Western blotting revisited: critical perusal of underappreciated technical issues.

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DOI 10.1002/prca.201400118

Proteomics Clin. Appl. 2015, 9, 396–405

REVIEW

Western blotting revisited: Critical perusal of underappreciated technical issues Thomas A. Gorr1,2 and Johannes Vogel1 1 2

¨ ¨ Institute of Veterinary Physiology, Vetsuisse Faculty, University of Zurich, Zurich, Switzerland Center for Pediatrics and Adolescent Medicine, Clinic IV: Division of Pediatric Hematology and Oncology, University Medical Center Freiburg, Freiburg, Germany

The most commonly used semiquantitative analysis of protein expression still employs protein separation by denaturing SDS-PAGE with subsequent Western blotting and quantification of the resulting ODs of bands visualized with specific antibodies. However, many questions regarding this procedure are usually ignored, although still in need of answering: Does isolation or separation procedure harm the integrity or affect modifications (e.g., phosphorylation) of the protein of interest? Does denaturation reduce binding of antibodies used for detection? Should denaturation be performed or should a native gel be run? How can artificial degradations or aggregations be distinguished from biological relevant ones? If the antibody detects multiple bands (which is not uncommon), which one(s) should be taken into account for quantification and why? Which loading control protein should be chosen and is it really “housekeeping” and how can this be verified? Is the image acquisition system linear and does it come with a sufficient dynamic range? How to account and control for background staining? This article is intended to address these questions and raise the readers awareness to possible Western blot alternatives in the attempt of minimizing possible pitfalls that might loom anywhere from protein isolation to acquisition of final quantitative data.

Received: August 27, 2014 Revised: December 19, 2014 Accepted: January 14, 2015

Keywords: 2D-PAGE / Antibodies / Densitometry / Protein quantification / Proteomics

1

Introduction

Several different methods can be used to quantify proteins in biological samples—all of them with advantages and disadvantages. However the standard SDS-PAGE/Western blot, which includes a 1D separation of dissolved polypeptides according to their molecular mass, is used by many laboratories to investigate or demonstrate expression changes of a given protein between control states and experimental conditions. This technique is still the method of choice for basic research, but also a useful tool in clinical applications. In principle, any biological sample can be examined by Western blot, ranging from cell or tissue extracts to body fluids, such as plasma or serum or urine. Correspondence: Professor Johannes Vogel, Institute of ¨ Veterinary Physiology, Vetsuisse Faculty, University of Zurich, ¨ Winterthurerstr. 260, CH-8057 Zurich, Switzerland E-mail: [email protected] Fax: +41 44 6358932 Abbreviations: PSR, protein synthesis rate; PTMs, posttranslational modifications; SDS-PAGE, sodium dodecyl sulfatepolyacrylamide gel electrophoresis C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

To sensitize the reader to potential problems associated with protein identification and quantification, we will subsequently discuss some of the hazards related to the highly variable structural and functional properties of proteins. We will also address potential pitfalls during protein isolation and separation, as well as those associated with band detection and quantification methods in Western blot experiments. A lack of awareness or attention to such issues makes comparisons of studies difficult.

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Problem 1: The variable nature of proteins

Protein diversity is driven through three main processes: first, at DNA level through differences in sequence (i.e., gene duplicates (paralogs), allelic forms, gene polymorphisms), second, at precursor messenger RNA (pre-mRNA) or mRNA level (i.e., alternative splicing), and, finally, at the level of protein themselves (i.e., covalent PTMs). For example, according to nomenclature rules by the International Union of Pure and Applied Chemistry (IUPAC), first published in 1964 and eventually revised and adopted in 1976, the term www.clinical.proteomics-journal.com


