A Defined Methodology for Reliable Quantification of Western Blot Data
Chemiluminescent western blotting has been in common practice for over three decades, but its use as a quantitative method for measuring the relative expression of the target proteins is still debatable. This is mainly due to the various steps, techniques, reagents, and detection methods that are used to obtain the associated data. In order to have confidence in densitometric data from western blots, researchers should be able to demonstrate statistically significant fold differences in protein expression. This entails a necessary evolution of the procedures, controls, and the analysis methods. We describe a methodology to obtain reliable quantitative data from chemiluminescent western blots using standardization procedures coupled with the updated reagents and detection methods.
KeywordsWestern blot Densitometry Protein expression
Western blotting has been a staple in life science labs for several decades—ever since researchers published the first detailed description of this protein detection technique in 1979 . This multistep method determines the presence or absence, size, and modification or degradation states of target proteins, as well enables the quantitation of proteins from complex protein mixtures or homogenates [2, 3, 4]. However, there are many potential stumbling blocks in this procedure that can preclude reliable results. These include challenges related to every step of the western blotting procedure, from sample preparation, normalization, SDS–PAGE gel loading, protein transfer, primary and secondary antibody selection, incubations, and washes, detection method selection to densitometric analysis. A recent report predicts that approximately 25 % of the accepted papers include at least one inappropriately manipulated figure and many of these are associated with western blotting . This underlines the negative perception by which the scientific community views the western blot data. Thus, editors and reviewers of scientific journals are looking at western blot results, particularly at the densitometric analysis to determine the fold differences in protein expression, with greater scepticism, often requesting the raw data files.
Chemiluminescent western blot data, derived from film-based detection, poses distinct challenges in producing quantifiable, reproducible data. These problems stem from a low-dynamic range of detection and the difficulty in accurately determining the limit of detection [6, 7]. The scientific community has largely ignored these challenges, mostly because of a common misperception that film produces the highest quality data from western blots. However, unless the experiments are performed with a deep understanding of these limitations, this method of detection is an approximation at the best and often nonquantitative if used inappropriately. By contrast, the rapid evolution of affordable and highly sensitive gel and blot imaging technologies coupled with new reagents gives researchers the means to produce truly quantitative western blot data—as long as the process is carried out with proper technique, validation, and controls. These new tools and techniques eliminate the limitations associated with film-based detection and meet the journal reviewers’ demands for quantifiable protein expression data.
Here, we will demonstrate how standardized protein samples, when processed with film versus digital imaging methods and different normalization approaches, produce vastly different results. Based on our findings, we propose a rigorous and simple methodology to produce high quality, reproducible, and quantitative western blot data.
Materials and Methods
Protein Sample Preparation and Separation
A mixture containing a lysate from HeLa cells and purified yeast alcohol dehydrogenase (ADH) (Sigma Aldrich) in Laemmli buffer (Bio-Rad) was used as the starting material for separation on Criterion TGX AnykD Stain-Free gels (Bio-Rad). All sample wells were loaded with 20 μl of the protein mixture with separation using the Criterion Dodeca gel apparatus (Bio-Rad) for 1 h at 200 V.
All gels were imaged using the stain-free application on the ChemiDoc MP (Bio-Rad) imager immediately after the protein separation and prior to western blotting.
Western Blot Transfer and Total Protein Imaging
Protein gels were blotted using the Trans-Blot Turbo transfer apparatus and PVDF Midi transfer packs (Bio-Rad). Membranes were immediately transferred to a blocking buffer for fluorescent western blotting (Rockland), and incubated with a gentle agitation for 1 h at room temperature. During blocking and after uniform wetting in blocking buffer, the membranes were imaged for the total protein transferred using the stain-free application on the ChemiDoc MP imager.
Antibody Incubation and Chemiluminescent Detection
Membranes were incubated overnight with gentle agitation at 4 °C in 30 ml of blocking buffer with a mixture containing anti-yeast ADH rabbit polyclonal Ab (ABCAM) (1:5000 dilution) and anti-human GAPDH mouse monoclonal Ab (Rockland) (1:10000 dilution). These blots were washed five times for 3 min in Tris-Buffered Saline with Tween-20 (TBST; 500 mM NaCl, 20 mM Tris–Cl, pH 7.5, 0.05 % (w/v) Tween 20), and incubated in 40 ml of a mixture containing goat anti-mouse HRP Ab (Bio-Rad) (1:50000 dilution) and goat anti-rabbit HRP Ab (Bio-Rad) (1:50000 dilution) in blocking buffer for 1 h with gentle agitation at room temperature. This was followed by five 3-min washes in TBST at room temperature and incubation in Clarity western ECL substrate chemiluminescent detection reagent (Bio-Rad) for 5 min prior to image acquisition.
Blot Imaging and Densitometric Analysis
Results and Discussion
Lysis of cells or tissue
Quantification of lysate total protein concentration
Equal loading and separation of samples using SDS–PAGE
Complete transfer of proteins separated on the gel to nitrocellulose or PVDF membrane
Determination of proper dilutions for primary and secondary antibodies
Optimal detection of the chemiluminescent signal
Quantification of densitometric data
Thus, a methodology to test and confirm the quality of each step should be employed.
