is dominant. Therefore, DNA electrophoresis is typically performed at a slightly alkaline pH to ensure full ionization of the phosphate residues.

3.1. THE GEL MATRIX: AGAROSE There are two types of gel matrix in common use in DNA laboratories: agarose and polyacrylamide. Agarose is a polysaccharide commercially derived from seaweed. The agarose polymer consists of multiple agarbiose molecules linked together into linear chains, with an average molecular weight of 120,000 daltons. The agarbiose subunit is a disaccharide consisting of P-d-galactose and 3,6-anhydro-a-l-galactose (Fig. 1) (6). The partially purified material, agar, consists of noncharged polymer chains, agarose, and negatively charged chains. The negative charges are typically the result of sulfate (-SO4) residues. In general, the more highly purified the agarose, the lower the sulfate concentration, the higher the quality of the separation, and the higher the price. Agarose is supplied as a white, nonhydroscopic powder. A gel is prepared by mixing agarose powder with buffer, boiling the mixture, pouring the molten gel into a casting tray, and cooling. During this process, the agarose chains shift from existing in solution as random coils to a structure in which the chains are bundled into double helices. The average pore size for agarose gels are typically in the range 100-300 nm3. Agarose gels are used at concentrations near 1% (wiv) for separating DNA fragments in the 1- to 20-kb size range. Examples of applications common in the DNA diagnostic laboratory include restriction digestion analysis of large plasmids and Southern transfer analysis of genomic DNA.

Fig. 2. Schematic demonstrating the polymerization of acrylamide and bis-acrylamide monomers.

Agarose Gel Electrophoresis of DNA

To pour a gel, agarose powder is mixed with electrophoresis buffer to the desired concentration, then heated in a microwave oven until completely melted. Most commonly, ethidium bromide is added to the gel (final concentration 0.5 ug/ml) at this point to facilitate visualization of DNA after electrophoresis. After cooling the solution to about 60C, it is poured into a casting tray containing a sample comb and allowed to solidify at room temperature or, if you are in a big hurry, in a refrigerator.

After the gel has solidified, the comb is removed, using care not to rip the bottom of the wells. The gel, still in its plastic tray, is inserted horizontally into the electrophoresis chamber and just covered with buffer. Samples containing DNA mixed with loading buffer are then pipeted into the sample wells, the lid and power leads are placed on the apparatus, and a current is applied. You can confirm that current is flowing by observing bubbles coming off the electrodes. DNA will migrate towards the positive electrode, which is usually colored red.

The distance DNA has migrated in the gel can be judged by visually monitoring migration of the tracking dyes. Bromophenol blue and xylene cyanol dyes migrate through agarose gels at roughly the same rate as double-stranded DNA fragments of 300 and 4000 bp, respectively.

When adequate migration has occured, DNA fragments are visualized by staining with ethidium bromide. This fluorescent dye intercalates between bases of DNA and RNA. It is often incorporated into the gel so that staining occurs during electrophoresis, but the gel can also be stained after electrophoresis by soaking in a dilute solution of ethidium bromide. To visualize DNA or RNA, the gel is placed on a ultraviolet transilluminator. Be aware that DNA will diffuse within the gel over time, and examination or photography should take place shortly after cessation of electrophoresis.

Migration of DNA Fragments in Agarose

Other Considerations

Agarose gels, as discussed above provide the most commonly-used means of isolating and purifying fragments of DNA, which is a prerequisite for building any type of recombinant DNA molecule.

By varying buffer composition and running conditions, the utility of agarose gels can be extended. Examples include:

  • Pulsed field electrophoresis is a technique in which the direction of current flow in the electrophoresis chamber is periodically altered. This allows fractionation of pieces of DNA ranging from 50,000 to 5 millon bp, which is much larger than can be resolved on standard gels.
  • Alkaline agarose gels are prepared with and electrophoresed in buffers containing sodium hydroxide. Such alkaline conditions are useful for analyzing single-stranded DNA.

Finally, if you haven’t had an opportunity to run agarose gels, try out the virtual agarose electrophoresis lab.

Optimizing Gel Electrophoresis: Ampere, Volts and Buffer concentrations - Biology

The name, &lsquowestern&rsquo blot, was first coined by Dr. Burnette in 1981 after the eponymous southern blot for DNA and consequent coinage of the northern blot in 1977 for RNA.[1][2] The western blot (WB) is an effective and widely utilized immunoassay that confers selective protein expression analysis. WB selects for an individual protein amongst a potentially significant milieu via leveraging the specificity of antigen (Ag)-antibody (Ab) binding. Of notable interest is its application in clinical pathology, wherein a western blot detects the presence of a pathologically relevant protein from a patient sample. This review will discuss the biochemical principles, clinical significance, and troubleshooting aspects of this technique. 

Specimen Requirements and Procedure

Principles of Western Blot

Western blot relies on the principles of equal loading of proteins, separation of proteins by molecular weight, electrophoretic transfer to a suitable membrane, and probing of antibodies.

