Protein Protocols & Applications

What is a protein?

The word protein is derived from the Greek proteios, meaning “of the first rank”. The term was coined in 1838 by the Swedish scientist Jöns Berzelius, to reflect the importance of this group of molecules.

A stretch of DNA called a gene carries the information required to build a protein. It is believed that there are between 20,000 and 25,000 genes in the human genome (1), but over 1 million proteins in the human proteome (2), making proteins the most abundant class of all biological molecules. The difference between the number of genes and proteins is due the fact that one gene is able to give rise to more than one protein, and that once produced, proteins can be chemically modified (usually by other proteins) to change their properties and activities.

The building blocks of proteins are amino acids. There are twenty naturally occurring amino acids (see table The naturally occurring amino acids) from which all natural proteins are constructed. All twenty are based on a common structure and differ in the chemical properties of their so-called side-chains. Some (e.g., tryptophan and phenylalanine) are strongly hydrophobic, while others (e.g., lysine and aspartic acid) carry an ionic charge at physiological pH, making them hydrophilic. Amino acids are linked together by peptide bonds to form protein chains. The sequence of amino acids in a protein and the way the protein chain is folded determine its properties.

The advances made in molecular biology over the past few decades have greatly improved the study of proteins. Previously, the only way to obtain a specific protein was to purify it from the natural source, a procedure that was often extremely inefficient and time-consuming. With the advent of recombinant molecular biological techniques it is possible to clone the DNA that encodes the protein of interest into an expression vector and express the protein in bacteria, often E. coli. The universality of the genetic code that translates a DNA sequence into a protein allows proteins from any organism to be expressed quickly and in large amounts.

This section describes procedures for expression, analysis, detection, and assay of proteins.

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Expression of proteins in E. coli

Expression of a recombinant protein can be approached in general by constructing a plasmid that encodes the desired protein, introducing the plasmid into the required host cell, growing the host cells and inducing protein expression, and then lysing the cells, purifying the protein, and performing SDS-PAGE analysis to verify the presence of the protein (see figure Generation of recombinant proteins).

The protocols and recommendations given in the DNA section of this online guide for the handling and transformation of E. coli are also valid for the production of recombinant proteins. With careful choice of host strains, vectors, and growth conditions, most recombinant proteins can be cloned and expressed at high levels in E. coli. Optimal growth and expression conditions for the protein of interest should be established with small-scale cultures before large-scale protein purification is attempted. 

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Basic principles

Culture media

The media of choice for the growth of E. coli cells containing an expression plasmid are LB medium and its modifications, 2x YT, or Super Broth, each containing the relevant selective antibiotic(s). Initially, it is advisable to try expression in all three media in parallel, and to do a time course analysis to monitor growth and expression after induction. Striking differences between the level of expression in different media and at different times are often observed.

Maintenance of the expression plasmid

Poor plasmid maintenance in the cells can lead to low expression levels. Ampicillin is an unstable antibiotic and is rapidly depleted in growing cultures due in part to the b-lactamase secreted by resistant bacterial cells. 

It is important to check plasmid levels by plating cells from the expression culture on plates with and without ampicillin. If the stability of the expression construct is a problem, the cultures should be grown in the presence of 200 µg/ml ampicillin, and the level should be maintained by supplementing ampicillin during long growth periods. Alternatively, the cultures may be grown in the presence of carbenicillin, a more stable b-lactam, at 50 µg/ml (see Antibiotics in the DNA section of this Protocols and Applications Guide).

Small-scale expression cultures

Small-scale expression and purification experiments are highly recommended and should be performed before proceeding with a large-scale preparation. In many cases, aliquots of the cells can be lysed in a small volume of sample buffer and analyzed directly by SDS-PAGE. The use of small expression cultures provides a rapid way to judge the effects of varied growth conditions on expression levels and solubility of recombinant proteins. Expression levels vary between different colonies of freshly transformed cells, and small-scale preparations permit the selection of clones displaying optimal expression rates.  

Induction of protein expression

The method used for induction of protein expression is dependent on the plasmid vector and E. coli strain used. Protein expression can be induced by a raising of the incubation temperature or by the addition of an inducing chemical such as isopropyl-b-D-thiogalactoside (IPTG) to the culture medium. Details of induction methods and the plasmids they relate to can be found in standard molecular biology texts (3, 4).

Time-course analysis of protein expression

To optimize the expression of a given protein construct, a time-course analysis by SDS-PAGE (see Protein analysis: SDS-PAGE) of the level of protein expression is recommended. Intracellular protein content is often a balance between the amount of soluble protein in the cells, the formation of inclusion bodies, and protein degradation. By checking the protein present at various times after induction, the optimal induction period can be established (see figure Time-course analysis of protein expression).

Colony blots

We recommend the colony-blot procedure (see Preparation of colony blots and figure Colony-blot procedure) to identify clones expressing a protein and to distinguish semi-quantitatively between expression rates. This can be an advantage for selecting clones after transformation, since freshly transformed colonies may differ significantly in their expression rates. Using this method, colonies subsequently found to be expressing proteins at rates as low as 0.1 to 0.5 mg/liter are easily distinguished from colonies that do not express protein.

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Preparation of colony blots

Chemiluminescent substrates are not recommended for use with colony blots.

Materials

Prepare the buffers, reagents, and agar plates. See tables 10% SDS, Denaturing solution, Neutralization solution, and 20x SSC. See also Agar plates.