Validation of the Loaded and Transferred Proteins
The handling, storage, and lysate preparation of tissue and cell culture specimens can be complicated by the nature of the sample itself and by the large array of reagents and equipment available to prepare them. Careful consideration should be given to determine the best approach to prepare samples in order to get a reliable end result. Since most lysis buffers contain detergents such as Triton X-100 or sodium dodecyl sulphate (SDS), a detergent-compatible protein assay should be chosen.
Following the transfer, the membranes were imaged to assess the transfer efficiency and lane-to-lane consistency of the transferred proteins, which retain the fluorescence imparted by the stain-free imaging process (Fig. 2c). Although, the linear dynamic range was much narrower for the transferred protein, a very good correlation (R2 = 0.9971) to the two-fold dilutions was obtained in the typical range (between about 10 and 70 μg) of total protein load for most western blotting techniques (Fig. 2d). The reduced sensitivity of the fluorescent signal from the membrane versus the gel is the result of some protein transferring through the membrane (over-transfer) and much higher background due to the autofluorescence of the PVDF membrane under UV illumination during stain-free imaging. The combined effects of over-transfer and variable transfer efficiency support the use of densitometric data produced from the membrane for normalization as opposed to the gel.
The Quantitative Linear Dynamic Range: Contrasting Detection Platforms
For the endogenous protein GAPDH, both film and the ChemiDoc MP were only linear for the lowest three dilutions. This can be attributed to very high abundance of this protein in the HeLa cell lysate (Fig. 3). In this case, more dilutions and a longer exposure time would extend the linear dynamic range, but these data show that to use GAPDH as a reliable and quantifiable loading control, a total protein load of not more than about 0.5 μg is required (Fig. 3).
For the film images, a slight decline in the density was observed with increasing protein load within the plateau region (Fig. 3b). This can be attributed to the saturation of background between and around the bands of interest. Given that the ADH and GAPDH bands of interest are already fully saturated at 4.4 μg of total protein load, the contribution of the increasing background density (that saturates quickly with the film) results in a net decrease in background-subtracted density for these proteins.
Optimization of Protein Loading for Quantitation
All blot and gel detection systems generate a two-dimensional image, and signal intensity forms the third dimension of information (Fig. 1). Therefore, if excessive protein is loaded in the gel lane such that the width of the gel has been filled, the detector (whether film, camera, or scanner) will only capture the signal from the protein that resides near the surface of the gel. The same effect is observed during blotting where the protein transferred from an overloaded gel lane will form a layer on top of the protein already bound to the surface of the transfer membrane such that the primary antibodies will only bind to the surface layer of the transferred protein [10, 11]. The plateau observed for the same blot using both film- and camera-based detection is not due to the saturation of the detector, but rather is the result of protein saturation within the gel itself and consequently of the blot itself with the multiple layers of target protein bound to the membrane (Fig. 3).
Since many labs typically load a specific, fixed amount of protein (typically between 10 and 50 μg) into the gel lanes without determining the optimal protein load, there is a strong potential for gel saturation, particularly for loading controls such as GAPDH, beta-actin, and tubulin. Although, this results in consistent band densities among the sample lanes, these data are often the result of overloading the gel for these abundant proteins such that the densities obtained are far outside the linear dynamic range in the plateau. The best way to avoid this issue is to produce a standard curve from a two-fold serial dilution series over 12 dilutions starting from about 80 μg of a pooled lysate from representative samples across the experimental conditions . A separate standard curve of band density versus protein load should be run to validate each primary antibody for western blot as shown in Fig. 3. In this fashion, the linear dynamic range of detection for each antibody can be determined, and the associated dilution factor required for individual sample loading can be obtained by diluting the samples to the mid point of the standard curve. In the present case, we determined that the linear dynamic range of ADH is between 0.04 and 2.5 ng of the purified protein (Fig. 4b, c).
Loading Controls for Quantitation
The transfer efficiency of protein to a blotting membrane can be inconsistent across the gel, resulting in a gradual two- to four-fold increase or decrease in the signal between the lanes. Furthermore, the preparation and quantitation of sample lysates for the concentrations coupled with their physical pipetting into the lanes of a protein gel can also lead to inconsistent densitometric data. Loading controls are useful to normalize these technical artifacts and become increasingly important when measuring small differences in protein expression between samples.
The most common loading controls include housekeeping proteins, such as GAPDH, beta-actin and tubulin, which are constitutively expressed proteins that maintain cell viability. However, these proteins are generally highly expressed in samples and are frequently overloaded in the gel lane with the target protein such that they would not serve to normalize the loading [12, 13]. Although, the densitometric data would be consistent, this data could be an artifact of overloading as observed in our experiments where GAPDH levels were reaching a plateau above 0.5 μg of HeLa lysate (Fig. 3). Furthermore, the housekeeping proteins themselves can be variably expressed between the experimental conditions, thereby eliminating their usefulness for the normalization of western blots [14, 15, 16, 17]. In order to avoid these issues, the total lane density of transferred protein on the membrane is being used for the normalization in lieu of the traditional loading controls [18, 19, 20, 21, 22].