Equal Loading of Proteins

Proper sample preparation for subsequent electrophoresis is crucial for downstream analysis. Western blot samples are first prepared by protein extraction with specialized cell lysis buffers and protease and phosphatase inhibitors (PPIs). There are numerous methods of extraction, and proper selection depends on the sample type. For example, most tissue preparation is by homogenization or sonication however, osmotic shock or detergent lysis is more suited for easily lysed cells such as erythrocytes or cultured cells. Furthermore, the cell lysis buffer used in extraction should align with target protein cellular localization.[3] For example, radioimmunoprecipitation assay buffer (RIPA) is more adept for nuclear and mitochondrial proteins. Although rare, some antibodies will not be able to detect denatured samples. As such, gentle buffers without detergents are required. PPIs are used to maintain the structure and phosphorylation status of the target protein from the activity of endogenous phosphatases upon cell lysis and exogenous phosphatases in the lysis microenvironment. Collectively, this information underscores the need to tailor protein extraction to sample type and the target protein.

There must be an equal concentration of proteins per western blot sample. Intuitively, this is imperative for a valid experiment as unequal proteins per lane can skew the analysis. By conducting a Bradford assay, a colorimetric protein assay that exploits a dye&rsquos interaction with protein, protein concentration is quantifiable.[4] In brief, the dye Coomassie Brilliant Blue G-250 complexes with proteins to change color, and this absorbance shift gets recorded by a spectrophotometer. Thus, by running this assay with known protein standards, a linear regression standard curve is generated to calculate unknown protein extract concentrations.

All western blot samples have three things: protein extract, cell lysis buffer, and Laemmli (sample) buffer. Protein extract gets normalized with cell-lysis buffer to the desired protein concentration, and there is an addition of an equal volume of Laemmli (sample) buffer. Therefore, there is always a 1 to 1 volume ratio of normalized protein and Laemmli buffer in a western blot sample. Laemmli buffer (60mM Tris-HCl pH 6.8 20% glycerol 2% SDS 4% beta-mercaptoethanol 0.01% bromophenol blue) is unique to western blot sample preparation as each reagent is purposeful for SDS-PAGE.[5][6] Glycerol adds density to samples, so they drift into the loading wells. Bromophenol blue (BPB) is a nonreactive reagent that serves as a dye front for electrophoresis. SDS is a potent anionic detergent that coats denatured proteins with an equal anion to mass ratio this masks proteins' charge, shape, and size characteristics and renders them solely a function of molecular weight. Beta-mercaptoethanol (BME) is a reducing agent that acts on disulfide bonds in the absence of BME, proteins with disulfide bonds retain some shape and do not electrophorese consummately by molecular weight. Tris-HCl pH 6.80 works in conjunction with the discontinuous buffer system, explained in further detail below. Prepared samples are heated before loading to further denature proteins to their respective primary structure. Thus, proteins undergo electrophoresis by their monomeric weight. 

Separation of Proteins by Molecular Weight

The separation of proteins by weight is possible due to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) --particularly its combinatory use of a detergent and a discontinuous buffer system.

Typically, PAGE is an analytical biochemistry method used to separate contents such as nucleic acids and proteins by electrophoretic mobility in a chemically inert gel however, by adding SDS, a potent anionic detergent, all denatured proteins will be coated with an equal charge to mass ratio. Therefore, the rate of protein migration is proportional to weight. Indeed, larger proteins travel slower in comparison to smaller proteins due to retarding properties of the porous gel. A gel matrix is formed from the polymerization of acrylamide and crosslinking of N, N'-methylenebisacrylamide. This matrix creates a molecular sieve that imbues retarding properties. The pore size of this sieve is alterable by adjusting the percentage of polyacrylamide/ N, N'-methylenebisacrylamide as they are inversely proportional.[7] Two different sized sieves are used in PAGE: a stacking gel and a resolving gel. As the name suggests, the stacking gel stacks proteins into a narrow band to allow proteins to enter the resolving gel at the same time, which is made possible due to its bigger pore size and acidity. With its much smaller pore size and basicity, the resolving gel is where the separation of proteins occurs.

The Laemmi discontinuous buffer system is most commonly used in SDS-PAGE. This system utilizes running buffer (25mM Tris 192mM glycine 0.1% SDS pH

8.30) as electrode buffer and Tris-HCl to buffer an acidic stacking gel (pH

6.80) and a basic resolving gel (pH

8.80). The deliberate use of varied pHs exploits glycine&rsquos charge properties. In an acidic environment, glycine is a zwitterion, but in a basic environment, it is a glycinate anion. Thus, when electrophoresis starts, the current quickly draws glycinate into the stacking gel. The acidic gel protonates glycine to its zwitterionic form, thereby severely impeding its mobility.

In contrast, the chloride from the Tris-HCl buffer in the gel disassociates from its counter ion and migrates quickly to the anode. Proteins lie in between a trailing front of glycine and a leading front of chloride this results in all proteins arriving at the resolving gel simultaneously, a vital component for subsequent separation. The basicity of the resolving gel reforms conjugate glycinate anions at the stacking-resolving gel interface. From this interface, glycinate anions quickly migrate past the protein front. The proteins now hit the resolving gel in narrow bands without a zone of high voltage previously formed from the leading and trailing ions in the stacking gel. Thus, this allows proteins to migrate down the resolving gel slower, which induces separation of proteins due to the higher concentration of polyacrylamide (Figure 1a). 

