Peer Reviewed: Detecting Proteins Separated by 2-D Gel

Detecting

Proteins Separated by 2-D Gel Electrophoresis

Thierry Rabilloud Bio Energétique Cellulaire et Pathologique Département de Biologie Moléculaire et Structurale (France)

48 A

A N A LY T I C A L C H E M I S T R Y / J A N U A R Y 1 , 2 0 0 0

etecting proteins in a two-dimensional (2-D) gel is a way of differentiating the proteins (i.e., the analytes) from the background medium (i.e., the polyacrylamide gel) based on the differences in chemical reactivity of the proteins and the medium. Detection is achieved by binding a label—such as a radioactive amino acid, a fluorophore, or a dye molecule—to the proteins or to their microenvironment in the gel. (In the context of this article, “dye” refers to a nonfluorescent molecule detected by absorption of visible light.) In choosing the chemical entity to bind to the proteins, the key issues of sensitivity (detection threshold), linearity, homogeneity (i.e., variation from one protein to another), and reproducibility must be considered. Homogeneity and reproducibility are generally not so important for 1-D electrophoresis, in which detection is carried out in parallel for the different samples loaded on the lanes of the 1-D gels. The situation is quite different in 2-D gels, in which there is only one sample per gel because the abundance of different protein spots separated on one gel have to be compared with different gels. Another consideration is that 2-D electrophoresis is used more and more as a micropreparative tool to isolate small quantities of proteins and make them amenable to characterization by various methods such as MS or Edman sequencing. Consequently, the detection methods used must be chemically compatible with the characterization process. For example, acylation of amines, which is a very convenient way of grafting a label to proteins, is completely unacceptable for Edman sequencing because it induces N-terminal blocking. Moreover, very important constraints are imposed by the separation method itself. Two-dimensional electrophoresis uses isoelectric focusing (IEF) in polyacrylamide gels as the first dimension and sodium dodecyl sulfate electrophoresis in polyacrylamide gels (SDS PAGE) as the second dimension. Consequently, the label can be bound to the proteins either by noncovalent interactions or by covalent bonds at four stages of the separation—before IEF, between IEF and SDS electrophoresis (second dimension), after SDS electrophoresis, or after transfer of the separated proteins onto an inert membrane (blotting). At each of these stages, three major types of labels can be used: radioactive labels, organic dyes, and fluorophores. Another type of staining uses transition metal or heavy metal ions, but is limited to post 2-D electrophoresis detection, because this protein metal-ion affinity is not compatible with electrophoretic separation conditions. In this article, a

D

“combined rationale” approach, based on the type of label and the point of label-fixation, presents the most important detection methods currently in use, their main features (sensitivity, linearity, reproducibility, and interfacing with subsequent analysis methods), and future prospects.

Affixing the label before IEF At this stage, labeling is theoretically very convenient because it can be done in a minimal volume with a high concentration of reactants; any unreacted label will be removed during the separation. However, the IEF step is typically performed under strongly denaturing conditions (i.e., chaotropes and detergents), so that noncovalent binding will be disrupted. Consequently, only covalent binding of the label can be used at this stage. Unfortunately, covalent labeling of proteins prior to IEF must comply with very stringent requirements imposed by the chemistry of the proteins and the process of IEF. In fact, IEF separates the proteins according to the pI, which is the pH at which the positive (i.e., amine groups of lysine and the amino terminus and guanidinium groups of arginine) and negative (i.e., carboxyl groups of the C-terminus, aspartate, glutamate, posttranslationally added phosphate, and sulfonic or sulfate groups) charges of the protein compensate exactly. Consequently, any labeling method that alters the charge state of the protein, either by removing a charged group or by adding a spurious one, is not compatible with 2-D electrophoresis. For example, amine acylation is quite convenient, but it should be avoided because the positive charge of the amine is lost. Amine or thiol alkylation with reactive acidic dyes such as Remazol must also be avoided because the proteins will gain additional negative charges. These constraints strongly limit the scope of labeling reactions that can be applied before IEF. In addition, because the number of chemically reactive sites on protein molecules is limited, only very sensitive labels must be introduced by covalent grafting. Thus, covalent grafting of dyes prior to IEF is not used because of poor sensitivity (1) and because most dyes are not electrically neutral. Labels are therefore limited to neutral fluorophores and radioactive compounds. However, covalent labeling prior to IEF also offers the possibility of multiplexing, which involves labeling several different samples with different probes, mixing the labeled samples prior to 2-D electrophoresis, and analyzing the relative intensities of the different labels within a single 2-D gel. This process alleviates reproducibility problems often encountered in the analysis of multiple 2-D gels.