The HeLa lysate total protein load of 0.34 μg (lane groups 5 and 6) was well below the dynamic range and even below the detectable limit for stain-free imaging (Fig. 5a), but within the linear dynamic range for GAPDH (Fig. 3). This makes GAPDH an ideal loading control for highly abundant proteins which require much lower sample loading. It is worth pointing out that in these cases, stain-free imaging cannot be used for normalization. Care must be taken to ensure that the amount of lysate loaded in this case is within the linear dynamic range of both the loading control and the target protein to ensure accurate, quantifiable, and normalized densitometric data.
Accurate Quantitation Using the Linear Dynamic Range
The 0.03-fold difference in HeLa lysate loading among lane groups 2, 5, and 6 (i.e., between 11 and 0.34 μg) was calculated to be only about 0.20–0.26-fold by relative band density of GAPDH (Figs. 6, 7, panel D contrast lane group 2 with 5 and 6 for “Fold GAPDH”). This can be explained by the fact that the total protein load of 0.34 μg is within the linear dynamic range of density for GAPDH but is in the plateau at 11 μg (Fig. 3). Thus, the densitometric fold difference is much smaller than expected because the two points are not within the linear range. This further underscores the importance of ensuring that the samples are diluted such that the loading control is well within the linear dynamic range of detection.
For ADH, the correlation between relative density and fold difference in ng quantity of spiked-in ADH protein load between all the lane groups was excellent when using the ChemiDoc MP imager (Fig. 6c), but poor with film (Fig. 7c). This is due to the different linear dynamic ranges obtained between film and the ChemiDoc MP for ADH (Fig. 4). All the loaded amounts of ADH (i.e., 0.17–1.39 ng) were within the linear dynamic range of detection for the ChemiDoc MP (Fig. 4b). However, this was not the case for the film, where the relative densities for ADH were measured from one value within the linear dynamic range (0.17 ng) and two values within the plateau (0.69 and 1.39 ng) (Figs. 3, 4).
Accurate quantitation of relative protein expression is possible only by following appropriate experimental procedures to determine the linear and quantitative dynamic range for each target protein under a given set of experimental conditions. We show that only by producing a two-fold dilution series of the protein lysate can the linear range of quantitation be determined, and this is entirely dependent on the abundance of each target or loading control protein in the sample. Thus, each antibody in a given study should first be tested with a dilution series of a pooled protein lysate from the study samples. This will ensure that the appropriate dilutions of samples are used for accurate and normalized quantitation of the target proteins by means of densitometric analysis.
Film does not always offer the dynamic range to quantify the full range of protein expression among samples, and this can become further complicated by the need to objectively determine the saturation point of the target protein bands. The advent of cooled CCD camera technologies has permitted the automated and precise determination of the camera CCD saturation point of chemiluminescence, and permits a broader dynamic range than possible with film. Coupled with the appropriate use of sophisticated software analysis tools, accurate densitometric analysis of western blots is indeed possible.
Since many of the traditionally used loading controls are highly abundant and usually loaded in saturating quantities in the typical range of sample loading (10–70 μg per lane), they cannot be used for accurate normalization. Since many labs are publishing small changes (between two- and four-fold) among samples from western blots, accurate normalization becomes critical to ensure that the reported changes are real. The use of stain-free gel technology permits accurate normalization in the range of 10–70 μg of the total protein load, and therefore provides an excellent solution for normalization between the lanes of western blots by total protein transferred.
To conclude, we propose a rigorous methodology of validating sample loading, standardizing antibody dilutions, determining the dynamic range with a sensitive, camera-based imaging system, and use of a stain-free technology to get high-quality and reliable quantitative data from western blots.
Conflict of Interest
The authors declare no Conflict of Interest
- 2.Bio-Rad Laboratories (2011) Protein Blotting Guide, Interactive PDF. Review B, Bulletin 2895.Google Scholar
- 3.Lin, J. C., Choi, E. I., & Pagano, J. S. (1985). Qualitative and quantitative analyses of Epstein–Barr virus early antigen diffuse component by western blotting enzyme-linked immunosorbent assay with a monoclonal antibody. Journal of Virology, 53, 793.Google Scholar
- 6.Schiavini, D. G., Puel, J. M., Averous, S. A., & Bazex, J. A. (1989). Quantitative western immunoblotting analysis in survey of human immunodeficiency virus-seropositive patients. Journal of Clinical Microbiology, 27(9), 2062–2066.Google Scholar
- 7.Elbaggari, A. McDonald, K., Alburo, A., Martin, C. (2008) Imaging of chemiluminescent western blots: comparison of digital imaging and X-ray film. Bio-Rad Tech Note 5809.Google Scholar
- 10.Dubitsky, A. and Perreault, S. A Model for Binding of DNA and Proteins to Transfer Membranes. Pall Corporation Tech Note.Google Scholar
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