isozyme/isoenzyme, coined by Hunter and Markert in 1957 [1], applies to multiple forms of enzymes that catalyze essentially the same reaction within a single species, and yet, arise from genetically determined differences in primary structure (i.e., from paralogs, alleles, polymorphisms), but not from a modification of the same primary sequence (i.e., PTMs; see IUPAC: http://www.chem.qmul.ac.uk /iubmb/ misc/isoen.html). Due to the high sensitivity and specificity of enzyme catalytic reactions, the characterization of isoenzymes based on their catalytic properties is still used (see review in [2]). With regard to the standard SDS-PAGE Western blot, protein diversity is easily displayed in form of extra bands detected by the same antibody once protein separation is achieved. These extra bands can occur either due to PTMs or artifactual chemical modifications (see below) induced during storage, protein isolation, and separation. Be this as it may, with the advent of 2D electrophoresis techniques [3], and particularly the coupling of mass sepctrometry analytics with 2D electrophoresis and LC (reviewed in [4]), the diversity of proteins reached unanticipated heights. Today it is well appreciated that the human genome, as a result of (i) the allelic variations behind isozymes or (noncatalytical) isoforms, (ii) posttranscriptional (pre-)mRNA splicing events affecting 40– 60% of all genes (see [2] and references cited therein), and (iii) several hundred to, perhaps, >1000 of covalent PTM categories, is able to generate hundreds of thousands of different gene products from an estimated 22 500 open reading frames [5]. Clearly, the old one-gene/one-protein doctrine has yielded a huge underestimation of the omic-scale translational capacity in humans and other mammals. In the characteristic 2D gels of proteomic studies, solubilized proteins are being separated in accordance to their pI through iso-electric focussing (first dimension), and their molecular mass during the following SDS-PAGE (second dimension). Here, the same protein is often detected in form of multiple spots of similar appearance in a relatively restricted portion of the gel, and, in many cases, in the form of a train of spots. To account for this spot multiplicity, Jungblut et al. introduced the term protein species in 1996 [6, 7] to define proteins bearing any chemical modification, including those derived after alternative splicing and PTMs [2]. Thus, a protein species describes any individual member of a family of single gene-derived proteins whose protein chain length (due to alternative splicing or proteolytic processing events) and/or covalent PTMs sets it apart from other family members [8]. This means that iso(en)zymes and protein isoforms represent a subset of protein species, since they are also chemically different. In contrast, two proteins with different PTMs represent different protein species, but not different isoforms [8]. PTMs are extremely important covalent modifications as they can adjust the physical and chemical properties, conformation, stability, and activity of a protein, and that way, alter the function of the polypeptide (see [9, 10] for review). In that sense, PTMs can offer valuable clues regarding the activity state of the respective signaling pathway containing this par C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

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Figure 1. Expression and phosphorylation status of the facultative inhibitor of cap-dependent mRNA translation, 4EBP1, as obtained by Western blotting. Cytosolic extracts of human hepatoma (Hep3B, lanes 1–3), breast carcinoma (MCF7, lanes 4–6), and cervical carcinoma cell lines (HeLa, lanes 7–9) subjected to 16 h of normoxic (N, air), hypoxic (H, 1% O2 ), and near-anoxic (A, 0.2% O2 ) atmospheres. To slow ATP-costly protein synthesis during stress conditions, such as severe O2 deprivation, 4EBP1 needs to be hypophosphorylated to hinder, through direct binding, the rate-limiting and cap-binding initiation factor eIF4E from associating with partner proteins as translation-promoting eIF4F complex (see text for more background). Blotting 4EBP1 with an antibody for all phosphospecies (top panel, 4EBP1 (53H11) rabbit mAb #9644, Cell Signaling Technology (CST), Danvers, USA) allows discrimination between hyperphosphorylated (␥/␤ bands) and hypophosphorylated protein (␣ band). In the shown cancer cells responses to O2 deprivation range from slight (HeLa) to pronounced (Hep3B) shifts from hyper- to hypophosphorylated 4EBP1 species (increasing ␣ band intensity), relative to normoxia. These changes are even more clearly elucidated with an antibody (i.e., nonphospho-4EBP1 (Thr46; 87D12) rabbit mAb #4923; CST) specifically detecting the active, eIF4E-scavenging species of 4EBP1 that is dephosphorylated at Thr46 (second panel from top, slight and massive increase of this species in hypoxic/anoxic HeLa and Hep3B, respectively), and an antibody (i.e., phospho4EBP1 (Thr70) Antibody #9455; CST) directed against inactive 4EBP1, phosphorylated at Thr70 (third panel from top, diminished signals in hypoxic/anoxic HeLa and Hep3B). However, breast carcinomas notoriously show loss of this O2 -dependent regulation of protein synthesis [52]. In line with this, MCF7 breast carcinoma cells display, relative to normoxic controls, elevation of both hypophospho (␣ band with total antibody; band with Thr46dephospho antibody) and hyperphospho signals (bands with Thr70-phospho antibody) of 4EBP1 upon O2 deprivation. Loading control, ␣-tubulin (bottom panel).