A guide

to choosing the right stain, dye, and fluorescent or radioactive label.

J A N U A R Y 1 , 2 0 0 0 / A N A LY T I C A L C H E M I S T R Y

49 A

Covalent fluorescent labeling Intense light absorption, high quantum yield, large Stokes shifts, and limited fading are additional chemical constraints that further limit the number of labels. For example, fluorescein derivatives, which have an excellent light absorption and quantum yield, cannot be used prior to IEF because the fluorescein molecule carries negative charges. Thus, pre-IEF fluorescent labeling has been limited for a long time to thiol alkylation with electrically neutral probes (2). Unfortunately, these probes generally offer limited sensitivity because they have limited light absorption and poor quantum yields in the water-containing solvents required by polyacrylamide gels. Using cyanine-based probes (3), in which attachment to the protein molecules is achieved via amine acylation, improves the process. In this case, the positive charge lost on the protein is compensated by the positive charge brought by the quaternary ammonium of the cyanine probe. Although sensitivity is much greater than with other probes, the cyanine probe approach is not without drawbacks. For example, only trace labeling is achievable—at most, one probe–molecule per protein molecule. At higher grafting ratios, the protein probe complex becomes insoluble. Sensitivity is therefore seriously limited. Because of the 4

pI

8 MW 200 kDa

70 kDa

46 kDa

30 kDa

20 kDa

13 kDa

FIGURE 1. Separation of 100 µg of total cell extract by 2-D electrophoresis and direct fluorescence detection without fixation. The second dimension is run in 0.05% SDS. Note the elongated spots in the second dimension (arrowheads) and the gray crescent caused by ampholytes in the gel (arrow).

50 A


large number of amino groups generally present on proteins, very small amounts of the label must be used to prevent multiple labeling. Consequently, most of the protein molecules will not bear any fluorophores. This scenario is in sharp contrast with DNA labeling with attomole detection limits (4), in which there is exactly one fluorescent molecule per DNA molecule. Another problem is that, even if the number of charges present on the proteins is preserved, labeling with large fluorophore molecules (three aromatic cycles or more) induces a minor detectable (but unpredictable) perturbation in the migration of the protein in the SDS dimension. Therefore, the position of the bulk of unlabeled, invisible proteins is often offset from the position of the fluorescent spot (3), which creates problems when the spots of interest must be excised for subsequent analysis. Last but not least, this scheme of charge compensation during labeling suffers from artifacts over the complete pH range, especially with basic proteins. For acidic and neutral proteins, the pI is quite remote from the pKa of the amino group of lysines. Consequently, these amines can be considered as fully protonated and can be replaced by any group bringing a positive charge with a high pKa without altering the pI to a detectable extent. For basic proteins, however, the pI is in the range of the lysine pKa, which means that all lysine amino groups will not bear a positive charge at the pI. Thus, introducing a true positive charge or group with a much higher pKa will definitely alter the pI. This problem also extends to thiol-reactive probes. Thiols are, in fact, weak acids (pKa~10), and thiol alkylation with basic proteins removes these negative charges and alters the pI.

Radioactive labeling Because of safety problems, radioactive labeling is best performed prior to IEF. In fact, chemical radioactive labeling of a protein can be performed in a few tens of microliters, versus a few milliliters IEF and the SDS PAGE step or a fraction of a liter after SDS PAGE. Radioactive labeling can be done very efficiently by introducing radioactive metabolites to the source of the sample. In this case, there are no problems of pI modification because the amino acids themselves are labeled. In vitro labeling is therefore rather simple (5), whereas in vivo is much less convenient and, of course, impossible for human tissues. In vitro labeling after protein extraction has been proposed (6). In contrast with fluorescent labeling, very high grafting schemes with small groups (e.g., methyl groups) can be done without noticeably modifying the pI or the apparent molecular weight of the proteins, even for basic proteins (6). One problem with radioisotopes is detecting those that emit low-level radiation, such as tritium, which is by far one of the most versatile and convenient labels. This limitation was overcome by fluorography (7 ), in which a scintillator is