The samples run in their respective lanes alongside a molecular weight marker, often called a protein ladder. For example, a typical setup would have the ladder in the first lane and the samples in the remaining lanes. The ladder establishes standard molecular weight bands that are then used to read the relative weight of proteins. 

Electrophoretic Transfer (Blotting)

Blotting is the electrophoretic transfer of gel contents onto a suitable membrane in a western blot, the contents are proteins. There are multiple methods of blotting in addition to multiple types of membranes. Although various transfer systems exist (wet, semi-dry, fast), the main principle of electrophoretic transfer remains the same. Like electrophoresis, negatively charged samples migrate toward an anode however, in blotting, a transfer sandwich is used with a slightly modified electrode buffer. Towbin buffer (25 mM Tris 192 mM glycine 20% methanol pH 8.3) is the standard transfer buffer, although small tweaks to this buffer are possible for the target protein.[8] Methanol is important in blotting as it increases the hydrophobicity of proteins and facilitates the release of SDS, both of which increase the adsorption of proteins onto the membrane. From cathode to anode, the sandwich organizes as filter paper, polyacrylamide gel, membrane, and filter paper. In a wet transfer system, fiber pads or sponges are placed superficially on each side. The sandwich gets subjected to a perpendicular current that drives gel contents onto the membrane. (Figure 1b). Equilibration of sandwich contents in transfer buffer is crucial for increasing transfer efficiency it prevents the drying of both the gel and membrane, washes electrophoretic contaminants off the gel, and reforms the original gel size. As electrophoresis runs, voltage increases temperatures, and this increases gel size. Thus cold transfer buffer shrinks to the proper size. Interestingly, methanol in transfer buffer also serves to cool the gel during equilibration.   

Each transfer system has its advantages, and selecting one largely depends on the target protein and lab workflow. Among varied transfer apparatuses, the two most commonly used are wet and semi-dry. It is essential to consider that between wet and semi-dry systems, the main differences are the volume of transfer buffer used and transfer time. Wet transfer uses a tank transfer system that requires a large volume of transfer buffer, whereas semi-dry transfer systems typically require only dampening of the sandwich. Semi-dry systems are also time-efficient as blotting usually finishes within an hour, but a low voltage gets applied overnight in a wet transfer. While semi-dry transfer seems to be the better option as there is a significant reduction in both the volume of transfer buffer and length of transfer time, it has its limitations. Large proteins such as membrane receptors do not blot well, and overall transfer efficiency is lower. Wet-transfer shines in its ability to yield high efficiency across a wide range of protein sizes, thus offering the most flexibility. 

When Drs. Burnette and Towbin published their seminal studies electrophoretic transfer was carried out on nitrocellulose membranes. They remained the gold standard until the advent of polyvinylidene difluoride (PVDF) membranes. Concisely, PVDF membranes outcompete nitrocellulose membranes in their protein binding capacity, chemical resistance, and enhanced transfer efficiency in the presence of SDS. PVDF's higher adsorption of proteins and its chemical resistance allows for stripping and reprobing of membranes. Also, by inserting a small percentage of SDS in the transfer buffer, transfer efficiency markedly improves. However, noted protein sensitivity from PVDF could also increase background signal for analysis. Methanol in transfer buffer can shrink nitrocellulose membranes and precipitate out large proteins. Both types of membranes come in different pore sizes, and membrane pore size is directly related to protein weight. Smaller proteins require smaller pore sizes, although a pore size of 0.45 microns is suitable for most proteins. Recent years have seen the development of unique membranes such as those used for near-infrared detection systems. As such, the type of membrane chosen should reflect the target protein and downstream detection systems.

Antibody Probing

Upon completing the electrophoretic transfer, proteins are now on the membrane, and two antibodies serve for probing and analysis. The primary antibody that binds a specific region on the target protein is used to detect its presence on the membrane. The secondary antibody conjugates with a component used for analysis. This antibody indirectly binds the target protein by binding to the constant regions of the primary antibody (Figure 1c). 

Since membranes have a high affinity for protein, before probing, membranes are incubated in a buffer to coat the remaining surface area. This &lsquoblocking&rsquo buffer includes a protein with a minimal binding affinity to the target protein and, consequently, the antibody. Typically, blocking buffer proteins include either casein from powdered milk or bovine serum albumin (BSA). Although casein is cheaper and suitable for most proteins, BSA is considered a better choice when the target protein is phosphorylated. There is cross-reactivity from casein and phosphorylation-specific primary antibodies. After blocking, the membrane is washed with TBS-T, a mixture of Tris-buffered saline, and Tween 20. Tween 20 is a nonionic detergent that helps remove peripherally bound proteins on the membrane.

Probing both primary and secondary antibodies is done by incubating the membrane in a probing buffer of either the primary or secondary antibody in TBS-T. The membrane is first incubated in the primary probing buffer typically overnight in a cold room, and washed again with TBS-T. The membrane is then incubated with the secondary probing buffer for about an hour and then washed as well. These washing steps are crucial to reduce background noise in the analysis. After probing and washing, the membrane is ready to be read.