ticular polypeptide [9, 10]. As an example, Fig. 1 documents O2 -mediated changes on the phosphorylation status of the facultative translation inhibitor factor 4E binding protein 1 (4EBP1) that affect the localization of the protein within the gel. Stress conditions, such as severe O2 deprivation, inactivate the mammalian target of rapamycin kinase complex 1 (mTORC1) in multiple cancer backgrounds, thus triggering hypophosphorylation in the mTORC1 effector 4EBP1. The resulting slowdown in the translation of capped mRNAs by hypophosphorylated 4EBP1 is believed to foster energy conservation as key adaptation of tumor cells toward developing www.clinical.proteomics-journal.com

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T. A. Gorr and J. Vogel


Figure 2. Effect of storage conditions on human plasma and urine proteome. The shown samples had been analyzed with capillary electrophoresis (CE) directly coupled with MS (CEMS). CE migration time in minutes (xaxis) is plotted against the m/z ratio (yaxis). Panel (A) depicts fresh plasma and panels (B) and (C) plasma stored at 4⬚C for 2 h or at 4⬚C for 2 h followed by freezing/thawing, respectively. Additional signals not found in the freshly analyzed sample are marked with boxes, whereas arrows indicate considerable signal broadening as a result of plasma sample storage. This suggests numerous degradation fragments of high molecular weight proteins, slightly varying in molecular weight as well as small fragments with low m/z ratio. Panels (D) to (F) show urine samples where (D) depicts the fresh sample and (E) and (F) urine samples stored under the same conditions as the plasma samples shown in (B) and (C). Note that the urine proteome remains completely unaffected by the storage conditions. Reprinted as relabeled grayscale image with permission from [21]. Copyright (2009) American Chemical Society.

hypoxia tolerance and radioresistance (see [11–13] for details). As illustrated in the figure, it is changes in the Western blot banding pattern that can be used to quantify the fraction of the phosphorylated form of a given effector protein (i.e., 4EBP1) as a proxy of the signaling activity of a considered pathway (i.e., mTORC1). If molecular mass changes occurring as a result of PTMs are too small to result in extra bands, they can also be detected with antibodies specific for a given alteration (Fig. 1). Due to the magnified resolution power of IEF-based protein separations (e.g., in 2D approaches), this technique raises the risk of detecting false-positive or spurious spots that will only lead to an exaggerated notion of protein diversity that is, however, irrelevant to the sample distinction at hand [14]. Spurious spots in IEF are known to result from the SH-group mediated aggregation of reduced, but nonalkylated, polypeptides; an unwanted side effect particularly aggravated in the alkaline pH region of the gel [15]. Another cause of misleading bands or spots after IEF separations, also prevented by SH-group alkylation, can stem from an increasing loss of H2 S groups from cysteine residues by ␤-elimination (also called desulfuration), which can culminate in serious degradation of the polypeptide [15, 16]. Redox artifacts, such as salt-induced electrolytic reduction of protein disulfide bonds and carboxylic acids, have further been reported from 2D separations [17]. Lastly, multiple IEF bands of one and the same heme-binding protein can result from pI-differing frac C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

tions with fully ferrous, fully ferric, and mixed ferrous/ferric hemes [18, 19]. To, thus, keep protein isolation results reproducible and yields high, formation of such artifacts needs to be countered through protein extraction at low temperatures with pH7–8 buffers containing reducing plus alkylating agents and protease inhibitors (below). All these points stress the necessity to actually consider multiple bands seen with an antibody in Western blots, or streaks of spots in 2D gels, when determining the total protein amount since these variant signals may well include native, modified, oxidized, aggregated, as well as partially degraded forms of the protein. Another common cause for additional bands is poor quality of the antibodies used (see below). Thus, if multiple bands/spots are excluded from quantification, the scientific rationale for it should clearly be stated.

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Problem 2: Introducing artifacts during the protein isolation procedure

As hinted above, proteolysis or degradation are other main sources of protein band/spot artifacts in addition to oxidative processes that can markedly be affected or initiated by events already during storage, but also during protein extraction (Fig. 2). Extraction of proteins from cells or tissues by cell lysis or disruption and homogenization of the tissue at hand per se can introduce degradation, aggregation, or other www.clinical.proteomics-journal.com