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Preparation: Prepare LB medium according to the composition given in the table “LB media”. Just before autoclaving, add 15 g agar per liter and mix. After autoclaving, swirl the medium gently to distribute the melted agar evenly throughout the solution. Take care that the hot liquid does not boil over when swirled.

Tip: Cool autoclaved agar medium to below 50°C (when you can hold it comfortably) before adding heat-sensitive antibiotics and nutrients. Mix thoroughly to obtain an even concentration throughout the medium before pouring.

Tip: Pour plates in a laminar-flow hood or, if no hood is available, on a cleaned bench surface next to a Bunsen. Use 30–35 ml medium per standard 90 mm petri dish (~30 plates per liter of medium).

After pouring plates, any air bubbles may be removed by passing the flame of a Bunsen burner briefly over the surface. Do not linger with the flame as this may destroy antibiotics in sections of the plates.

Dry plates either directly after solidification or just before use by removing the lids and standing the plates in a laminar-flow hood for 1 hour. Alternatively, if you do not have access to a hood, plates can be dried with the covers slightly open in a 37°C incubator for 30 min, or left upside down with lids on at room temperature for 2–3 days.

Tip: Store plates inverted at 4°C in a dark room or wrapped in aluminum foil to preserve light-sensitive antibiotics. Do not store for longer than 3 months as antibiotics may degrade.

Procedure

1. Plate freshly transformed cells on LB-agar plates containing the appropriate antibiotics, and incubate overnight.
   Tip: After spreading the transformation mix, dry the plates inverted with the lids slightly open until small wrinkles develop on the surface of the agar. To prevent smearing, incubation should not be started until all of the liquid has been absorbed into the agar.
   Tip: To avoid expression of toxic proteins in the absence of an inducer (a result of “leaky” promoters) and to maintain plasmid stability, incubation can be carried out at 30°C.
   Tip: If the expressed protein is not toxic and the plasmids are stable, incubation can be carried out at 37°C, but care should be taken that the colonies do not become too large.
2. Remove the plates from the incubator, open lids slightly, and allow any condensation to dry for 10 min.
3. Place a dry, numbered nitrocellulose filter on the agar surface in contact with the colonies, taking care not to introduce air bubbles.
   Tip: Hold the filter on opposite sides with blunt-ended forceps, and lower gently onto the agar surface, making contact first along the middle and then lowering (but not dropping) the sides.
   Tip: Number filters with a water-resistant marking pen or pencil. 
4. Using a syringe needle, pierce the filter and agar at asymmetric positions to facilitate proper alignment following detection. Grip filter on the sides with blunt-ended forceps, and peel it off in one movement. 
5. Transfer filter (colony side up) to a fresh LB-agar plate and induce expression, e.g., by using a plate containing antibiotics and 250 µM IPTG (see “Induction of protein expression”). Avoid introducing air bubbles.
   Tip: Hold the filter on opposite sides with blunt-ended forceps, and lower gently onto the agar surface, making contact first along the middle and then lowering (but not dropping) the sides. 
6. Incubate plates for 4 h at 37°C. Place master plates in a 30°C incubator for 4 h to allow colonies to regrow.
7. Prepare a set of polystyrene dishes for colony lysis and binding of protein to the filters. Each dish should contain a sheet of Whatman 3MM paper soaked with the following solutions:
   Dish 1. 10% SDS solution
   Dish 2. Denaturing solution
   Dish 3. Neutralization solution
   Dish 4. Neutralization solution
   Dish 5. 2x SSC
   Note: Discard excess fluid so that paper is moist but not wet. Excess liquid promotes colony swelling and diffusion and will result in blurred signals. 
8. Place the nitrocellulose filters (colony side up) on top of the paper in each of these dishes, taking care to exclude air bubbles (colonies above air bubbles will not lyse properly and will generate a higher background in the final staining step). Incubate sequentially in the dishes (prepared in step 7), at room temperature as follows:
   Dish 1. 10% SDS solution 10 min
   Dish 2. Denaturing solution 5 min
   Dish 3. Neutralization solution 5 min
   Dish 4. Neutralization solution 5 min
   Dish 5. 2x SSC 15 min
9. Continue with the protocol for immunodetection using a chromogenic substrate (see “Immunodetection using a chromogenic method”).
   Tip: Due to the problem of high background, protocols using chemiluminescent substrates are not recommended for detection after colony blotting.
   Note: At times there is only a slight difference between colonies which express protein and those that do not.
   Tip: Shorter staining times are required with this procedure. A 2–3 min staining time is usually sufficient, but it is very important to monitor color development at this stage.
   Tip: If it is still difficult to differentiate between positive clones and background, the cause of the high background should be determined. 

The following controls should be included:

• A plate of host bacteria without the expression plasmid
• A plate of host bacteria harboring the expression plasmid without the insert 
• A colony-blot treated only with secondary antibody prior to detection 
• A positive control expressing the protein of interest, if possible

Growth of standard expression cultures (100 ml)