precipitated in the 2-D gel prior to drying. The scintillator converts the low-energy radiation of tritium, which does not penetrate the dry polyacrylamide layer, into visible light that is detected on autoradiographic film. Best results are obtained with 2,5-diphenyloxazole dissolved in DMSO (7 ) or acetic acid (8). Linearity is another important problem. With classical film detection, linearity was obtained with preflashing (9). This procedure was, however, difficult to standardize, and linearity was achieved over a limited range. A major improvement in sensitivity and linearity was afforded by filmless detection methods, such as phosphor storage (10). In this technique, the ß-radiation induces an energy change in a europium salt, which, upon excitation by a laser, is converted into visible light and quantified with a photomultiplier. Compared with autoradiographic film, sensitivity is increased 20- to 100-fold. The major improvement is in linearity, providing a linear dynamic range of 4 orders of magnitude. These methods, however, are not very efficient for tritium. Another new detection method is based on amplification detectors similar to those used in high-energy physics. Here again linearity is obtained over a wide range, with a 100-fold increase in sensitivity compared with film detection (11). Moreover, dual-isotope detection can be easily carried out, provided that the two radiation energies differ sufficiently (e.g., 3H versus 14C or 35S; 14C or 35S versus 32P). Furthermore, sensitive detection of tritium can be easily carried out provided that the radiation is not absorbed by the separation medium (12). Using blots or ultrathin gels is therefore highly recommended. The main drawback of this detection method is that the gel must be in the detector during the data-acquisition time, thereby precluding any parallel detection by films or multiple phosphor storage plates. Compared with film detection, these new methods suffer from a lack of resolution because they are limited to the 200- to 400-µm, while a 50- to 100-µm resolution is required for the correct analysis of 2-D gels. Radioactive detection probably offers the best S/N and the best ultimate sensitivity. With the latest developments, it is ideally suited for dual labeling techniques by allowing duplex analysis of samples on the same gel or for determining amino acid ratios. However, limitations in scope (particularly for tissues), the time needed for detection (usually several days), and safety regulations confine this method to specialized niches.

micelles), which can seriously limit the performance of noncovalently bound labels. One successful example of noncovalent labeling after IEF is the staining of proteins with the dye Coomassie Blue during migration in the SDS dimension (13). The main limitations of this method are a heavily stained background, which makes quantitation difficult, and poor detection limits (several micrograms). Covalent labeling is preferred at this stage. Dye labeling is typically excluded because of poor sensitivity. Because of the volumes required, radioactive labeling is also not typically used, although there are some exceptions (14). Thus, fluorescent labeling dominates at this stage and protein nucleophiles (amines and thiols) are the preferred targets for grafting the labels. A major problem arises from the presence of carrier ampholytes. These compounds are used in IEF to form the

Fluorescent labeling dominates at this stage, and protein nucleophiles are the preferred targets for grafting the labels.

Between IEF and SDS PAGE Compared with pre-IEF labeling, labeling between the IEF and SDS PAGE stages offers much greater flexibility, because the charge-state of the proteins can be altered provided that the apparent molecular weight is kept constant. However, SDS PAGE is done in a special environment (SDS

pH gradient or to smooth conductivity when immobilized pH gradients are used. Ampholytes are usually polyamino polycarboxylic acids, although some also contain phosphate or sulfate groups. Ampholytes are present in large excess over the proteins. If these compounds contain available amino groups, they will consume most of the label and thereby dramatically decrease the efficiency and sensitivity of protein labeling. Most carrier ampholytes are made by grafting carboxylic, phosphonic, or sulfonic acids to the polyamine, for example, pentaethylene hexamine. These compounds have a low molecular mass and migrate with the SDS front during the second dimension, so they do not interfere significantly with the detection process, except with very low molecular weight proteins. Another group of ampholytes is made by polymerizing amino acids and amines with epichlorohydrin. These ampholytes offer lower conductivity and greater diversity than the others, resulting in a smoother and more precise pH gradient. (Ampholyte-based pH gradients are stepwise gradients—one step per chemical of given pI. When the number of species per pH unit increases, the gradient becomes smoother.) Their molecular mass and binding to SDS is higher than that of the other ampholytes, especially in the neutral or basic ranges, and, as a result, the polymerized compounds migrate in the gel and not just with the SDS front. Because they are made from amino acids as proteins are, these polymerized compounds can be detected by most


