Anal. Chem. 2000, 72, 2519-2525
Automated Ultra-Thin-Layer SDS Gel Electrophoresis of Proteins Using Noncovalent Fluorescent Labeling Zsolt Csapo,† Arpad Gerstner,‡ Maria Sasvari-Szekely,‡ and Andras Guttman*,†
Genetic BioSystems, Inc., San Diego, California 92121, and Institute of Medical Chemistry, Molecular Biology and Pathobiochemistry, Semmelweis University Medical School, Budapest, Hungary
Ultra-thin-layer SDS gel electrophoresis in conjunction with automated laser-induced fluorescence detection is a novel and powerful method for the analysis of fluorophore-labeled proteins. The technique described in this paper employs instant, noncovalent fluorophore labeling by the addition of a fluorescent staining dye to the sample proteins either during or immediately prior to the sample loading process. Thus, the method does not require timeconsuming post- or preseparation staining/labeling. By combining the multilane format of SDS polyacrylamide slab gel electrophoresis and the high separation efficiency of capillary SDS gel electrophoresis, ultra-thin-layer SDS gel electrophoresis features rapid, high-throughput, and high-resolution analysis of proteins in the molecular mass range of 14-116 kDa. The good heat dissipation inherent to the ultrathin format enables the use of agarose and agarose-based composite separation matrixes, which can be easily replaced within the separation platform. Labeling efficiency as a function of the concentration of the staining dye, SDS, and proteins is thoroughly discussed. Detection sensitivity of the method was found to be at the lowfemtomole level (1.25 ng/band), determined by analyzing a set of serial dilutions of standard proteins. Practical example of molecular mass determination and characterization of a complex protein mixture are also shown. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is one of the most extensively used bioanalytical methods for the separation, identification, and characterization of proteins and protein mixtures.1 This simple technique proved to be very useful for protein molecular mass assessment and purity evaluation. Most proteins, when boiled in the presence of SDS and a reducing agent, such as dithiothreitol, are fully denatured and, regardless of their size and shape, bind a close to constant amount of SDS per unit weight (1.4 g of SDS/g of protein).2 The binding characteristics is described as mostly hydrophobic * Corresponding author. Present address: Novartis Agricultural Discovery Institute 3115 Merryfield Row, San Diego, CA 92121. Tel: (858) 812-1052. Fax: (858) 812-1097. e-mail:
[email protected]. † Genetic BioSystems, Inc.. ‡ Semmelweis University Medical School. (1) Andrews, A.T. Electrophoresis; Clarendon Press: Oxford University Press: New York, 1986. (2) Weber, K; Osborn, M. J. Biol. Chem. 1969, 244, 446-4412. 10.1021/ac991501+ CCC: $19.00 Published on Web 04/26/2000
© 2000 American Chemical Society
interactions.3 Sodium dodecyl sulfate molecules cover the proteins and mask their intrinsic charge, resulting in SDS-protein complexes of approximately constant charge per unit mass and similar denatured random coil shape,4 leading to practically identical free solution mobilities. However, when electrophoresis is carried out in a sieving matrix, consisting of a hydrophilic sieving gel or polymer network such as polyacrylamide5 or agarose6 with a given pore size, separation of the SDS-protein complexes can be accomplished on the basis of their size. In SDS gel electrophoresis of non-posttranslationally modified proteins, the mobility values are proportional to the logarithm of the effective molecular radius, thus, to the logarithmic molecular mass of the polypeptide chain.7 Posttranslational modifications, such as glycosylation, phosphorylation, etc., change the constant mass-to-charge ratio of the SDS-protein complexes. Therefore, the above-described mobility versus mass relationship is altered. It is important to note here, that although SDS-PAGE is widely used due to its simplicity, it is usually a time-consuming process and difficult to automate. Visualization of the separated protein bands in slab gel electrophoresis is usually performed after the completion of the separation process (postelectrophoresis), either in situ within the separation matrix itself or after a transfer of the separated proteins onto polymeric membrane support materials (blotting).8 Both methods require extra time (up to several hours) after the separation process is completed. Currently, the most commonly used postseparation in situ staining methods are Coomassie Brilliant Blue (CBB)9 staining and silver staining.10 CBB staining is inexpensive, but suffers from low sensitivity (0.1-0.2 µg/band) and long processing times (several hours). Silver staining is up to 100 times more sensitive than CBB staining, but still requires several labor-intensive and time-consuming steps, as well as chemicals that are relatively unstable and toxic. Moreover, silver (3) Ganzler, K; Greeve, K. S.; Cohen, A. S.; Karger, B. L.; Guttman, A.; Cooke, N. C. Anal. Chem. 1992, 64, 2665-2671. (4) Reynolds, J. A.; Tanford, C. Proc. Natl. Acad. Sci. U.S.A. 1970, 66, 10021007. (5) Chrambach, A. The Practice of Quantitative Gel Electrophoresis; VCH Publishers: Deerfield Beach, FL, 1985. (6) Wu, M.; Kusukawa, N. BioTechniques 1998, 24, 676-678. (7) Shapiro, A. L.; Vinuela, E.; Maizel, J. V. Biochem. Biophys. Res. Commun. 