As mentioned earlier, the secondary antibody conjugates with a component-specific to the type of analysis. Autoradiography was a common way to visualize bands but has declined in its popularity due to hazards associated with this method. It uses a radiolabeled isotope conjugated to the secondary antibody. More commonly, a chemiluminescence method is used. This method uses substrates that react with an enzyme-conjugated secondary antibody. These enzymes are either horseradish peroxidase (HRP) or alkaline phosphatase (AP). The enzyme-mediated reaction produces light that is then recorded with an imaging system. More recently, secondary antibodies have been conjugated with fluorophores that are capable of being detected without the need for substrates. This fluorescence-based detection is gaining popularity due to its capability of probing two target proteins via secondary antibodies with different wavelength fluorophores this is a selective advantage for relative protein expression analysis as housekeeping proteins are visible alongside a protein of interest.

The visualization of bands can serve different analytical purposes. Simply, the presence of bands can verify the expression of a protein, whereas the density of bands can show comparative relative protein expression. A housekeeping protein is also probed to evaluate relative protein expression. A housekeeping protein is a ubiquitous protein that constitutively expresses in all cells. By normalizing the band densities of the target protein with those of the housekeeping protein, a statistically significant difference between sample types can be measured.[9][10]

Clinical Significance

As highlighted earlier, a western blot has a considerable amount of steps. This lengthy process drives up the time and cost needed for accurate results. However, unlike an ELISA, the western blot is less likely to give false-positive results, which is especially true in the diagnosis of HIV.[11]

Western blotting is used to detect anti-HIV antibodies in human serum and urine samples. The protein samples from a known HIV-infected individual get separated by electrophoresis and then blotted on the nitrocellulose membrane. Then a specific antibody is affixed to detect the protein. The western blot is usually performed after the ELISA test to confirm the diagnosis of HIV.[12] It is far more sensitive than the ELISA test. More recently, in commercial HIV western blot kits, viral proteins come affixed to the membrane.  Antibodies from human urine or serum samples bind to these proteins, and anti-HIV antibodies are used to detect bands alongside quality controls. 

The western blot is also useful in detecting Lyme disease and atypical and typical bovine spongiform encephalopathy.[13][14]

Quality control and Lab Safety

Quality Controls

Like any experiment, quality controls should be used to validate findings. In a western blot, a positive control, negative control, loading control, and a no first-degree A-B control are all effective in achieving and maintaining robust experiments.

Controls are dedicated lanes wherein the sample is altered specifically for the control type. A positive control is a sample known to contain the target protein, whereas a negative control is known not to contain the target protein. This can be as general as different organ types or as specific as different cellular localization. For example, if an analysis of the expression of a nuclear protein is the aim, and subcellular fractioning is done to isolate this region, a negative control evaluates the quality of fractioning, non-specific binding of antibodies, and a false-positive. Positive controls are powerful in verifying that the workflow is well-optimized even in the absence of bands in sample lanes. Also, a positive control can verify a negative result.

Loading control is a housekeeping protein such as alpha-tubulin or beta-actin. Probing with antibodies specific for a housekeeping protein checks for an equal amount of proteins per sample.

A nonspecific secondary antibody can yield false positives. The specificity of a secondary antibody is evaluated by not incubating a membrane strip with the primary antibody.


Unfortunately, there are many error arms in this method due to many steps and a lengthy workflow. Discussing every error, its cause, and the solution is outside the scope of this review. The most common issues and their troubleshooting will be of focus below.

Smiling of Bands

When bands are not migrating equally down the gel, this pattern can exaggerate to a smiley pattern. This indicates that the gel has air bubbles, voltage is too high, or the volume of the loading sample is too large. Air bubbles within the gel can distort the migration of bands. A constant voltage during electrophoresis is directly proportional to resistance, and since resistance and temperature are directly linked, a high voltage increases the temperature in the electrophoresis tank. Heat pockets and an overall increase in the temperature of running buffer can also alter migration. Before the buffer can warm up, a high voltage at the start of electrophoresis will rush bands and cause nonlinear migration. A large volume of loading samples can cause spillover into other lanes, and these large bands can skew into another lane.

Absence of Bands

If the detection system shows no signal across all lanes except the ladder, there are a multitude of possible causes. It is best to first localize wherein the workflow that the error occurred. Typically, the most common culprits are poor transfer efficiency or poor probing.

By staining the membrane with Ponceau S, a membrane-safe red dye, bands can be visualized. If bands are well illustrated on the membrane, particularly in the area where target protein is expected to be, it indicates that transfer efficiency is not likely the cause. If there are no bands, transfer settings must be altered. A washout of proteins can occur in which the proteins from the membrane migrate to the filter paper. This is due to transfer time being too high reducing voltage and or transfer time can prevent washout. A poor transfer can also occur if little to no proteins were adsorbed on the membrane. To understand the directionality of transfer, the gel can be stained to reveal bands. A significant visualization of bands can suggest the actual transfer was poor rather than a high voltage or time. Rechecking the quality of transfer buffer, increasing transfer settings, and ensuring proper contact of gel and membrane can resolve this issue. If the target protein is small, a semi-dry system may be preferable.

If a positive control lane is used and there is an absence of bands, this can be due to a poor detection kit, poor antibodies, or even an incorrect antibody concentration. Antibody concentration is optimized by running titration experiments.