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modifications that complicate final detection or quantification of the protein of interest. Almost all biological samples contain proteases or peptidases and consequently appropriate inhibitors of these enzymes should always be added to the sample or extraction buffer (see below). Preferred samples in the clinical context are body fluids because they can be obtained with minimal invasion (plasma/serum) or noninvasively (urine). However, blood clotting as a proteolytic process may induce protein degradation and unwanted by-products. Therefore, the human proteome consortium recommends analyzing blood as plasma rather than serum [20]. Nevertheless, plasma obtained under these optimized conditions retains some proteolytic activity and should consequently be processed as quickly as possible [21]. In contrast, the urine proteome is much more stable as it may not contain significant concentrations of proteases. In line with this notion it could be shown that storage of urine samples at room temperature or at 4⬚C with or without several freeze/thaw cycles for 6–24 h did not harm significantly the integrity of the proteins contained in that body fluid [22, 23] (Fig. 2). To minimize the introduction of artifacts during sample preparation, a fresh, unprocessed sample would be ideal as in an ELISA-based analysis of a specific protein of interest. Proteomics, in contrast, aim to detect/quantify the changes of entire protein mixtures. In such settings, nonprotein molecules, such as lipids or carbohydrates (e.g., from cell membranes), may interfere with the subsequent separation and detection of the proteins in Western blots or 2DE. To keep the probability of artifact introduction low, unwanted low-molecular weight components in the sample should be removed in as few steps as possible. Importantly, conditions for protein extraction or purification must be adjusted according to the nature of the samples and proteins of interest as well as the assays to be run [24,25]. A distinction of particular relevance in this regard is whether native or denatured proteins are going to be examined. If proteins are needed with their natural fold and assembly still intact (native state), extraction should (1) comprise lysis protocols based on physical means and/or nondenaturing detergents (i.e., nonionic detergents TritonTM X-100 or Tween series; zwitterionic detergents, such as CHAPS), (2) be carried out at 4⬚C or on ice to slow proteolytic processes and further the solubility of the majority of harvested proteins, (3) be done swiftly in a nonstop fashion until proteolysis inhibitors can be added to the extraction medium, and (4) completely avoid denaturing agents, such as ionic detergents (i.e., SDS), strong chaotropes (i.e., urea, guanidium chloride), cross-linking compounds (e.g., formaldehyde), acids, and heat.

enzymatic activity assays, native PAGE, coimmunoprecipitation, electrophoretic mobility shift assays, and spectroscopic analysis of chromophore-bound polypeptides. However, routine isolations and subsequent electrophoretic, Western blot, and MS analyses are primarily conducted with denatured protein samples. This time, denaturing agents, such as SDS, are intentionally added to the extraction buffer for a maximal yield of soluble proteins. Lysis of cells and disruption of tissues can be accomplished by physical means of disintegration, such as rotating blades (e.g., blenders), liquid homogenization (e.g., Dounce homogenizer, French Press), and cell shearing by sonication or repeated freeze-thaw cycles. Additionally, tissue samples are routinely frozen in liquid nitrogen and crushed by manual grinding with mortar and pestle. More recently, detergentbased lysis methods have become the norm. The amphipathic nature of detergent molecules, that is, the fact that they contain both a nonpolar “tail” with aliphatic or aromatic character and a polar, in some cases ionic, “head,” allows an efficient release of soluble proteins and dispersion of membrane proteins and lipids. In order to reduce the complexity of whole-cell samples, methods to purify or enrich specific subcellular compartments are commonly used, for example, plasma membrane fraction, mitochondria, nuclei, synaptic vesicles, etc. We routinely subfractionate cell extracts in form of cytosolic and nuclear compartments according to [26]. Whether one uses whole cell extracts or subfractioned compartments, a cocktail of proteolytic inhbitors should be added to the extraction buffer to further (in addition to 4⬚C/ice conditions) slow degradative processes and maximize the yield of intact polypeptides. Typically, commercial inhibitor cocktails contain 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride, aprotinin and leupeptin (serine proteases), E-64 and leupeptin (cysteine proteases), pepstatin A (aspartic acid proteases), as well as bestatin (amino peptidases), or any subset thereof, to provide protection from degradation by indicated protease and peptidase systems. To prevent the dephosphorylation activity of serine/threonine phosphatases plus tyrosine phosphatases, cocktails may additionally contain sodium fluoride, beta-glycerophosphate, sodium pyrophosphate, and sodium orthovanadate, respectively. If the goal is to isolate proteins with a particularly fast turnover (e.g., hypoxia-inducible transcription factor alpha subunits are degraded with t1/2 < 5 min once hypoxic cell cultures are processed under the reoxygenating conditions of room air [27]), addition of the freshly prepared serine protease inhibitor PMSF (dissolved in methanol; unstable in water/buffer) to the inhibitor mix just prior to cell lysis might also be advisable. Commercial cocktail formulations are available with or without the cation chelating metalloproteinase inhibitor ethylenediaminetetraacetate.