1967, 28, 815-820. (8) Wortj, P. J.; Romano, A. J. Chromatogr., A 1995, 698, 123-143. (9) Chrambach, A.; Reisfeld, R. A.; Wyckoff, M.; Azccari, J. Anal. Biochem. 1967, 20, 150-155. (10) Oakley, B. R.; Kirsh, D. R.; Morris, N. R. Anal. Biochem. 1980, 105, 361363.
Analytical Chemistry, Vol. 72, No. 11, June 1, 2000 2519
staining tends to be protein selective as the staining intensity and color is very dependent on the sequence of the polypeptide chain and the degree of glycosylation. Recently, various high-sensitivity fluorescent labeling methods were introduced to visualize proteins, separated by SDS gel electrophoresis. For postseparation fluorophore staining, dyes such as ethidium bromide11 and Nile red12 were reported to provide the sensitivity level comparable to that attained by silver staining and most importantly were not particularly protein selective. The novel Sypro dyes13 provide even better detection limits of as low as 1-2 ng of protein/band, but the postseparation staining process still require 30-60 min. However, as this dye appear to bind to the detergent coat surrounding the proteins in denaturing SDS gels, staining in such gels is not strongly selective for particular polypeptide sequences.14 Postelectrophoresis in situ protein staining is by far the most frequently used visualization method in SDS slab gel electrophoresis; however, a few procedures have been described for labeling proteins prior to the separation process. Such preseparation covalent fluorophore labeling was recently reported by Hunt and Nasabeh15 using 5-TAMRA.SE, as a very sensitive approach for monitoring consistency of biotechnology products, requiring 2 h of derivatization time. Fluorescent dye-labeled/stained proteins are mostly visualized by UV or visible light illumination using the proper filters for CCD camera archiving systems or with laser-induced fluorescent scanning detection techniques.16 Automated, ultra-thin-layer gel electrophoresis of biopolymers is a recently developed method that offers an alternative approach to conventional slab gel electrophoresis with the advantages of automation, fast separation, and the capability of real time detection during the separation process, i.e., no time-consuming postseparation visualization step is required.17 This novel technique combines the benefits of the multilane format of slab SDS gel electrophoresis and the speed as well as the high detection and separation efficiency inherent to capillary electrophoresis. The SDS-protein complexes are labeled instantly prior to or during the injection process with the appropriately chosen fluorophore. The fluorescently labeled sample components are then separated by ultra-thin-layer gel electrophoresis and detected in real time with an integrated scanning laser-induced fluorescence/avalanche photodiode detection system, eliminating the need of the lengthy postseparation staining/visualization process. The electrophoresis separation can be carried out in agarose or composite agaroselinear polymer additive (e.g., linear polyacrylamide, poly(ethylene oxide), derivatized celluloses, etc.) gels. The sieving matrix is filled into the separation platform in a melted form at 50-60 °C and used after solidification at room temperature. The use of agarosebased composite separation matrixes also enables easy replacement of the gel within the separation platform by simply pumping fresh melted agarose or agarose-based composite gels into the
cassette for each analysis. Sample loading could be accomplished by conventional well loading or by recently introduced membranemediated loading technology.18 In this paper, we report the first use of a novel, automated separation and visualization method for SDS-protein complexes. Ultra-thin-layer composite agarose-linear polyacrylamide gel electrophoresis is employed in conjunction with noncovalent, instant fluorophore labeling of the SDS-protein complexes by applying a high-sensitivity fluorophore staining dye during the sample-loading process. Labeling efficiency as a function of SDS, protein, and dye concentration is thoroughly discussed, and a practical example of molecular mass determination and characterization of a complex protein mixture are also demonstrated.