51 A

protein-detection methods if present in the gel (Figure 1). Carrier ampholytes must be removed from the gel by a fixation process, which eliminates them by diffusion while precipitating the proteins within the gel. This process is usually achieved with several aqueous-acid alcohol baths or strongly acidic baths (e.g., 10% trichloroacetic acid in water), allowing subsequent reaction of the protein with the fluorescent probe (15). However, this process can lead to protein losses because one cannot be sure that the proteins will be resolubilized at the SDS PAGE step. This drawback can be alleviated by using thiol-reactive

Molecules that are fluorescent only after coupling are generally chosen, such as 2-methoxy-2,4-diphenyl furanone (16), because any fluorescent molecule remaining in the gel will induce a very high background. Most detection methods for labeling protein molecules after 2-D electrophoresis rely on noncovalent interactions. Although the contrast is theoretically lower than with covalent bonding, the much higher number of bound sites compensates easily. In addition, methods using only noncovalent interactions are generally reversible and therefore compatible with most characterization methods (MS, Edman sequencing), which is not the case with covalent grafting, especially with amine-reactive probes. In fact, the numerous amino groups present on the proteins make grafting of probes a random process. Thus, lysine side chains are modified to a variable extent. Moreover, because these lysine sites are often used for peptide generation, internal Edman sequencing, and MS, artifacts of long and complicated peptides are produced from covalently modified proteins—regardless of the grafting reaction. The only exceptions are trace labeling and lysine-reductive methylation. Detection of proteins after 2-D electrophoresis can be carried out directly in the polyacrylamide gel or on blotting membranes. Blots are frequently used, especially if further analysis is planned after separation. Blotting offers an important concentration effect. Proteins spread throughout the thickness of the polyacrylamide gel are concentrated at the surface of the blotting membrane and therefore are readily accessible to reagents. Moreover, blotting affords a cleanup process for the proteins.

Protein detection after 2-D electrophoresis can be carried out directly in the polyacrylamide gel or on blotting membranes. probes, for example, maleimido fluorescein, which itself has drawbacks. First, conjugation with these probes often leads to apparent molecular weight modification. Second, proteins without cysteines will go unnoticed with that labeling scheme. Third, sensitivity is not as good because only a few fluorophore molecules are grafted per protein. Finally, the hardware must be considered. The optical apparatus in DNA sequencing is fixed, and the labeled bands pass through. In 2-D electrophoresis, with point light excitation (e.g., laser excitation), a scanning device must be moving either the gel or the optical device (or both) so that all the gel will be scanned. This requirement is the case with laser fluorescence scanners. Such scanners are best used with probes having an excitation maximum close to an available laser wavelength (e.g., fluorescein and rhodamine derivatives). In another setup, the illumination is done in the gel plane and the fluorescent light is collected by a camera, usually a chargecoupled device camera (15, 16). In this case, molecules excited by UV are preferred, because the excitation light can be very efficiently filtered before the camera optics. The UVexcited probes are generally less sensitive and more prone to photobleaching than visible light-excited probes. In addition to the hardware problems, the interaction between the fluorophore and the protein residues or the SDS bound to the protein is still unknown and may also give rise to energy-transfer phenomena and thus to a large decrease in fluorescence intensity.

After SDS PAGE The problems discussed earlier for covalent grafting of the label, such as low number of sites and volume problems, also apply to labeling after 2-D electrophoresis. Consequently, only old methods with limited sensitivity include this scheme.