1. Inoculate 10 ml culture medium containing relevant antibiotics in a 50 ml flask. Grow the cultures overnight at 37°C.
2. Inoculate 100 ml prewarmed medium (with antibiotics) with 5 ml of the overnight culture and grow at 37°C with vigorous shaking until an OD600 of 0.6 is reached (30–60 min).
   Tip: Take a 1 ml sample immediately before induction. This sample is the noninduced control. Pellet cells and resuspend them in 50 µl 5x SDS-PAGE sample buffer (see table 5x SDS-PAGE sample buffer). Store at –20°C until SDS-PAGE analysis. 
3. Induce expression (e.g., by adding IPTG to a final concentration of 1 mM). 
4. Incubate the cultures for an additional 4–5 h.
   Tip: Collect a second 1 ml sample. This sample is the induced control. Pellet cells and resuspend them in 100 µl 5x SDS-PAGE sample buffer. Freeze and store the sample at –20°C until SDS-PAGE analysis.
   Tip: If expressing a protein for the first time, take a 1 ml sample every hour and treat as above to produce a time-course of expression. 
5. Harvest the cells by centrifugation at 4000 x g for 20 min. 
6. Freeze the cells in dry ice–ethanol or liquid nitrogen, or store cell pellet overnight at –20°C

Culture growth for preparative production (1 liter)

1. Inoculate 20 ml culture medium containing the relevant antibiotics. Grow overnight at 37°C with vigorous shaking.
2. Inoculate a 1 liter culture 1:50 with the noninduced overnight culture. Grow at 37°C with vigorous shaking until an OD₆₀₀ of 0.6 is reached.
   Tip: Take a 1 ml sample immediately before induction. This sample is the noninduced control. Pellet cells and resuspend in 50 µl 5x SDS-PAGE sample buffer (see table 5x SDS-PAGE sample buffer). Store at –20°C until SDS-PAGE analysis. 
3. Induce expression (e.g., by adding IPTG to a final concentration of 1 mM). 
4. Incubate the culture for an additional 4–5 h.
   Tip: Collect a second 1 ml sample. This is the induced control. Pellet cells in a microcentrifuge tube and resuspend in 100 µl 5x SDS-PAGE sample buffer. Store at –20°C until SDS-PAGE analysis. 
5. Harvest the cells by centrifugation at 4000 x g for 20 min. Freeze the cells in dry ice–ethanol or liquid nitrogen, or store cell pellet overnight at –20°C.

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Protein purification

The expression and purification of recombinant proteins facilitates production and detailed characterization of virtually any protein. Although a wide variety of heterologous expression systems have been developed and are currently used to produce recombinant proteins, the purification of the proteins obtained can still be problematic. Classical purification procedures can be employed, but in most cases recombinant DNA techniques permit the construction of fusion proteins in which specific affinity tags are added to the protein sequence of interest; the use of these affinity tags simplifies the purification of the recombinant fusion proteins by employing affinity chromatography methods. Ideally a tag should be small, and have a minimal effect on the structure, activity, and properties of the recombinant protein. Different affinity tags have different sizes and properties. The 6xHis tag has a size of just 0.84 kDa, compared to 26 kDa for the glutathione S-transferase tag, 30 kDa for protein A, and 40 kDa for maltose-binding protein. The FLAG tag consists of just 8 amino acids, but is highly immunogenic, which means that the FLAG tag must be removed before a recombinant protein can be used to produce antibodies. In contrast, the His tag has extremely low immunogenicity and rarely interferes with protein structure or function, making His-tagged proteins suitable for all kinds of downstream applications without cleaving the tag.

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Protein analysis: SDS-PAGE

Principle of SDS-PAGE analysis

SDS polyacrylamide gel electrophoresis (SDS-PAGE) involves the separation of proteins based on their size. By heating the sample under denaturing and reducing conditions, proteins become unfolded and coated with SDS detergent molecules, acquiring a high net negative charge that is proportional to the length of the polypeptide chain. When loaded onto a gel matrix and placed in an electric field, the negatively charged protein molecules migrate towards the positively charged electrode and are separated by a molecular sieving effect. After visualization by a protein-specific staining technique, the size of a protein can be estimated by comparison of its migration distance with that of a standard of known molecular weight. It is also possible to blot the separated proteins onto a positively charged membrane and to probe with protein-specific antibodies in a procedure termed western blotting (see Western transfer).

Acrylamide concentration

The concentration of acrylamide used for the gel depends on the size of the proteins to be analyzed. Low acrylamide concentrations are used to separate high molecular weight proteins, while high acrylamide concentrations are used to separate proteins of low molecular weight (see table Compositions and separation properties of SDS-PAGE gels). Improved resolution of protein bands is achieved by the use of a discontinuous gel system having stacking and separating gel layers.

Preparation of dilute or salt-containing samples for SDS-PAGE

Acid precipitation of proteins (see below) can be carried out prior to SDS-PAGE analysis in order to concentrate dilute samples or to remove high concentrations of salts that may interfere with the SDS-PAGE procedure.

TCA precipitation of proteins

1. Dilute samples to 100 µl; add 100 µl 10% trichloroacetic acid (TCA).
2. Leave on ice for 20 min; centrifuge for 15 min in a microcentrifuge.
3. Wash pellet with 100 µl of ice-cold ethanol, dry, and resuspend in 5x SDS-PAGE sample buffer (see “see table 5x SDS-PAGE sample buffer). Boil for 7 min at 95°C, and then load samples immediately onto a gel for SDS-PAGE.

Separation of proteins by SDS-PAGE

Materials required

• Gel apparatus and electrophoresis equipment
• 30% acrylamide/0.8% bis-acrylamide stock solution (see table 30% acrylamide/0.8% bis-acrylamide stock solution)
• 2.5x separating gel buffer (see table 2.5x separating gel buffer)
• 5x stacking gel buffer (see table 5x stacking gel buffer)
• TEMED (N,N,N’,N’-tetramethylethylenediamine)
• 10% ammonium persulfate
• Butanol
• 5x electrophoresis buffer (see table 5x electrophoresis buffer)
• 5x SDS-PAGE sample buffer (see table 5x SDS-PAGE buffer)
• Protein samples

IMPORTANT: Acrylamide is a potent neurotoxin and is absorbed through the skin. Take appropriate safety measures particularly when weighing solid acrylamide/bis-acrylamide, and also when working with the solutions and gels.