(11) Vincent, A.; Scherrer, K. Mol. Biol. Rep. 1979, 5, 209-214. (12) Daban, J. R.; Bartolome, S.; Samso, M. Anal. Biochem. 1991, 199, 169174. (13) Haugland, R. Ph. In Handbook of fluorescent probes and research chemicals; Spence, M. T. Z., Ed.; Molecular Probes, Inc., Eugene, OR, 1996. (14) Steinberg, T. H.; Haugland, R. P.; Singer, V. L. Anal. Biochem. 1996, 239, 238-245. (15) Hunt, G.; Nasabeh, W. Anal. Chem. 1999, 71, 2390-2397. (16) Southerland, J. C. In Advances in Electrophoresis; Chrambach, A., Dunn, M. J., Radola, B. J., Eds.; VCH Publishers: Weinheim, Germany, 1993. (17) Guttman, A. LC/GC Mag. 1999, 17, 1020-1026.
ν ) µPEK[L-]m
2520
Analytical Chemistry, Vol. 72, No. 11, June 1, 2000
THEORY The staining dye (complexing ligand, L-), binds the high local concentration SDS moiety of the SDS-protein complex (Pn-), and due to its negative charge at the separation pH of 8.4,19 it increases the ionic charge of the resulting complex (PLm(n+m)-):
Pn- + mL- h PLm(n+m)-
(1)
K ) [PLm(n+m)-]/[Pn-][L-]m
(2)
where K is the formation constant of the complex, m is the number of the negatively charged ligand molecules in the complex, and n is the total number of negative charges on the SDS-protein complex (as a first approximation, let us consider that in most instances it is close to equal to the number of SDS molecules in the complex). The electrophoretic velocity (v) of the polyiondye complex in SDS gel electrophoresis is described as
ν ) l/tM ) µPERP
(3)
where l is the effective length of the separation platform (from the injection to the detection point), tM is the migration time of the SDS-protein-staining dye complex, µP is the electrophoretic mobility of the polyion, and E is the applied electric field strength. The molar ratio of the complexed dye in the SDS-protein - dye complex, RP, is given by
RP )
[PLm(n+m)-] K[Pn-][L-]m ) cP cP
(4)
where cP is the total concentration of the polyion. Combining eqs 3 and 4 with the assumption that cP ≈ [Pn-] (since the limited amount of fluorescent staining dye binds only a fraction of the SDS-protein complex) results in
(5)
Equation 5 suggests the extent of the increase in electrophoretic velocity when the negatively charged staining dye binds the SDSprotein complex. Please note that one should also take into (18) Cassel, S.; Guttman, A. Electrophoresis 1998, 19, 1341-1346. (19) Molecular Probes, Inc. Eugene, OR. Personal communications, 1999.