52 A


Noncovalent detection with organic dyes Staining with noncovalent dyes occurs after electrophoresis, generally after fixing to remove compounds that interfere with dye binding, such as SDS and carrier ampholytes. The staining process is generally regressive, that is, the gel is first saturated with the dye solution and then destained by a process that takes advantage of the higher affinity of the dye for the proteins over the acrylamide gel. Many dyes have been described for staining proteins, but the Brilliant Blue G and R dyes (also known as Coomassie Blue dyes) are the most popular. Because of their high extinction coefficients and high affinities for proteins, detection limits are ~1 µg (17 ). Moreover, Brilliant Blue stains proteins uniformly, except for proteins with extreme pIs (often not analyzed by 2-D gels) and glycoproteins. The main limitation of staining with dyes is sensitivity. In addition, regressive staining makes batch-to-batch reproducibility difficult to achieve. To overcome this drawback, a staining scheme should be used in which the proteins are pro-

gressively stained, while the background is kept clear. In this case, the gel is stained to the end point (saturation of the proteins with the dye), which dramatically increases reproducibility; the cost is a much longer staining time. A simple but efficient way to achieve progressive staining is to use a dilute solution of dye and to stain for a long time (18). Although very efficient, a large volume of staining solution relative to gel volume must be used so that enough dye molecules are present to saturate the proteins in the gel. This approach is therefore not compatible with staining a large series of gels, which is often encountered in 2-D electrophoresis. A very elegant progressive staining approach, colloidal staining, has been devised by Neuhoff’s group (19). In this approach, Brilliant Blue G forms microprecipitates, which are excluded from the polyacrylamide gel in acidic media containing ammonium sulfate. The concentration of the free dye is very low so that background staining is minimal. However, the microprecipitates act as a reservoir of dye molecules, so that enough dye is present to occupy all the binding sites on all the proteins, provided that staining is long enough to reach steady state (48 hours). In addition to very good reproducibility, an increase in sensitivity is obtained (Figures 2a and 2b). In fact, the presence of ammonium sulfate in the colloidal staining solution increases the strength of hydrophobic interactions, resulting in better sensitivity (~100 ng). Staining on blots with organic dyes is done almost exclu(a)

(b)

(c)

4 (d)

(e)

pI

sively by regressive staining. Because most dyes bind weakly to neutral membranes (nitrocellulose and polyvinylidene difluoride) and the concentration effect afforded by the blotting process, high-nanogram sensitivities are easily reached (20). This method does not work with cationic blotting membranes (charged nylon) because they bind anionic dyes so strongly that staining is not practical.

Fluorescence detection Noncovalent, environment-sensitive, fluorescent probes can also be used. Generally, these probes are nonfluorescent in water but highly fluorescent in apolar media, such as detergent, in which they take advantage of SDS binding to proteins to build a fluorescence-promoting environment at the protein sites in the gel. Typical probes of this type are naphthalene derivatives (21), Sypro dyes (22), and Nile Red (23). This method has limited contrast. Ideally, detergent should be present with the proteins in the gel, but without micelles. Lowering the SDS concentration in the gel to avoid micelles (23) often leads to vertical streaking (Figure 1). Consequently, excess SDS in the gel must be washed off prior to staining with the fluorophore. However, the washing process before and during staining must be controlled to maximize reproducibility and sensitivity because excess washing will remove too much SDS from the proteins. Noncovalent fluorescent detection is also feasible on blots. In this case, lanthanide complexes are used (24). An advantage of lanthanide complexes is that time-lapse detection with a very good S/N is possible. In the case of polyvinylidene diflouride membranes, which are not wettable with water or water alcohol mixtures with low alcohol concentration, a fluorescent probe with a weak affinity for proteins but with very good spectral characteristics (e.g., fluorescein, rho8 damine) can be used. It will not bind MW 200 kDa to the unwetted blotting membrane, only to the wetted protein sites (25). Although very high sensitivities have been claimed for these fluorescence methods with 2-D gels (26 ), it appears that the true limits are in the high-nanogram range, similar to colloidal Coomassie Blue (Figure 2). The situation is better on blots with detec10 kDa tion limits of ~10 ng.

FIGURE 2. Comparison of sensitivities for different staining methods for 400 µg of mitochondrial proteins separated by 2-D electrophoresis.