Tip: Use only high-quality reagents and water for SDS-PAGE. Gel buffers and self-prepared acrylamide/bis-acrylamide stock solutions should be filtered, degassed, and stored at 4°C.

1. Assemble gel plates with spacers according to the manufacturer’s instructions. 
   Tip: The plates should be thoroughly cleaned and dried before use.
2. Mark the level to which the separating gel should be poured — a few millimeters below the level where the wells will be formed by the comb.
3. Mix the following in a beaker or similar vessel (for a 12% acrylamide 8 x 8 or 8 x 10 cm, 1 mm thick, minigel). 
   2.2 ml 30% acrylamide/0.8% bis-acrylamide stock solution
   2.2 ml 2.5x separating gel buffer
   1.1 ml distilled water
   5 µl TEMED
   The volumes of acrylamide/bis-acrylamide solution and water should be adjusted according to the percentage acrylamide required (dependent on the size of protein to be separated; see table Compositions and separation properties of SDS-PAGE gels).
   Tip: The size of the gel apparatus used will determine the volumes of gel solutions necessary.
4. Just before pouring, add 50 µl 10% ammonium persulfate, and mix well. Pour the gel between the assembled gel plates to the level marked in step 2. Overlay with butanol.
   Tip: Water can be used instead of butanol when using apparatus that may be damaged by the use of butanol — see the manufacturer’s instructions.
   Tip: As soon as ammonium persulfate is added, the gel should be poured quickly before the acrylamide polymerizes.
   Tip: Prepare ammonium persulfate solution freshly each time it is required
5. After polymerization is complete (around 20 min), pour off butanol, rinse with water and dry.
   Tip: Water remaining on the plates can be removed using pieces of filter paper.
6. For the stacking gel, mix the following:
   0.28 ml 30% acrylamide/0.8% bis-acrylamide stock solution
   0.33 ml 5x stacking gel buffer
   1 ml distilled water
   2 µl TEMED
7. Just before pouring, add 15 µl 10% ammonium persulfate, and mix well. Pour on top of the separating gel. Insert comb, avoiding introduction of air bubbles.
   Tip: As soon as ammonium persulfate is added the stacking gel should be poured quickly, before the acrylamide polymerizes.
   Tip: With a marker pen, mark and/or number the positions of the wells before removing the comb. This aids loading of samples.
8. After the stacking gel polymerizes (around 10 min), the gel can be placed in the electrophoresis chamber. Fill the chamber with electrophoresis buffer and remove the comb.
9. Before loading, add 1 volume 5x SDS-PAGE sample buffer to 4 volumes of protein sample (i.e., add 2 µl sample buffer to 8 µl sample giving a final volume of 10 µl). Vortex briefly and heat at 95°C for 5 min.
   Tip: During heating at 95°C, release pressure build up in tubes by briefly opening lids, or piercing a small hole in the lid with a needle. After heating, samples should be briefly centrifuged and vortexed.
10. Load samples and run gel. For electrophoresis conditions refer to the recommendations provided by the manufacturer of the apparatus.
    Tip: Before loading the samples, rinse out wells with 1x electrophoresis buffer using a suitable syringe and needle.
    Tip: Load empty wells with 1x SDS-PAGE sample buffer to ensure that sample lanes do not spread out. Tip: Ensure that the electrodes are correctly connected. The proteins will migrate towards the positive (labeled +, usually red) electrode.
    Tip: Running the gel until the bromophenol blue dye reaches the bottom edge usually gives a satisfactory spread of protein bands.

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Visualization of proteins in SDS-PAGE gels

Visualization of protein bands is carried out by incubating the gel with a staining solution. The two most commonly used methods are Coomassie and silver staining. Silver staining is a more sensitive staining method than Coomassie staining, and is able to detect 2–5 ng protein per band on a gel. Many protocols are available but in order to increase reproducibility, use of a commercially available kit is recommended. Silver staining of proteins depends on the reaction of silver with sulfhydryl or carboxyl moieties in proteins and is therefore not quantitative, with some proteins being poorly stained by silver. In addition, after silver staining the protein becomes oxidized and cannot be used for further applications, such as sequencing. Coomassie staining, though less sensitive, is quantitative and Coomassie-stained proteins can be used for downstream applications.

Coomassie staining

Materials required

• Coomassie staining solution (see table Coomassie staining solution)
• Destaining solution (see table Destaining solution) 
• SDS polyacrylamide gel containing separated proteins (see Separation of proteins by SDS-PAGE)

1. Incubate the gel in Coomassie staining solution for between 30 min and 2 h with gentle shaking.
   Tip: Coomassie Brilliant Blue R reacts nonspecifically with proteins.
2. Gently agitate the stained gel in destaining solution until the background becomes clear (1–2 h).
   Tip: A folded paper towel placed in the destaining bath will soak up excess stain and allow the reuse of destaining solution.

After destaining the proteins appear as blue bands against a clear gel background. Typically, bands containing 50 ng protein can be visualized.

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Western blotting

Following electrophoresis, proteins in a polyacrylamide gel can be transferred to a positively charged membrane (e.g., Schleicher and Schuell BA85) in a buffer-tank–blotting apparatus or by semi-dry electroblotting as described below.