account the mass increase of the SDS-protein-dye complex due to the additional mass of the labeling fluorophore that somewhat decreases this resulting electrophoretic velocity in eq 5. EXPERIMENTAL SECTION Chemicals. In all the experiments, 1% low electroendosmosis (EEO; -mr ) 0.06) Amresco’s Agarose-III (Solon, OH) and 2% linear polyacrylamide (MW 700 000-1 000 000) (Polysciences, Inc., Warrington, PA) were dissolved in 50 mM Tris, 50 mM N-tris(hydroxymethyl)methyl-4-aminobutanesulfonic acid (TAPS) buffer (pH 8.4), containing 0.05% SDS (referred to as TTS). Tris, TAPS, SDS, and mercaptoethanol were obtained from ICN (Costa Mesa, CA), all in electrophoresis grade. The fluorescent staining dye Sypro Red (SR) was purchased from Molecular Probes (Eugene, OR) in 5000× concentration. Standard proteins of R-lactalbumin (MW 14 200), trypsin inhibitor (MW 20 100), carbonic anhydrase (MW 29 000), ovalbumin (MW 45 000), bovine serum albumin (MW 66 000), phosphorylase B (MW 97 400), and β-galactosidase (MW 116 000) were obtained from Sigma Chemical Co. (St. Louis, MO), dissolved in double-deionized water (18 MΩ) in the final concentration of 1 mg/mL, and stored at -20 °C until use. Instrumentation. The automated ultra-thin-layer (190 µm) agarose gel electrophoresis system equipped with real time laserinduced fluorescent detection was described earlier.20 This particular system employed a fiber-optic bundle-based scanning illumination/detection unit, using a 532-nm frequency-doubled green Nd:YAG laser excitation source and avalanche photodiode (APD) detection with a 585 ( 25 nm wide-band interference filter. A lens set scanned across the multilane separation platform at various distances from the injection site by means of a translation stage and collected the emitted fluorescent light.21 Effective separation lengths of 2.5, 3.5, and 5.5 cm (measured from the loading well to the scanning lens set) were used. The analog signal from the APD was digitized in a microcontroller and acquired by a personal computer. Integrated optical density (IOD) values of the separated bands were obtained by using the Gel-Pro Analyzer software package from Media Cybernetics (Silver Spring, MD). The ultra-thin-layer separation platform was an 18 cm × 7.5 cm × 190 µm float glass cartridge with built-in 15-mL plastic buffer reservoirs at both ends. The inside surfaces of the cassettes were covalently coated by linear polyacrylamide, according to the procedure of Hjerten22 to minimize electroosmotic flow (EOF) at the separation pH of 8.4. Procedures. An appropriate amount of agarose powder was suspended in TTS buffer and boiled repetitively in a microwave oven until clear. Linear polyacrylamide (LPA, MW 700 0001 000 000) was added to the melted agarose in 2% final concentration. The composite matrix was then kept at 60 °C for 10 min before use. The preheated separation cassettes (45-50 °C) were filled with melted agarose-LPA composite mixture, and after several minutes of cooling/solidification, the gel-filled cassette was ready to be used. Preheating the separation cassette helped to prevent premature solidification of the freshly poured gel. After each run, the used gels were replaced in the separation cassette by simply pumping through fresh, melted composite matrix. In all the separations, the applied voltage was 420 V, generating 5-mA (20) Trost, P.; Guttman, A. Anal. Chem. 1998, 70, 3930-3935. (21) Guttman, A. Trends Anal.Chem. 1999, 12, 694-699. (22) Hjerten, S. J. Chromatogr. 1985, 347, 191-198.
Figure 1. Relationship between the integrated optical density (IOD) values of the bovine serum albumin bands and the staining dye concentration using noncovalent fluorescent labeling in automated ultra-thin-layer SDS gel electrophoresis.