Detection by differential salt binding

Gels were stained with (a) Brilliant Blue G in acid alcohol medium, (b) colloidal Brilliant Blue G, (c) imidazole zinc, (d) silver, and (e) Sypro Orange (detected with a fluorescence laser scanner). Homologous spots are marked with arrows.

Differential salt binding and staining methods are based on protein-


53 A

bound salts (e.g., dodecyl sulfate or heavy cations), which are chemically less active than the free salt in the gel. Consequently, precipitation of proteins bound to these salts is slower than in the gel background. This kinetic difference can be used to devise negative stains in which the proteins stay translucent while the background becomes opaque due to salt precipitation. In these staining protocols, the limiting factor is the affinity of the proteins for one of the ions. The higher and the more general the affinity, the better the stain. All negative stains use heavy divalent cations (copper, zinc) for forming a precipitate with dodecyl sulfate (27 ). Submicrogram sensitivities are easily obtained within minutes, and destaining is easily achieved by using divalent chelators such as EDTA. Increased sensitivity can be achieved by modifying a zinc stain in which the precipitated salt is no longer zinc dodecyl sulfate but a complex salt of zinc and imidazole (28, 29). The protocol is simple and clearly the method of choice, but optimal results depend on careful timing. With the zinc imidazole stain, detection sensitivities approach the nanogram range (Figures 2c and 2d). Proteins precipitated as colorless zinc salts in the gel can be stored for several days at 4 °C. The drawbacks of zinc imidazole staining, which are common to all methods of this type, are difficult reproducibility and very poor linearity. The technique cannot be used as a general tool for studying quantitative variations on 2-D gels, and detection is often poor for glycoproteins and lowmolecular-weight proteins. Last, the protein zones are not easily recognized by the human eye, which can lead to difficulties in spot excision for further protein analysis. Because of the low SDS content in protein blots, completely different schemes are used for staining blotted proteins. The metal ion is bound to the blotted proteins and made visible with an organic complexant. Low-nanogram sensitivities have been reported (30), and the stain is reversible by adding EDTA.

bind to silver. Silver reduction is then primed where the sensitizer is bound (i.e., at the protein level) and a positive image is obtained due to autocatalysis (31). Silver-staining protocols are divided into two families. In one, the silvering agent is silver nitrate and the developer is formaldehyde in an alkaline carbonate solution. In the second, the silvering agent is a silver–ammonia complex and the developer is formaldehyde in dilute citric acid. Within each family, various protocols have been described, differing mainly in the fixative (e.g., with or without aldehydes) and the sensitizer. These differences can lead to widely differing results in terms of sensitivity, reproducibility, and protein-toprotein response, and recent protocols clearly perform much better than older ones (32). Kits are also commercially available, but none of them are as sensitive as good, homemade protocols. High-performance silver-staining methods use at least one solution with a limited shelf life, which makes these homemade protocols unmarketable. In addition to the detection threshold, there are some criteria for selecting the appropriate method. Protocols using fixation with an aldehyde are more sensitive, more uniform from protein to protein, and more reproducible. However, aldehyde fixation precludes any further analysis of the stained protein. Protocols using short steps (≤1 min) are less reproducible than protocols using longer steps only. Protocols using ammoniacal silver are slightly more sensitive and give darker hues than those based on silver nitrate. They are also more sensitive for basic proteins but less sensitive for very acidic proteins. They are, however, more prone to negative staining phenomena (hollows or doughnuts), especially with aldehyde-free fixatives. In addition, adequate results are obtained only when thiosulfate is included in the gel matrix during polymerization (33), so that ready-made gels cannot be used. Some gel systems (e.g., Tricine-based systems) give unacceptable backgrounds with ammoniacal silver. Last but not least, the performances of ammoniacal silver methods strongly depend on the ammonia–silver ratio (34 ), which is tedious to control. The sensitivity of silver staining with modern protocols is in the low-nanogram range both for silver nitrate and silver– ammonia, with minimal protein-to-protein variation. This is 100-fold better than classical Brilliant Blue staining, 10-fold better than colloidal Brilliant Blue staining, and approximately 2-fold better than zinc staining—plus it gives a much better contrast (Figure 2). Because protein are fixed before staining and formaldehyde is used for development, interfacing silver-stained gels with identification methods is difficult. Fixation with aldehydes precludes any further analysis. When fixation is avoided, good results are often obtained with methods in which the proteins

Most staining methods have reached a plateau close to their theoretical maximum.