4 Stored aliquots of extracts should also not be subjected to any of the agents listed under “d.” Once native proteins are isolated, functional assays can be carried out, such as C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

Problem 3: Loading control

To correct for loading errors or uneven transfer efficiencies, usually normalization to so-called “housekeeping proteins,” www.clinical.proteomics-journal.com

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such as glyceraldehyde 3-phosphate dehydrogenase (GAPDH), ␤-actin or ␣-tubulin, is done in Western blot experiments. However, during the last decade it has become increasingly evident that in many circumstances genes classically used as “housekeeping” genes may not be appropriate at the protein level as expression can be markedly affected in many pathological situations or in response to biological stimuli [28–30]. This applies not only to the transcriptional level as shown, for example, in a recent qRT-PCR study of the postnatal development of the rat retina where mitogen-activated protein kinase 1, generally not considered a housekeeping gene, was found to be a much better internal control than 18S rRNA or ␤-actin [29]. More importantly, expression of “housekeeping” proteins can vary considerably between different cell lines and tissues or as function of experimental conditions [31, 32], and can even be affected just by the density of cultured cells [33]. Thus, in principle one needs to validate, for each experimental design, which candidate housekeeping gene really is constantly expressed and not affected by the different experimental conditions considered. In addition, it is necessary to know the linear concentration range of the “housekeeping” as well as the target protein to avoid errors introduced when concentrating or diluting the loaded sample (cf. below 8, Problem 7). Practical strategies for overcoming this dilemma may include the use of multiple reference genes, once they have been tested for their stable expression in the considered experimental conditions [29], and assessing the linearity of the concentration and OD in the desired concentration range of the sample [31]. Alternatively, exact quantification of the loaded protein amount (Fig. 3), and the addition of a reference marker of known concentration [34], allows omission of housekeeping proteins altogether [30]. To avoid tedious assays for finding the proper loading control, absolute quantification of the loaded protein amount can be used, provided that the total protein synthesis rate (PSR) of the cells/organs remains unaffected by the particular conditions tested (i.e., many hypoxic or anoxic cell cultures aim to lower their energy expenditure by a marked slow-down of total PSR; see Fig. 1 and [31]). Yet, if PSR is a constant across conditions, this approach might well prove to be superior to internal loading controls. Of course, the quality of absolute protein controls stays and falls with the accuracy of the protein concentration measurements [30] (Fig. 3).

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Problem 4: Membrane transfer

Successful and quantitative detection of proteins by Western blotting also relies on the quality of protein transfer from the gel to the blotting membrane. The traditional and widely used Laemmli system for protein separation in SDSPAGE/Western blotting [35,36] can result in band distortion, loss of resolution, and spurious bands due to protein deamination and alkylation at the high pH (9.5) of the separating gel, reoxidation of reduced disulfide bonds due to the varying redox state of the gel, as well as cleavage of aspartyl-prolyl C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim


peptide bonds due to heating the proteins in Laemmli buffer (pH 5.2) [37]. The Bis-Tris gel system, which operates at a neutral pH (7.0), provides significant benefits over the Laemmli system as it improves protein stability, minimizes protein modifications, maintains proteins in their reduced states, and prevents aspartyl-prolyl cleavage during electrophoresis. Moreover, the Bis-Tris gel system allows higher resolution and separation, because it produces sharper bands, thus, increasing sensitivity and providing more reliable data. In addition, the electrophoresis run time is reduced to 35 min [37,38]. With focus on the blot procedure, the transfer is generally achieved by electroblotting because this method is fast and ensures a rather complete transfer of proteins. The translocation of polypeptides by electroblotting includes actually three different methods that can be classified as wet, semidry, and dry transfer depending on the amount of transfer buffer required. The traditional wet transfer offers high efficiency and is, therefore, good practice. However, it takes considerable time and effort. For convenience and to save time, semidry blotting has been introduced, however, often at the expense of transfer quality. A more recently introduced method offering highquality transfer in conjunction with speed and convenience R Dry Blotting System from Life Technologies. is the iBlot Apart from the transfer method itself, a considerable number of factors can further influence the membrane transfer of proteins. First, transfer is more complete and faster if thinner gels are used. Yet, thin gels easily crack while handling. Therefore, gels with a thickness of 0.5 mm may be a practical compromise. Second, transfer efficacy depends on the type of gel used. The most common SDS-PAGE gels contain the proteins as anions and thus the membrane has to be mounted between gel and anode. In other settings using urea gels, the membrane should be placed between gel and cathode. Another inherent problem involves the potential hindrance of protein movement by cross-linking agents, such as acrylamide. While polymerized acrylamide forms, of course, the basis for the separation of proteins, one should consider the lowest cross-linker concentration possible to still yield the required resolution and keeping interference effects during protein transfer to a minimum. Especially high molecular weight proteins are affected by this problem that may, if necessary, be solved using heat, special transfer buffer compositions, or even partial in-gel protein digestion prior to membrane transfer [39–42]. On the other hand, the risk of washing off particularly small proteins/peptides from the membrane during the subsequent staining procedure can be reduced through a slight glutaraldehyde fixation of the proteins once they have been transferred to the membrane [43]. Third, the type of membrane material used will determine the transfer and later detection efficiency. The most commonly used membranes are made of nitrocellulose (NC), polyvinylidine diflouride (PVDF), and nylon. All these membranes have a microporous surface that provide (i) a large volume to surface area ratio, (ii) high binding capacity, (iii) possibility of storage of immobilized proteins, (iv) ease of www.clinical.proteomics-journal.com


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Figure 3. Comparison of total protein determination versus b-actin or b-tubulin as loading controls. (A) Image of a protein dilution series (1, 5, 10, 20, 30, 40 ␮g) of whole brain homogenate and visualization of ␤-actin and ␤-tubulin using the LICOR system and its Odyssey fluorescence-imaging scanner. (B) Quantitative densitometry of this dilution series shows the limited linear ranges of ␤-actin (black circle) and ␤-tubulin (open triangle). Note that the tubulin signal appears to saturate at less than 10 ␮g of brain homogenate. (C) At a lower concentration range (0.5, 1, 2, 4, 6, 8, 10,12, and 14 ␮g), linearity of ␤-tubulin appears to be better as demonstrated by the densitometry data (D). (E) Total protein stain of dilution series (2, 10, 20, 40, 80 ␮g) of a BSA standard (2 ␮g/mL). Imaging of this dilution series demonstrates a broad concentration range without saturation at a single protein concentration as shown by the graphical representation of the ODs of the respective bands (F). This demonstrates wide linear detection with high correlation (0.998) proving total protein determination as a useful method for controlling protein load. (G) Total protein stain of a whole brain homogenate dilution series (1, 5, 10, 20, 30, 40 ␮g). (H) Correlation between the total OD of the total protein stain (gray line) and the bicinchoninic acid assay (BCA, black line) for the shown protein dilution series demonstrates the broad concentration range detectable linearly without saturation and with high correlation (0.996 and 0.979, respectively), validating the use of total protein measurements as a ‘‘loading control’’ for quantitative Western blotting. Reprinted as grayscale image from [30] with permission (doi:10.1371/journal.pone.0072457.g004).

processing by allowing solutions to penetrate into the membrane and interact with the bound proteins, (v) lack of interference with the detection strategy, and (vi) reproducible detection. NC binds proteins most likely by hydrophobic, noncova C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

lent interactions. It can be handled with all required chemicals to detect and visualize the bound proteins; however, because it is relatively fragile it cannot be repeatedly stripped and reprobed. In contrast to NC membranes, PVDF membranes www.clinical.proteomics-journal.com

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can be used more easily for post-Western blot analysis, such as N-terminal sequencing or proteolysis-peptide sequencing. Nylon membranes have a much better mechanical stability and about six times higher protein-binding capacity than NC membranes. The latter however causes, as a side effect, higher unspecific binding of antibodies as well as dyes (Coomassie blue, Ponceau S, fast green, or toluidine blue) used for specific band detection or general protein visualization, respectively.