Multiple Bands

Only a single row of bands should be visualized in detection. The presence of multiple bands suggests the non-specific binding of antibodies. Polyclonal antibodies typically produce this result, as well as a high antibody concentration. As mentioned earlier, titration experiments should be performed to optimize detection.

High Background With or Without Splotches

Poor membrane blocking, excessive antibody concentration, and a dry membrane can result in high background signals. Increasing the period for blocking or changing the type of protein used in blocking buffer may solve this. Titration experiments have to be run for antibody optimization. Insufficient washing can result in high background signal and are a major cause for splotches on the membrane. Membranes must be maintained wet throughout the experiment, and a dry membrane can give high background signals. 

Standard lab safety rules apply. If casting gels, acrylamide is a potent neurotoxin however, it is chemically inert once polymerized. Careful handling of this reagent is a must, and proper precautionary measures should be met before handling it.

Enhancing Healthcare Team Outcomes

Interprofessional healthcare team members involved in treating and managing conditions where western blot testing applies need to understand the examination results. Lab techs and nursing staff who take and prepare the samples must be well-trained to ensure the samples are of appropriate quality for accurate testing. Interprofessional training, coordination, and communication will improve the clinical validity of western blot results and result in improved patient care and outcomes. [Level 5]

Western Blot Doctor™ — Protein Band Appearance Problems


The Western Blot Doctor is a self-help guide that enables you to troubleshoot your western blotting problems. In this section, you can find solutions to issues related to protein band appearance.

Other sections in the Western Blot Doctor:

Problems and Solutions

Click on the thumbnail that is most representative of your own blot to discover the probable causes and find specific solutions to the problem.

Problem: Inconsistent control protein levels among samples

  • Check that total protein levels are consistent:
    • Initial sample quantitation (O.D., weight, cell count, etc.)
    • Check concentration of protein samples (e.g., using Bradford or Lowry protein assays)
    • To determine whether the changes in loading control levels are due to differences in the amount of sample loaded, or if the differences are caused by variations in expression of the loading control proteins, use total protein stains (e.g., Ponceau S, Coomassie, colloidal gold, or SYPRO Ruby) to visualize proteins on gels and blots before and after transfer to determine relative protein loading. If planning to use the blot in downstream steps, make sure that your stain can be removed or is compatible with antibody detection. For example, Coomassie and colloidal gold are not compatible with downstream steps (see Bio-Rad Protein Stains and the Protein Stain Selection Guide)
      • To determine if there is residual, untransferred protein remaining on the gel, use a total protein stain on the gel after transfer
      • To verify protein transfer, stain the membrane with Ponceau S after blotting
      • Visualize total protein on gels and blots using Bio-Rad&rsquos Stain-Free Gels featuring our proprietary Stain-Free Technology. In addition to providing visual verification of transfer at every step, stain-free imaging enables total protein normalization in each lane, eliminating the need for housekeeping proteins as loading controls
      • Check that loading control expression is consistent across conditions using a secondary loading control. If loading control expression varies with experimental conditions, try using another loading control
      • Try total protein normalization using stain-free technology instead of normalizing to a single housekeeping protein. For more information see the following:
        • Applications & Technologies: Stain-Free Imaging Technology
        • Trends in Protein Separation and Analysis &mdash the Advance of Stain-Free Technology Bioradiations, Sept. 2014
        • A Method for Greater Reliability in Western Blot Loading Controls: Stain-Free Total Protein Quantitation Bulletin 6360

        Problem: Ghost protein bands

        • Decrease total protein loaded for samples
        • Check concentration of protein samples (e.g., using Bradford or Lowry protein assays) before loading the gel
        • Optimize sample loading see Determining the Appropriate Sample Load for Western Blots Protocol
        • Decrease concentration of primary and/or secondary antibodies
        • Optimize your primary and/or secondary antibody concentrations using a checkerboard screening protocol
        • Use a shorter exposure time
        • Use multi-acquisition feature on data acquisition software
        • Use a less sensitive detection substrate
        • Reduce incubation time with detection substrate
        • When constructing the blotting sandwich, do not readjust the blot after the gel has come in contact with the membrane, as this can lead to ghosting on the blotting membrane.

        Problem: Swirls or missing bands bands appear diffuse on blot

        Foam pads for Bio-Rad wet tank blotting systems:

        • Criterion&trade Blotter (1704086)
        • Trans-Blot ® Plus System (1703995)
        • Trans-Blot System (1703914)
        • Mini Trans-Blot System (1703933)

        Problem: Blurry bands

        • Repeat gel electrophoresis at lower voltage
        • Run at lower voltage for entire run
        • Run at lower voltage until proteins begin to enter the resolving gel, then increase voltage for remainder of run
        • Carefully remove air bubbles between the gel and membrane before protein transfer
        • Prepare fresh running buffer or use premixed commercial buffers (see our selection of Buffers and Reagents)

        Problem: Bands are curved (smiling) not straight

        • Check and optimize gel electrophoresis conditions. Consult your instruction manual or the Electrophoresis Guide
        • Reduce voltage during electrophoresis
        • Run gel at 4°C. Place electrophoresis cell in a 4°C cooler during run. Use chilled buffers, a cooling coil, or a &ldquoblue ice&rdquo cooling insert