Detection by metal ion reduction Metal ion reduction (silver staining) is one of the most complex protein-detection processes, the driving force of which is the affinity of proteins for the silver cation. However, many substances (e.g., SDS, chloride, and amino acids) present in SDS gels also show a high affinity for silver, so they must be removed prior to contacting proteins with silver ion. A fixation step is therefore required. What makes a silver stain highly sensitive is the strong autocatalytic character of silver reduction. This condition is achieved by using a very weak developer (dilute solutions of formaldehyde) and sensitizers between fixation and silver impregnation. Sensitizers bind to proteins and react with or 54 A


are digested or hydrolyzed (e.g., peptide mass fingerprinting). This tendency can be partly attributed to the fact that silver staining is a surface stain, which means that the proteins below the surface are not damaged by the staining process. Protocols that include either silver nitrate (35) or silver ammonia (36) have been successfully used with peptide mass fingerprinting. However, proteins stained with silver sometimes give disappointing or no results. Destaining silver-stained gels with ferricyanide and thiosulfate greatly improves subsequent analysis with MS (37 ). Disappointing results are probably due to the staining process and the possible presence of nanogram amounts of proteins in the silver-stained spot, which can give a positive result with MS (38) but not with most other protein-identification methods. Silver-staining of blots by the same protocols used with gels has been reported (20), but it is seldom used because of lower sensitivity than gel staining.

(3) (4) (5) (6) (7) (8) (9) (10) (11)

What’s next?

(18) (19) (20) (21) (22) (23)

Most staining methods have reached a plateau close to their theoretical maximum. For dye staining, the maximum occurs when the dye is bound to all available sites on the protein. Colloidal Brilliant Blue staining has come close to achieving its maximum. A maximum also seems to have been reached for silver staining—no increase in sensitivity has been reported in the last five years, suggesting that all bound silver ion that is reducible under contrasting conditions is reduced with the current protocols. Any additional increases in sensitivity of silver staining will be based on different principles than those currently in use, requiring serious methodological development. The method with the most potential for improvement is fluorescence. In addition to the problems already described for probe design, hardware problems related to the high spatial resolution required for correct 2-D gel analysis need to be solved. Methods that use sophisticated hardware for fluorescence and radioactivity detection also need dedicated hardware for excising the detected spots for further analysis. This issue can be elegantly solved by simultaneous detection and analysis by IR matrix-assisted laser desorption/ionization (MALDI) MS of proteins electroblotted from a 2-D gel (39). By combining this approach with labeling of metabolites containing stable isotopes, data on the quantity and synthesis rate can be obtained in a single step. For the time being, this approach is plagued by the insufficient speed of MALDI time-of-flight instruments (in shots per second) and by the enormous amount of data generated by an object the size of a 2-D gel with a 100-µm resolution. High-speed MS may make this method an attractive choice.

References (1) (2)

Bosshard, H. F.; Datyner, A. Anal. Biochem. 1977, 82, 327–333. Urwin, V. E.; Jackson, P. Anal. Biochem. 1993, 209, 57–62.

(12) (13) (14) (15) (16) (17)

(24) (25) (26) (27) (28) (29) (30) (31) (32) (33) (34) (35) (36) (37) (38) (39)