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Problem 5: Antibodies

As already mentioned above, additional bands seen in a Western blot can be due to physiological diversity (i.e., isoforms, species) or artifactual degradation or aggregation due to suboptimal sample handling. A third reason for extra bands is nonspecific antibody binding. In principle, the primary as well as the secondary antibody (or both) can result in unspecific bands. Generally, we all rely on commercially available antibodies hoping that they have been tested sufficiently for our application and that we can trust the supplier’s data sheets. However, at least some simple verification of the specificity of the antibody used lies in the users best own interest. First, one should be aware that polyclonal antibodies derived from immunized animals will detect a multitude of antigenic epitopes, be it on the targeted or on other (nonspecific) proteins on the blot. Thus, whenever possible, the use of affinity purified or monoclonal antibodies is recommended. Specificity of the desired secondary antibody can simply be assessed by omitting the primary antibody, which should result in an unstained membrane. Specificity of the primary antibody can be tested by probing the membrane with an irrelevant “nonimmune” primary antibody or even better by using extracts from cell that do not contain the target protein (i.e., from cells treated with small interfering ribonucleic acid, or knockdown clones) or from tissue samples of knockout animals. If available, recombinant peptides from the targeted protein can also be used to see if their excess inclusion during staining is able to quench the signal. Finally, it is worth mentioning that nonspecific results can be obtained from detection reagents, for example, avidin or streptavidin binding to endogenous biotin [44–46]. Thus, one should use blocking agents to endogenous biotin before adding the secondary antibody when using avidin/streptavidin-based detection systems. If the antibodies do not perform as expected, one has to browse (unfortunately costly and time-consuming) possible alternative antibodies with better performance or consider using an antibody-independent approach, such as silver-stained gels in conjunction with a small interfering ribonucleic acid driven knockdown of the targeted transcript and/or through subsequent MS-based peptide identification of likely candidate bands/spots if no a priori sequence data are available. C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim


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Problem 6: Image acquisition

We previously analyzed the problem of image acquisition and quantification of band intensities (cf. also 9, Problem 8) [47]. Briefly, the main problems arise when digitizing X-ray films. The employed acquisition system should have a linear dynamic range that at least is greater than that of the film, no (or at least disabled) automatic gain control and allow shading correction. However, the best way to avoid the pitfalls inherent with the image acquisition step from films is simply to omit it and directly acquire the images using desktop darkrooms or phosphoimagers. The most advanced way is fluorescence “in-gel” band detection, as it avoids not only image acquisition from a film, but also membrane transfer. Moreover, this technique allows simultaneous visualization of two to three separate proteins on the same gel without additional experimental time required. As the detection is performed within the infrared spectrum, the background is extremely low and generally also avoids the need for background correction (cf. last section), resulting in a signalto-noise ratio considerably better than ECL detection (biosupport.licor.com/docs/AppNote_InGelWesternDet_98812954.pdf). Finally, the fluorescent detection also provides a much higher dynamic range than ECL [48, 49].

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Problem 7: Linearity of the signal

Another significant problem associated with antibodydependent protein detection in Western blots is the linearity between protein concentration and the final optical density (OD) signal. Linearity between protein concentration and OD with corresponding slopes must be obtained for the target and loading control protein(s). Unfortunately target and loading control protein often show pronounced differences in their concentration, especially when low abundance proteins are to be investigated. To bring this problem to the reader’s attention, it was shown for ␤-actin signals, perhaps the most commonly used loading control protein, to not allow discrimination of different protein concentrations. Even diluting the antibody and/or shortening, the incubation time had little effect on its signal intensity [28]. The abundance of endogenous ␤-actin protein in many source materials is so high that the OD signal all too easily will reach saturating intensities. Other control proteins, such as GAPDH, may perform better [28], but the concentration of this “housekeeping” proteins as well as other ones in the loaded sample is also not linearly correlated with the OD of the final blot image and the limited linear range of each of these correlations is found at different concentration ranges [31]. Although other proteins so far not tested in this regard may behave differently, they most likely also fail to display an entirely linear correlation between their signal intensities and concentrations. Consequently, when attempting to quantify ODs from Western blot experiments, linearity should be ascertained for the target as well as www.clinical.proteomics-journal.com

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Table 1. Checklist to avoid pitfalls in Western blot experiments

Step

Things to care/think about

Recommendations

Protein isolation

Proteolysis, dephosphorylation, native or denaturizing gel

Loading

Rationale behind choosing a certain “housekeeping” gene. Is expression of this gene affected by the experimental condition

Membrane transfer

Is transfer efficient? Are the bands well (enough) separated

Antibody staining

Specificity, background staining, and linearity of the antibodies used

Signal linearity

Is the OD of the blot image linearly correlated to the protein concentration?