        Problem: Broad or misshapen bands

        • Electrophoresis artifacts may occur as a result of poor gel polymerization, inappropriate running conditions, contaminated buffers, sample overload, etc. Consult your instruction manual for more details, and see the Electrophoresis Guide for guidance in all aspects of protein electrophoresis
        • Check the salt concentrations of the samples, especially when running salt-precipitated samples. When possible, maintain similar salt contents in all wells. Wells with higher salt levels tend to expand when next to wells with less salt due to osmosis. High salt differentials (especially between sample and buffers) can also cause larger band distortion. Be careful when running salt-precipitated samples
        • High-salt samples can often be desalted using Bio-Gel ® P6 columns

        Problem: White (negative) bands on film using ECL method

        • Load less sample
        • Repeat with dilution series of sample
        • Optimize the sample loading see Determining the Appropriate Sample Load for Western Blots Protocol
        • Reduce/optimize the antibody concentrations using checkerboard screening protocols

        Problem: No bands

        • Confirm protein transfer by staining the membrane with Ponceau S and/or the gel with Coomassie Blue Dye, or use Bio-Rad&rsquos Stain-Free Gels to verify transfer using our unique stain-free imaging technology
        • Note how well any prestained molecular weight markers have transferred onto the blot
        • Optimize and check transfer conditions and setup (especially orientation to electrodes)
        • Repeat using two membranes in case protein has transferred through the first membrane (over-transfer is especially likely with low-MW proteins)
        • Try lower concentration of blocking agent
        • Try alternate blocking agents
        • Use azide-free buffers
        • Use fresh detection reagents
        • Add fresh peroxide to substrate buffer
        • Retrace steps to check compatibility between primary and secondary antibodies
        • Reprobe with correct secondary or strip blot and reprobe if necessary
        • Repeat experiment with the correct antibody combination
        • Increase the antibody concentration 2&ndash4 times higher than initial trial
        • Use a checkerboard screening protocol
        • Increase length of incubation
        • Reduce stringency of wash steps
        • Lower temperature, reduce detergent concentration, reduce ionic strength
        • Test and optimize antibody on dot blots
        • Try an alternate antibody. Bio-Rad now offers high-quality antibodies for all applications
        • Check datasheet for recommended conditions
        • Test and optimize antibody on dot blots
        • Validate target and antibody combinations using checkerboard screening protocols
        • Try an alternate antibody
        • Test on a dot blot at several concentrations
        • Freeze aliquots of antibody and only thaw one at a time as needed for blots store thawed aliquots at 4°C
        • Use fresh aliquots of antibody that have been stored at &ndash20ºC or below
        • If storing an antibody for a very long period of time, store at &ndash80ºC
        • Avoid repeated freeze-thaw cycles
        • For more information, see Tips for Caring for Your Antibody
        • Use another source of target protein
        • Include a positive control in experiment (all PrecisionAb Antibodies include a postitive control lysate)
        • Check concentration of protein samples (e.g., using Bradford or Lowry protein assays)
        • Optimize sample loading see Determining the Appropriate Sample Load for Western Blots Protocol
        • Increase the amount of source material
        • Fractionate or concentrate the sample using one or more of these techniques:


        SACK #10: Post-sequencing reaction

        How to clean my sequenced templates?

        10-20 bp of sequence data, but also make it difficult for the basecalling software to interpret post-'blob' peaks.

        a) Ethanol precipitation: inexpensive, simple, and generates good quality data (Protocol_A [plates].docx) (Protocol_B [tubes].docx click Foil cap.jpg and 96-place racks.jpg to view two of the tools used in these protocols. For further information, click EDTA vs. sodium acetate for DNA precipitation? & 'Dump-&-Blot', pipette, or 'spin-out' ethanol?.

        b) ZR DNA Sequencing Clean-up Kit (available directly from Zymo Research or its distributor,Genesee Scientific): For those who prefer column technologies to Ethanol-cleanups, this Zymo product can be an acceptable alternative and the manufacturer claims that the columns can be regenerated up to 10X with 0.1% HCl. However, in our hands, parallel cleanups of split reactions resulted in much lower signal intensities from the Zymo column (vs. our Ethanol-EDTA protocol), which could cause problems with basecalling of weaker sequencing reactions. Further, although the protocol claims that samples can be eluted in 20% formamide and run directly on the sequencer, we Strongly Discourage that practice as water in formamide leads to the formation of ions that compete for injection on the capillaries. Instead, after elution, samples should be dried and resuspended in pure HiDi formamide. Alternatively, under some circumstances, the samples can be left in water see When to resuspend in water vs. formamide? for further information.

        c) Commercial 'gel-filtration' columns: more expensive. Some people attempt (as we did at one time) to reduce costs by reusing these columns after "rinsing" them and storing them with new 'buffer'. However, although this process can be done without carry-over of sequence from one use to the next, reuse of the columns does lead to unincorporated dye terminator peaks

        70 bp into the DNA sequence. In addition, after the first use, small amounts of the 'gel' can bypass the membrane on a plate and end up in the sample well this gel will bind the dye-labeled products, leading to poor signal intensities unless the 'purified' samples are allowed to resuspend for several hours at 4 o C. As such, we have stopped using the commercial plates altogether, preferring to use the inexpensive and highly effective EtOH-EDTA option instead.

        d) CleanSEQ by Agencourt: costs

        SACK #12: Data analysis

        How long does it take to obtain my sequence data?