current. An aluminum heat sink was employed to hold the separation cartridge in horizontal position and to dissipate extra Joule heat. The temperature of the heat sink was regulated by a thermostated air bath with a precision of (1 °C. The actual separation temperature was measured at the middle of the heat sink by a thermocouple. The protein samples were diluted to 0.025-0.8 mg/mL with 50 mM Tris, 50 mM TAPS, 0.05-1% SDS, and 10% sucrose containing buffer and boiled in a water bath for 5 min after the addition of 2.5% 2-mercaptoethanol. The samples were then cooled to room temperature and fluorescently labeled by complexation with Sypro Red dye (λex ) 540 nm, λem ) 625 nm), which was added to the sample in final concentrations of 0.1×-4× prior to loading. Samples (0.2-0.5 µL) were injected into preformed loading wells (2.5 × 4 × 0.19 mm). RESULTS AND DISCUSSION Effects of Labeling Conditions. Initial efforts were focused on understanding of the effects of staining dye, SDS, and sample protein concentrations on noncovalent fluorophore labeling of SDS-protein complexes. First, the effect of staining dye concentration on the resulting fluorescent signal was investigated. Fluorescent staining dye was added to 0.1 mg/mL BSA sample in final concentrations of 0.25×, 0.5×, 1×, 2×, and 4× using the 5000× stock solution. Figure 1 shows the IOD values of the separated and detected bands as the function of staining dye concentration. Each data point in the figure represents the average of six experiments and the error bars correspond to the SD values of the data. The plot in Figure 1 exhibits a quasi-sigmoid relationship between the staining dye concentration and the fluorescent signal (IOD). With staining dye concentrations at or below 0.5× added to the sample, the resulting signal was at a quite low level and apparently did not show any dye concentration dependency. Increasing the staining dye concentration from 0.5× to 2×, the IOD response of the detected bands increased linearly (r2 ) 0.998). With further addition of staining dye to the sample (from 2× to 4×), the IOD response increased another 50%, which was still significant but not linear as was observed in the interval of 0.5×-2.0×. It is also important to note that higher than 4× staining dye concentration in the sample caused a significant Analytical Chemistry, Vol. 72, No. 11, June 1, 2000
2521
Figure 2. Integrated optical density (IOD) values of the bovine serum albumin bands and the SDS front as a function of surfactant concentration in automated ultra-thin-layer SDS gel electrophoresis with noncovalent fluorescent labeling.
increase in the lane background that eventually became so strong that it interfered with band identification. On the basis of the above results, we concluded that the optimal labeling dye concentration was between 0.5× and 2× for noncovalent staining of BSA in automated ultra-thin-layer SDS gel electrophoresis. As reported earlier, the surfactant (SDS) concentration in polyacrylamide slab gels has a significant effect on postseparation protein staining, especially with such binding fluorophores as the Sypro dyes.23 When the SDS concentration is above its critical micelle concentration, cmc ) 8.3 mM (∼0.24%, in water and somewhat less in buffer soutions), the major portion of the staining dye gets attached to the micelles. The bar diagram in Figure 2 shows the integrated optical density values of the BSA bands and the SDS front of the same lanes, obtained by automated ultrathin-layer SDS gel electrophoresis. When the regular amount of 1% SDS (> cmc) was used in the sample buffer, apparently a large amount of staining dye got attached to the SDS micelles. This resulted in a very intensive SDS-dye front, even interfering with the lower molecular mass protein bands of 0.9). While the dilution responses for TRI and CBA were still adequate (r2 > 0.5), the dilution linearity response of PHB was poor. The
Figure 5. Comparison of the band intensities of the seven-protein mixture of R-lactalbumin (ALA), trypsin inhibitor (TRI), carbonic anhydrase (CBA), ovalbumin (OVA), bovine serum albumin (BSA), phosphorylase B (PHB), and β-galactosidase (BGA) (0.2 mg/mL each) injected in a set of serial double dilutions (1-, 2-, 4-, 8-, 16-, and 32fold) and detected at 2.5 (lower panel) and 5.5 cm (upper panel) from the loading site. Arrows at 16 min show the time when the scanning distance was changed without interrupting the application of the electric field strength. Separation conditions: gel, 1% agarose, 2% linear polyacrylamide (LPA, MW 700 000-1 000 000) in 50 mM Tris, 50 mM TAPS, 0.05% SDS (pH 8.4); separation buffer, 50 mM Tris, 50 mM TAPS, 0.05% SDS (pH 8.4); separation voltage, 420 V; current, 5 mA; gel thickness, 190 µm; temperature, 25 °C; sample loading, 0.2 µL into 2.5 × 4 × 0.19 mm injection wells. Sample buffer contained 0.05% SDS and 1× Sypro Red. Table 1. Integrated Optical Density (IOD) Data of All the Bands (Upper Panel) Separated in Figure 5 dilution factor (×)
BGA
PHB
protein (IOD) BSA OVA CBA