Image acquisition

When using X-ray films: automatic gain control, dynamic range of the imaging system. Dynamic range of the X-ray film. Shading correction Background correction algorithm? Which bands to include? Sample tool size? Which parameter

Cell/tissue lysis at +4⬚C/on ice; add protease (e.g., aprotinin, leupeptin, pepstatin A) and phosphatase (e.g., sodium fluoride, beta-glycerophosphate, sodium orthovanadate) inhibitors, and others depending on the protein of interest (e.g., PMSF for hypoxia inducible transcription factor 1␣ detection) Check for constant expression of the “housekeeping” gene in your setting. When possible try absolute protein quantification and avoid loading control genes. In the latter case check if total protein expression is not affected by your setting For efficient transfer prefer wet transfer or the R iBlot Dry Blotting System. Try replacing the Laemmli buffer with the Bis-Tris gel system. If the equipment is present, use “in-gel” staining with fluorescing antibodies and omit the membrane transfer If you doubt the specificity of the antibody or cannot control excessive background staining, try other antibodies or use cells/tissue from knockdown experiments Use dilution series to check for linearity in the desired concentration range. Check for the dynamic range and linearity of the image acquisition system and/or X-ray film Switch whenever possible to direct image acquisition that avoids the need of X-ray films. Dynamic range of infrared fluorescence systems is much higher than that of ECL-based detection First try to get rid of as much of the background as possible. Then use baseline subtraction as correction algorithm. Explain why you exclude bands and if you cannot take all into account. Tool size should be 30% of the lane width. Use integral under the OD profile or peak height

Quantification of band intensities

control proteins within the experimental setting by carrying out gel electrophoresis of a dilution series of either polypeptide and/or the sample extract [28, 31]. However, even when the ideal housekeeping gene was loaded at a concentration in the linear range, or when the total protein concentration was measured at maximum accuracy to avoid housekeeping proteins altogether, ECL detection of Western blot signals still provides only semiquantitative expression data due to the nonlinearity of the cumulative luminescence and limited quantitative reproducibility [50]. Therefore, new fluorescencebased detection methods have been developed to improve the classical ECL-based Western blotting for a more reliable and reproducible quantification of protein expression after SDSPAGE [51]. These techniques exhibit better linearity between signal intensity and protein content as well as a higher dynamic range and therefore allow absolute quantification of the expression level of the target protein without loading control. As a consequence, however, exact measurement of the loaded protein amount is imperative [30] (Fig. 3). Finally, lin C 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

earity and dynamic range of the image acquisition system should be known and fit to the experiments as limitations at that point also might distort the correlation between protein concentration and OD signal.

9

Problem 8: Quantification of band intensities

Even with the best antibody and image acquisition, often more than one band/spot is visible (see above), leading to the question which signal(s) should be taken for analysis and which should be excluded. Generally, taking only the band with the expected molecular mass (or for 2D techniques: pI and molecular mass) might be misleading. Additional bands can arise from natural (cf. Problem 1) or artificially introduced protein modifications (cf. Problems 1, 2, and 5), which are not always distinguishable. Ideally the origin of the different bands is known or can be determined, for example, by www.clinical.proteomics-journal.com

404


employing Western blots with PTM-specific antibodies or by MS-guided peptide identification. In addition, parallel knockdown approaches might provide clarity as to the specificity of antibody-detected signals. If certain signals are being ignored, it needs to be stated why band(s) are excluded from the quantification and others not. If this is not possible, we feel that one should take all bands into account that have been detected by the antibody. Next, in classical ECL Western blotting one needs to correct for background staining. The most potent algorithm appears to be the baseline subtraction algorithm [47] that is applied individually to each single lane after acquiring its OD profile. This profile is then also used to quantify the band intensities, best as integral under the curve (profile) or, as second choice procedure, as peak height. The width of the sampling tool used for acquisition of the lane’s OD profile should be about 30% of the lane width and centered to the lane. Band volume as well as band intensity are poor measuring parameters and should not be taken [47]. As a final note, journals should demand entire blot images to be presented in publications (at least as Supporting Information data) to exclude unjustified cropping or removal of extra bands.

10

Summarizing conclusion

In everyday laboratory life, classical Western blotting is still a very common and useful technique and it is unlikely that this will change in near future. In principle, this technique is simple, relatively cheap, and produces data that are easily interpreted and, in theory at least, reproduced. However, as always, one should be critical in one’s own data identification and quantification and know where problems along a Western blot experiment can arise (cf. checklist in Table 1). This helps to identify artifacts, to optimize the workflow in the laboratory, and, most importantly, to produce reliable data. The authors have declared no conflict of interest.

11

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