With the semi-dry electroblotting method, the gel and membrane are sandwiched between two stacks of filter paper that have been pre-wet with transfer buffer. The membrane is placed near the anode (positively charged), and the gel is placed near the cathode (negatively charged). SDS-coated, negatively charged proteins are transferred to the membrane when an electric current is applied. With the tank-blotting method, a blotting cassette is submerged in a tank for blotting (see figure Tank- and semi-dry blotting methods).Tank blotting can be performed over extended periods since the buffer capacity is far greater than that with semi-dry transfer systems. Results obtained with the tank-blotting method are typically better, with more efficient transfer, particularly of large proteins. Transfer efficiency can be checked by staining proteins on the membrane using Ponceau S (see Ponceau S staining). Once transferred to the membrane, the proteins can be probed with epitope-specific antibodies or conjugates.

Materials required

• Transfer apparatus 
• Filter paper (e.g., Whatman 3MM) 
• Positively-charged membrane (e.g., Schleicher and Schuell BA85) 
• SDS polyacrylamide gel containing separated proteins (see Separation of proteins by SDS-PAGE) 
• Transfer buffer (semi-dry or tank-blotting) (see table Semi-dry transfer buffer or Tank-blotting transfer buffer)

1. Cut 8 pieces of filter paper and a piece of membrane to the same size as the gel.
   Tip: To avoid contamination, always handle the filter paper, membrane, and gel with gloves.
2. Incubate membrane for 10 min in semi-dry or tank-blotting transfer buffer.
3. Soak filter paper in semi-dry or tank-blotting transfer buffer. Proceed to step 4 if performing semi-dry transfer or step 5 if performing tank blotting.
4. Semi-dry transfer: Avoiding air bubbles, place 4 sheets of filter paper on the cathode (negative, usually black), followed by the gel, the membrane, 4 sheets of filter paper, and finally the anode (positive, usually red).
5. Tank-blotting: Avoiding air bubbles, place 4 sheets of filter paper on the fiber pad, followed by the gel, the membrane, 4 sheets of filter paper, and finally the second fiber pad.
   Tip: Air bubbles may cause localized nontransfer of proteins. They can be removed by gently rolling a Pasteur pipet over each layer in the sandwich.
6. Carry out the transfer procedure. For current, voltage, and transfer times specific to your apparatus, consult the manufacturer’s instructions.
   Tip: Time of transfer is dependent on the size of the proteins, percentage acrylamide, and gel thickness. Transfer efficiency should be monitored by staining (see below). The field strength required is determined by the surface area and thickness of the gel: 0.8 mA/cm² is a useful guide (1 h transfer).
7. After transfer, mark the orientation of the gel on the membrane.

Ponceau S staining

1. Incubate membrane in Ponceau S staining solution (see table Ponceau S staining solution) with gentle agitation for 2 min.
2. Destain in distilled water until bands are visible.
3. Check that proteins of different sizes have been transferred uniformly to the membrane. Hydrophobic proteins may be more efficiently transferred by increasing the percentage of methanol in the transfer buffer.
4. Mark membrane using a suitable pen (i.e., one not containing water-soluble ink) or pencil, or cut as desired.

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Dot blots

Dot blotting is a simple, convenient method for detection of proteins in crude lysates or solutions without the need for separation by SDS-PAGE. This method is especially useful as a simple control because it avoids problems that may be due to the western transfer process. Any components that interfere with binding or bind nonspecifically, however, will not be spatially separated from the protein and will interfere with the intensity of signals. Suitable controls should always be employed to compensate for this.

Materials required 

• Nitrocellulose membrane (e.g., Schleicher and Schuell BA85) 
• Protein samples 
• Dilution buffer for native or denaturing conditions (see table Dilution buffer for denaturing conditions or Dilution buffer for native conditions)

1. Dilute protein samples in buffer to final protein concentrations of 1–100 ng/µl.
   Tip: The protein of interest is diluted in dilution buffer for denaturing conditions, dilution buffer for native conditions, or another preferred buffer.
2. Apply 1 µl samples of diluted protein directly onto membrane. It is also possible to use crude cell lysate and apply 1 µl samples with an estimated concentration of 1–100 ng/µl protein.
   Note: Under native conditions especially, the antibody epitope must be at least partially exposed to allow antibody binding. In most cases diluting the protein with buffer containing denaturing reagents will increase epitope exposure and give better results.
   Tip: To differentiate between nonspecific and positive signals, an extra sample containing 1 µl of a cell extract of the host strain without plasmid (or other suitable control) should also be applied to the membrane and treated together with the protein of interest.
3. After applying the samples, the membrane should be dried for a short time at room temperature before proceeding with the detection process.
   Tip: For larger sample volumes, suitable equipment is available from several suppliers.
4. Proceed with immunodetection (see Immunodetection using a chemiluminescent detection method or Immunodetection using a chromogenic detection method).