Unlii, M.; Morgan, E. M.; Minden J. S. Electrophoresis 1997, 18, 2071–2077. Smith, L. M., et al. Nature 1986, 321, 674–679. O’Farrell, P. H. J. Biol. Chem. 1975, 250, 4007–4021. Rabilloud, T.; Hubert, M.; Tarroux P. J. Chromatogr. 1986, 351, 77–89. Bonner, W. M.; Laskey, R. A. Eur. J. Biochem. 1974, 46, 83–88. Skinner, M. K.; Griswold, M. D. Biochem. J. 1983, 209, 281–284. Laskey, R. A.; Mills, A. D. Eur. J. Biochem. 1975, 56, 335–341. Johnston, R. F.; Pickett, S. C.; Barker, D. L . Electrophoresis 1990, 11, 355–360. Charpak, G.; Dominik, W.; Zaganidis, N. Proc. Natl. Acad. Sci. U.S.A. 1989, 86, 1741–1745. Tribollet, E.; Dreifuss, J. J.; Charpak, G.; Dominik, W.; Zaganidis, N. Proc. Natl. Acad. Sci. U.S.A. 1991, 88, 1466–1468. Schägger, H.; Aquila, H.; Von Jagow, G. Anal. Biochem. 1988, 173, 201–205. Boxberg, Y. V. Anal. Biochem. 1988, 169, 372–375. Urwin, V. E.; Jackson, P. Anal. Biochem. 1991, 195, 30–37. Jackson, P.; Urwin, V.; Mackay, C. D. Electrophoresis 1988, 9, 330–339. Fazekas de St. Groth, S.; Webster, R. G.; Datyner, A. Biochim. Biophys. Acta 1963, 71, 377–391. Chen, H.; Cheng, H.; Bjerknes, M. Anal. Biochem. 1993, 212, 295–296. Neuhoff, V.; Arold, N.; Taube, D.; Ehrhardt, W. Electrophoresis 1988, 9, 255–262. Christiansen, J.; Houen, G. Electrophoresis 1992, 13, 179–183. Horowitz, P. M.; Bowman, S. Anal. Biochem. 1987, 165, 430–434. Steinberg, T. H.; Haugland, R. P.; Singer, V. Anal. Biochem. 1996, 239, 238–245. Alba, F. J.; Bermudez, A.; Bartolome, S.; Daban, J. R. Biotechniques 1996, 21, 625–626. Lim, M. J.; Patton, W. F.; Lopez, M. F.; Spofford, K. H.; Shojaee, N.; Shepro, D. Anal. Biochem. 1997, 245, 184–195. Coull, J. M.; Pappin, D. J. C. J. Prot. Chem. 1990, 9, 259–260. Steinberg, T. H.; Jones, L. J.; Haugland, R. P.; Singer, V. Anal. Biochem. 1996, 239, 223–237. Dzandu, J. K.; Johnson, J. F.; Wise, G. E. Anal. Biochem. 1988, 174, 157–167. Ortiz, M. L.; Calero, M.; Fernandez-Patron, C.; Castellanos, L.; Mendez, E. FEBS Lett. 1992, 296, 300–304. Fernandez-Patron, C., et al. Electrophoresis 1998, 19, 2398–2406. Patton, W. F.; Lam, L.; Su, Q.; Lui, M.; Erdjument-Bromage, H.; Tempst, P. Anal. Biochem. 1994, 220, 324–335. Rabilloud, T.; Vuillard, L.; Gilly, C.; Lawrence, J. J. Cell. Mol. Biol. 1994, 40, 57–75. Rabilloud, T. Electrophoresis 1992, 13, 429–439. Hochstrasser, D. F.; Merril, C. R. Appl. Theor. Electrophoresis 1988, 1, 35–40. Eschenbruch, M.; Burk, R. R. Anal. Biochem. 1982, 125, 96–99. Shevchenko, A.; Wilm, M.; Mann, M. Anal. Chem. 1996, 68, 850–858. Rabilloud, T., et al. Electrophoresis 1998, 19, 1006–1014. Gharahdaghi, F.; Weinberg, C. R.; Meagher, D. A.; Imai, B. S.; Mische, S. M. Electrophoresis 1999, 20, 601–605. Figeys, D.; Gygi, S. P.; Zhang, Y.; Watts, J.; Gu, M.; Aebersold, R. Electrophoresis 1998, 19, 1811–1818. Eckerskorn, C., et al. Anal. Chem. 1997, 69, 2888–2892.

Thierry Rabilloud is CNRS senior research fellow at CEA Grenoble. His research interests include proteomics and 2-D electrophoresis and their application to cell biology. Address correspondence about this article to him at DBMS/BECP, CEA Grenoble, 17 rue des Martyrs, F38054 Grenoble, France ([email protected]). J A N U A R Y 1 , 2 0 0 0 / A N A LY T I C A L C H E M I S T R Y

55 A

Peer Reviewed: Detecting Proteins Separated by 2-D Gel

Recommend Documents