        2 hr for a single run (16 samples)

        6 hr for a half-plate (48 samples) and,

        12 hr for a full plate (96 samples).

        b) Modified run modules: as little as

        6 hr for a full plate (e.g., when selecting "M.L.=400" -- which typically generates

        500 bp of data). For further information, see 'Run times' under Why does the location of my samples in a 96-well plate matter?

        c) Turn-around Time: Typically, results are available within 1-3 working days of submission.

        How do I obtain my sequence data?

        a) Genomic Facility website: When your sequences are ready, you will receive an automated email directing you to 'login', click on 'Test Results', and then click on the link to your 'Data' file.

        b) 'Data' files: “zipped”, reducing

        30 Mb of data from a full plate (96 samples) to

        10 Mb after 60 days, all files are archived.

        How to analyze my data?

        a) Format: Results are available as text files and electropherograms.

        i) Text files, accessible through DNAStar, BioEdit, or Notepad.

        ii) Electropherograms, accessible with freeware (Software links):

        (1) Sequence Scanner v1.0 – excellent quality-control tool
        (2) BioEdit – excellent alignment tool and,
        (3) Chromas LITE.

        b) Reanalysis: The 3130XL’s Sequencing Analysis Software can sometimes improve your results however, such analyses are performed only after a specific request. Use Sequence Scanner to determine which sequences are worth having reanalyzed.

        Why analyze data with Sequence Scanner?

        For the purpose of answering your research questions, you may analyze your electropherograms with any software of your choosing. In fact, many other programs are superior to Sequence Scanner in terms of manipulating sequence data. However, few &ndash if any &ndash can match Sequence Scanner (freeware) for investigating the quality of your sequence data and for helping to trouble-shoot sequencing problems. Go to 'Links' to download Sequence Scanner or other DNA analysis software.

        When to have my data reanalyzed with Sequencing Analysis v5.2?

        .70/sample, but yields extremely clean DNA and exceptional sequence reads. Please see us prior to using this method. There are now other similar products that are reported to be just as effective, but much less expensive.

        e) BigDye® XTerminator™ Purification Kit (ThermoFisher Part #s 4376484-4376486): As of June 2021, List price ranges from $2.35/rxn (1000 rxn kit) to 1.53/rxn (2500 rxn kit). Vendor claims complete removal of dye blobs and better sequencing results overall method requires the addition of only two reagents (sequentially or premixed), followed by vortexing for 30 minutes.

        In our tests, XTerminator worked well and was extremely simple to use (samples are never dried nor resuspended). However, our EtOH-precipitation cleanup gave the same quality of sequence and an additional 70-100 bp of sequence (>900 bp vs.

        830 bp for BDx). Nevertheless, given that there are absolutely no losses of sequenced product or of signal strength, this method may be an excellent choice if your reactions typically have low signal strength.

        Caution : Use of BDx requires a special run module on the 3130xl to prevent damage to the array also, certain precautions must be observed during the cleanup itself. Please see us prior to using this method.

        EDTA vs. sodium acetate for DNA precipitation?

        Although we recommend using EDTA, you may use sodium acetate to precipitate your sequenced DNA. However, in our experience, labs that use sodium acetate have greater problems removing the unincorporated dye terminators than do labs that use EDTA.

        As a result, samples from lab using sodium acetate tend to have a massive peak in the first 50-80 bp of each electropherogram, obliterating

        10-20 bp of data. Further, because the basecalling software scales all peaks to the highest observed peak, this dye terminator peak also causes the real peaks to be 'squashed' &ndash making it hard for the software to decipher the trace data.

        ABI's product insert (pdf) for BigDye Terminator Mix v3.1 (cms_041329.pdf) lists two ethanol precipitation methods: EtOH&ndashEDTA and, EtOH&ndashEDTA&ndashsodium acetate. Both methods have benefits and detriments, and neither uses sodium acetate without EDTA.

        'Dump-&-Blot', pipette, or 'spin-out' ethanol?

        10-20 bp of data. With exceptionally high UDT levels, secondary peaks appear about 40 bp later &ndash sometimes even tertiary peaks will form. Second, the software scales all peaks to the highest observed peak thus, your real peaks will be 'squashed', making it hard for the software to decipher the trace data &ndash especially towards the end of your sequence. The methods below are ranked in order of their ability to fully remove the UDT's.

        How to resuspend cleaned, sequenced templates?

        a) Dry: Ensure that samples are completely dry (i.e., all water and ethanol have been removed). If using a thermalcycler for drying samples and ending the dry cycle with a 'hold', ensure that the 'hold' temperature is &ge25 o C (i.e., it needs to be several degrees ABOVE room temperature) holds at low temperatures can lead to the room's humidity condensing in the samples, which may degrade the BigDye. particularly the G-nts.

        b) Resuspend: To wells with samples, add 15 μl of Hi-Di formamide. For further information, see What kind of formamide should I use? and Why keep formamide 'dry'?.

        i) Do NOT create ‘Sets of 16’ by putting formamide in blank wells we may need them!
        ii) Seal samples lightly vortex (optional, do so only if seal integrity is definitely ensured) and, briefly centrifuge (store at 4 o C or freeze).

        c) Options: See When to resuspend in water vs. formamide? for further information.