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Protein detection: Specific antibody-mediated detection of proteins on a membrane

Working with antibodies

Antibodies are proteins synthesized by an animal in response to the presence of a foreign substance (antigen). By injecting an antigen into an animal, after a certain time, antibodies of a class termed IgG (immunoglobulin G) that react specifically with the introduced protein can be harvested from the animal’s serum. Each antibody has a specific affinity for a particular region of the antigen. This region is termed an epitope. The antibody–epitope interaction can be utilized for highly specific and sensitive detection of a protein that has been immobilized on a membrane, in a process termed immunodetection. The antibody that binds specifically to the protein of interest is termed the primary antibody and is often obtained from rabbits or mice. The primary antibody is applied to the membrane and allowed to bind to the target protein. In order to locate the primary antibody (and therefore the protein of interest), a secondary antibody is required. The secondary antibody recognizes and binds to all IgG antibodies from another animal species. It is important that the secondary antibody used in an experiment is directed against IgGs from the species of origin of the primary antibody. For example, if the primary antibody was generated in a mouse, then a goat anti-mouse secondary antibody can be used for detection. The secondary antibody is usually chemically coupled to a reporter, which allows detection and visualization of the antibody. Fluorescing molecules, or enzymes that produce colored or luminescent reaction products, are typically used as reporter groups. A primary antibody chemically coupled to a reporter enzyme is termed a conjugate, and can be used for direct detection without the use of a secondary antibody. 

Detection of a protein on a membrane

After protein transfer from an SDS-PAGE gel to a membrane (see Western transfer), the remaining protein-free sites on the membrane must be blocked. This prevents the primary or secondary antibody from binding directly to the membrane and giving rise to a high background signal. Several blocking reagents are in common use, including nonfat dried milk, BSA, and casein. After blocking, the primary antibody is added and allowed to bind to the protein (see figure Immunodetection of a protein immobilized on a membrane). After washing (which removes nonspecifically bound antibody), the secondary antibody is added, to detect where the primary antibody has bound. After another wash step, the location of the secondary antibody, (and therefore the primary antibody and the protein of interest) is determined by adding a substrate for the enzyme conjugated to the secondary antibody. Substrates are available that give rise to a colored compound (chromogenic detection), or to the emission of light (chemiluminescent detection), at the reaction site. The use of an antibody that reacts specifically with an epitope commonly introduced into a recombinant protein eliminates the need for a protein-specific antibody, and allows the use of one antibody for the detection of all proteins containing this feature. Coupling a reporter enzyme directly to such antibodies eliminates the need for a secondary antibody, and delivers significant time savings. Detailed information on immunodetection procedures can be found in current molecular biology manuals (3, 4).

Immunodetection using a chemiluminescent method

Materials required

• Western blot (see Western transfer) 
• TBS buffer (see table TBS buffer) 
• TBS-Tween/Triton buffer (see table TBS-Tween/Triton buffer) 
• Blocking buffer (see table Blocking buffer)
• Primary (protein-specific) antibody stock solution 
• Secondary antibody-enzyme conjugate stock solution 
• Secondary antibody dilution buffer (see table Secondary antibody dilution buffer) 
• Chemiluminescent substrate

Note: For chemiluminescent detection, BSA does not sufficiently block nonspecific binding of the secondary antibody to the membrane, so milk powder should be used to dilute the secondary antibody. However, in some cases, dilution of antibody in a buffer containing milk powder can lead to reduced sensitivity. If this is the case, the primary antibody should be diluted in BSA solution, and the secondary antibody in milk powder solution.

Perform all incubation and wash steps on a rocking platform or orbital shaker.

1. Wash membrane twice for 10 min each time with TBS buffer at room temperature.
2. Incubate membrane for 1 h in blocking buffer at room temperature.
   Tip: Seal the vessel used for incubation with plastic film to prevent the membrane from drying out.
3. Wash membrane twice for 10 min each time in TBS-Tween/Triton buffer at room temperature.
4. Wash membrane for 10 min with TBS buffer at room temperature.
5. Incubate membrane with primary antibody solution (1/1000–1/2000 dilution of primary antibody stock solution in blocking buffer) at room temperature for 1 h.
   Tip: Make sure that the membrane is fully coated by the antibody solution. Do not allow the membrane to dry out.
   Tip: To reduce the volume of antibody required, the membrane can be sealed in a plastic bag.
6. Wash membrane twice for 10 min each time in TBS-Tween/Triton buffer at room temperature.
7. Wash membrane for 10 min in TBS buffer at room temperature.
8. Incubate the membrane with a dilution of secondary antibody in 10% nonfat dried milk in TBS for 1 h at room temperature. Dilute the secondary antibody according to the manufacturer’s recommendations.
   Tip: Ensure that your secondary antibody is directed against the species of origin of your primary antibody!
   Tip: Milk powder is needed to reduce background because BSA does not block sufficiently for the very sensitive chemiluminescent detection method.
   Tip: Use the lowest recommended concentration to avoid false signals.
9. Wash 4 times for 10 min each time in TBS-Tween/Triton buffer at room temperature.
10. Perform chemiluminescent detection reaction, cover the membrane with thin plastic wrap, and expose to X-ray film according to the manufacturer’s recommendations.
    Tip: Ensure that you use the correct chemiluminescent detection substrate, i.e., an AP substrate for AP conjugates, or an HRP substrate for HRP conjugates!
    Tip: Blots can be wrapped in plastic wrap and stored at 4°C. Protocols exist for stripping the blot, which can subsequently be reprobed with a different antibody (5).

Immunodetection using a chromogenic method

Materials required

• Western blot (see Western transfer) 
• TBS buffer (see table TBS buffer) 
• TBS-Tween/Triton buffer (see table TBS-Tween/Triton buffer) 
• Blocking buffer (see table Blocking buffer) 
• Primary (protein-specific) antibody stock solution 
• Secondary antibody-enzyme conjugate stock solution 
• Chromogenic substrate (see table Chromogenic substrates for immunoblotting procedures)

Note: 3% BSA (w/v) can be used as both the blocking buffer and the secondary antibody dilution buffer for chromogenic detection. 