32 16 8 4 2 1 r2
2142 1601 1338 920 536 180 0.947
1241 585 1845 1024 632 335 0.125
8473 5161 1948 2042 2485 2219 0.969
6778 3922 1580 1032 547 43 0.995
8180 6484 7642 7659 6025 2647 0.523
TRI
ALA
3049 3447 2700 1799 2179 3674 0.699
732 74 34 24 17 18 0.927
differences in absolute responses across the protein mixture showed quite a variation, suggesting the need of optimization of the labeling parameters for individual proteins or protein mixtures. The varying responses among the labeling of individual proteins observed in Figure 5 were probably caused by the various amounts of SDS they bound, due to glycosylation differences. Since carbohydrate moieties do not bind SDS (the labeling dye binds the SDS shell only) glycosylated parts do not complex with the dye, resulting in variability of the detection signal. This is similar to that observed previously with silver staining. Please note that, in an industrial setting, where speed, ease of use, and automation are the major requirements for evaluating a large number of similar sample mixtures, the method suggested in this paper can be easily optimized and applied. Another important feature of the consecutive double-distance detection shown in Figure 5 is the apparently high migration time fidelity of the detected bands at the 5.5-cm distance (measured and calculated average migration times of all seven proteins in the first five lanes of Figure 5 are depicted in Table 2, showing Analytical Chemistry, Vol. 72, No. 11, June 1, 2000
2523
Table 2. Measured (2.5 and 5.5 cm) and Calculated (5.5 cm) Migration Times of the Individual Proteins in the Seven-Protein Test Mixture Derived From the Separations Shown in Figure 5 protein
2.5 cm (measd)
5.5 cm (calcd)
5.5 cm (measd)
error (%)
ALA PHB BSA OVA CBA TRI ALA
8.95 9.32 10.17 10.99 12.10 13.30 14.32
19.69 20.50 22.37 24.18 26.61 29.27 31.50
19.60 20.32 22.18 24.05 26.35 29.22 31.48
0.46 0.88 0.85 0.54 0.98 0.17 0.06
Figure 7. Standard curve of logarithmic molecular mass vs electrophoretic mobility (derived from the data of lane B in Figure 6) for molecular mass estimation.
Figure 6. Analysis of phosphorylase B (PHB) (lane A) and the separation of a five-protein test mixture of R-lactalbumin (ALA), carbonic anhydrase (CBA), ovalbumin (OVA), bovine serum albumin (BSA), and β-galactosidase (BGA) (lane B). Conditions are the same as in Figure 5; effective separation length, 3.5 cm; protein concentration, 0.2 mg/mL each. Sample buffer contained 0.05% SDS and 1× Sypro Red.
an average error of 0.562%). This actually means that one can accurately predict the migration time at any distance from the loading well (5.5 cm in this instance) based on the mobility values calculated from the migration time results of the 2.5-cm detection. This would allow precise collection of any detected proteins for further characterization by microsequencing or mass spectrometry. A fraction collection device is being developed in our laboratory to exploit this feature of our automated ultra-thin-layer SDS gel electrophoresis system. Molecular Mass Estimation and Application. Figure 6 shows the separation of a five-protein test mixture of ALA, CBA, OVA, BSA, and BGA (lane B) and the analysis of the molecular mass of PHB (lane A). All proteins were noncovalently labeled by the fluorophore staining dye just prior to the loading process and detected in real time during the electrophoresis separation. The actual separation distance in this case was 3.5 cm, to obtain high resolution but still rapid analysis time. The standard curve for molecular mass estimation was constructed by plotting the logarithmic molecular masses of the five proteins in the standard test mixture against their electrophoretic mobilities (Figure 7). Best fitting was attained by applying a second-order polynomial function, which resulted an extremely high confidence level (r2 ) 0.9999) for this relationship. This is in contrast to previous reports on the linear relationship between the logarithmic molecular mass and electrophoretic mobility. The slight curvature of this calibration plot is probably caused by the noncovalent attachment of the negatively charged staining dye that increases the charge and, concomitantly, the overall electrophoretic velocity of the complex (eq 5). Lane A in Figure 6 depicts the automated 2524 Analytical Chemistry, Vol. 72, No. 11, June 1, 2000
Figure 8. Analysis of commercial low fat milk (lane A) and the fiveprotein test mixture (lane B) by automated ultra-thin-layer SDS gel electrophoresis using noncovalent fluorophore staining. Separation conditions: the five-protein test mixture is the same as in Figure 6; gel, 1% agarose, 2% LPA (MW 700 000-1 000 000) in 50 mM Tris, 50 mM Taps, 0.05% SDS (pH 8.4); separation buffer, 50 mM Tris, 50 mM Taps, 0.05% SDS (pH 8.4); separation voltage, 420 V; current, 5 mA; effective separation length, 3.5 cm; gel thickness, 190 µm; temperature, 25 °C; sample loading, 0.5 µL into 2.5 × 4 × 0.19 mm injection wells. Sample buffer contained 0.05% SDS and 1× Sypro Red.