        When to resuspend in water vs. formamide?

        With good technique, signal strengths should be well above the minimum threshold when samples are resuspended in formamide. However, if signal strength is a problem even after thoroughly troubleshooting your situation, consider resuspending your samples in water as this can increase signal strength by &ge10X. Nevertheless, please note the ' cautions ' listed below:

        Caution 1: Samples should be overlain with mineral oil to prevent oxidation and sample evaporaton. Without the oil overlay, your samples may degrade substantially if they are not run soon after being resuspended with the overlay, sequenced DNA is nearly as stable in water as in formamide. Further, freezing your resuspended samples until they are run on the 3130XL instrument may help to limit the degradation.

        Caution 2: If omitting the oil overlay, resuspend in &ge20-µl of water to minimize the likelihood that evaporation will reduce sample volumes below the minimum required for the capillary array pins to make good contact with the samples. Poor contact will lead to reduced signal no contact will, of course, lead to failed injections. ABI specifies a minimum of 7-µl, but notes that even a slight tilt to the autosampler can mean that >7-µl will be required in some wells thus, the true minimum is more like 10-µl. Do NOT assume that covering samples with the septa mat will deal with this problem water evaporates easily through the septa mat, even when inside the 3130xl sequencer. Evaporation rates vary among sequencers (perhaps due to their location relative to room air ducts) however, in our facility, for a single full 96-well plate, a starting volume of 20-µl has been sufficient. As it takes

        12-hours to run a full 96-well plate (using the standard run module for POP7 and a 50-cm array), samples resuspended in water (without the oil overlay) should be run first if >1 plate is loaded onto the instrument.

        Caution 3: If considering this option (i.e., water resuspension), please note that excessively strong signal reduces read length.

        Caution 4: One purpose of resuspending samples in formamide is to maintain the DNA in a denatured state. So far, we have not noticed any problem with using water instead of formamide however, resuspension in water might be inappropriate if your templates are capable of forming substantial secondary structure.

        What kind of formamide should I use?

        When used to resuspend DNA sequence products, high quality formamide is essential for reproducible data thus, we strongly recommend using ABI's Hi-Di formamide. Formamide purchased from other commercial suppliers is often contaminated with water and undesirable organic and inorganic ions. In addition, formamide is often supplied in glass bottles, which (when opened) exposes the formamide to the air and allows the formamide to absorb water. Minerals may also leach from the glass into the formamide.

        To give you a sense of the degree of purity needed, here is a brief description of the process of preparing raw formamide for use in DNA sequencing. First, the raw (prior to deionization) formamide must be &ge99.5% purity, have low water content, be packed under an inert gas, and have a conductivity of

        100 µSiemens/cm. Impurities (such as ammonium and formate ions) are removed by passing the raw formamide through a mixed-bed resin containing specific strong ion-exchange functional groups (cationic and anionic). Finally, alkaline EDTA (200 mM, to minimize the addition of water) is added to the deionized formamide to stabilize it and to facilitate the electrokinetic injection of DNA. For further details, see Why keep formamide 'dry'? and review ABI Publication 4315832C.

        Why keep formamide 'dry'?

        Store formamide in the freezer to prevent absorption of water when expecting to use formamide repeatedly during the day, you may keep a small amount in the refrigerator. Water reacts slowly with formamide to produce formic acid (methanoic acid) and ammonia. As shown in Formamide_water.jpg, the ionic products of this reaction cause two problems:

        Why restrict freeze-thaw cycles for formamide?

        Store formamide in the freezer to prevent absorption of water when expecting to use formamide repeatedly during the day, you may keep a small amount in the refrigerator. Water reacts slowly with formamide to produce formic acid (methanoic acid) and ammonia.

        Excerpt from page 82 of the DNA Fragment Analysis by CE Chemistry Guide.pdf (PN 4474504, Revision B):


        Fried, M. & Crothers, D.M. Equilibria and kinetics of Lac repressor-operator interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res. 9, 6505–6525 (1981).

        Garner, M.M. & Revzin, A. A gel-electrophoresis method for quantifying the binding of proteins to specific DNA regions: application to components of the Escherichia coli lactose operon regulatory system. Nucleic Acids Res. 9, 3047–3060 (1981).

        Fried, M.G. Measurement of protein-DNA interaction parameters by electrophoresis mobility shift assay. Electrophoresis 10, 366–376 (1989).

        Garner, M.M. & Revzin, A. The use of gel-electrophoresis to detect and study nucleic acid-protein interactions. Trends Biochem. Sci. 11, 395–396 (1986).

        Fried, M.G. & Crothers, D.M. Equilibrium studies of the cyclic-AMP receptor protein-DNA interaction. J. Mol. Biol. 172, 241–262 (1984).

        Lohman, T.M., Dehaseth, P.L. & Record, M.T. Pentalysine-deoxyribonucleic acid interactions: a model for the general effects of ion concentrations on the interactions of proteins with nucleic acids. Biochemistry 19, 3522–3530 (1980).

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