Perform all incubation and wash steps on a rocking platform or orbital shaker. 

1. Follow steps 1–7 of the protocol Immunodetection using a chemiluminescent method.
2. Incubate the membrane with secondary antibody solution diluted in 3% BSA (w/v) in TBS for 1 h at room temperature. Dilute according to the manufacturer’s recommendations.
   Tip: Ensure that your secondary antibody is directed against the species of origin of your primary antibody.
   Tip: Use the lowest recommended concentration to avoid false signals.
3. Wash 4 times for 10 min each time in TBS-Tween/Triton buffer at room temperature.
4. Stain with AP or HRP staining solution until the signal is clearly visible (approximately 5–15 min). Do not shake blots during color development.
   Tip: Ensure that you use the correct chromogenic detection substrate, i.e., an AP substrate for AP conjugates, or an HRP substrate for HRP conjugates.
5. Stop the chromogenic reaction by rinsing the membrane twice with water.
6. Dry the membrane and photograph as soon as possible as the colors will fade with time. The product formed when using HRP is particularly unstable.  

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Protein assay: ELISA

Enzyme-linked immunosorbent assay (ELISA) is a method that is analogous to immunodetection of proteins on a membrane, and is used for the quantitative assay of proteins in solution. In an ELISA, proteins are immobilized on a solid support (e.g., the wells of a 96-well plate) and used as capture molecules to bind the protein that is being assayed. After a wash step to remove nonspecifically bound material, a secondary antibody — specific for the protein being assayed — is added. This secondary antibody is usually conjugated to an enzyme that allows its detection by chromogenic or chemiluminescent methods.

In one type of ELISA assay, an antibody that binds an epitope on a target protein is immobilized, and a test solution added. The immobilized antibody will capture any target protein present in the sample. A wash step removes nonspecifically bound material, and subsequently a second antibody is added that reacts with a second epitope on the protein. Alternatively, a protein can be immobilized on a solid support, and antibodies reacting with the protein can be detected and quantitated in a test solution by the addition of a secondary antibody that reacts with the primary antibody (see figure Two common types of ELISA).

Coating 96-well microplates with protein for ELISA

This procedure is used to immobilize proteins onto the inner surfaces of 96-well microplates. The proteins can then be assayed using a primary and secondary antibody in a process analogous to that used for detection of proteins on western blots.

Important notes before starting

• The ease with which proteins bind to polystyrene plates is very much dependent on the particular protein. Optimization of binding conditions is necessary. Refer to the manufacturer’s instructions. 
• As a starting point, three buffers at different pH values should be compared. 
• Binding may be carried out at 4–37°C. Successful binding may depend on the stability of the protein.

Materials required 

• Suitable 96-well microplates

Coating buffers

PBS, pH 7.2 (see table PBS, pH 7.2) 
50 mM sodium carbonate, pH 9.6 (see table Sodium carbonate, pH 9.6) 
50 mM sodium carbonate, pH 10.6 (see table Sodium carbonate, pH 10.6) 
Microplate blocking buffer(see table Microplate blocking buffer)  

1. Serially dilute the protein to be immobilized in coating buffer(s).
2. Add 200 µl of the protein solution to each well, and incubate overnight at 4°C.
3. Wash wells 4 times with PBS. Soak wells for 10–60 s per wash, and dry the wells by tapping the plate on paper towels.
4. Block wells with 250 µl of microplate blocking buffer for 2 h at room temperature (20–25°C) on a shaker platform.
   Tip: After blocking, plates can be dried overnight at 20–25°C, but sensitivity of the assay will be reduced.
   Tip: After drying, it may be possible to store the plates at 4°C for a period of time before use, but this will depend on the specific protein to be assayed.
5. Wash wells 4 times with PBS. Soak wells for 10–60 s per wash, and dry the wells by tapping the plate on paper towels.
6. Proceed with the protocol Assay of proteins with a protein specific antibody.

Assay of proteins with a protein specific antibody

Important notes before starting

• Binding of detection antibodies should be carried out for at least 1 h at room temperature. If the concentration of the protein to be detected is very low or if the epitope is partly hidden, incubation times of 2–4 h or overnight may increase sensitivity. 
• Best results will be obtained if all steps are carried out on a shaker. If there is no shaker available, incubation times should be increased (up to 2–3 h at room temperature or overnight at 4°C) or the incubation temperature should be raised to allow sufficient diffusion of molecules. 
• Antibody dilution depends on the individual antibody used. Please refer to manufacturer’s recommendations or begin at concentrations useful for western-blot or dot-blot analyses and try further dilutions. Usually primary monoclonal antibody at 0.1 µg/ml to 1 µg/ml will yield satisfactory results. Each antibody should be titrated over this range of concentrations to determine the optimal dilution.
• Suitable negative controls are essential. Assays should always be performed in parallel with samples without any proteins (lysis/dilution buffer alone, reagent blank) and with samples similar to those assayed but lacking the target protein (e.g., lysate from E. coli transformed with vector lacking the protein-encoding insert). These controls should be incubated with antibodies and the remaining assay components. Note: This protocol is intended to be used as an example. Optimal conditions for each individual protein and antibody should be determined. 
• If establishing a new assay system, the binding of the protein or antibody to the solid support should be optimized first (incubation time and amounts of protein). Primary antibody or other secondary components of the assay should be optimized afterwards.