ultra-thin-layer SDS gel electrophoresis analysis of phosphorylase B using noncovalent fluorescent labeling. On the basis of the calibration curve in Figure 7, we estimated the molecular mass of the phosphorylase B band in lane A to be 97.250 kDa, representing only a 0.25% error compared to the literature value (MW 97 40024). Rapid separation of a complex protein mixture by automated ultra-thin-layer SDS gel electrophoresis is demonstrated in Figure 8, in which, lane A depicts the analysis of commercial low fat milk using noncovalent, instant fluorophore labeling. Assignment of the separated bands was accomplished by their molecular masses derived from a calibration curve, using the procedure described above. In this case, the calibration curve was plotted based on the electrophoretic mobility values of the five-protein test mixture (lane B). The molecular masses attained were then compared to (24) Sigma Catalog, Sigma Chemical Co., St Louis, MO, 1997; p 1895.
available literature values,25 and some of the basic milk proteins such as R-lactalbumin (band 1), β-lactoglobulin (band 2), and casein (band 3) were recognized. The faint band at 16 min in Figure 8 has not been identified. CONCLUSION SDS gel electrophoresis is one of the most frequently used tools for the separation of complex protein mixtures. With the recently increasing interest of proteome analysis, SDS gel electrophoresis became even more important, since it is used almost exclusively as the second separation dimension in two-dimensional (2D) electrophoresis. Although the method is informative, it is cumbersome and time-consuming and lacks of automation. To the best of our knowledge, this is the first report describing an automated, high-performance SDS gel electrophoresis system, that is based on electric field-mediated separation of SDS-protein complexes using an ultra-thin-layer platform, along with noncovalent, instant fluorophore labeling. Integrated fiber-optic bundlebased scanning laser-induced fluorescence detection technology readily provided real time detection of the separated proteins with high sensitivity. We have demonstrated rapid separations of noncovalent fluorophore stained proteins in the molecular mass range of 14-116 kDa in ∼15 min. The efficiency of noncovalent labeling as a function of staining dye and SDS concentrations was thoroughly investigated and 0.5× and 0.05% were found to be the (25) Righetti, P. G.; Tudor, G.; Ek, K. J. Chromatogr. 1981, 220, 115-194.
optimums, respectively. The limit of detection (LOD) of our automated system was as low as 1.25 ng (41 fmol for carbonic anydrase). Practical examples of molecular mass determination and characterization of the complex protein mixture of low fat milk were also demonstrated. Besides the obvious advantage of the significant time saving of our method by avoiding long post- or preseparation staining/ labeling processes, ultra-thin-layer SDS gel electrophoresis offers full automation for detecting and visualizing the separated proteins. It also requires 3-4 orders of magnitude less staining dye for visualization, compared to regular postseparation staining methods. One of our future plans is to further develop this methodology toward protein labeling after the first dimensional isoelectric focusing step of two-dimensional electrophoresis (i.e., during the buffer equilibration step) in order to able to automate band visualization as well as to enhance the speed, detection limit, and specificity of proteome analysis. ACKNOWLEDGMENT This work was supported by the NIH-SBIR Grant 1R43CA80569 and the U.S.sHungarian Science and Technology Joint Research Fund, Project JF 654/96. The stimulating discussions with Drs. Aran Paulus and Ernst Gassmann are also highly appreciated. Received for review December 18, 1999. Accepted March 13, 2000. AC991501+
Analytical Chemistry, Vol. 72, No. 11, June 1, 2000
2525