Materials required 

• 96-well microplates coated with protein (see “Coating 96-well microplates with protein for ELISA”) 
• PBS/BSA (see table PBS/BSA) 
• PBS (see table“PBS, pH 7.2) 
• Anti-target–protein antibody 
• Secondary-antibody conjugate 
• Substrate for alkaline phosphatase or horseradish peroxidase or one of the alternative substrates for horseradish peroxidase as described in tables Phosphate–citrate buffer, pH 5.0, Substrate for alkaline phosphatase: p-Nitrophenyl Phosphate (NPP), Substrate for horseradish peroxidase: 2,2’Azino-bis[3-Ethylbenz-thiazoline-6-Sulfonic Acid] ABTS, Alternative substrate for horseradish peroxidase: o-Phenylenediamine (OPD), and Alternative substrate for horseradish peroxidase: 3,3’,5,5’-Tetramethylbenzidine (TMD. Further details about substrates can be found in the table Details of protein substrates for protein assay procedures.  

1. Add 200 µl of anti-target–protein antibody diluted 1/2000 in PBS/BSA. Cover plate, and incubate for 1–2 h at RT.
   For higher sensitivity, antibody binding can be performed overnight at 4°C.
2. Wash wells 4 times with PBS-Tween. Soak wells for 10–60 s per wash, and dry the wells by gently tapping the plate on paper towels after the wash.
3. Dilute secondary antibody in PBS/BSA according to the manufacturer’s recommendations. Add 200 µl of the diluted antibody to each well, and incubate at room temperature for 45 min.
4. Wash wells 4 times with PBS-Tween. Soak wells for 10–60 s per wash, and dry the wells by gently tapping the plate on paper towels.
5. Add 200 µl of substrate solution, and monitor color development in a microplate reader.
   Note: Substrate solution should always be prepared immediately before use.
   Tip: Monitor color development over a period of 45 min, or add 50 µl stopping reagent after a specific time and measure product. When testing a new assay system, a time-course of color development should be carried out to determine optimal development time and temperature.
   Tip: If the reaction is stopped the signal will increase slightly, depending on the substrate used, and the color will be stable for a certain period of time.

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Quantifying proteins using the Bradford method

The Bradford method is a quantitative protein assay method, based on the binding of a dye, Coomassie Brilliant Blue, to a protein sample, and comparing this binding to a standard curve generated by the reaction of known amounts of a standard protein, usually BSA.

For this assay, protein samples should be diluted in an appropriate buffer (generally the same buffer in which they are dissolved). The BSA standard curve should be prepared using the same buffer.

Materials required

• BSA standard solution (1 mg/ml) 
• Bradford assay dye reagent (available commercially, e.g., Bio-Rad Protein Assay Dye Reagent Concentrate, cat. no. 500-0006) 
• Protein dilution buffer

Note: Proteins should be diluted in the buffer in which they are dissolved. Use the same buffer to prepare the standard curve.

Note: Concentration of the BSA standard solution should be measured photometrically. A 1 mg/ml solution of BSA should have an A₂₈₀ of 0.66.

1. Prepare a standard curve by pipetting together carefully the solution volumes listed in the table “Standard curve samples for Bradford protein assay” corresponding to 0, 0.05, 0.1, 0.2, 0.3, 0.4, and 0.5 mg/ml BSA. The end volume of all samples should be 200 µl.
2. Dilute an aliquot of the dye reagent concentrate 1:5 with distilled water. Pipet 20 µl of each BSA dilution into a plastic cuvette, add 1 ml diluted dye reagent, mix well, and incubate for 5 min at room temperature. 
   Note: Stored in the dark, diluted dye reagent is stable for 2 weeks. Write the date of preparation on the bottle and cover with aluminum foil.
3. Measure the OD at 595 nm for each sample, and plot the standard curve.
4. Prepare a second standard curve by pipetting together carefully the solution volumes listed in the table Standard curve samples for Bradford protein assay corresponding to 0, 0.2, 0.4, 0.6, 0.8, and 1.0 mg/ml BSA. The end volume of all samples should be 200 µl.
5. Dilute an aliquot of the dye reagent concentrate 1:5 with distilled water. Pipet 20 µl of each BSA dilution into a plastic cuvette, add 1 ml diluted dye reagent, mix well, and incubate for 5 min at room temperature.
6. Measure the OD at 595 nm for each sample, and plot the standard curve.
7. Prepare dilutions of the protein sample of interest and test 20 µl aliquots in duplicate as above. It is important that the protein samples to be tested are handled in exactly the same manner as the samples used in generating the standard curves. Calculate the protein concentration of the test samples by comparing the OD₅₉₅ with the standard curve(s), taking into account dilution factors (see figure Standard curve generated using the Bradford method).

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References

1. International Human Genome Sequencing Consortium (2004) Finishing the euchromatic sequence of the human genome. Nature. 431, 931.
2. Jensen, O.N. (2004) Modification-specific proteomics: Characterization of post-translational modifications by mass spectrometry. Curr. Opin. Chem. Biol. 8, 33.
3. Ausubel, F.M., et al. (1999) Current Protocols in Molecular Biology. New York: John Wiley and Sons.
4. Sambrook, J. and Russell, D. (2001) Molecular Cloning: A Laboratory Manual. 3rd ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
5. Kaufmann S.H., Ewing, C.M., and Shaper, J.H. (1987) The erasable Western blot. Anal. Biochem. 161, 89.

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